The Purple Phototrophic Bacteria
Advances in Photosynthesis and Respiration VOLUME 28 Series Editor : GOVINDJEE University of Illinois, Urbana, Illinois, U.S.A.
Consulting Editors: Julian EATON-RYE, Dunedin, New Zealand Christine H. FOYER, Newcastle upon Tyne, U.K. David B. KNAFF, Lubbock, Texas, U.S.A. Anthony L. MOORE, Brighton, U.K. Sabeeha MERCHANT, Los Angeles, California, U.S.A. Krishna NIYOGI, Berkeley, California, U.S.A. William PARSON, Seatle, Washington, U.S.A. Agepati RAGHAVENDRA, Hyderabad, India Gernot RENGER, Berlin, Germany
The scope of our series, beginning with volume 11, reflects the concept that photosynthesis and respiration are intertwined with respect to both the protein complexes involved and to the entire bioenergetic machinery of all life. Advances in Photosynthesis and Respiration is a book series that provides a comprehensive and state-of-the-art account of research in photosynthesis and respiration. Photosynthesis is the process by which higher plants, algae, and certain species of bacteria transform and store solar energy in the form of energy-rich organic molecules. These compounds are in turn used as the energy source for all growth and reproduction in these and almost all other organisms. As such, virtually all life on the planet ultimately depends on photosynthetic energy conversion. Respiration, which occurs in mitochondrial and bacterial membranes, utilizes energy present in organic molecules to fuel a wide range of metabolic reactions critical for cell growth and development. In addition, many photosynthetic organisms engage in energetically wasteful photorespiration that begins in the chloroplast with an oxygenation reaction catalyzed by the same enzyme responsible for capturing carbon dioxide in photosynthesis. This series of books spans topics from physics to agronomy and medicine, from femtosecond processes to season long production, from the photophysics of reaction centers, through the electrochemistry of intermediate electron transfer, to the physiology of whole organisms, and from X-ray crystallography of proteins to the morphology or organelles and intact organisms. 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, respiration and related processes. For other titles published in this series, go to www.springer.com/series/5599
The Purple Phototrophic Bacteria Edited by
C. Neil Hunter University of Sheffield, United Kingdom
Fevzi Daldal University of Pennsylvania, USA
Marion C. Thurnauer Argonne National Laboratory, USA and
J. Thomas Beatty University of British Columbia, Canada
Library of Congress Control Number: 2008932524
ISBN 978-1-4020-8814-8 (HB) ISBN 978-1-4020-8815-5 (e-book) Published by Springer, P.O. Box 17, 3300 AA Dordrecht, The Netherlands. www.springer.com
Cover: Four aspects of purple phototrophic bacteria, from one of their habitats through to atomic resolution structures, are superimposed on a map derived from the genome sequence of Rhodopseudomonas palustris CGA009 supplied by Professor Caroline Harwood, University of Washington, Seattle, USA. Top left. Purple sulfur bacteria (Amoebobacter purpureus) on the shoreline of Mahoney Lake, British Columbia, Canada. Image from Professor J.T. Beatty. Top right. Rhodobacter capsulatus streaked out on an agar plate. Image from Professor J.T. Beatty. Bottom right. Model of a spherical chromatophore vesicle from Rhodobacter sphaeroides constructed by the in silico combination of atomic force microscopy, linear dichroism, electron microscopy, and X-ray crystallography data. Image from Dr. Melih Sener and Professor Klaus Schulten, prepared using VMD (Humphrey et al. (1996) J Mol Graphics 14: 33–38). Bottom left. Structure of the reaction center complex from Rhodobacter sphaeroides showing the subunits and the pathway of electron transfer between cofactors. See Fig. 1, Chapter 20. Image from Professor Colin Wraight, prepared using VMD. The camera ready text was prepared by Lawrence A. Orr, Center for Bioenergy & Photosynthesis, Arizona State University, Tempe, Arizona 85287-1604, USA.
Printed on acid-free paper
All Rights Reserved © 2009 Springer Science + Business Media B.V. No part of this work may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, microfilming, recording or otherwise, without written permission from the Publisher, with the exception of any material supplied specifically for the purpose of being entered and executed on a computer system, for exclusive use by the purchaser of the work.
This book is dedicated to Roderick K. Clayton, a pioneer of photosynthesis research
From the Series Editor Advances in Photosynthesis and Respiration Volume 28: The Purple Phototrophic Bacteria I am delighted to announce the publication, in the Advances in Photosynthesis and Respiration (AIPH) Series, of The Purple Phototrophic Bacteria. Four distinguished authorities from three countries (UK, USA and Canada) have edited this Volume: C. Neil Hunter, the Chief Editor, Fevzi Daldal, Marion C. Thurnauer and J. Thomas Beatty. This book is produced as a sequel to Volume 2 of the Series (Anoxygenic Photosynthetic Bacteria), published in 1995, and edited by Robert E. Blankenship, Michael T. Madigan and Carl E. Bauer. Published Volumes (2008–1994) • Volume 27 (2008) Sulfur Metabolism in Phototrophic Organisms, edited by Christiane Dahl, Rüdiger Hell, David Knaff and Thomas Leustek, from Germany and USA. 24 Chapters, 551 pp, Hardcover. ISBN: 978-4020-6862-1 • Volume 26 (2008): Biophysical Techniques In Photosynthesis, Volume II, edited by Thijs Aartsma and Jörg Matysik, both from The Netherlands. 24 Chapters, 548 pp, Hardcover. ISBN: 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. 37 Chapters, 603 pp, Hardcover. ISBN: 978-140204515-8 • Volume 24 (2006): Photosystem I: The LightDriven Plastocyanin: Ferredoxin Oxidoreductase, edited by John H. Golbeck, from USA. 40 Chapters, 716 pp, Hardcover. ISBN: 978-140204255-3 • Volume 23 (2006): The Structure and Function of Plastids, edited by Robert R. Wise and J. Kenneth Wise, from USA. 27 Chapters, 575 pp, Softcover: ISBN: 978-1-4020-6570-6, Hardcover, ISBN: 978-1-4020-4060-3
• Volume 22 (2005): Photosystem II: The LightDriven Water:Plastoquinone Oxidoreductase, edited by Thomas J. Wydrzynski and Kimiyuki Satoh, from Australia and Japan. 34 Chapters, 786 pp, Hardcover. ISBN: 978-1-4020-4249-2 • Volume 21 (2005): Photoprotection, Photoinhibition, Gene Regulation, and Environment, edited by Barbara Demmig-Adams, William W. III Adams and Autar K. Mattoo, from USA. 21 Chapters, 380 pp, Hardcover. ISBN: 97814020-3564-7 • Volume 20 (2006): Discoveries in Photosynthesis, edited by Govindjee, J. Thomas Beatty, Howard Gest and John F. Allen, from USA, Canada and UK. 111 Chapters, 1304 pp, Hardcover. ISBN: 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. 31 Chapters, 820 pp, Hardcover. ISBN: 978-1-4020-3217-2 • Volume 18 (2005): Plant Respiration: From Cell to Ecosystem, edited by Hans Lambers and Miquel Ribas-Carbo, from Australia and Spain. 13 Chapters, 250 pp, Hardcover. ISBN: 978-14020-3588-3 • Volume 17 (2004): Plant Mitochondria: From Genome to Function, edited by David Day, A. Harvey Millar and James Whelan, from Australia. 14 Chapters, 325 pp, Hardcover. ISBN: 978-14020-2399-6 • Volume 16 (2004): Respiration in Archaea and Bacteria:Diversity of Prokaryotic Respiratory Systems, edited by Davide Zannoni, from Italy. 13 Chapters, 310 pp, Hardcover. ISBN: 97814020-2002-5 • Volume 15 (2004): Respiration in Archaea and Bacteria: Diversity of Prokaryotic Electron Transport Carriers, edited by Davide Zannoni, from Italy. 13 Chapters, 350 pp, Hardcover. ISBN: 978-1-4020-2001-8 • Volume 14 (2004): Photosynthesis in Algae,
edited by Anthony W. Larkum, Susan Douglas and John A. Raven, from Australia, Canada and UK. 19 Chapters, 500 pp, Hardcover. ISBN: 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. 17 Chapters, 544 pp, Hardcover. ISBN: 97807923-6335-4 • Volume 12 (2003): Photosynthetic Nitrogen Assimilation and Associated Carbon and Respiratory Metabolism, edited by Christine H. Foyer and Graham Noctor, from UK and France. 16 Chapters, 304 pp, Hardcover. ISBN: 978-07923-6336-1 • Volume 11 (2001): Regulation of Photosynthesis, edited by Eva-Mari Aro and Bertil Andersson, from Finland and Sweden. 32 Chapters, 640 pp, Hardcover. ISBN: 978-0-7923-6332-3 • Volume 10 (2001): Photosynthesis: Photobiochemistry and Photobiophysics, by Bacon Ke, from USA. 36 Chapters, 792 pp, Softcover: ISBN: 978-0-7923-6791-8. Hardcover: ISBN: 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. 24 Chapters, 644 pp, Hardcover. ISBN: 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 UK and USA. 20 Chapters, 420 pp, Hardcover. ISBN: 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. 36 Chapters, 760 pp, Hardcover. ISBN: 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. 15 Chapters, 332 pp, Hardcover. ISBN: 978-0-7923-5173-3 • Volume 5 (1997): Photosynthesis and the Environment, edited by Neil R. Baker, from UK. 20 Chapters, 508 pp, Hardcover. ISBN: 97807923-4316-5 • Volume 4 (1996): Oxygenic Photosynthesis: The Light Reactions, edited by Donald R. Ort,
and Charles F. Yocum, from USA. 34 Chapters, 696 pp, Softcover: ISBN: 978-0-7923-3684-6. Hardcover: ISBN: 978-0-7923-3683-9 • Volume 3 (1996): Biophysical Techniques in Photosynthesis, edited by JanAmesz and Arnold J. Hoff, from The Netherlands. 24 Chapters, 426 pp, Hardcover. ISBN: 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. 62 Chapters, 1331 pp, Hardcover. ISBN: 978-0-7923-3682-8 • Volume 1 (1994): The Molecular Biology of Cyanobacteria, edited by Donald R. Bryant, from USA. 28 Chapters, 916 pp, Hardcover. ISBN: 978-0-7923-3222-0 Further information on these books and ordering instructions can be found at under the Book Series ‘Advances in Photosynthesis and Respiration.’ Table of Contents of Volumes 1–25 can be found at . Special discounts are available to members of the International Society of Photosynthesis Research, ISPR (). (See http://www.springer.com/ispr for more information.) About Volume 28: The Purple Phototrophic Bacteria The Purple Phototrophic Bacteria has 48 authoritative Chapters, and is authored by 116 international authorities from 13 countries (Australia; Canada; Czech Republic; France; Germany; Israel; Italy; Japan; Netherlands; Poland; Russia; UK; and USA). It is a truly international book and the Chief Editor of this volume, C. Neil Hunter, and his three coeditors, Fevzi Daldal, Marion C. Thurnauer and J. Thomas Beatty, deserve our thanks and our congratulations for compiling this updated survey of these interesting and important organisms. The Purple Phototrophic Bacteria is a comprehensive survey of all aspects of these fascinating bacteria, most metabolically versatile organisms on Earth. This volume is organized into the following sections: Physiology, Evolution and Ecology; Biosynthesis of Pigments, Cofactors and Lipids; Antenna Complexes: Structure, Function and Orgaviii
nization; Reaction Centre Structure and Function; Cyclic Electron Transfer Components and Energy Coupling Reactions; Metabolic Processes; Genomics, Regulation and Signaling; and New Applications and Techniques. This book is a compilation of 48 chapters, written by leading experts who highlight the huge progress made in spectroscopic, structural and genetic studies of these bacteria since 1995, when the last such book was published (Anoxygenic Photosynthetic Bacteria, Volume 2 in the Advances in Photosynthesis and Respiration Series, edited by Robert E. Blankenship, Michael T. Madigan and Carl E. Bauer; Kluwer Academic Publishers (now Springer), Dordrecht). This new volume is similarly intended to be the definitive text on these bacteria for many years to come, and it will be a valuable resource for experienced researchers, Ph.D. students, and advanced undergraduates in the fields of ecology, microbiology, biochemistry and biophysics. Scientists interested in future applications of these bacteria which could harness their potential for nanotechnology, solar energy research, bioremediation, or as cell factories, will also find the book useful. The readers can easily find the titles and the authors of the individual chapters in the Table of Content of this book. Instead of repeating this information here, I prefer to thank each and every author by name (listed in alphabetical order) that reads like a ‘Who’s Who’ in the field of purple phototrophic bacteria:
Loach; Chris Mackenzie; Chris Madigan; Michael T. Madigan; Pier Luigi Martelli; Bernd Masepohl; Shinji Masuda; Donna L. Mielke; Osamu Miyashita; Mamoru Nango; Robert A. Niederman; Wolfgang Nitschke; Vladimir I. Novoderezhkin; Dror Noy; U. Mirian Obiozo; Melvin Okamura; Ozlem Onder; Miroslav Papiz; Pamela S. ParkesLoach; William W. Parson; Marcela Ávila Pérez; Tomáš Polívka; Oleg G. Poluektov; Pu Qian; Jason Raymond; Bruno Robert; Brigitte R. Robinson; Simona Romagnoli; Johannes Sander; Carsten Sanders; Hugo Scheer; Simon Scheuring; Barbara Schoepp-Cothenet; Klaus Schulten; Melih K. Şener; Aaron Setterdahl; James P. Shapleigh; James N. Sturgis; Wesley D. Swingley; F. Robert Tabita; Shinichi Takaichi; Banita Tamot; Serdar Turkarslan; Lisa M. Utschig; Rienk van Grondelle; Michael A. van der Horst; Luuk J. van Wilderen; André Verméglio; Paulette M. Vignais; Martin J. Warren; Arieh Warshel; JoAnn C. Williams; Robert D. Willows; Colin A. Wraight; Jiang Wu; Vladimir Yurkov; Davide Zannoni; and Jill Helen Zeilstra-Ryalls. It is a pleasure to note that the following 27 participants in Volume 2 have also contributed to Volume 28; they are (alphabetically): James P. Allen; Judith P. Armitage; Carl E. Bauer; J. Thomas Beatty; Robert E. Blankenship; Richard J. Cogdell; Fevzi Daldal; Harry A. Frank; Caroline S. Harwood; C. Neil Hunter; J. Baz Jackson; Pierre Joliot; Gabriele Klug; Robert G. Kranz; Paul A. Loach; Michael T. Madigan; Melvin Okamura; Pamela S. Parkes-Loach; William W. Parson; Hugo Scheer; F. Robert Tabita; Rienk van Grondelle; André Verméglio; Paulette M. Vignais; Arieh Warshel; JoAnn C. Williams; and Davide Zannoni. It is remarkable that all three editors of Volume 2 (Carl E. Bauer, Robert E. Blankenship (Chief Editor), and Michael T. Madigan) are authors in Volume 28; and 3 of the 4 editors of Volume 28 (J. Thomas Beatty; Fevzi Daldal; and C. Neil Hunter (Chief Editor)) were authors in Volume 2. As Volume 28 is a sequel to Volume 2, it is beneficial for the readers of the new volume to consult and cite chapters in the earlier volume; I present below authors, titles of chapters and page numbers of all the chapters in that book. Please note that this volume was published by Kluwer Academic Publishers which was later acquired by Springer, the publishers of the current volume.
Maxime T.A. Alexandre; James P. Allen; Judith P. Armitage; Herbert Axelrod; Carl E. Bauer; Christoph Benning; Edward A. Berry; Robert E. Blankenship; Francesca Borsetti; Paula Braun; Per A. Bullough; Rita Casadio; Madhusudan Choudhary; Toh Kee Chua; Richard J. Cogdell; Jason W. Cooley; Julius T. Csotony; Christiane Dahl; Fevzi Daldal; Evelyne Deery; Takehisa Dewa; Timothy J. Donohue; Katie Evans; Boris A. Feniouk; Leszek Fiedor; Anthony Fordham-Skelton; Harry A. Frank; Elaine R. Frawley; Mads Gabrielsen; Alastair T. Gardiner; Toni Georgiou; Marie-Louise Groot; Marilyn R. Gunner; Wolfgang Haehnel; Deborah K. Hanson; Caroline S. Harwood; Klaas J. Hellingwerf; Johnny Hendriks; Theresa Hillon; Jonathan Hosler; Li-Shar Huang; C. Neil Hunter; Kouji Iida; J. Baz Jackson; Pierre Joliot; Michael R. Jones; Deborah O. Jung; Wolfgang Junge; Samuel Kaplan; John T. M. Kennis; Gabriele Klug; Hans Georg Koch; Jürgen Köhler; David M. Kramer; Robert G. Kranz; Alison M. Kriegel; Philip D. Laible; Jérôme Lavergne; Dong-Woo Lee; Paul A. ix
Complete List of Chapters in Anoxygenic Photosynthetic Bacteria, edited by R.E. Blankenship, M.T. Madigan and C.E. Bauer, Kluwer Academic Publishers, 1995
in purple bacteria, pp 349–372 Chapter 18: H.A. Frank and R.L. Christensen (1995) Singlet energy transfer from carotenoids to bacteriochlorophylls, pp 373–384 Chapter 19: A. Freiberg (1995) Coupling of antennas to reaction centers, pp 385–398 Chapter 20: R.E. Blankenship, J.M. Olson and M. Mette (1995) Antenna complexes from green photosynthetic bacteria, pp 399–435 Chapter 21: P.A. Loach and P.S. Parkes-Loach (1995) Structure-function relationships in core light-harvesting complexes (LHI) as determined by characterization of the structural subunit and by reconstitution experiments, pp 437–471 Chapter 22: C.N. Hunter (1995) Genetic manipulation of the antenna complexes of purple bacteria, pp 473–501 Chapter 23: C.R.D. Lancaster, U. Ermler and H. Michel (1995) The structures of photosynthetic reaction centers from purple bacteria as revealed by X-ray crystallography, pp 503–526 Chapter 24: N.W. Woodbury and J.P. Allen (1995) The pathway, kinetics and thermodynamics of electron transfer in wild type and mutant reaction centers of purple nonsulfur bacteria, pp 527–557 Chapter 25: W.W. Parson and A. Warshel (1995) Theoretical analyses of electron-transfer reactions, pp 559–575 Chapter 26: M.Y. Okamura and G. Feher (1995) Proton-coupled electron transfer reactions of QB in reaction centers from photosynthetic bacteria, pp 577–594 Chapter 27: M. Volk, A. Ogrodnik and M.E. Michel-Beyerle (1995) The recombination dynamics of the radical pair P+H– in external magnetic and electric fields, pp 595–626 Chapter 28: W. Mäntele (1995) Infrared vibrational spectroscopy of reaction centers, pp 627–647 Chapter 29: H. Scheer and G. Hartwich (1995) Bacterial reaction centers with modified tetrapyrrole chromophores, pp 649–663 Chapter 30: U. Feiler and G. Hauska (1995) The reaction center from green sulfur bacteria, pp 665–685 Chapter 31: J. Amesz (1995) The antenna-reaction center complex of heliobacteria, pp 687–697 Chapter 32: R. Feick, J.A. Shiozawa and A. Ertlmaier (1995) Biochemical and spectroscopic properties of the reaction center of the green filamentous bacterium, Chloroflexus aurantiacus, pp 699–708 Chapter 33: R.G. Kranz and D.L. Beckman (1995) Cytochrome biogenesis, pp 709–723
Chapter 1: J.F. Imhoff (1995) Taxonomy and physiology of phototrophic purple bacteria and green sulfur bacteria, pp 1–15 Chapter 2: M.T. Madigan and J.G. Ormerod (1995) Taxonomy, physiology and ecology of heliobacteria, pp 17–30 Chapter 3: B.K. Pierson and R.W. Castenholz (1995) Taxonomy and physiology of filamentous anoxygenic phototrophs, pp 31–47 Chapter 4: H. Van Gemerden and J. Mas (1995) Ecology of phototrophic sulfur bacteria, pp 49–85 Chapter 5: R.W. Castenholz and B.K. Pierson (1995) Ecology of thermophilic anoxygenic phototrophs, pp 87–103 Chapter 6: K. Shimada (1995) Aerobic anoxygenic phototrophs, pp 105–122 Chapter 7: D.E. Fleischman, W.R. Evans and I.M. Miller (1995) Bacteriochlorophyll-containing rhizobium species, 123–136 Chapter 8: M.O. Senge and K.M. Smith (1995) Biosynthesis and structures of the bacteriochlorophylls, pp 137–151 Chapter 9: S.I. Beale (1995) Biosynthesis and structures of porphyrins and hemes, pp 153–177 Chapter 10: J.F. Imhoff and U. Bias-Imhoff (1995) Lipids, quinones and fatty acids of anoxygenic phototrophic bacteria, pp 179–205 Chapter 11: J. Weckesser, H. Mayer and G. Schulz (1995) Anoxygenic phototrophic bacteria: Model organisms for studies on cell wall macromolecules, pp 207–230 Chapter 12: G. Drews and J.R. Golecki (1995) Structure, molecular organization, and biosynthesis of membranes of purple bacteria, pp 231–257 Chapter 13: J. Oelze and J.R. Golecki (1995) Membranes and chlorosomes of green bacteria: Structure, composition and development, pp 259–278 Chapter 14: A. Verméglio, P. Joliot and A. Joliot (1995) Organization of electron transfer components and supercomplexes, pp 279–295 Chapter 15: W.S. Struve (1995) Theory of electronic energy transfer, pp 297–313 Chapter 16: H. Zuber and R.J. Cogdell (1995) Structure and organization of purple bacterial antenna complexes, pp 315–348 Chapter 17: V. Sundström and R. Van Grondelle (1995) Kinetics of excitation transfer and trapping x
Chapter 34: T.E. Meyer and T.J. Donohue (1995) Cytochromes, iron-sulfur, and copper proteins mediating electron transfer from the Cyt bc1 complex to photosynthetic reaction center complexes, pp 725–745 Chapter 35: K.A. Gray and F. Daldal (1995) Mutational studies of the cytochrome bc1 complexes, pp 747–774 Chapter 36: W. Nitschke and S.M. Dracheva (1995) Reaction center associated cytochromes, pp 775–805 Chapter 37: Z. Gromet-Elhanan (1995) The protontranslocating F0F1 ATP synthase-ATPase complex, pp 807–830 Chapter 38: J.B. Jackson (1995) Proton-translocating transhydrogenase and NADH dehydrogenase in anoxygenic photosynthetic bacteria, pp 831–845 Chapter 39: D.C. Brune (1995) Sulfur compounds as photosynthetic electron donors, pp 847–870 Chapter 40: R. Sirevåg (1995) Carbon metabolism in green bacteria, pp 871–883 Chapter 41: F.R. Tabita (1995) The biochemistry and metabolic regulation of carbon metabolism and CO2 fixation in purple bacteria, pp 885–914 Chapter 42: M.T. Madigan (1995) Microbiology of nitrogen fixation by anoxygenic photosynthetic bacteria, pp 915–928 Chapter 43: P.W. Ludden and G.P. Roberts (1995) The biochemistry and genetics of nitrogen fixation by photosynthetic bacteria, pp 929–947 Chapter 44: D. Zannoni (1995) Aerobic and anaerobic electron transport chains in anoxygenic phototrophic bacteria, pp 949–971 Chapter 45: J. Mas and H. Van Gemerden (1995) Storage products in purple and green sulfur bacteria, pp 973–990 Chapter 46: J. Gibson and C.S. Harwood (1995) Degradation of aromatic compounds by nonsulfur purple bacteria, pp 991–1003 Chapter 47: J.P. Armitage, D.J. Kelly and R.E. Sockett (1995) Flagellate motility, behavioral responses and active transport in purple non-sulfur bacteria, pp 1005–1028 Chapter 48: J.C. Williams and A.K.W. Taguchi (1995) Genetic manipulation of purple photosynthetic bacteria, pp 1029–1065 Chapter 49: M. Fonstein and R. Haselkorn (1995) Physical mapping of Rhodobacter capsulatus: Cosmid encyclopedia and high resolution genetic map, pp 1067–1081 Chapter 50: M. Alberti, D.H. Burke and J.E. Hearst (1995) Structure and sequence of the photosynthesis
gene cluster, pp 1083–1106 Chapter 51: J.L. Gibson (1995) Genetic analysis of CO2 fixation genes, pp 1107–1124 Chapter 52: A.J. Biel (1995) Genetic analysis and regulation of bacteriochlorophyll biosynthesis, pp 1125–1134 Chapter 53: G.A. Armstrong (1995) Genetic analysis and regulation of carotenoid biosynthesis: Structure and fuction of the crt genes and gene products, pp 1135–1157 Chapter 54: J.A. Shiozawa (1995) A foundation for the genetic analysis of green sulfur, green filamentous and heliobacteria, pp 1159–1173 Chapter 55: P.M. Vignais, B. Toussaint and A. Colbeau (1995) Regulation of hydrogenase gene expression, pp 1175–1190 Chapter 56: R.G. Kranz and P.J. Cullen (1995) Regulation of nitrogen fixation genes, pp 1191–1208 Chapter 57: J.T. Beatty (1995) Organization of photosynthesis gene transcripts, pp 1209–1219 Chapter 58: C.E. Bauer (1995) Regulation of photosynthesis gene expression, pp 1221–1234 Chapter 59: G. Klug (1995) Post-transcriptional control of photosynthesis gene expression, pp 1235–1244 Chapter 60: R.C. Fuller (1995) Polyesters and photosynthetic bacteria: from lipid cellular inclusions to microbial thermoplastics, pp 1245–1256 Chapter 61: E.R. Goldman and D.C. Youvan (1995) Imaging Spectroscopy and combinatorial mutagenesis of the reaction center and light harvesting II antenna, pp 1257–1268 Chapter 62: Michiharu Kobayashi and Michihiko Kobayashi (1995) Waste remediation and treatment using anoxygenic phototrophic bacteria, pp 1269–1282
Future AIPH 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): • C-4 Photosynthesis and Related CO2 Concentrating Mechanisms (Editors:Agepati S. Raghavendra and Rowan Sage); • Photosynthesis: Perspectives on Plastid Biology, Energy Conversion and Carbon Metabolism (Editors: Julian Eaton-Rye and Baishnab Tripathy); xi
• Abiotic Stress Adaptation in Plants: Physiological, Molecular and Genomic Foundation (Editors: Ashwani Pareek, Sudhir K. Sopory, Hans J. Bohnert and Govindjee); • The Chloroplast: Biochemistry, Molecular Biology and Bioengineering (Editors: Constantin Rebeiz, Hans Bohnert, Christoph Benning, Henry Daniell, Beverley R. Green, J. Kenneth Hoober, Hartmut Lichtenthaler, Archie R. Portis and Baishnab C. Tripathy); • Photosynthesis In Silico: Understanding Complexity from Molecules to Ecosystems (Editors: Agu Laisk, Ladislav Nedbal and Govindjee); and • Lipids in Photosynthesis: Essential and Regulatory Function (Editors: Hajime Wada and Norio Murata).
Readers are encouraged to send their suggestions for these and future Volumes (topics, names of future editors, and of future authors) to me by E-mail (gov@ illinois.edu) or fax (1-217-244-7246). In view of the interdisciplinary character of research in photosynthesis and respiration, it is my earnest hope that this series of books will be used in educating students and researchers not only in Plant Sciences, Molecular and Cell Biology, Integrative Biology, Biotechnology, Agricultural Sciences, Microbiology, Biochemistry, and Biophysics, but also in Bioengineering, Chemistry, and Physics. I take this opportunity to thank and congratulate C. Neil Hunter, Fevzi Daldal, Marion Thurnauer and Thomas Beatty for their editorial work. My special thanks go to C. Neil Hunter, the Chief Editor of Volume 28, for his painstaking work not only in editing, but also in organizing this book for Springer, and for his highly professional dealing with the typesetting process and his help in preparing this editorial. I particularly thank all the 116 authors (see the list above) of this book: without their authoritative chapters, there would be no such Volume. I give special thanks to Larry Orr for typesetting this book: his expertise has been crucial in bringing this book to completion. We owe Jacco Flipsen, Noeline Gibson and André Tournois (of Springer) thanks for their friendly working relation with us that led to the production of this book. Thanks are also due to Jeff Haas (Director of Information Technology, Life Sciences, University of Illinois at Urbana-Champaign, UIUC), Evan DeLucia (Head, Department of Plant Biology, UIUC) and my dear wife Rajni Govindjee for constant support.
In addition to these contracted books, the following topics, among others, are under consideration: • • • • • • • • • • •
Cyanobacteria Genomics, Proteomics and Evolution Biohydrogen Production ATP Synthase and Proton Translocation Interactions between Photosynthesis and other Metabolic Processes Carotenoids II Green Bacteria and Heliobacteria Ecophysiology Photosynthesis, Biomass and Bioenergy Global Aspects of Photosynthesis Artificial Photosynthesis
May 14, 2008 Govindjee Series Editor, Advances in Photosynthesis and Respiration University of Illinois at Urbana-Champaign, Department of Plant Biology, 265 Morrill Hall, 505 South Goodwin Avenue, Urbana, IL 61801-3707, USA E-mail:
[email protected]; URL: http://www.life.uiuc.edu/govindjee
xii
Govindjee, Series Editor
Govindjee, born in 1932, obtained his B.Sc. (Chemistry, Biology) and M.Sc. (Botany, Plant Physiology) in 1952 and 1954, from the University of Allahabad, India, and his Ph.D. (Biophysics, under Eugene Rabinowitch), in 1960, from the University of Illinois at Urbana-Champaign (UIUC), IL, U.S.A. He is best known for his research on excitation energy transfer, light emission, primary photochemistry and electron transfer in Photosystem II (PS II). His research, with many collaborators, has included the discovery of a short-wavelength form of chlorophyll (Chl) a functioning in the Chl b-containing system, now called PS II, and of the two-light effects in Chl a fluorescence and in NADP (nicotinamide adenine dinucleotide phosphate) reduction in chloroplasts (Emerson Enhancement). Further, he has worked on the existence of different spectral fluorescing forms of Chl a and the temperature dependence of excitation energy transfer down to 4 K; basic relationships between Chl a fluorescence and photosynthetic reactions; the unique role of bicarbonate on the acceptor side of PS II, particularly in protonation events involving the QB binding region; the theory of thermoluminescence in plants; 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 against excess light. His research on photosynthetic bacteria
included the observation of the absence of the Emerson Enhancement Effect (1960s); measurements on the lifetime of bacteriochlorophyll fluorescence (1970s); and the use of the bacterial reaction center structure in homology modeling of Photosystem II, particularly on its electron acceptor side (1990s). His current focus is on the ‘History of Photosynthesis Research,’ in ‘Photosynthesis Education’, and in the ‘Possible Existence of Extraterrestrial Life.’ He has served on the faculty of the UIUC for ~40 years. Since 1999, he has been Professor Emeritus of Biochemistry, Biophysics and Plant Biology at the same institution. His honors include: Fellow of the American Association of Advancement of Science; Distinguished Lecturer of the School of Life Sciences, UIUC; Fellow and Lifetime member of the National Academy of Sciences (India); President of the American Society for Photobiology (1980–1981); Fulbright Scholar and Fulbright Senior Lecturer; Honorary President of the 2004 International Photosynthesis Congress (Montréal, Canada); the 2006 Recipient of the Lifetime Achievement Award from the Rebeiz Foundation for Basic Biology; and the 2007 Recipient of the ‘Communication Award’ of the International Society of Photosynthesis Research (ISPR), presented to him at the 14th International Congress on Photosynthesis, held in Glasgow, Scotland, U.K.
xiii
Contents From the Series Editor
vii
Contents
xv
Preface
xxxi
Author Index
xxxvii
Color Plates
CP1–CP16
Part 1: Physiology, Ecology and Evolution 1
An Overview of Purple Bacteria: Systematics, Physiology, and Habitats Michael T. Madigan and Deborah O. Jung Summary I. Introduction II. Systematics of Purple Bacteria III. Physiology of Purple Bacteria IV. Habitats of Purple Bacteria V. Purple Bacteria in Extreme Environments VI. Final Remarks Acknowledgments References
2
1–15 2 2 3 4 7 9 12 12 12
Evolutionary Relationships Among Purple Photosynthetic Bacteria and the Origin of Proteobacterial Photosynthetic Systems 17–29 Wesley D. Swingley, Robert E. Blankenship and Jason Raymond Summary I. Introduction II. The Alphaproteobacteria III. Aerobic Purple Bacteria IV. The Photosynthesis Gene Cluster and its Role in Evolution V. Proteobacterial Comparative Genomics: Photosynthetic versus NonPhotosynthetic Proteins VI. Origin and Evolution of Proteobacterial Phototrophy VII. Origin and Evolution of Proteobacterial Carbon-fixation VIII. Future Directions: High-Throughput Sequencing and Metagenomics Acknowledgments References
xv
17 18 18 19 20 21 22 24 27 28 28
3
New Light on Aerobic Anoxygenic Phototrophs Vladimir Yurkov and Julius T. Csotonyi
31–55
Summary 31 I. Introduction 32 II. Morphological Diversity, Taxonomic Nuances, Phylogeny and Evolution 34 III. Nutritional Versatility and Peculiarities of Carbon Metabolism 40 IV. Photosynthetic Pigment Composition and Synthesis Reveal Surprises 41 V. The Mysterious Photosynthetic Apparatus of Aerobic Anoxygenic Phototrophs 44 VI. Speculation on Ecological Roles 47 VII. Concluding Remarks and Perspectives 51 Acknowledgments 52 References 52
Part 2: Molecular Structure and Biosynthesis of Pigments and Cofactors 4
Biosynthesis of Bacteriochlorophylls in Purple Bacteria Robert D. Willows and Alison M. Kriegel
57–79
Summary I. Introduction II. δ-Aminolevulinate to Protoporphyrin IX III. Protoporphyrin IX to Bacteriochlorophyll a and b IV. Concluding Remarks Acknowledgments References
5
Vitamin B12 (Cobalamin) Biosynthesis in the Purple Bacteria Martin J. Warren and Evelyne Deery
57 58 59 65 75 75 75
81–95
Summary I. Background II. Function of Cobalamin III. B12 Biosynthesis in Rhodobacter capsulatus and Rhodobacter sphaeroides IV. Summary of Events Required for Cobalamin Biosynthesis V. Biosynthesis of Precorrin-2 and Its Onward Route Towards Siroheme and Cobalamin VI. Control and Regulation of Cobalamin Biosynthesis Acknowledgments References
6
Distribution and Biosynthesis of Carotenoids Shinichi Takaichi Summary I. Introduction II. Carotenogenesis III. Carotenoids in Purple Bacteria Acknowledgments References
xvi
81 81 82 83 84 86 92 92 92
97–117 97 98 101 111 114 114
7
Membrane Lipid Biosynthesis in Purple Bacteria Banita Tamot and Christoph Benning Summary I. Introduction II. Fatty Acids III. Phosphoglycerolipids IV. Glycoglycerolipids V. Betaine Lipid VI. Ornithine Lipid VII. Lipid Function VIII. Perspectives Acknowledgments References
119–134 119 120 122 123 126 128 129 130 131 131 132
Part 3: Antenna Complexes: Structure, Function and Organization 8
Peripheral Complexes of Purple Bacteria Mads Gabrielsen, Alastair T. Gardiner and Richard J. Cogdell Summary I. Introduction II. Structure III. The Biology of Purple Bacterial Antenna Complexes IV. Final Remarks Acknowledgments References
9
Reaction Center-Light-Harvesting Core Complexes of Purple Bacteria Per A. Bullough, Pu Qian and C. Neil Hunter Summary I. Introduction II. The Building Blocks: The α and β Polypeptides of Light-Harvesting Complex 1, and PufX III. Circles, Arcs and Ellipses — The Light-Harvesting 1 Complex IV. Monomeric Reaction Center-Light-Harvesting 1 Complexes V. Monomeric Reaction Center-Light-Harvesting 1-PufX Complexes VI. Dimeric Reaction Center-Light-Harvesting 1-PufX Complexes VII. The Biogenesis of Core Complexes Acknowledgments References
xvii
135–153 135 136 136 146 151 151 151
155–179 155 156 156 160 162 166 168 174 175 175
10 Structure-Function Relationships in Bacterial Light-Harvesting Complexes Investigated by Reconstitution Techniques Paul A. Loach and Pamela S. Parkes-Loach
181–198
Summary I. Introduction II. Reversible Dissociation of Core Light-Harvesting 1 Complexes to a Subunit Form (B820) III. Reversible Dissociation of B820 to its Fundamental Components IV. Cofactor Requirements V. Cofactor – Protein Interactions VI. Cofactor – Cofactor Interactions VII. Protein–Protein Interactions VIII. Effect of PufX on Reconstitution of Light-Harvesting 1 Complexes IX. In vitro versus In vivo Assembly of Complexes X. Reconstitution of the Reaction Center Acknowledgments References
11 Spectroscopic Properties of Antenna Complexes from Purple Bacteria Bruno Robert
181 182 182 183 184 184 188 188 191 192 193 195 195
199–212
Summary 1. Introduction II. The Different Spectral Forms of Antenna Proteins from Purple Bacteria III. Antenna Proteins from Purple Bacteria: Variations Around a Structural Theme IV. The Role of Ground State Interactions in Tuning the Antenna Absorption Transition V. Excitonic Interactions and Disorder in Light-Harvesting Complexes VI. Chromophore Interactions in Light-Harvesting Proteins: Additional Effects VII. Conclusions Acknowledgments References
12 Energy Transfer from Carotenoids to Bacteriochlorophylls Harry A. Frank and Tomáš Polívka
201 203 205 207 208 209 209
213–230
Summary I. Introduction II. Carotenoid Excited States III. Energy Transfer in Light-Harvesting 2 Complexes IV. Energy Transfer in Light-Harvesting 1 Complexes and Reaction Centers V. Outlook Acknowledgments References
xviii
199 200 200
213 214 215 216 226 227 227 227
13 Spectroscopy and Dynamics of Excitation Transfer and Trapping in Purple Bacteria 231–252 Rienk van Grondelle and Vladimir I. Novoderezhkin Summary 232 I. Introduction 232 II. Structure and Exciton Spectra of Light-Harvesting 1 and 2 Bacterial Antenna Complexes 235 III. Equilibration Dynamics 239 IV. Competition of Intraband B800-800 and Interband B800-850 Energy Transfer in the Light-Harvesting 2 Complex 241 V. Energy Trapping in the Core Reaction Center-Light-Harvesting 1 Complex 243 VI. Slow Conformational Motions and Excitation Dynamics in the B850-LightHarvesting 2 Complex 244 VII. Concluding Remarks 247 Acknowledgments 248 References 248
14 Organization and Assembly of Light-Harvesting Complexes in the Purple Bacterial Membrane 253–273 James N. Sturgis and Robert A. Niederman Summary I. Introduction II. Themes and Variations — Structural Variability of Complexes in Native Membranes III. Principles of Photosynthetic Unit Organization IV. Proposals for the Functional Organization of Photosynthetic Units V. In Vivo Assembly of Light-Harvesting Complexes VI. Perspectives for the Next Ten Years References
15 From Atomic-Level Structure to Supramolecular Organization in the Photosynthetic Unit of Purple Bacteria Melih K. ùener and Klaus Schulten
254 254 255 257 263 267 269 270
275–294
Summary 275 I. Introduction 276 II. Components of the Photosynthetic Unit 278 III. Quantum Physics of Light-Harvesting and Excitation Energy Transfer 279 IV. Effect of Thermal Disorder on the Spectra of Light-Harvesting Complexes 282 V. Physical Constraints Shaping the Structure of Individual Light-Harvesting Complexes 284 VI. Supramolecular Organization of the Photosynthetic Unit 286 VII. An Atomic-level Structural Model for a Photosynthetic Membrane Vesicle 287 VIII. Light-Harvesting and Excitation Transfer across a Photosynthetic Membrane Vesicle 287 IX. Conclusions 289 Acknowledgments 290 References 290
xix
Part 4: Reaction Center Structure and Function 16 Structural Plasticity of Reaction Centers from Purple Bacteria Michael R. Jones
295–321
Summary I. The Plastic Purple Reaction Center II. Biochemical and Genetic Alteration of Polypeptide Composition III. Cofactor Exclusion IV. Cofactor Replacement V. Helix Symmetrization VI. Water and Other Unexpected Things in Electron Density Maps VII. Building New Functionality VIII. Conclusions Acknowledgments References
295 296 297 299 302 307 310 312 313 314 314
17 Structure and Function of the Cytochrome c2:Reaction Center Complex from Rhodobacter sphaeroides Herbert Axelrod, Osamu Miyashita and Melvin Okamura Summary I. Introduction II. History III. Structure of the Cytochrome c2:Reaction Center Complex IV. Electron Transfer Reactions V. Effects of Mutation VI. Mechanism of Inter-Protein Electron Transfer Acknowledgments References
18 Directed Modification of Reaction Centers from Purple Bacteria JoAnn C. Williams and James P. Allen Summary I. Introduction II. Properties of the Cofactors III. Electron Transfer Concepts IV. Pathways of Electron Transfer V. Conclusions Acknowledgments References
xx
323–336 323 324 324 325 327 329 332 333 333
337–353 337 338 338 343 346 349 349 349
19 Mechanism of Charge Separation in Purple Bacterial Reaction Centers William W. Parson and Arieh Warshel Summary I. The Reaction Sequence and Kinetics II. Energies of the Radical-Pair Intermediates III. Unusual Features of the Charge-Separation Reactions IV. Theories of Electron-Transfer Reactions Acknowledgments References
20 The Acceptor Quinones of Purple Photosynthetic Bacteria— Structure and Spectroscopy Colin A. Wraight and Marilyn R. Gunner Summary I. Introduction II. The Acceptor Quinone Reactions III. Structural Features of the Acceptor Quinone Binding Sites IV. Spectroscopy of the Acceptor Quinones V. Functionality of the Two Quinone Positions in the QB Site VI. Conclusions Note Added in Proof Acknowledgments References
355–377 355 356 357 360 363 370 370
379–405 379 380 382 383 389 396 398 398 398 399
Part 5: Cyclic Electron Transfer Components and Energy Coupling Reactions 21 Biogenesis of c-type Cytochromes and Cytochrome Complexes Carsten Sanders, Serdar Turkarslan, Ozlem Onder, Elaine R. Frawley, Robert G. Kranz, Hans Georg Koch and Fevzi Daldal Summary I. Introduction II. Maturation of c-type Cytochromes: Ccm-system I and Ccs-system II III. Biogenesis of Cytochrome Complexes Acknowledgments References
xxi
407–423
407 408 409 415 421 421
22 Structural and Mutational Studies of the Cytochrome bc1 Complex 425–450 Edward A. Berry, Dong-Woo Lee, Li-Shar Huang and Fevzi Daldal Summary I. Introduction II. Structural and Mutational Studies III. Conclusions and Perspectives Acknowledgments References
23 The Cytochrome bc1 and Related bc Complexes: The Rieske/ Cytochrome b Complex as the Functional Core of a Central Electron/Proton Transfer Complex David M. Kramer, Wolfgang Nitschke and Jason W. Cooley
425 426 427 446 447 447
451–473
Summary I. Introduction II. Structures of the Cytochrome bc1 and Related Rieske/Cytochrome b Complexes III. Catalysis in the Rieske/Cytochrome b Complexes: The General Q-cycle Framework IV. Phylogeny and Evolution V. The ‘Third’ Redox Subunit is a Phylogenetic Marker VI. The Rieske/Cytochrome b Complex, a Primordial Enzyme VII. The Molecular Mechanism of the Qo Site: Avoiding Q-Cycle Short Circuits VIII. The Quinone Reduction Site, Qi of the Cytochrome bc1 Complexes IX. The Quinone Reduction Site,Qi of the Cytochrome b6 f and Related Complexes X. The Functional Mechanism of the Rieske/Cytochrome b Complexes is Conserved Acknowledgments References
452 452 453 455 456 458 459 460 467 468 469 469 469
24 Proton Translocation and ATP Synthesis by the FoF1-ATPase of Purple Bacteria Boris A. Feniouk and Wolfgang Junge
475–493
Summary I. Introduction II. Structure and Rotary Catalysis III. Proton Translocation and its Coupling to ATP Synthesis/Hydrolysis III. Role of Proton Translocation in the Regulation of ATP Synthase Acknowledgments References
475 476 476 478 486 487 488
xxii
25 Proton-Translocating Transhydrogenase in Photosynthetic Bacteria J. Baz Jackson and U. Mirian Obiozo
495–508
Summary I. Introduction II. An Overview of the Main Structural Features of Transhydrogenase III. Distribution of Transhydrogenase Among Species IV. Phylogenetic Relationships between Transhydrogenases from Different Species V. The Function of Transhydrogenase in Photosynthetic Bacteria VI. The Mechanism of Coupling Between Hydride Transfer and Proton Translocation in Transhydrogenase VII. Conformational Changes in the Coupling Reactions of Transhydrogenase Acknowledgments References
495 496 496 496 498 500 501 504 505 506
26 Functional Coupling Between Reaction Centers and Cytochrome bc1 Complexes 509–536 Jérôme Lavergne, André Verméglio and Pierre Joliot Summary I. Introduction II. Structure of the Protein Complexes III. The Electron Donors to the Reaction Center IV. Kinetics of P+ Reduction by Mobile Cytochromes V. Donor Side Shuttling and Turnover of the Cytochrome bc1 Complex VI. Quinone Reactions VII. Supramolecular Organization in Rhodobacter sphaeroides and Rhodobacter capsulatus VIII. Quinone Confinement in Rhodobacter sphaeroides IX. Quinone Traffic in the PufX– Mutant of Rhodobacter sphaeroides X. The Supercomplex Model: Difficulties and Alternative Possibilities XI. Mitochondrial Supercomplexes XII. Diffusion and Confinement of Cytochrome c2: Possible Mechanisms XIII. Diffusion and Confinement of Quinones: Possible Mechanisms XIV. Conclusions References
509 510 512 514 515 517 518 519 522 522 524 526 527 528 529 530
Part 6: Metabolic Processes 27 Respiration and Respiratory Complexes 537–561 Davide Zannoni, Barbara Schoepp-Cothenet and Jonathan Hosler Summary I. Aerobic Respiration II. Respiration Utilizing Substrates other than Oxygen III. Respiration versus Photosynthesis: Which One Came First? IV. Respiration and Photosynthesis are Intermingled Acknowledgments References
xxiii
538 538 546 553 553 555 555
28 Carbon Dioxide Metabolism and its Regulation in Nonsulfur Purple Photosynthetic Bacteria Simona Romagnoli and F. Robert Tabita
563–576
Summary I. Introduction II. Regulation of cbb Gene Expression Acknowledgments References
29 Degradation of Aromatic Compounds by Purple Nonsulfur Bacteria Caroline S. Harwood
563 564 564 575 575
577–594
Summary I. Introduction II. Biochemical Themes III. Species that Degrade Aromatic Compounds IV. Aerobic Degradation of Aromatic Compounds by Rhodopseudomonas palustris V. Anaerobic Benzoate Degradation VI. The Molecular Regulation of Anaerobic Aromatic Compound Degradation VII. Comparative Aspects Acknowledgments References
30 Metabolism of Inorganic Sulfur Compounds in Purple Bacteria Johannes Sander and Christiane Dahl Summary I. Introduction II. Sulfur Oxidation Capabilities of Purple Bacteria III. Sulfur Oxidation Pathways IV. Sulfate Assimilation V. Conclusions Acknowledgments References
31 Dissimilatory and Assimilatory Nitrate Reduction in the Purple Photosynthetic Bacteria James P. Shapleigh Summary I. Introduction II. Denitrification III. Assimilation of Nitrogen IV. Conclusion Acknowledgments References
577 578 578 579 579 580 589 590 591 591
595–622 596 596 596 602 612 615 616 616
623–642 623 624 624 636 638 639 639
xxiv
32 Swimming and Behavior in Purple Non-Sulfur Bacteria Judith P. Armitage
643–654
Summary I. Introduction II. Swimming III. Behavioral Responses Acknowledgments References
643 644 644 646 653 653
33 Metals and Metalloids in Photosynthetic Bacteria: Interactions, Resistance and Putative Homeostasis Revealed by Genome Analysis Francesca Borsetti, Pier Luigi Martelli, Rita Casadio and Davide Zannoni
655–689
Summary I. Introduction II. Classification of Metals and Metalloids by their Toxicity or Essentiality to Bacterial Cells (Groups I, II and III). III. Metal Toxicity, Tolerance and Resistance: Generalities IV. General Features on Microbial Metal Resistance/Tolerance Mechanisms V. On the Bacterial Interactions with Metals and Metalloids VI. Metal(loid)s Homeostasis in Phototrophs as Revealed by Genome Analysis VII. Concluding Remarks Acknowledgments References
656 656 656 657 658 666 675 679 682 682
Part 7: Genomics, Regulation and Signaling 34 Purple Bacterial Genomics 691–706 Madhusudan Choudhary, Chris Mackenzie, Timothy J. Donohue and Samuel Kaplan I. II. III. IV.
Introduction Genome Architecture and Characteristics Gene Homologs and Metabolic Versatility Variation in Transcriptional Regulation and Adaptation to Changing Environments V. Transposons and Genomic Rearrangements VI. Circadian Clock and Gas Vesicle Proteins VII. Inorganic Compounds as Reducing Power VIII. Genomic Insights into the Photosynthetic Lifestyle Acknowledgments References
xxv
692 692 696 701 702 702 703 703 704 704
35 Regulation of Gene Expression in Response to Oxygen Tension 707–725 Carl E. Bauer, Aaron Setterdahl, Jiang Wu and Brigitte R. Robinson Summary I. Introduction II. RegB/RegA Two-Component Signal Transduction System III. Aerobic repression by CrtJ IV. Regulation by Fnr Acknowledgments References
36 Regulation of Genes by Light Gabriele Klug and Shinji Masuda
707 707 708 716 721 722 722
727–741
Summary I. Introduction II. Photoreceptors in Purple Photosynthetic Bacteria III. Light-Dependent Responses that Do Not Depend on Photoreceptors IV. Concluding Remarks Acknowledgments References
37 Regulation of Hydrogenase Gene Expression Paulette M. Vignais
727 728 728 735 737 737 737
743–757
Summary I. Introduction II. Regulation of Hydrogenase Gene Expression: Signaling and Transcription Control References
38 Regulation of Nitrogen Fixation Bernd Masepohl and Robert G. Kranz
743 744 744 755
759–775
Summary I. Nitrogen Fixation in Purple Nonsulfur Bacteria II. Three Regulatory Levels of Nitrogen Fixation and Molecular Mechanisms Studied in Rhodobacter capsulatus III. Other Factors that Feed into the Nitrogen Regulatory Circuitry IV. Regulation in Other Purple Photosynthetic Bacteria V. Future Perspectives Acknowledgments References
xxvi
760 760 762 768 770 771 772 772
39 Regulation of the Tetrapyrrole Biosynthetic Pathway Jill Helen Zeilstra-Ryalls
777–798
Summary I. Introduction II. Tetrapyrrole Biosynthesis Genes III. Comparing and Contrasting Oxygen Control of Tetrapyrrole Biosynthesis Genes in Species of Rhodobacter IV. Other Aspects of Transcriptional Regulation of Tetrapyrrole Biosynthesis Genes in Rhodobacter Species V. A Genomics Perspective on the Regulation of Tetrapyrrole Biosynthesis in Other Purple Anoxygenic Photosynthetic Bacteria Note Added in Proof Acknowledgments References
40 Bacteriophytochromes Control Photosynthesis in Rhodopseudomonas palustris Katie Evans, Toni Georgiou, Theresa Hillon, Anthony Fordham-Skelton and Miroslav Papiz I. II.
777 778 778 779 789 789 795 795 795
799–809
Introduction Bacteriophytochrome Gene Organization and Regulation of Photosynthesis in Rhodopseudomonas palustris III. Phytochrome Domain Organization IV. Bilin Chromophore Photo-conversion V. Chromophore Binding Domain of Deinococcus radiodurans Bacteriophytochrome VI. Small Angle X-ray Scattering Solution Structure of Bph4 from Rhodopseudomonas palustris VII. Conclusions Acknowledgments References
800 800 803 804 805 805 807 807 807
41 Photoreceptor Proteins from Purple Bacteria 811–837 Johnny Hendriks, Michael A. van der Horst, Toh Kee Chua, Marcela Ávila Pérez, Luuk J. van Wilderen, Maxime T.A. Alexandre, Marie-Louise Groot, John T. M. Kennis and Klaas J. Hellingwerf Summary I. Introduction II. Light, Oxygen, or Voltage Domains III. The BLUF Domain Containing Family of Photoreceptors IV. Comparison Between LOV and BLUF Domains V. The Xanthopsins VI. Bacteriophytochromes VII. Concluding Remarks Acknowledgments References
xxvii
811 812 813 816 822 823 827 831 832 832
Part 8: New Applications and Techniques 42 Foreign Gene Expression in Photosynthetic Bacteria Philip D. Laible, Donna L. Mielke and Deborah K. Hanson Summary I. Introduction II. Design of a Rhodobacter-Based System for the Expression of Membrane Proteins for Structural and Functional Studies III. Production of Foreign Membrane Proteins in Rhodobacter IV. Optimization and Generalization of Heterologous Expression in Rhodobacter V. Advantages Afforded by Rhodobacter VI. Perspectives Acknowledgments References
43 Assembly of Bacterial Light-Harvesting Complexes on Solid Substrates Kouji Iida, Takehisa Dewa and Mamoru Nango Summary I. Introduction II. Atomic Force Microscopy Imaging of Reassociated Bacterial LightHarvesting Complex 1 on a Mica Substrate III. Conductivity of the Bacterial Reaction Center on Chemically Modified Gold Substrates Using Conductive Atomic Force Microscopy IV. Molecular Assembly of Photosynthetic Antenna Core Complex on an Amino-terminated Indium Tin Oxide Electrode V. Concluding Remarks Acknowledgments References
44 Optical Spectroscopy of Individual Light-Harvesting Complexes from Purple Bacteria Jürgen Köhler Summary I. Introduction II. The Experimental Setup III. Energy Transfer, Excitons, Strong and Weak Coupling IV. Spectroscopy of Single Light-Harvesting 2 Antenna Complexes V. Energy Transfer in a Single Photosynthetic Unit Acknowledgements References
xxviii
839–860 840 840 841 847 850 852 856 856 856
861–875 861 862 864 865 869 873 873 873
877–894 877 877 879 880 882 889 891 891
45 De novo Designed Bacteriochlorophyll-Binding Helix-Bundle Proteins Wolfgang Haehnel, Dror Noy and Hugo Scheer
895–912
Summary I. Introduction II. Chlorophyll Structures and Interactions with Natural Proteins III. Challenges in Designing de novo Chlorophyll- and Bacteriochlorophyllbinding Proteins IV. Modular Organized Chlorophyll Proteins Based on Branched Fourhelix Bundle Proteins V. Incorporating Chlorophylls and Bacteriochlorophylls into Self-assembling Protein Maquettes VI. From Water-soluble to Amphiphilic Chlorophyll- and Bacteriochlorophyllprotein Maquettes Acknowledgments References
46 Design and Assembly of Functional Light-Harvesting Complexes Paula Braun and Leszek Fiedor Summary I. Introduction II. Design of Model Light-Harvesting Proteins III. Assembly of Functional Light-Harvesting 1 Complexes IV. Conclusions and Prospects Acknowledgments References
895 896 897 899 901 904 905 907 907
913–940 914 914 916 924 935 935 936
47 The Supramolecular Assembly of the Photosynthetic Apparatus of Purple Bacteria Investigated by High-Resolution Atomic Force Microscopy 941–952 Simon Scheuring Summary I. Introduction II. Atomic Force Microscopy Analysis of the Complexes of the Bacterial Photosynthetic Apparatus III. Conclusions IV. Feasibilities, Limitations and Outlook Acknowledgments References
xxix
941 942 943 949 950 951 951
48 Protein Environments and Electron Transfer Processes Probed with High-Frequency ENDOR Oleg G. Poluektov and Lisa M. Utschig
953–973
Summary I. Introduction II. Low Temperature Interquinone Electron Transfer in the Photosynthetic Reaction Center. Characterization of QB– States III. Electron Transfer Pathways and Protein Response to Charge Separation IV. Concluding Remarks Acknowledgments References
Index
953 954 956 963 968 969 969
975–1013
xxx
Preface This book follows the tradition initiated in 1963, then subsequently extended in 1978 and 1995, of summarizing our knowledge of the photototrophic bacteria in a single volume. The first book was Bacterial Photosynthesis (Howard Gest, Anthony San Pietro and Leo P.Vernon, eds), 1963, Antioch Press, Yellow Springs, Ohio. Fifteen years later, Roderick K. Clayton and William R. Sistrom edited The Photosynthetic Bacteria (1978, Plenum Press, New York), an indispensable book for any scientist in the field at that time. In 1995, Robert Blankenship, Michael Madigan and Carl Bauer edited the most complete survey of the subject, Anoxygenic Photosynthetic Bacteria. By that time, the book had been taken under the wing of the Advances in Photosynthesis Series, initiated by Govindjee, as Volume 2 (Kluwer Academic Publishers, Dordrecht), and now, in 2008, we come to volume 28, entitled The Purple Phototrophic Bacteria (Springer, Dordrecht). The word “phototrophic” is used because it has become clear that many purple bacteria are incapable of photosynthesis (synthesis of all cellular carbon components from CO2), although they are capable of obtaining energy in the form of ATP from light (using light-harvesting and reaction center complexes that are homologous to those of purple photosynthetic bacteria). This latest survey of the field is restricted to the purple bacteria, and does not attempt to cover, for example, the green sulfur bacteria. There have been so many exciting developments since 1995 that the editors felt that it was sufficiently challenging to summarize thirteen years of research on the purple bacteria. This proved to be the case, and 48 chapters and more than 1000 pages were necessary to encompass the depth and breadth of the progress made in studying these fascinating organisms. Since 1995, there has been an explosion of information available from genome sequencing and related projects. The first 3-D structure of the bacterial cytochrome bc1 complex has been determined, and more structural information has been obtained for light-harvesting and reaction center membrane-protein complexes. Site-directed modifications of light-harvesting and reaction center complexes have been correlated with altered spectroscopic properties, even on a femtosecond timescale. Spectroscopic methods in general have advanced considerably since the last volume in 1995; an important example is the development of single
molecule approaches. New theoretical frameworks have had to keep pace with such technical developments. Through the use of atomic force microscopy it has been possible to examine the organization of clusters of individual light-harvesting and reaction center complexes in their native membranes. We are now beginning to see how the properties of native and modified photosynthetic complexes can be harnessed on the nanoscale for the design of biologically-inspired energy and electron transfer devices. In addition to these advances, we are reminded that many new phototrophic bacteria are still being discovered, and although it has been 45 years since the first book on these bacteria appeared, there is much that we do not know or understand. Recent publications indicate that purple phototrophic bacteria are ubiquitous on Earth, and raise questions about their contributions to global cycles of elements. Certainly, the extraordinary metabolic versatility of the purple bacteria, and their amenability to investigation by genetic, biochemical and biophysical techniques, will ensure that despite the inevitably cyclical and variable nature of science funding, there will always be compelling reasons to carry out research on the purple bacteria. Since 1995, it has become ever easier to obtain information online, and it could be argued that the need for a book on this topic is not as compelling as it might have been thirty years ago. However, there is still much value in having a hard copy of the diverse collection of information represented by the 48 chapters of this book, compiled at this point in time and which can be held in the hand. We have attempted to impart some coherence to this project, a process helped by using the organization of the 1995 volume as a starting point. The editors have adopted a pragmatic approach to the issue of taxonomy and, in the light of the ever-changing nature of specific bacterial names, a rigid policy of using only the most recent ones has not been enforced. Thus, Rhodopseudomonas acidophila is now Rhodoblastus acidophilus, for example, but the former name still predominates in the book. Helpful lists of purple phototrophic bacteria, as well as lists of genes, enzymes, pathways, and many more attributes of these bacteria, appear throughout this book. In addition, a section is included at the end of the present volume on new applications and techniques, with the hope that perhaps some of these will form the basis of a xxxi
fifth book, several years from now. This new volume is intended to be a resource for present and future researchers in the fields of ecology, microbiology, biochemistry and biophysics, some of whom might be interested in harnessing the potential of these bacteria as cell factories, or for bioremediation, nanotechnology or solar energy research. We hope that this book will help to attract a new generation of scientists to this field. We thank the authors of all the chapters for entering into the spirit of this project, which is intended to create a lasting work of reference, and a milestone in the field, a staging post on a journey that still has a long way to run.
We are grateful to various individuals who have offered advice, including Govindjee, (the Series Editor), John Golbeck (Editor of the book on Photosystem I, Volume 24 in the Series), Robert Blankenship (one of the Editors of Volume 2 in the Series) and Hugo Scheer (one of the Editors of Volume 25 that focused on Chlorophylls and Bacteriochlophylls), but above all we thank Larry Orr, whose guidance has underpinned the progress of this book. Finally the editors thank their respective families and members of their laboratories for their patience throughout the editing process. May 14, 2008 C. Neil Hunter Krebs Institute for Biomolecular Research Department of Molecular Biology and Biotechnology University of Sheffield Firth Court, Western Bank Sheffield S10 2TN U.K. Email: c.n.hunter@sheffield.ac.uk Fevzi Daldal Department of Biology University of Pennsylvania Philadelphia, PA 19104 U.S.A. Email:
[email protected] Marion C. Thurnauer Argonne National Laboratory Chemistry Division, E125 9700 S. Cass Avenue Argonne, Illinois 60439 U.S.A. Email:
[email protected] J. Thomas Beatty Dept. of Microbiology & Immunology University of British Columbia Rm. 4556, 2350 Health Sciences Mall Vancouver, BC, V6T 1Z3 Canada Email:
[email protected] xxxii
C. Neil Hunter was born in 1954 in Yorkshire, U.K., and is a Professor in the Department of Molecular Biology and Biotechnology at the University of Sheffield, U.K. He obtained his B.Sc. Degree at the University of Leicester in 1975, where the lectures of Professor Hans Kornberg and a laboratory project with Professor Peter Henderson inspired a lifelong interest in metabolism and bioenergetics. Wanting to learn about photosynthesis, Hunter moved to Bristol University, where the laboratories of Professors Trevor Griffiths, Owen Jones and Tony Crofts were studying many aspects of this topic, from pigment biosynthesis through to electron transport. Hunter studied for his Ph.D. under the guidance of Professor Owen Jones on the topic of membrane assembly in bacterial photosynthesis, and a sabbatical visit of Professor Robert Niederman to the Jones laboratory in Bristol led, in 1978, to a postdoctoral fellowship at Rutgers University, New Jersey. Hunter’s interests in light-harvesting complexes and membrane assembly developed further at this stage, but research in this
area was hampered by a lack of molecular genetic tools to study Rhodobacter sphaeroides. After returning to Bristol to learn about molecular biology in the laboratory of Professor Geoff Turner, it was possible eventually to use transposon Tn5 mutagenesis to gain access to the genes encoding the enzymes for bacteriochlorophyll and carotenoid biosynthesis and to develop a toolkit for site directed mutagenesis of photosynthetic complexes. In 1984, Hunter was appointed to a Lectureship at Imperial College, London, attached to the group of Professor James Barber. He returned to his native Yorkshire in 1988 to a Senior Lectureship at Sheffield University. In 1996 Hunter was awarded a D. Sc. by Bristol University. He is now the Krebs Professor of Biochemistry at Sheffield, and continues to apply a combination of molecular genetic, biochemical, structural and spectroscopic approaches to dissect the pathways for bacteriochlorophyll and carotenoid biosynthesis, and to investigate the assembly, structure, function and organization of bacterial photosynthetic membranes.
xxxiii
Fevzi Daldal is a Professor in the Department of Biology, University of Pennsylvania. He obtained his BS/MS degrees in 1974 at the Institut National des Sciences Appliquées (INSA) de Lyon, France, where he was first introduced into bacterial research in François Stoeber’s group. He followed his graduate studies on the genetics of Escherichia coli cell division at the Université Louis Pasteur, Strasbourg, France, under the guidance of Drs. Raymond Minck (Director of the Microbiology Institute ) and Maxime Schwartz (who was at the Pasteur Institute in Paris, France), obtaining his doctorate in 1977. From 1969 to 1977, his undergraduate and graduate studies were supported by scholarships from the French Government. In 1978, he joined, as a postdoctoral fellow, Dr. Dan G. Fraenkel at Harvard Medical School, Microbiology and Molecular Genetics Department to work on E. coli intermediary metabolism and on the phosphofructokinase II enzyme. After joining the Cold Spring Harbor Laboratory as a Scientist in 1983, Daldal started his studies with purple non sulfur bacteria Rhodobacter species, focusing on cy-
xxxiv
tochromes, which led him to discover the membrane attached electron carrier cytochrome cy, to co-discover the cytochrome cbb3 oxidase as well as novel genes involved in cytochrome biogenesis. His main work on the cytochrome bc1 complex included many studies, extending from the isolation of the structural genes to the crystallization of the enzyme complex to address detailed structure-function aspects using molecular genetics, biochemistry and biophysical approaches with Rhodobacter. Daldal’s current research is centered on the structure, function, regulation, proteomics and biogenesis of membrane cytochromes in Rhodobacter species. His contributions have been recognized by his election as a Member to the Turkish Academy of Science (2000), the American Academy of Microbiology (2005), the American Association for the Advancement of Science (2006) and Chair of the Gordon Conference on Molecular and Cellular Bioenergetics (2007). Daldal was an Editor of the Archives of Microbiology (2000–2006), and currently is a member of the Editorial Board of the Journal of Bacteriology, and a reviewer for many journals.
Marion C. Thurnauer received a B.A.(1968), M.S.(1969),and Ph.D. (1974) in Chemistry, all from the University of Chicago, Illinois, USA. Her thesis research, under the guidance of Gerhard Closs, was in the area of photochemistry; in particular, she studied magnetic interactions in radical pairs using electron paramagnetic resonance (EPR) spectroscopy. In 1974, she became ‘hooked’ on photosynthesis research after accepting a postdoctoral position at the Argonne National Laboratory (ANL), Lemont, IL, USA, to study magnetic resonance properties of excited triplet states of chlorophylls in photosynthetic units and in low temperature solutions. During that period, she and her mentors James R. Norris and Joseph J. Katz demonstrated that the unique electron spin polarization observed in the EPR spectrum of the triplet state of the primary electron donor in the reaction center was due to radical pair induced intersystem crossing. In 1977, she was promoted to staff member at ANL where she served as Group Leader of the Photosynthesis Research Group (1993–1995) and Director of the ANL Chemistry Division (1995–2003). Currently, she is Argonne Distinguished Fellow, Emeritus. Her research involved studies of sequential electron transfer in natural photosynthetic systems of photosynthetic bacteria and oxygenic photosynthesis, using primarily time-resolved EPR methods. She and her
colleagues demonstrated the spin correlated radical pair nature of the transient oxidized primary electron donor and reduced quinone acceptor in photosystem I, and, by comparison in purple photosynthetic bacteria. Her work (together with her colleagues) also extended to development of time-resolved magnetic resonance techniques, particularly for application to study photochemical energy conversion. Collaborating with Tijana Rajh, her research also included EPR studies of photoexcited surface modified nanocrystalline metal oxide colloids to mimic the energy transduction of natural photosynthesis. She served as Chair of the Gordon Research Conference on Photosynthesis: Biophysical Aspects (1994); she has organized several symposia and workshops on magnetic resonance and photosynthesis. During 2002–2003, she served on the Editorial Board of Biophysical Journal. In 2002, she was awarded the 2002 Francis P. Garvan-John M. Olin Medal by the American Chemical Society. Other honors include: the University of Chicago Award for Distinguished Performance at Argonne in 1991; the Agnes Fay Morgan Research Award in 1987; elected as a Fellow of the American Association for the Advancement of Science in 1998. She was the first recipient of the University of Chicago-Argonne Pinnacle of Education Award in 2007.
xxxv
J. Thomas Beatty is a Professor in the Department of Microbiology and Immunology at the University of British Columbia, Canada. He obtained the B.S. degree at the University of Washington in 1976, where a research project under the supervision of James T. Staley sparked Beatty’s interest in phototrophic bacteria. Beatty went on to graduate studies (M.A. in 1978; Ph.D. in 1980) under the guidance of Howard Gest at Indiana University, Bloomington, IN, USA, where he studied the metabolism of purple and green bacteria. Beatty’s postdoctoral research (1980–1983), in Stanley N. Cohen’s laboratory in the Department of Genetics, at the Stanford University School of Medicine, California, USA, included the discovery of differential degradation of light-harvesting 1 (LH1) and reaction center (RC) mRNA segments in the purple bacterium Rhodobacter capsulatus as a process that underlies the relative amounts of LH1 and RC complexes in the cell membrane. After taking up a faculty position at the University of British Columbia in 1983, Beatty has contributed to diverse areas of photosynthesis research: transcrip-
xxxvi
tional regulation of photosynthesis gene expression and elucidation of ‘superoperons’; discovery of the PufX protein and its role in quinone translocation as part of the RC/LH1/PufX core complex; isolation and characterization of new species of phototrophic bacteria; mechanisms of proton translocation into the RC; assembly of photosynthetic complexes; genomics and proteomics of purple bacteria. Beatty’s current research continues on the problems mentioned above, as well as on the ‘gene transfer agent’ (GTA) of Rba. capsulatus. His contributions have been recognized by a prize from the American Society for Microbiology (2002) and a Killam Research Fellowship from the Canada Council for the Arts (2008); he has been featured in popular publications such as the National Geographic and in several newspaper articles. Beatty was an Editor of FEMS Microbiology Letters, and a member of the Editorial Boards of Applied and Environmental Microbiology and the Journal of Bacteriology for many years. He is at present an Editor of Photosynthesis Research.
Author Index Alexandre, Maxime T. A. 811–837 Allen, James P. 337–353 Armitage, Judith P. 643–654 Axelrod, Herbert 323–336
Hillon, Theresa 799–809 Hosler, Jonathan 537–561 Huang, Li-Shar 425–450 Hunter, C. Neil 155–179
Bauer, Carl E. 707–725 Benning, Christoph 119–134 Berry, Edward A. 425–450 Blankenship, Robert E. 17–29 Borsetti, Francesca 655–689 Braun, Paula 913–940 Bullough, Per A. 155–179
Iida, Kouji 861–875
Casadio, Rita 655–689 Choudhary, Madhusudan 691–706 Chua, Toh Kee 811–837 Cogdell, Richard J. 135–153 Cooley, Jason W. 451–473 Csotony, Julius T. 31–55
Kaplan, Samuel 691–706 Kennis, John T. M. 811–837 Klug, Gabriele 727–741 Koch, Hans Georg 407–423 Köhler, Jürgen 877–894 Kramer, David M. 451–473 Kranz, Robert G. 407–423; 759–775 Kriegel, Alison M. 57–79
Dahl, Christiane 595–622 Daldal, Fevzi 407–423; 425–450 Deery, Evelyne 81–95 Dewa, Takehisa 861–875 Donohue, Timothy J. 691–706
Jackson, J. Baz 495–493 Joliot, Pierre 509–536 Jones, Michael R. 295–321 Jung, Deborah O. 1–15 Junge, Wolfgang 475–493
Laible, Philip D. 839–860 Lavergne, Jérôme 509–536 Lee, Dong-Woo 425–450 Loach, Paul A. 181–198
Evans, Katie 799–809 Feniouk, Boris A. 475–493 Fiedor, Leszek 913–940 Fordham-Skelton, Anthony 799–809 Frank, Harry A. 213–230 Frawley, Elaine R. 407–423 Gabrielsen, Mads 135–153 Gardiner, Alastair T. 135–153 Georgiou, Toni 799–809 Groot, Marie-Louise 811–837 Gunner, Marilyn R. 379–405 Haehnel, Wolfgang 895–912 Hanson, Deborah K. 839–860 Harwood, Caroline S. 577–594 Hellingwerf, Klaas J. 811–837 Hendriks, Johnny 811–837
Mackenzie, Chris 691–706 Madigan, Michael T. 1–15 Martelli, Pier Luigi 655–689 Masepohl, Bernd 759–775 Masuda, Shinji 727–741 Mielke, Donna L. 839–860 Miyashita, Osamu 323–336 Nango, Mamoru 861–875 Niederman, Robert A. 253–273 Nitschke, Wolfgang 451–473 Novoderezhkin, Vladimir I. 231–252 Noy, Dror 895–912 Obiozo, U. Mirian 495–493 Okamura, Melvin 323–336 Onder, Ozlem 407–423
xxxvii
Papiz, Miroslav 799–809 Parkes-Loach, Pamela S. 181–198 Parson, William W. 355–377 Pérez, Marcela Ávila 811–837 Polívka, Tomáš 213–230 Poluektov, Oleg G. 953–973
Tabita, F. Robert 563–576 Takaichi, Shinichi 97–117 Tamot, Banita 119–134 Turkarslan, Serdar 407–423
Qian, Pu 155–179
van Grondelle, Rienk 231–252 van der Horst, Michael A. 811–837 van Wilderen, Luuk J. 811–837 Verméglio, André 509–536 Vignais, Paulette M. 743–757
Raymond, Jason 17–29 Robert, Bruno 199–212 Robinson, Brigitte R. 707–725 Romagnoli, Simona 563–576 Sander, Johannes 595–622 Sanders, Carsten 407–423 Scheer, Hugo 895–912 Scheuring, Simon 941–952 Schoepp-Cothenet, Barbara 537–561 Schulten, Klaus 275–294 Şener, Melih K. 275–294 Setterdahl, Aaron 707–725 Shapleigh, James P. 623–642 Sturgis, James N. 253–273 Swingley, Wesley D. 17–29
Utschig, Lisa M. 953–973
Warren, Martin J. 81–95 Warshel, Arieh 355–377 Williams, JoAnn C. 337–353 Willows, Robert D. 57–79 Wraight, Colin A. 379–405 Wu, Jiang 707–725 Yurkov, Vladimir 31–55 Zannoni, Davide 537–561; 655–689 Zeilstra-Ryalls, Jill Helen 777–798
xxxviii
Color Plates
Fig. 1. Schematic comparison of photosynthesis gene clusters (PGC) from various purple bacteria. Lines represent transpositions of syntenous gene regions between two PGC. Numbers annotated above the Rhodospirillum rubrum PGC represent the location on the genome of each separate PGC segment. Genes colored in green represent BChl biosynthesis genes, orange represent carotenoid biosynthesis genes, red represent structural genes of the photosynthetic apparatus, blue represent regulatory genes and other colors represent unique genes. The black and blue lines with arrows represent inversion of the genes contained between the lines and the red lines represent shifts in position without inversions. See Chapter 2, p. 20.
C. Neil Hunter, Fevzi Daldal, Marion C. Thurnauer and J. Thomas Beatty (eds): The Purple Phototrophic Bacteria, pp. CP1–CP16. © 2009 Springer Science + Business Media B.V.
Color Plates
Fig. 2. Photosynthetic and habitat diversity of aerobic anoxygenic phototrophs (AAP) A–E. In vivo absorbance spectra of cells grown under different physiological conditions, with numerals above peaks denoting LH and RC wavelengths. Insets, photographs of liquid cultures. A. Roseicyclus mahoneyensis, illuminated (dashed line) and dark (solid line). B,C. Dashed line, rich organic medium (3 g·l–1 organics); solid line, minimal acetate (1 g·l–1) medium (B. Roseococcus thiosulfatophilus, C. Erythrobacter litoralis). D. Erythromicrobium ramosum, rich organic medium, dark (dashed line) and oligotrophic medium, light:dark regimen of 12h:12h, illuminated with diffuse ambient sunlight (solid line). E. Citromicrobium bathyomarinum strain JF1, rich organic medium (dashed line), minimal glucose medium (dotted line), compared with Erythrobacter litoralis (solid line). F. Hypothetical electron transfer system of aerobic anoxygenic photosynthesis, showing electron flow through major carriers: P870, special pair of BChl in RC (photoexcited state indicated by ‘*’); BChl-BPh, accessory BChl and bacteriophaeophytin in the RC; QA, quinone primary electron acceptor; Cyt bc1, cytochrome bc1 complex; Cyt c2, cytochrome c2. The symbol ‘+’ indicates that the midpoint potential of QA in all tested AAP is positive and higher than in anaerobic phototrophs. G–I. Extreme environment habitats of AAP. G. Hydrothermal vent field in Eastern Pacific Ocean, showing smoker chimney. H. Hypersaline spring system (East German Creek) in Manitoba, Canada: spring pool in foreground and playa in background, with white patches of salt precipitates. I. Cyanobacterial-Thiothrix mat development in thermal springs of Banff National Park, Alberta, Canada. See Chapter 3, p. 39.
CP2
Color Plates
Fig. 3. Representations of the β/α-protomer building blocks that comprise LH2. The β-chains are shown in blue and the α-chains in green. The BChls are shown in red and the carotenoid in pale blue. a) A view of the α/β-protomer with the pigments removed. The main BChl liganding amino acid side chains are shown. b) A view of how two adjacent β/α-protomers interact with each other, again with the pigments removed. c) Similar to b) but looking down onto the two adjacent α/β-protomers from the C-terminal side of the complex. d) The same view as a) but with the pigments included. e) A view of the α/β-protomer from Phaeospirillum (Phs.) molischianum. The main differences between d) and e) are the altered orientation of the B800 bacteriochlorin ring and the different organization of the C-terminal region of the α-apoprotein. f) A view of the α/β-protomer from Rps. acidophila strain 7050. Note the different amino acids shown at the C-terminal end of the α-apoprotein that are responsible for the altered absorption properties of this B800-820 complex. The structures shown in a–d are representations of the structure of the B800-850, LH2 complex from Rps. acidophila strain 10050. See Chapter 8, p. 138.
Fig. 4. The LH2 complex from Rhodopseudomonas (Rps.) acidophila strain 10050. a) A view looking down on the top of the complex from the presumed periplasmic surface of the membrane. (b) A side view looking from within the presumed photosynthetic membrane with the periplasmic side of the complex uppermost. In both panels the LH2α polypeptide is green, LH2β is blue, B850 BChls are purple, B800 BChls are brown and the carotenoids are red. See Chapter 8, p. 140.
CP3
Color Plates
Fig. 5. (A) Dynamics of the transient absorption (TA) spectrum measured upon 1017 nm excitation of the LH1 of Blc. viridis at 77 K (Monshouwer et al., 1998). (B) Measured (points) and calculated (solid lines) red-shift of the TA spectrum between 0 and 400 fs upon 1017 nm excitation of the LH1 of Blc. viridis at 77K (Novoderezhkin and van Grondelle, 2002). (C) Measured (points) and modeled (solid line) TA kinetics at 1050 nm traces upon 1055 nm excitation of the LH1 of Blc. viridis at 77K (Novoderezhkin et al., 2000). See Chapter 13, p. 240.
Fig. 6. Coherent dynamics of the density matrix for a single LH2 complex, i.e., calculated without relaxation of populations and coherences for one realization of the disorder corresponding to a blue-shifted (top), an intermediate (middle), and a red-shifted (bottom) FL spectrum (Novoderezhkin et al., 2006). The initial population corresponds to the thermal equilibrium at room temperature (shown in the inserts by blue, green, and red for the blue-shifted, intermediate, and red-shifted spectra, respectively), initial coherences have arbitrary fixed phases. The figure shows the diagonal elements of the density matrix in the site representation ρ(n,n) as a function of time. The color scale is used to indicate the absolute values of the density matrix from zero (blue) to the maximal value (red). See Chapter 13, p. 247.
CP4
Color Plates
Fig. 7. Structural models of monomeric and dimeric core complexes. In each case a side view and a space-filling representation is used. For the RC the complete structure is present (shown in blue), but for the LH1 (red) and helix W/PufX components (green) only the transmembrane helices are shown. a. The monomeric RC-LH1-W core complex from Rhodopseudomonas palustris (Roszak et al., 2003). b. The dimeric RC-LH1-PufX core complex of Rhodobacter (Rba.) sphaeroides. The coordinates of the LH1 αβ pairs were adopted from Roszak et al. (2003), fitted into the tilted 3-D dimer structure from single particle analysis (Chapter 9, Bullough et al.), and adjusted according to the cryo-EM projection map in Qian et al. (2005). The transmembrane domain of PufX was taken from Tunnicliffe et al. (2006). See Chapter 9, p. 172.
Fig. 8. Organization of the photosynthetic unit (PSU) from the supramolecular architecture to individual chlorophylls and the energy transfer network. The high-light adapted vesicle model for Rba. sphaeroides (Şener et al., 2007) is shown. The three different constituent proteins and their BChls are colored as follows: LH2: green, LH1: red, RC: blue. From left to right three different rendering styles are featured. On the left, the light-harvesting proteins are shown in the tube representation (backbone only, no pigments); in the middle, the proteins are rendered transparently while the BChls are visible represented by their porphyrin rings; on the right, only the BChls are shown but this time together with their respective electronic couplings (Eq. 1). For simplicity only the strongest couplings are shown. The vesicle construction depicted here is based on atomic force microscopy data on intracytoplasmic membranes (Bahatyrova et al., 2004). Figure made with VMD (Humphrey et al., 1996). See Chapter 15, p. 288.
CP5
Color Plates
Fig. 9. Structure of the Cyt c2 : RC complex. A) Side view of the complex. The Cyt c2 is bound with heme positioned directly over the BChl2. A small region of close contact is indicated by the circle. A conformational change, indicated by the arrow, is due to crystal packing interactions with an adjacent bound Cyt. B) View of the structure in the region between the cofactors. The heme and BChl are separated by the planar aromatic ring of Tyr (L162). The closest distance between the conjugated rings in the cofactors is 14.2 Å as shown by the long arrow. A strong tunneling pathway between the two cofactors is indicated by the short arrows. C) Open book view of the interface between the complexes. The two proteins are separated by rotating around the center line. The central region of shortrange contacts (indicated by the circle) is surrounded by charged residues involved in long-range electrostatic interactions. Interacting residues are color coded. The contact between the heme (green) and the ‘hot spot’ Tyr L162 (green) is shown by the arrow. Hydrophobic residues Leu (M191), Val (M192) (black) on the RC interact with Phe (C102) on the Cyt (black). Hydrogen bonds (cyan) are formed between atoms on RC and Cyt. A cation-pi interaction is formed between Tyr (M295) and Arg (C32). Electrostatic interactions occur between negatively charged groups on the RC (red) and positively charged groups on the Cyt (violet). Other groups in van der Waals contact are shown in yellow. Modified from Axelrod et al. ( 2002). See Chapter 17, p. 326.
CP6
Color Plates
Fig. 10. The cytoplasmic surface of a model LM preparation of the Rba. sphaeroides RC. (a) The LM heterodimer is shown as a solid object, with the L- and M-polypeptides shown in yellow and white, respectively, and the quinones shown as spheres with green carbons and red oxygens. The view is along the symmetry axis, and parts of the QA quinone are visible from the outside of the complex. (b) View as in (a) but with amino acids overlaying the quinone sites shown in stick format, revealing the underlying quinones (carbons in yellow (L) or white (M), oxygens in red and nitrogens in blue). In both panels the dotted ovals show the approximate positions of the QA and QB quinones. See Chapter 16, p. 298.
Fig. 11. Region around the HB site in the wild-type and AM149W RC complexes from Rba. sphaeroides. The protein is shown as a solid object, with the L- and M-polypeptides shown in yellow and white, respectively, and bound waters shown in cyan. The BB and QB cofactors are shown as green and beige solid objects, respectively. In (a) the HB cofactor is shown as blue sticks and the phytol chain of PB as red sticks. In (b) the phytol chain of PB in the normal conformation is shown as red sticks and the phytol chain of PB in the alternate conformation is shown as orange sticks. See Chapter 16, p. 302.
CP7
Color Plates
Fig. 12. The reaction center (RC) complex from Rhodobacter sphaeroides comprises three subunits, a heterodimer of similar, but non-identical L (yellow) and M (blue) subunits, and subunit H (green), which caps LM on the cytosolic side of the membrane. The LM dimer binds all the cofactors, while subunit H stabilizes the structure and is involved in H+-ion uptake and transfer associated with electron transfer to the quinones. The L and M subunits and all associated cofactors are arranged around a quasi-2-fold rotational symmetry axis, normal to the plane of the membrane and passing through the primary donor (P), the special pair dimer of bacteriochlorophylls (BChl), and a ferrous (Fe2+) iron midway between the two quinones. Electron transfer proceeds from the excited singlet state of the primary donor (P*), via the A-branch of cofactors — monomer BChl (BA) and BPhe (HA), bound to the L subunit — to the primary quinone, QA, which is bound in a fold of the M subunit. From QA– the electron crosses the symmetry axis to the secondary quinone, QB, bound in a similar fold in the L subunit. (Figure prepared in VMD.) See Chapter 20, p. 381.
Fig. 13. The quinone binding sites — QA (top) and QB (bottom) — are related by the two-fold, rotational symmetry axis of the reaction center, which passes through the iron atom of the acceptor quinone complex. Both quinones are hydrogen bonded through the carbonyl oxygens — hydrogen bonds shown in purple for QA (top) and in green for QB (bottom). The binding sites are predominantly hydrophobic, with the notable exception of GluL212, AspL213 and SerL223 in the QB site, which constitute terminal components of the H+ delivery pathway to QB. Note that the orientations of the methoxy groups of QA and QB do not follow the two-fold rotational symmetry. (Figure prepared in VMD.) See Chapter 20, p. 385.
CP8
Color Plates
Fig. 14. The Ccm-system I components for c-type Cyt maturation. All components of the c-type Cyt maturation, except the thiol-disulfide oxidoreductase DsbA, are located in the cytoplasmic membrane. Both apoCyt c and heme follow different routes to the heme ligation core complex, composed of CcmI, CcmH and CcmF. Apocyt c is translocated via the Sec pathway; its cysteine thiols in the conserved CXXCH motif are first oxidized by the DsbA-DsbB pathway and then reduced by the Cyt c maturation specific CcdA-CcmG and/or CcmH thio-reductive pathway. CcmI is involved in delivering apoCyt c to the core heme ligation complex via its different domains. Heme is translocated across the membrane, possibly via ABC–type transporter CcmABCD and is covalently attached to the conserved His residue of the heme chaperone CcmE. CcmC is involved in attaching heme to CcmE and CcmD enhances holo-CcmE production. CcmA and CcmB promote release of holo-CcmE from CcmC and CcmD. Upon formation of the thioether bonds between the apoCyt c and the heme vinyls, catalyzed by the CcmH-CcmI-CcmF complex, mature holoCyt c is released. See Chapter 21, p. 410.
Fig. 15. The Ccs-system II components for c-type Cyt maturation. Similarly to system I, all components of the system II c-type Cyt maturation pathway are located in the cytoplasmic membrane, with the exception of DsbA. The apoCyt c is translocated across the membrane by the Sec machinery and is oxidized by DsbA, then reduced by CcsX. The transmembrane proteins CcsB and CcsA form a complex that represents the system II synthetase (heme ligation core complex). The reduced apoCyt c is likely bound by the periplasmic domain of CcsB. Heme is translocated by the CcsBA complex, probably through CcsA and binds to the CcsA periplasmic WWD domain before ligation to the reduced apoCyt c. Following covalent ligation of the heme vinyl groups to the reduced cysteines of the apoCyt c CXXCH motif, the holoCyt c folds into its mature form with release from the complex. See Chapter 21, p. 412.
CP9
Color Plates
Fig. 16. Overall structure of the Rhodobacter (Rba.) capsulatus Cyt bc1. Dimeric Cyt bc1 is depicted in the membrane, indicating the likely positions of the boundaries of the membrane lipid bilayer (A). The lower panels show a monomeric Cyt bc1 with the Fe/S protein ED in the b (B) or c1 (C) positions. The substrate Cyt c2 (1C2R.pdb) is modeled into its likely reactive position based on the yeast co-crystal structure (1KYO.pdb). The regions of the Cyt bc1 structure that differ significantly from the mitochondrial enzyme are colored in red, otherwise Cyt b is shown in different shades of blue, Cyt c1 in green, the Fe/S protein in yellow, and Cyt c2 in magenta. Hemes are shown in red, and the [2Fe2S] cluster is shown as a spacefilled structure in yellow and orange. The Qo site inhibitor stigmatellin locks the Fe/S protein ED in the b position. See Chapter 22, p. 427.
Fig. 17. Stereoview of structural details of Q sites in the Rba. capsulatus Cyt bc1. Panel A, helices α-a, A, D, and E making the Qi pocket are colored in light pink, light pink, wine, and green, respectively. Heme bH is shown in gray and quinone bound to the Qi site in yellow as ball-and-stick models. Panels B and C, the surface of Cyt b, showing interaction of the Fe/S protein ED with the ‘binding crater’ at the Qo site when the ED is in the ‘b’ (B) or ‘c1’ (C) positions. Cyt b is depicted using space-filling representation with different regions colored as described below. The white outline demarks the position of the Fe/S protein ED. Four regions surrounding the binding crater, the cd1-cd2 helices, ef-loop, gh loop and α-ef2 portion are indicated in green, magenta, yellow and cyan, respectively. At the bottom-center of the binding crater stigmatellin is shown in red. The side chains of the H135 and H156 residues of the Fe/S protein subunit are shown as ball-and-stick models. Atoms of Cyt b that are in contact (i. e., less than 3.9 Å) with the Fe/S protein ED are colored in gray, and Y302 (light green, with gray OH atom) in the α-ef helix and K329 (lime green with gray Ce and Nz atoms) between the α-ef 2 and F helices are indicated by arrows. See Chapter 22, p. 434.
CP10
Color Plates
Fig. 18. Similarities and differences in the electron transfer chains and core structures of the cyt bc1 and cyt b6 f complexes. The gross functional monomeric structures of the cyt b (blue), cyt c (red) and the ISP (yellow) coordinating portions of the cyt bc1- (left) or cyt b6 ftype (right) complexes are illustrated side-by-side as matching ribbon structures with the cofactors coordinated in each drawn as space filled spheres (colored as the individual subunits to which they belong). The glaring differences of the presence of an extra c-type heme (heme ci) adjacent to the Qi (or QN) site of the cyt b6 f, as well as the presence of the non-redox active chlorophyll molecule (dark green) in this same complex, are readily seen in these side-by-side illustrations. However, in each case, a similar high (red) and low potential (blue) chain of cofactors facilitating bifurcated electron flow following QH2 oxidation is easily envisioned. Both types of complex are shown with the inhibitor, stigmatellin (light green), bound at the Qo (or Qp) site as well as amino acids, pictured as stick models (blue), thought to be important for Qi (or QN) site substrate binding to aid in visually identifying the two spatially distinct Q binding sites. The illustrations have been worked up from the Rhodobacter (Rba.) capsulatus and Mastigocladus laminosus derived atomic coordinate files 1ZRT.pdb and 1VF5.pdb, respectively. See Chapter 23, p. 454.
CP11
Color Plates
Fig. 19. Side reactions that bypass the Q-cycle. Reactions shown are those that differ from the Q-cycle in the fate of the SQ generated at the Qo site. Details are given in the text. See Chapter 23, p. 461.
(a)
(b)
Fig. 20. (a) The hydride-transfer site in 2OO5.pdb. The dI(B) component is shown in blue and dIII in green. The nucleotides are in standard atom colors (except that the C4 atoms are in yellow and are linked by the pink dashed line). H-bonds are show as green dashed lines. (b) The loop E ‘lid’ of dIII and the ‘mobile loop’ of dI closed down over the hydride transfer site of 2OO5.pdb. The nucleotides (NADP+ in dIII, and the NADH analog, H2NADH, in dI) are shown in a space-filling format. The dI(B) polypeptide is shown in green (with its mobile loop in cyan), and the dI(A) polypeptide in yellow. The dIII polypeptide is shown in pink (with its loop E in magenta). Also shown are the RQD loop of dI and helix D/loop D of dIII (see text). See Chapter 25, p. 504.
CP12
Color Plates
Fig. 21. In response to a short flash of light photosynthetic reaction centers generate transmembrane voltage and release quinol (QH2); subsequent oxidation of QH2 by the cytochrome bc1 complex is coupled to proton pumping into the chromatophore lumen. The resultant ~ + drives protons through the ATP synthase and powers ATP synthesis. See Chapter 24 (inset from Fig 3A), page 483. ∆µ H
CP13
Color Plates (i) (a)
(ii)
(iii)
(b)
(c)
(i)
(ii)
Fig. 22. Localization of cytoplasmic chemosensory proteins and loss of localization and position in PpfA mutant. (a) Rba. sphaeroides with a soluble chemoreceptor TlpC-GFP fusion and McpG-GFP, expressed from genomic replacements. (b) Fluorescence images of cells of Rba. sphaeroides expressing both (i) CheW2-YFP and (ii) CheW4-CFP from genomic replacements showing separation of the two chemotaxis pathways; (iii) overlay image. (c)(i) Cephalexin treated Rba. sphaeroides with genomically expressed TlpT-CFP showing localization at single cell distances through the filament. (ii) Cephalexin treated Rba. sphaeroides ∆PpfA showing loss of localization of the chemosensory clusters. The scale bars are 2 µM. See Chapter 32, p. 650.
Fig. 23. BLUF photocycle and active site structure. Panel a: active site receptor state. Panel b: active site signaling state. Panel c: typical photocycle of BLUF-domains. Panels a and b were prepared using the program PyMOL (http://www.pymol.org). Color coding for panels a and b: Grey, Carbon; Blue, Nitrogen; Red, Oxygen; Orange, Phosphorus; Green, hydrogen bond.The structure coordinate file of the BLUF domain of AppA from Rba. sphaeroides (PDB ID: 1YRX) (Anderson et al., 2005) was used to create both panels. Note that for panel b the positions of the side-chain nitrogen and oxygen of Q63 were manually switched to indicate the Q-flip. See Chapter 41, p. 818.
CP14
Color Plates
Fig. 24. The X-ray structure of the chromophore binding domain (CBD) from Deinococcus (D.) radiodurans (Wagner et al., 2005). The PAS-2 (blue) and GAF (yellow) domains are aligned, relative to one another, by a trefoil knot formed from a loop of the GAF domain (yellow) and residues 1-35 (green) N-terminal to PAS-2. Biliverdin IXα (BV-purple) is covalently bound to Cys24 and is located in a pocket within the GAF domain formed from a β-sheet and two α-helices. See Chapter 40, p. 806.
Fig. 25. A small angle X-ray scattering (SAXS) ab-initio envelope (transparent orange) of Bph4 from Rhodopseudomonas palustris (Evans et al., 2006). The structure is a homodimer and is modelled with the crystal structures of the CBD (green) (Wagner et al., 2005) and histidine kinase (HK, cyan) (Marina et al., 2005). The positions of the BV (purple), phosphorylating histidine (red) and ATP (orange) are shown. There is no high resolution structure of a PHY domain that can be fitted, but it is assumed to occupy the envelope between CBD and the HK domains. See Chapter 40, p. 807.
CP15
Color Plates
Fig. 26. Top:Helical net representation of shielding helices Si and binding helices Bj. The amino acids that were varied (Xk for Si and Zl for Bj) are emphasized in gray, the ligating histidine of the binding helices in green. The tables contain the amino acids of the individual helices at the varied positions. Long, solid arrows indicate the N→C direction of the peptide chains, dashed arrows the dipole moments of salt bridges. Bottom: photograph of a cellulose membrane carrying 38 modular proteins, after incubation with Ni-BPheide in buffer/ DMF and washing in buffer. Columns 2-7 correspond to identical binding helices Bj (j = 7 – 12), rows 1 - 6 to identical shielding helices Si (i = 1 - 6). The empty spots in columns 1 and 8 correspond to modular proteins that have been punched out for mass spectrometry; the two isolated spots in the rightmost column were not used for the analyses. See Chapter 45, p. 902.
Fig. 27. Stages of LH1 formation in the presence of carotenoids in vitro. See Chapter 46, page 934.
CP16
Chapter 1 An Overview of Purple Bacteria: Systematics, Physiology, and Habitats Michael T. Madigan* and Deborah O. Jung Department of Microbiology, Southern Illinois University, Carbondale, IL 62901, U.S.A.
Summary .................................................................................................................................................................. 2 I. Introduction......................................................................................................................................................... 2 II. Systematics of Purple Bacteria .......................................................................................................................... 3 A. Purple Sulfur Bacteria .......................................................................................................................... 4 B. Purple Nonsulfur Bacteria .................................................................................................................... 4 III. Physiology of Purple Bacteria ............................................................................................................................ 4 A. Purple Sulfur Bacteria .......................................................................................................................... 4 B. Purple Nonsulfur Bacteria .................................................................................................................... 6 1. Photoheterotrophy....................................................................................................................... 6 2. Dark Growth ................................................................................................................................ 7 3. Nitrogen Fixation ......................................................................................................................... 7 IV. Habitats of Purple Bacteria................................................................................................................................. 7 A. Purple Sulfur Bacteria .......................................................................................................................... 7 1. Blooms in Stratified Lakes........................................................................................................... 7 2. Microbial Mats ............................................................................................................................. 8 B. Purple Nonsulfur Bacteria .................................................................................................................... 9 1. Sewage ....................................................................................................................................... 9 2. Purple Nonsulfur Bacteria in Waste Lagoons ............................................................................. 9 V. Purple Bacteria in Extreme Environments.......................................................................................................... 9 A. Thermophilic Purple Bacteria ............................................................................................................. 10 1. Thermochromatium tepidum .................................................................................................... 10 2. Other Thermophilic Purple Bacteria .......................................................................................... 10 B. Halophilic and Alkaliphilic Purple Bacteria ......................................................................................... 10 C. Acidophilic Purple Bacteria ................................................................................................................ 11 D. Purple Bacteria from Permanently Cold Habitats............................................................................... 11 E. Environmental Limits to Photosynthesis in Purple Bacteria ............................................................... 12 VI. Final Remarks .................................................................................................................................................. 12 Acknowledgments ................................................................................................................................................... 12 References .............................................................................................................................................................. 12
*Author for correspondence, email:
[email protected] C. Neil Hunter, Fevzi Daldal, Marion C. Thurnauer and J. Thomas Beatty (eds): The Purple Phototrophic Bacteria, pp. 1–15. © 2009 Springer Science + Business Media B.V.
2
Michael T. Madigan and Deborah O. Jung
Summary Anoxygenic phototrophic purple bacteria are a major group of photosynthetic microorganisms widely distributed in nature, primarily in aquatic habitats. Nearly 50 genera of these organisms are known and some have become prime model systems for the experimental dissection of photosynthesis. Purple sulfur bacteria differ from purple nonsulfur bacteria on both metabolic and phylogenetic grounds, but species of the two major groups often coexist in illuminated anoxic habitats in nature. Purple sulfur bacteria are strong photoautotrophs and capable of limited photoheterotrophy, but they are poorly equipped for metabolism and growth in the dark. By contrast, purple nonsulfur bacteria, nature’s preeminent photoheterotrophs, are capable of photoautotrophy, and possess diverse capacities for dark metabolism and growth. Several purple bacteria inhabit extreme environments, including extremes of temperature, pH, and salinity. Collectively, purple bacteria are important phototrophs because they (1) consume a toxic substance, H2S, and contribute organic matter to anoxic environments by their autotrophic capacities; (2) consume organic compounds, primarily non-fermentable organic compounds, in their roles as photoheterotrophs; and (3) offer scientists in the photosynthesis community a smörgasbord of molecular diversity for the study of photosynthesis. I. Introduction Anoxygenic phototrophic purple bacteria are a major group of phototrophic microorganisms that inhabit aquatic and terrestrial environments. Purple bacteria that inhabit oxic habitats and which carry out photosynthesis only aerobically are called ‘aerobic anoxygenic phototrophs’ and are covered in Chapter 3 (Yurkov and Csotonyi). The current chapter covers only the classical purple bacteria: purple sulfur bacteria and purple nonsulfur bacteria. Purple bacteria are photosynthetic gram-negative prokaryotes that convert light energy into chemical energy by the process of anoxygenic photosynthesis. Purple bacteria contain photosynthetic pigments–bacteriochlorophylls and carotenoids — and can grow autotrophically with CO2 as sole carbon source. Many genera of purple bacteria are known and the organisms share many basic properties with their nonphototrophic relatives. Some general characteristics of purple bacteria are listed in Table 1. Purple bacteria share with oxygenic phototrophic prokaryotes — the cyanobacteria — the ability to conserve energy by photophosphorylation. However, unlike cyanobacteria and aerobic anoxygenic phototrophs, photosynthesis in purple bacteria only occurs under anoxic (O2-free) conditions. This is also true of the other classical anoxygenic phototrophs: green sulfur bacteria, green nonsulfur bacteria, and Abbreviations: BChl – bacteriochlorophyll; LH – light-harvesting; Rba. – Rhodobacter; Rcy. – Rhodocyclus; Rfx. – Rhodoferax; Rps. – Rhodopseudomonas; Rsp. – Rhodospirillum; Tch. – Thermochromatium
the heliobacteria (Blankenship et al., 1995). Purple bacteria require anoxic conditions for phototrophic growth because pigment synthesis in these organisms is repressed by molecular oxygen (Cohen-Bazire et al., 1957). Thus, the competitive success of purple bacteria in nature requires both light and anoxic conditions. This combination is most commonly found in lakes, ponds, estuaries, and other aquatic environments where H2S is present (Pfennig, 1967, 1978a, 1989). Once these general conditions are met, the exact physiochemical nature of the habitat (sulfide concentration, pH, light quality and intensity, temperature) controls the abundance and diversity of purple bacteria that develop there (Pfennig, 1978a, 1989; Madigan, 1988). Purple bacteria participate in the anoxic cycling of carbon both as primary producers (CO2 fixation, photoautotrophy) and as light-stimulated consumers of reduced organic compounds (photoheterotrophy). In certain habitats particularly favorable for their development, purple bacteria have been shown to be significant primary producers (Czeczuga, 1968; Takahashi and Ichimura, 1968; Overmann et al., 1994, 1996, 1999). However, in most illuminated sulfidic habitats the role of purple bacteria as H2S consumers is probably more important than any contribution they make to primary production; H2S is a highly poisonous substance for plants and animals and also for many bacteria. The oxidation of sulfide by purple bacteria yields nontoxic forms of sulfur, such as elemental sulfur (S0) and sulfate (SO42–). Sulfide oxidation thus allows the upper waters of a lake to remain oxic and suitable for plants, animals, and aerobic bacteria.
Chapter 1
Overview of Purple Bacteria
3
Table 1. General properties of anoxygenic purple phototrophic bacteriaa Property
Examples
Groups/phylogeny
Purple sulfur bacteria (gammaproteobacteria); purple nonsulfur bacteria (alphaor betaproteobacteria) Allochromatium vinosum and Thiocapsa roseopersicina (purple sulfur bacteria); Rhodobacter capsulatus, Rhodobacter sphaeroides, Rhodospirillum rubrum, and Rhodopseudomonas palustris (purple nonsulfur bacteria)
Major species studied
Pigments/color of dense cell suspensions
BChl a or b; major carotenoids include spirilloxanthin, spheroidene, lycopene, and rhodopsin, and their derivatives; cell suspensions purple, purple-red, purpleviolet, red, orange, brown, or yellow brown (BChl a-containing species); green or yellow (BChl b-containing species)
Location of pigments in cells
Within intracytoplasmic membranes arranged as membrane vesicles, tubes, bundled tubes, or in stacks resembling lamellae
Absorption maxima of living cells
BChl a-containing species: near 800 nm, and anywhere from 815–960 nm; BChl b-containing species: 835–850 nm and 1010–1040 nm
Electron donors/sulfur globulesb
H2S, S0, S2O32–, H2, Fe2+; if S0 is produced from the oxidation of sulfide, the S0 is stored intracellularly only in certain purple sulfur bacteria (see Fig. 1a)
Photoheterotrophy/dark respiratory growth
Purple sulfur bacteria limited on both accounts; purple nonsulfur bacteria typically diverse on both accounts
a
All purple bacteria are gram-negative prokaryotes. All species contain peptidoglycan and an outer membrane containing lipopolysaccharide. bVirtually all purple bacteria are capable of autotrophic growth. When growing autotrophically, the Calvin cycle (reductive pentose phosphate cycle) is used as the mechanism for CO2 fixation
II. Systematics of Purple Bacteria Purple sulfur bacteria and purple nonsulfur bacteria were originally distinguished on physiological grounds based on their tolerance and utilization of sulfide. Purple sulfur bacteria were species that tolerated millimolar levels of sulfide and oxidized sulfide to sulfur globules stored intracellularly (Fig. 1a), while purple nonsulfur bacteria were species that did neither (van Niel, 1932, 1944). However, classic chemostat experiments by Hansen and van Gemerden (1972) showed that these criteria for classifying purple bacteria were not absolute. At low levels of sulfide, typically less than 0.5 mM, most species of purple nonsulfur bacteria will grow and in so doing, oxidize sulfide to S0, S4O62–, or SO42–. Nevertheless, an important distinction in the sulfide metabolism of purple sulfur and purple nonsulfur bacteria remains: any S0 formed by purple nonsulfur bacteria is not stored intracellularly, but instead is deposited outside the cell (Hansen and van Gemerden, 1972; Brune, 1995) (species of Ectothiorhodospiraceae are an exception here). Thus, when grown on sulfide, it is easy to differentiate a purple sulfur from a purple nonsulfur bacterium because of the microscopically obvious globules of S0 formed (Fig. 1a). Subsequent isolations of purple nonsulfur bacteria from highly sulfidic habitats have shown that
Fig. 1. Phase photomicrographs of phototrophic purple bacteria. (a) Cells of a strain of the purple sulfur bacterium Thermochromatium tepidum isolated from a New Mexico (USA) hot spring. Note the bright refractile intracellular sulfur globules (arrows). (b) Cells of Rhodobaca bogoriensis, an alkaliphilic purple nonsulfur bacterium isolated from Lake Bogoria (Kenya). If purple nonsulfur bacteria oxidize sulfide, any S0 they produce remains outside the cell. Cells of Tch. tepidum are about 1.5 µm wide and cells of Rbc. bogoriensis are about 0.8 µm wide.
many species of this group are actually quite sulfide tolerant. For example, both the marine species Rhodobacter (originally Rhodopseudomonas, Rps.) sulfidophilus and the cold-active species Rhodoferax antarcticus (isolated from the sulfidic bottom waters
4 of the permanently frozen freshwater Lake Fryxell, McMurdo Dry Valleys) can tolerate over 4 mM sulfide (Hansen and Veldkamp, 1973; Jung et al., 2004). This concentration of sulfide is toxic to many purple sulfur bacteria (Pfennig, 1967, 1978a, 1989)! The original classification of purple bacteria on the basis of sulfide metabolism has been supported by molecular criteria. Phylogenetic analyses of purple bacteria based on comparative 16S rRNA sequencing have shown that purple sulfur bacteria are species of gammaproteobacteria while purple nonsulfur bacteria are either alpha- or betaproteobacteria (Imhoff et al., 2005) (Tables 1 and 2).
Michael T. Madigan and Deborah O. Jung phic species (Imhoff et al., 2005). If one considers the fact that the pigments and photocomplexes in the different species of purple nonsulfur bacteria are very similar, this suggests that the acquisition of phototrophic capacity in purple nonsulfur bacteria has occurred by lateral gene transfer. Sequence analyses of photocomplex proteins have confirmed this (Nagashima et al., 1997). Table 2 also lists the three letter genus name abbreviations for both purple sulfur and nonsulfur bacteria; these abbreviations will be used throughout this book. III. Physiology of Purple Bacteria
A. Purple Sulfur Bacteria Over 25 genera of purple sulfur bacteria are now recognized, consisting of a variety of morphological types (Table 2). Purple sulfur bacteria include both species that store S0 inside the cell (family Chromatiaceae, see Fig. 1a), and those that produce extracellular S0 (Ectothiorhodospiraceae) (Table 1). [It should be noted that research on the mechanism of sulfide oxidation by Allochromatium vinosum has shown that the ‘intracellular sulfur’ produced by this organism actually accumulates in the periplasm rather than in the cytoplasm (Pattaragulwanit et al., 1998); this is likely true for all species of Chromatiaceae as well]. Most laboratory studies of purple sulfur bacteria have focused on Allochromatium and Thiocapsa species (Table 1) since these are the most easily grown. Many species of purple sulfur bacteria are ‘extremophilic’ species, including in particular, species that grow best at high salt and/or pH (Table 2). B. Purple Nonsulfur Bacteria Twenty genera of purple nonsulfur bacteria are now recognized (Table 2). Species of Rhodobacter and Rhodopseudomonas have been the workhorses for laboratory studies of anoxygenic photosynthesis (Table 1). But many other interesting species, some of which have one or more unusual metabolic features, are also known. For example, extremophilic species inhabiting hot, cold, salty, alkaline (Fig. 1b), and acidic environments have been isolated and will be discussed in a later section of this chapter (see section V). As shown in Table 2, all purple nonsulfur bacteria are proteobacteria, and phylogenetic trees show the various species to be closely related to nonphototro-
Purple bacteria are relatively easy to grow in laboratory culture; in most cases all that is needed is an anoxic mineral medium supplemented with either sulfide plus bicarbonate (photoautotrophic growth) or an organic compound (photoheterotrophic growth). Because of this, and because anoxygenic photosynthesis is a simpler form of photosynthesis than the oxygenic process, purple bacteria have emerged as ideal model systems for dissecting the physiology, biochemistry and molecular biology of photosynthesis. Moreover, anoxygenic photosynthesis preceded oxygenic photosynthesis on Earth by billions of years. Thus, studies of purple and other anoxygenic phototrophs have contributed in major ways to our understanding of the evolution of photosynthesis (Raymond et al., 2003; Chapter 2, Swingley et al.). A. Purple Sulfur Bacteria The physiology of purple sulfur bacteria is intimately linked to sulfide, and large populations of purple sulfur bacteria are observed in nature only in illuminated environments where sulfide is present (Pfennig, 1967, 1978a, 1989). This implies that the growth of purple sulfur bacteria in nature is primarily phototrophic. If growth is photoautotrophic, sulfide, thiosulfate or H2 are used as photosynthetic electron donors (Trüper and Fischer, 1982; Madigan, 1988; Brune, 1995). A few species can also use ferrous (Fe2+) iron as an electron donor, oxidizing it to ferric (Fe3+) iron (Ehrenreich and Widdel, 1994), and at least one species, a strain of Thiocapsa, can use nitrite (NO2–) as photosynthetic electron donor, oxidizing it to nitrate (NO3–. In addition to autotrophic growth, a few organic carbon sources are photoassimilated by purple sulfur
Chapter 1
Overview of Purple Bacteria
5
Table 2. Genera of anoxygenic phototrophic purple bacteria Taxonomy/Phylogeny Purple Nonsulfur Bacteria Alphaproteobacteria
Betaproteobacteria
Purple Sulfur Bacteria Gammaproteobacteria Family Chromatiaceae b
Family Ectothiorhodospiraceae b
Genus
Genus abbreviation a
Morphology
Rhodobaca c Rhodobacter Rhodovulum Rhodopseudomonas c Rhodoblastusc Blastochloris Rhodomicrobium Rhodobium Rhodoplanes Rhodocistac Rhodospirillum Phaeospirillum Rhodopila c Rhodospira Rhodovibrio c Rhodothallasium c Roseospira Roseospirillum Rhodocyclus Rhodoferaxc Rubrivivax
Rca. Rba. Rdv. Rps. Rbl. Blc. Rmi. Rbi. Rpl. Rcs. Rsp. Phs. Rpi. Rsa. Rhv. Rts. Ros. Rss. Rcy. Rfx. Rvi.
Cocci to short rods Rods Rods-Cocci Budding rods Budding rods Budding rods Budding rods Rods Rods Spirilla Spirilla Spirilla Cocci Spirilla Vibrio Spirilla Spirilla Spirilla Curled vibrios Rods, vibrios Rods, curved rods
Allochromatium Amoebobacter Chromatium Halochromatium c Isochromatium Lamprobacter Lamprocystis Marichromatium Rhabdochromatium Thermochromatium c Thioalkalicoccus c Thiobaca Thiocapsa Thiococcus Thiocystis Thiodictyon Thioflavicoccus Thiohalocapsac Thiolamprovum Thiopedia Thiorhodococcus Thiorhodovibrio Thiospirillum
Alc. Amb. Chr. Hch. Isc. Lpb. Lpc. Mch. Rbc. Tch. Tac. Tba. Tca. Tco. Tcs. Tdc. Tfc. Thc. Tlp. Tpd. Trc. Trv. Tsp.
Rods Cocci in plates or clumps Rods Rods Rods Rods Cocci in clusters Rods Rods Rods Cocci Rods Cocci Cocci Cocci to short rods Rods forming aggregates Cocci Cocci Cocci Cocci, often in plates Cocci Vibrios to spirilla Spirilla
Ectothiorhodospira c Ect. Vibrios to spirilla Hlr. Vibrios to spirilla Halorhodospira c Trs. Vibrios to spirilla Thiorhodospira c Ectothiorhodosinus Ets. Rods a Abbreviations in accordance with Imhoff and Madigan (2004). bSpecies of Chromatiaceae store sulfur from the oxidation of sulfide intracellularly (see Fig. 1A); species of Ectothiorhodospiraceae do not. c Contain one or more extremophilic species growing at an extreme of temperature, pH, or salinity greater than marine salinity
6 bacteria. Organic acids and fatty acids are the preferred substrates, but short-chain alcohols and even carbohydrates are used by certain species (Sojka, 1978). Photoheterotrophic growth of Allochromatium vinosum and other Allochromatium species that are capable of assimilatory sulfate reduction does not require sulfide. However, some purple sulfur bacteria will not grow without sulfide and are also nutritionally quite restricted. These include the large-celled Chromatium species such as Chromatium okenii and Chromatium weissei, and Thiospirillum (Trüper, 1978), as well as the thermophilic species Thermochromatium (Tch.) tepidum (Madigan, 1986) (Fig. 1a). Sulfide is required for growth of these species and the only organic compounds that are photoassimilated are acetate and pyruvate (Trüper 1981; Madigan, 1986). Dark growth of some purple sulfur bacteria is possible. For example, certain Chromatiaceae, including species of Allochromatium, Thiocystis, Amoebobacter, and Thiocapsa, can grow in darkness as either chemoorganotrophs or chemolithotrophs when the oxygen concentration is significantly reduced [microaerobic growth; Kämpf and Pfennig, 1980]. Thiocapsa roseopersicina and Thiocystis violacea are the most oxygen tolerant purple sulfur bacteria (Kondratieva et al., 1976; Kämpf and Pfennig, 1980); however, respiratory growth of these species is very slow compared with phototrophic growth. If one considers that dark growth of purple sulfur bacteria in nature puts them in direct competition with nonphototrophic bacteria as well as with purple nonsulfur bacteria, the ecological significance of dark metabolism by purple sulfur bacteria is probably minor. It is more likely that dark energy metabolism by purple sulfur bacteria helps these organisms survive intermittently oxygenated environments or is used as a means to generate ATP at night, rather than being a major means of supporting extended growth in nature (van Gemerden, 1968). B. Purple Nonsulfur Bacteria Purple nonsulfur bacteria are a physiologically versatile group of purple bacteria that can grow well both phototrophically and in darkness. Growth of some purple nonsulfur bacteria, for example, Rhodobacter (Rba.) capsulatus, is possible under phototrophic conditions with either CO2 or organic carbon, or in darkness by respiration, fermentation, or chemolithotrophy. This makes Rba. capsulatus probably the
Michael T. Madigan and Deborah O. Jung most metabolically versatile of all known bacteria (Madigan and Gest, 1979). Carbon metabolism in purple nonsulfur bacteria has been summarized in the excellent reviews by Tabita (1995) and by Gibson and Harwood (1995); see also Chapter 28, Romagnoli andTabita; and Chapter 29, Harwood. 1. Photoheterotrophy Under phototrophic (anoxic/light) conditions, typical purple nonsulfur bacteria can grow photoautotrophically with H2 or low levels of sulfide as electron donors; a few species can use S2O32– or Fe2+ as photosynthetic electron donors (Ehrenreich and Widdel, 1994; Brune, 1995). However, most purple nonsulfur bacteria grow best as photoheterotrophs in media containing a readily useable organic compound, such as malate or pyruvate, and ammonia as nitrogen source (Sojka, 1978). Yeast extract is a common addition to media formulated for purple nonsulfur bacteria (Biebl and Pfennig, 1981). Yeast extract is a source of B-vitamins, one or more of which are required by the majority of recognized species of purple nonsulfur bacteria. Requirements for thiamine, nicotinic acid, biotin, and p-aminobenzoic acid are the most common. Requirements for B-complex vitamins have never been observed in purple sulfur bacteria, although many species require vitamin B12, a growth factor required by only a handful of purple nonsulfur bacteria (Pfennig, 1978b; Siefert and Koppenhagen, 1982). However, beyond its role as a source of vitamins, yeast extract also stimulates the growth of purple nonsulfur bacteria because of its assortment of organic compounds that can fuel photoheterotrophic growth. Several individual organic compounds support photoheterotrophic growth of purple nonsulfur bacteria. Organic acids, amino acids, fatty acids, alcohols, carbohydrates, and even C-1 compounds are metabolized by different species (Sojka, 1978; Trüper and Pfennig, 1981). With minor exceptions, the citric acid cycle intermediates malate, succinate, and fumarate are universally used, as are pyruvate and acetate; many species also use ethanol, lactate, and propionate (Sojka, 1978; Trüper and Pfennig, 1981). A few purple nonsulfur bacteria photoassimilate aromatic compounds such as benzoate, hydroxy derivatives of benzoate, and cyclohexane carboxylate (Gibson and Harwood, 1995). Enrichment cultures employing benzoate as carbon source typically yield strains of Phaeospirillum (formerly Rhodospirillum)
Chapter 1
Overview of Purple Bacteria
fulvum or Rps. palustris (Gibson and Harwood, 1995). Benzene is not utilized by these or other purple nonsulfur bacteria, but at least one aromatic hydrocarbon, toluene, supports photoheterotrophic growth of certain strains of Blastochloris sulfoviridis (Zengler et al., 1999). Growth of purple nonsulfur bacteria on aliphatic hydrocarbons has not been described. 2. Dark Growth Many of the same organic compounds that are photoassimilated by purple nonsulfur bacteria can also be used as electron donors and carbon sources for dark respiratory growth. Oxygen tolerances for respiratory growth vary among species, but some, such as Rhodobacter species, can be grown with vigorous aeration (Madigan, 1988). Certain purple nonsulfur bacteria can grow under anoxic dark conditions by either fermentation or anaerobic respiration. For example, pyruvate (Uffen and Wolfe, 1970; Gurgen et al., 1976) and certain sugars (Madigan and Gest, 1978; Schultz and Weaver, 1982) support fermentative growth of some purple nonsulfur bacteria, most notably Rhodospirillum (Rsp.) rubrum and Rba. capsulatus. Extensive fermentative growth of Rba. capsulatus requires addition of an accessory oxidant such as dimethyl sulfoxide or trimethylamine-N-oxide (Madigan and Gest, 1978; Schultz and Weaver, 1982). Rba. sphaeroides is capable of true denitrification, reducing NO3– to N2 using nonfermentable carbon sources as electron donors (Satoh et al., 1976). Dark chemolithotrophic growth of certain species of purple nonsulfur bacteria is possible using H2 or S2O32– as electron donors. In Rba. capsulatus, chemolithotrophic growth on H2 occurs and the organism can be grown in a synthetic medium supplied with the gases H2, O2, and CO2 as electron donor, electron acceptor, and carbon source, respectively (Madigan and Gest, 1979). Whether chemolithotrophy is a significant growth strategy for purple bacteria in nature is unknown, but it is likely that the ability to conserve energy from the oxidation of inorganic electron donors gives purple bacteria an added physiological dimension in competition with nonphototrophic bacteria. 3. Nitrogen Fixation With only a couple of known exceptions, purple nonsulfur bacteria can fix nitrogen (N2 + 8H → 2NH3 + H2) (Madigan, 1995). The Rhodobacter species Rba.
7 capsulatus and Rba. sphaeroides grow most rapidly with N2 as sole nitrogen source and show the highest rates of in vivo nitrogenase activity (Madigan et al., 1984). Consequently, Rhodobacter species tend to dominate enrichment cultures for purple nonsulfur bacteria that employ nitrogen fixation as a selective condition. Because in general purple nonsulfur bacteria are excellent nitrogen-fixing bacteria (Madigan et al., 1984), it is likely that the capacity for diazotrophy confers a significant competitive advantage on them in anoxic environments that are limited in fixed nitrogen. IV. Habitats of Purple Bacteria A. Purple Sulfur Bacteria A detailed description of the major habitats of purple bacteria including a wealth of specific examples can be found in the reviews of Madigan (1988), Pfennig (1967, 1978a, 1989), and van Gemerden and Mas (1995). Large masses (blooms) of purple sulfur bacteria often develop in sulfidic aquatic ecosystems exposed to light. Although blooms of phototrophic sulfur bacteria may occur in shallow lagoons polluted by sewage (which triggers the activities of sulfate-reducing bacteria), densely stratified ‘plates’ of purple bacteria form only in the deep waters of lakes protected from excessive wind mixing and which contain sufficient sulfate to support sulfate reduction in the sediments. 1. Blooms in Stratified Lakes Intense microbial activity occurs in the sediments of productive stratified lakes. Organic material reaching the bottom waters is catabolized by fermentation, which releases a variety of reduced organic products, including lactate, ethanol, and fatty acids. In freshwater lakes containing low levels of sulfate, these fermentation products can be photoassimilated by purple nonsulfur bacteria or converted to methane by the cooperative interactions of syntrophic bacteria and methanogenic Archaea. Alternatively, if electron acceptors such as Fe3+ or NO3– are available, the fermentation products will fuel anaerobic respirations supported by these electron acceptors before extensive occurs. If sulfate is present, sulfate-reducing bacteria
8 will be active in the sediments forming sulfide; the sulfide diffuses upwards from the sediments into the water column forming a gradient. Sulfide triggers the growth of purple sulfur bacteria, which develop in specific zones of the water column where light and sulfide are optimal (Pfennig, 1967, 1975, 1978a). If cell numbers are sufficiently high, the lake water itself will become pigmented red, purple, or reddishbrown (Pfennig 1978a, 1989; Overmann et al., 1994, 1996, 1999). When this occurs, it is often possible to identify the major genera of purple bacteria present by simple microscopic examination. In many stratified lakes the bloom of purple bacteria consists of a mixture of species (Caldwell and Tiedje, 1975a,b), while in others, the bloom may contain only a single species (Overmann et al., 1994, 1996, 1999). A nice example of the layering of phototrophic purple bacteria in stratified lakes can be found in the work of Caldwell and Tiedje (1975a,b). These workers examined water samples collected at 1 m intervals from two eutrophic lakes — Wintergreen and Burke — located in southwest Michigan (USA). In these studies, done before the days of molecular microbial ecology, the purple sulfur bacteria could be adequately identified by their characteristic morphologies. Both lakes contained species of Thiopedia, Thiospirillum, Thiocystis, and Chromatium. However, in Burke Lake the Thiospirillum population dominated while in Wintergreen Lake, the Thiopedia population dominated. Although physiochemical profiles of the two lakes were not performed, the dominant population in each case was likely selected by the major characteristics — light, sulfide, pH, dissolved organic carbon, and the like — that defined each lake (Caldwell and Tiedje, 1975a,b). Both Burke and Wintergreen lakes also contained green sulfur bacteria in the waters beneath the purple sulfur bacteria. The green bacterial population contained phototrophic consortia (Overmann and Schubert, 2002) such as ‘Chlorochromatium’ (Caldwell and Tiedje, 1975a,b). Green bacteria can outcompete purple bacteria at low light intensities because they possess large antenna pigment structures called chlorosomes (Kimble and Madigan, 2002; Frigaard and Bryant, 2004). Green bacteria also use different regions of the spectrum than do purple bacteria and are typically much more sulfide tolerant. Thus, green bacteria can exist beneath a layer of purple bacteria in a water column, and this pattern is common in stratified lakes (Pfennig, 1967, 1975, 1978a, 1989).
Michael T. Madigan and Deborah O. Jung In hypersulfidic shallow lakes, such as those in the karstic Banyoles area of northern Spain, millimolar levels of sulfide are present and phototrophic sulfur bacteria bloom throughout the water column. In Lake Cisó, the best studied of these lakes, the entire water column is anoxic, and the lake turns bright red during an active bloom (Guerrero et al., 1985). Both green and purple sulfur bacteria are present in Lake Cisó but the lake is dominated by a Chromatium (probably Allochromatium) species during periods of the most extensive bloom. 2. Microbial Mats Purple sulfur bacteria are also common in microbial mats, including mats that form in marine or hypersaline environments (van Gemerden and Mas, 1995) and in the effluents of thermal springs (Castenholz and Pierson, 1995). Microbial mats are laminated organo-sedimentary structures composed primarily of filamentous cyanobacteria and anoxygenic phototrophs, such as Chloroflexus, but often contain purple bacteria as well. As new growth occurs from the top of the mat, the lower mat layers decompose and sulfate-reduction typically occurs. This supplies the sulfide necessary to trigger growth of purple bacteria, usually purple sulfur bacteria. Mat thickness can vary considerably. In siliceous alkaline hot spring microbial mats, mats can be 4–5 cm thick. Mats containing only purple bacteria, such as those of Tch. tepidum that form in the Mammoth hot springs of Yellowstone are much thinner, up to 0.5 cm in thickness (Ward et al., 1989). Thiocapsa and Allochromatium species are common inhabitants of marine microbial mats. These purple sulfur bacteria often form a dense pigmented layer between the cyanobacteria and the lower layers of the mat. In this niche purple bacteria oxidize sulfide that diffuses upwards from below before it reaches the cyanobacterial layers (van Gemerden and Mas, 1995). Thiocapsa roseopersicina, in particular, is very common in marine mats, probably because of its metabolic versatility. Besides its photoautotrophic and photoheterotrophic capacities, this purple sulfur bacterium can grow in darkness by heterotrophic and chemolithotrophic means (Kondratieva et al., 1976). This versatility allows Thiocapsa roseopersicina to take full advantage of the variable growth conditions that characterize different layers of microbial mats (van Gemerden and Mas, 1995).
Chapter 1
Overview of Purple Bacteria
B. Purple Nonsulfur Bacteria Purple nonsulfur bacteria occasionally form dense blooms in habitats where levels of sulfide are either low or undetectable. Purple nonsulfur bacteria are usually present in only low numbers in blooms of purple sulfur bacteria, probably because of their sulfide sensitivity. Instead of photoautotrophy, purple nonsulfur bacteria specialize in photoheterotrophy. Although this puts them in competition with heterotrophs for organic compounds, photoheterotrophic capacity likely confers a significant selective advantage on purple nonsulfur bacteria. This is because unlike heterotrophs, phototrophs do not need to conserve energy from the carbon sources they photoassimilate; carbon goes almost quantitatively into cell material. However, no known purple nonsulfur bacteria can hydrolyze major polymeric substances such as cellulose or starch, and so ultimately, the phototrophs depend on the heterotrophs to generate the lowmolecular-weight compounds they photoassimilate (Pfennig, 1978a). 1. Sewage Purple nonsulfur bacteria are present in sewage (Holm and Vennes 1970; Siefert et al., 1978). In a detailed study by Siefert et al. (1978), it was shown that the mean number of purple nonsulfur bacteria in a sewage plant in Göttingen, Germany was highest in the activated sludge stage of the treatment process; counts fluctuated between 105 and 106 cells ml–1 but were never greater than 106 cells ml–1 (as measured using plate counting techniques). Purple sulfur bacteria, on the other hand, were quantitatively insignificant in sewage and were detectable by culture only from activated sludge (~103 cells ml–1). A variety of purple nonsulfur bacteria were identified in the sewage plant, including Rba. sphaeroides and Rba. capsulatus, Rps. palustris and Rps. (now Blastochloris) viridis, Rhodocyclus (now Rubrivivax) gelatinosus and Rhodocyclus (Rcy.) tenuis and Rsp. photometricum. Rba. sphaeroides, Rubrivivax gelatinosus, Rps. palustris, and Rba. capsulatus made up the bulk of the purple bacteria present. Despite these relatively high numbers, it was concluded that purple nonsulfur bacteria probably played only a minor role in organic matter transformations in sewage compared with heterotrophic bacteria, which were present at 108 to 109 cells ml–1. Purple nonsulfur bacteria were also detectable in the strictly anaerobic (and dark)
9 sewage sludge digestor, but these likely represented only transient cells traveling through the system (Siefert et al., 1978). 2. Purple Nonsulfur Bacteria in Waste Lagoons Waste lagoons offer excellent conditions for growth of purple nonsulfur bacteria (Jones, 1956; Cooper et al., 1975; Kobayashi, 1975). For example, a pigmented bloom of purple nonsulfur bacteria was reported from the waste lagoon of a vegetable canning plant in Minnesota (USA); prolific growth leading to intensely red-colored lagoons occurred and the bloom was associated with a significant reduction in odor (Cooper et al., 1975). Rba. sphaeroides, Rba. capsulatus, and Rps. palustris were the key species in this bloom, and it is likely that their consumption of volatile fatty acids produced by fermentation led to the odor reduction observed. The morphologically and phylogenetically unique purple nonsulfur bacterium Rcy. purpureus, isolated by Norbert Pfennig over 30 years ago, was the dominant phototroph in a swine waste lagoon in Iowa (USA) (Pfennig, 1978b). This organism, of which only a single strain has ever been isolated, probably thrived on the combination of organic constituents present in the waste materials. However, it is of interest that Rcy. purpureus, one of the only purple nonsulfur bacteria to lack a nitrogenase system (Madigan et al., 1984; Madigan, 1995), was the dominant purple bacterium in this particular habitat and that it has never been reported from elsewhere. One would expect a swine waste lagoon to be high in amines and thus that nitrogen fixation would be unnecessary. It is thus possible that Rcy. purpureus is somehow selected for in otherwise suitable habitats for purple nonsulfur bacteria that are very high in ammonia and volatile amines. Since Rcy. purpureus is easily cultured, an enrichment study of its distribution in nature using media containing elevated levels of ammonia and amines could yield insight on its ecology. V. Purple Bacteria in Extreme Environments Purple bacteria have been isolated from extreme environments, including hot, cold, acidic, alkaline, and hypersaline (Madigan, 2003). Unfortunately, with a few exceptions, these inherently interesting ‘ex-
10 tremophilic’ purple bacteria have been little utilized in the study of photosynthesis thus far. However, the success of these unusual purple bacteria in their harsh habitats implies that they have evolved important solutions to photosynthesis under stress conditions. We can therefore learn much from studying them. For example, molecular adaptations linked to photosynthesis under extreme conditions should be relatively easy to identify in extremophilic purple bacteria by applying a bioinformatics/structural biology approach to their genomes. A. Thermophilic Purple Bacteria The first extremophilic purple bacteria were discovered in the 1960s and were either halophiles or acidophiles, including extremely halophilic species of the genus Ectothiorhodospira. Into the late 1970s, several new halophilic and haloalkaliphilic purple bacteria were discovered. In the 1980s, the first thermophilic purple bacterium was isolated, Tch. tepidum. Since then, a large diversity of alkaliphilic and halophilic purple bacteria has been isolated. Psychrophilic phototrophs have been described only very recently, with two representatives currently in culture. 1. Thermochromatium tepidum Purple bacteria were first identified in Yellowstone microbial mats over 30 years ago (Castenholz, 1977). But it was not until the 1980s that the purple sulfur bacterium Thermochromatium (originally Chromatium) tepidum was isolated in pure culture (Madigan, 1984, 1986). Tch. tepidum (Fig. 1a) is thermophilic (optimumtemperature ~50 ºC, maximum temperature 57 ºC) and produces a novel light-harvesting (LH) 1 photopigment complex that absorbs maximally near 920 nm (Garcia et al., 1986; Nozawa et al., 1986). The Tch. tepidum LH1 (core) antenna complex has been studied in connection with the mechanism of energy transfer to the reaction center. A biophysical conundrum exists with this photocomplex in that its absorption maximum is 50 nm to the red of that of the reaction center. Nevertheless, because there is a small overlap between spectra of the two components, efficient energy transfer occurs from the Tch. tepidum LH1 complex to the reaction center (Kramer and Amesz, 1996). Examples of long-wavelength-absorbing core antenna complexes even more spectacular than that of Tch. tepidum have been discovered, indicating that in purple bacteria, LH1 complexes
Michael T. Madigan and Deborah O. Jung that absorb very far to the red (963 nm) can still transfer energy to the reaction center (Permentier et al., 2001). The photosynthetic reaction center of Tch. tepidum is similar in most respects to that of other purple bacteria, except for its increased thermal stability (Nozawa and Madigan, 1991). To probe the mechanism behind this, the Tch. tepidum reaction center was crystallized. From this work, key substitutions were identified in the L and M subunits of the Tch. tepidum reaction center that are likely responsible for the thermostability of this photocomplex (Nogi et al., 2000). In addition to thermal stable photocomplexes, a thermophilic ribulose bisphosphate carboxylase (a key enzyme of the Calvin cycle) of the green plant type was characterized from Tch. tepidum and shown to be stable to at least 60 ºC (Heda and Madigan, 1988. 1989). 2. Other Thermophilic Purple Bacteria Other mildly thermophilic purple bacteria (optimum growth temperature ~40 ºC) have been cultured from hot spring microbial mats. These include the BChl b-containing species Rhodopseudomonas sp. strain GI, isolated from a New Mexico hot spring (Resnick and Madigan, 1989), and Rps. cryptolactis (Statwald-Demchick et al., 1990) and Rsp. centenum (Rhodocista centenaria) (Favinger et al., 1989), both isolated from a Thermopolis (Wyoming, USA) hot spring. Rhodocista centenaria in particular has been useful as a model organism for biochemical/genetic research on phototaxis and related issues of motility (see, for example McClain et al., 2002), and its genome has recently been sequenced (C. E. Bauer, personal communication). B. Halophilic and Alkaliphilic Purple Bacteria Several extremophilic purple bacteria are halophilic or haloalkaliphilic (Imhoff et al., 1978, 1979). These include purple sulfur bacteria such as Ectothiorhodospira, Halorhodospira, Halochromatium, Marichromatium, and Thiohalocapsa, and purple nonsulfur bacteria such as Rhodovibrio, Rhodothalassium, Rhodobium, Rhodovulum, and Roseospira. Collectively, these purple bacteria have salt optima that range from seawater salinities to over 20% NaCl (Imhoff, 2001). Interestingly, the Dead Sea purple nonsulfur bacterium Rhodovibrio sodomensis shows a distinct intermediate level salt requirement (optimum
Chapter 1
Overview of Purple Bacteria
at 8–11% NaCl) (Mack et al., 1993), which is very near that of its habitat. Some purple sulfur bacteria can grow in saturated salt solutions, making them the most halophilic of all known phototrophic bacteria (Imhoff, 2001). In the 1990s several new purple bacteria were isolated from low salinity soda lakes. Most of these differed dramatically from known halophilic or haloalkaliphilic species in that they required little if any NaCl for growth. These isolates are, however, strongly alkaliphilic (pH optima near 9) and phylogenetically distinct. These include purple nonsulfur bacteria such as Rhodobaca (Milford et al., 2000), and purple sulfur bacteria such as Thioalkalicoccus (Bryantseva et al., 2000), and Thiorhodospira (Bryantseva et al., 1999). Rhodobaca (Fig. 1b) is of particular interest because it lacks a peripheral (LH2) antenna complex, a rarity among purple nonsulfur bacteria (Glaeser and Overmann, 1999), and it produces several unusual carotenoids that render phototrophic cultures yellow in color (Takaichi et al., 2001). Rhodobaca shows various metabolic peculiarities as well, including an inability to fix N2 and to grow photoautotrophically (Milford et al., 2000); both of these properties are hallmarks of purple nonsulfur bacteria (Madigan, 1988). C. Acidophilic Purple Bacteria The list of acid-loving anoxygenic phototrophs is short, as only two genera (three species) are known. Rhodoblastus acidophilus (formerly Rhodopseudomonas acidophila) is common in mildly acidic environments, such as bogs, marshes, and acidic lakes. In the original description of this organism, several strains were described, some containing orange/brown carotenoids and others purple/red carotenoids. The strains are otherwise similar, and all show a lower limit for growth near pH 4 (Pfennig, 1969). A very similar organism is the species Rhodoblastus sphagnolica (Kulichevskaya et al., 2006). Rhodopila globiformis was isolated from acidic warm sulfide springs (pH 3.5–4) that flow out along the Gibbon River in Yellowstone National Park (Pfennig, 1974). Rhodpila globiformis-like organisms have also been obtained from springs that feed into Nymph Lake, a warm, acidic and sulfidic lake adjacent to the Gibbon River; the pH of the Nymph Lake springs is 3 (Madigan, 2003). Rhodopila globiformis has a low pH optimum for growth similar to that of Rhodoblastus species, but, if carefully tested, would likely be more
11 acid tolerant than Rhodoblastus species because of the more strongly acidic nature of its habitat. Phylogenetically, Rhodopila and Rhodoblastus are quite distinct. No acidophilic purple sulfur bacteria are known and this is likely because at acid pH, sulfide would exist exclusively as H2S, its most toxic form. D. Purple Bacteria from Permanently Cold Habitats Permanently cold environments are habitats for cold-active purple bacteria (Burke and Burton, 1988; Madigan, 1998). The Madigan laboratory has been studying phototrophs that inhabit lakes in the McMurdo Dry Valleys of Antarctica. These are closed basin lakes with a biology that is exclusively microbial, and they remain permanently frozen with ice covers of 4–7 m. Purple nonsulfur bacteria have been isolated from samples of microbial mats that develop along the edge of the lakes as well as from water under the ice. Molecular studies using pufM as a measure of the biodiversity of purple bacteria suggest that species related to known purple nonsulfur bacteria reside in these lakes (Karr et al., 2003). The purple bacterium Rhodoferax antarcticus inhabits both microbial mats and the water column of Lake Fryxell (77º S latitude), a lake supporting major sulfur cycling activities (Sattley and Madigan, 2006). Lake Fryxell is unmixed and weakly stratified, with saline bottom waters overlain by freshwater. A gradient of sulfide is present in Lake Fryxell from micromolar levels at a depth of 9 m to nearly 1.5 millimolar near the sediments (~18 m). Despite these nearly perfect conditions for the development of phototrophic sulfur bacteria, no evidence for purple sulfur bacteria has emerged from enrichment culture or nucleic acid probing studies using pufM (Achenbach et al., 2001; Karr et al., 2003). Instead, Rfx. antarcticus, a very sulfide tolerant purple nonsulfur bacterium (Jung et al., 2004), seems to dominate anoxygenic photosynthesis in Lake Fryxell. Although Rfx. antarcticus is not strongly psychrophilic (optimal growth occurs at 18 ºC and no growth occurs above 24 ºC), it is the first anoxygenic phototroph to show a distinct cold adaptation and growth at 0 ºC (Madigan et al., 2000). Also in the water column of Lake Fryxell, gas vesiculate purple nonsulfur bacteria are present, including a morphologically unique strain of Rfx. antarcticus (betaproteobacteria) and a rod-shaped organism related to Rhodobacter species (alphaproteobacteria). These gas vesiculate
12 purple bacteria position themselves in zones of the water column where photosynthesis can occur optimally (Karr et al. 2003; Jung et al., 2004). Biomass measurements have shown that they localize in the water column at a depth of 10 m and that cell numbers drop off sharply below this depth and are undetectable above this depth (Madigan, unpublished). E. Environmental Limits to Photosynthesis in Purple Bacteria Work with extremophilic purple bacteria has given a good indication of the limits to which photosynthesis can be pushed. Photosynthesis in purple bacteria can occur at temperatures up to at least 57 ºC and down to 0 ºC, pH values as low as 3 or as high as 11, and at salinities up to and including saturated solutions of NaCl (~32%). Although these are probably not the absolute physiochemical limits to photosynthesis in anoxygenic phototrophs, they are likely to be very close to the limits. It is notable that the green nonsulfur bacterium Chloroflexus aurantiacus, which contains a purple bacterial-type photosynthetic reaction center (Achenbach et al., 2001), can grow up to 70 °C (Castenholz and Pierson, 1995). This holds out hope that purple bacteria capable of growth at temperatures above 57 °C (the upper limit for Tch. tepidum, Madigan, 1986) may exist in nature. Further exploration for new purple bacteria in sulfidic hot springs with temperatures above 60 °C should answer this question. VI. Final Remarks Our understanding of purple bacteria goes far beyond what has been discussed here. Through the years purple bacteria have become increasingly important as research tools for the study of basic problems in photosynthesis and have contributed in major ways to our understanding of the biochemistry, genetics and evolution of photosynthesis. And importantly, the beautiful colors and metabolic versatility of purple bacteria continue to attract talented young people to the field. The reader will see the fruits of their labors as well as those of the more seasoned investigators in the following chapters where the biology of purple bacteria will unfold in a spectacular way.
Michael T. Madigan and Deborah O. Jung Acknowledgments Research in the Madigan laboratory is supported by the United States National Science Foundation, most recently by grant MCB0237576. MTM thanks all of his former students and postdoctoral colleagues who did research on purple bacteria in his laboratory. These include Rich Masters, Rick Stegeman, Glenn Wright, Carletta Ooten, Erin Mack, Joseph Mayers, Ghanshyam Heda,Vasiliki Karayiannis, Sol Resnick, Chad Rubin, Gerrit Hoogewerf, Ike Pantazopoulous, Amy Milford, Linda Kimble, Amy Stevenson, Terry Locke, Tiffany Full, Jen Carey, Elizabeth Karr, Jill Crespi, Mahmoud Tayah, Tom Wahlund, Matt Sattley, and Marie Asao. References Achenbach LA, Carey JR and Madigan MT (2001) Photosynthetic and phylogenetic primers for detection of anoxygenic phototrophs in natural environments. Appl Environ Microbiol 67: 2922–2926 Biebl H and Pfennig N (1981) Isolation of members of the family Rhodospirillaceae. In: Starr MP, Stolp H, Trüper HG, Balows A and Schlegel HG (eds) The Prokaryotes — a Handbook on Habitats, Isolation and Identification of Bacteria, pp 267–273. Springer-Verlag, New York Blankenship RE, Madigan MT and Bauer CE (1995) Anoxygenic Photosynthetic Bacteria (Advances in Photosynthesis and Respiration, Vol 2). Kluwer Academic Publishers, Dordrecht Brune DC (1995) Sulfur compounds as photosynthetic electron donors. In: Blankenship RE, Madigan MT and Bauer CE (eds) Anoxygenic Photosynthetic Bacteria (Advances in Photosynthesis and Respiration, Vol 2), pp 847–870. Kluwer Academic Publishers, Dordrecht Bryantseva IA, Gorlenko VM, Kompantseva EI and Imhoff JF (2000) Thioalkalicococcus limnaeus gen. nov., sp. nov., a new alkaliphilic purple sulfur bacterium with bacteriochlorophyll b. Int J Syst Bacteriol 50: 2157–2163 Bryantseva IA, Gorlenko VM, Kompantseva EI, Imhoff JF, Süling J and Mityushina L (1999) Thiorhodospira sibirica gen. nov., sp. nov., a new alkaliphilic purple sulfur bacterium from a Siberian soda lake. Int J Syst Bacteriol 49: 697–703 Burke CM and Burton HR (1988) Photosynthetic bacteria in meromictic lakes and stratified fjords of the Vestfold Hills, Antarctica. Hydrobiologia 165: 13–23 Caldwell DE and Tiedje JM (1975a) A morphological study of anaerobic bacteria from the hypolimnia of two Michigan lakes. Can J Microbiol 21: 362–376 Caldwell DE and Tiedje JM (1975b) The structure of anaerobic bacterial communities in the hypolimnia of several Michigan lakes. Can J Microbiol 21: 377–385 Castenholz RW (1977) The effect of sulfide on the blue-green algae of hot springs II. Yellowstone National Park. Microbial Ecology 3: 79–105 Castenholz RW and Pierson BK (1995) Ecology of thermophilic
Chapter 1
Overview of Purple Bacteria
anoxygenic phototrophs. In: Blankenship RE, Madigan MT and Bauer CE (eds) Anoxygenic Phototrophic Bacteria, pp 87–103. Kluwer Academic Publishers, Dordrecht Cohen-Bazire G, Sistrom WR and Stanier RY (1957) Kinetic studies of pigment synthesis by non-sulfur purple bacteria. J Cell Comp Physiol 49: 25–68 Cooper DE, Rands MB and Woo C-P (1975) Sulfide reduction in fellmongery effluent by red sulfur bacteria. J Water Pollution Control Fed 47: 2088–2100 Czeczuga B (1968) Primary production of the purple sulphuric bacteria, Thiopedia rosea Winogr. (Thiorhodaceae). Photosynthetica 2: 161–166 Ehrenreich A and Widdel F (1994) Anaerobic oxidation of ferrous iron by purple bacteria, a new type of phototrophic metabolism. Appl Environ Microbiol 60: 4517–4526 Favinger J, Stadtwald R and Gest H (1989) Rhodospirillum centenum, sp. nov., a thermotolerant cyst-forming anoxygenic photosynthetic bacterium. Ant van Leeuwenhoek 55: 291–296 Frigaard NU and Bryant DA (2004) Seeing green bacteria in a new light: Genomics-enabled studies of the photosynthetic apparatus in green sulfur bacteria and filamentous anoxygenic phototrophic bacteria. Arch Microbiol 182: 265–276 Garcia D, Parot P, Verméglio A and Madigan MT (1986) The light-harvesting complexes of a thermophilic purple sulfur photosynthetic bacterium Chromatium tepidum. Biochim Biophys Acta 850: 390–395 Gibson J and Harwood CS (1995) Degradation of aromatic compounds by nonsulfur purple bacteria. In: Blankenship RE, Madigan MT and Bauer CE (eds) Anoxygenic Photosynthetic Bacteria (Advances in Photosynthesis and Respiration, Vol 2), pp 991–1003. Kluwer Academic Publishers, Dordrecht Glaeser J and Overmann J (1999) Selective enrichment and characterization of Roseospirillum parvum, gen. nov. and sp. nov., a new purple nonsulfur bacterium with unusual light absorption properties. Arch Microbiol 171: 405–416 Griffin BM, Schott J and Schink B (2007) Nitrite, an electron donor for anoxygenic photosynthesis. Science 316: 1870 Guerrero R, Montesinos E, Pedrós-Alió C, Esteve I, Mas J, van Gemerden H, Hofman PAG and Bakker JF (1985) Phototrophic sulfur bacteria in two Spanish lakes: Vertical distribution and limiting factors. Limnol Oceanogr 30: 919–931 Gurgen V, Kirchner G and Pfennig N (1976) Fermentation of pyruvate by 7 species of phototrophic purple bacteria. Z Allg Mikrobiolo 16: 573–586 Hansen TA and van Gemerden H (1972) Sulfide utilization by purple nonsulfur bacteria. Arch Mikrobiol 86: 49–56 Hansen TA and Veldkamp H (1973) Rhodopseudomonas sulfidophila, nov. spec., a new species of the purple nonsulfur bacteria. Arch Mikrobiol 92: 45–58 Heda GD and Madigan MT (1988) Thermal properties and oxygenase activity of ribulose-1,5-bisphosphate carboxylase from the thermophilic purple bacterium, Chromatium tepidum. FEMS Microbiol Lett 51: 45–50 Heda GD and Madigan MT (1989) Purification and characterization of the thermostable ribulose-1,5-bisphosphate carboxylase/ oxygenase from the thermophilic purple bacterium Chromatium tepidum. Eur J Biochem 184: 313–319 Holm HW and Vennes JW (1970) Occurrence of purple sulfur bacteria in a sewage treatment lagoon. Appl Microbiol 19: 988–996 Imhoff JF (2001) True marine and halophilic anoxygenic photo-
13 trophic bacteria. Arch Microbiol 176: 243–254 Imhoff JF and Madigan MT (2004) International Committee on Systematics of Prokaryotes Subcommitteee on the taxonomy of phototrophic bacteria. Minutes of the meetings, 27 August 2003, Tokyo, Japan. Int J Syst Evol Microbiol 54: 1001–1003 Imhoff JF, Hashwa F and Trüper HG (1978) Isolation of extremely halophilic phototrophic bacteria from the alkaline Wadi Natrun, Egypt. Arch Hydrobiol 84: 381–388 Imhoff JF, Sahl HG, Soliman GSH and Trüper HG (1979) The Wadi Natrun: chemical composition and microbial mass developments in alkaline brines of eutrophic desert lakes. Geomicrobiol J 1: 219–234 Imhoff JF, Hiraishi A and Süling J (2005) Anoxygenic phototrophic bacteria. In: Brenner DJ, Krieg NR and Staley JT (eds) Bergey’s Manual of Systematic Bacteriology, 2nd ed, Vol 2, part A, pp 119–132. Springer, New York Jones BR (1956) Studies of pigmented non-sulfur purple bacteria in relation to cannery waste lagoon odors. Sewage Ind Wastes 28: 883–893 Jung DO, Achenbach LA, Karr EA, Takaichi S and Madigan MT (2004) A gas vesiculate planktonic strain of the purple non-sulfur bacterium Rhodoferax antarcticus isolated from Lake Fryxell, Dry Valleys, Antarctica. Arch Microbiol 182: 236–243 Kämpf C and Pfennig N (1980) Capacity of Chromatiaceae for chemotrophic growth. Specific respiration rates of Thiocystis violacea and Chromatium vinosum. Arch Microbiol 127: 125–135 Karr EL, Sattley WM, Jung DO, Madigan MT and Achenbach LA (2003) Remarkable diversity of phototrophic purple bacteria in a permanently frozen Antarctic lake. Appl Environ Microbiol 69: 4910–4914 Kimble-Long LK and Madigan MT (2002) Irradiance effects on growth and bacteriochlorophyll content of phototrophic heliobacteria, purple and green photosynthetic bacteria. Photosynthetica 40: 629-632 Kobayashi M (1975) Role of photosynthetic bacteria in foul water purification. Prog Water Technol 7: 309–315 Kondratieva EN, Zhukov VG, Ivanovsky RN, Petushkova YP and Monosov EZ (1976) The capacity of phototrophic sulfur bacterium Thiocapsa roseopersicina for chemosynthesis. Arch Microbiol 108: 287–292 Kramer H and Amesz J (1996) Antenna organization in the purple sulfur bacteria Chromatium tepidum and Chromatium vinosum. Photosynth Res 49: 237–244 Kulichevskaya IS, Guzev VS, Gorlenko VM, Liesack W and Dedysh SN (2006) Rhodoblastus sphagnicola sp. nov., a novel acidophilic purple non-sulfur bacterium from Sphagnum peat bog. Intl J Syst Evol Microbiol 56: 1397–1402 Mack EE, Mandelco L, Woese CR and Madigan MT (1993) Rhodospirillum sodomense, sp. nov., a Dead Sea Rhodospirillum species. Arch Microbiol 160: 363–371 Madigan MT (1984) A novel photosynthetic purple bacterium isolated from a Yellowstone hot spring. Science 225: 313–315 Madigan MT (1986) Chromatium tepidum sp. nov., a thermophilic photosynthetic bacterium of the family Chromatiaceae. Int J Syst Bacteriol 36: 222–227 Madigan MT (1988) Microbiology, physiology, and ecology of phototrophic bacteria. In: AJB Zehnder (ed) Biology of Anaerobic Microorganisms, pp 39–111, John Wiley & Sons, New York
14 Madigan MT (1995) Microbiology of nitrogen fixation by anoxygenic photosynthetic bacteria. In: Blankenship RE, Madigan MT and Bauer CE (eds) Anoxygenic Photosynthetic Bacteria (Advances in Photosynthesis and Respiration, Vol 2), pp 915–928. Kluwer Academic Publishers, Dordrecht Madigan MT (1998) Isolation and characterization of psychrophilic purple bacteria from Antarctica. In: Peschek GA, Löffelhardt W and Schmetterer G (eds) The Phototrophic Prokaryotes, pp 699–706. Plenum, New York Madigan MT (2003) Anoxygenic phototrophic bacteria from extreme environments. Photosynth Res 76: 157–171 Madigan MT and Gest H (1978) Growth of a photosynthetic bacterium anaerobically in darkness, supported by ‘oxidant-dependent’ sugar fermentation. Arch Microbiol 117: 119–122 Madigan MT and Gest H (1979) Growth of the photosynthetic bacterium Rhodopseudomonas capsulata chemoautotrophically in darkness with H2 as the energy source. J Bacteriol 137: 524–530 Madigan MT, Cox SS and Stegeman RA (1984) Nitrogen fixation and nitrogenase activities in members of the family Rhodospirillaceae. J Bacteriol 157: 73–78 Madigan MT, Jung DO, Woese CR and Achenbach LA (2000) Rhodoferax antarcticus sp. nov., a moderately psychrophilic purple nonsulfur bacterium isolated from an Antarctic microbial mat. Arch. Microbiol. 173: 269–277 McClain J, Rollo DR, Rushing BG and Bauer CE (2002) Rhodospirillum centenum utilizes separate motor and switch components to control lateral and polar flagellum rotation. J Bacteriol 184: 2429–2438 Milford AD, Achenbach LA, Jung DO and Madigan MT (2000) Rhodobaca bogoriensis gen. nov. and sp. nov., an alkaphilic purple nonsulfur bacterium from African Rift Valley soda lakes. Arch Microbiol 174: 18–27 Nagashima KVP, Hiraishi A, Shimada K and Matsuura K (1997) Horizontal transfer of genes coding for the photosynthetic reaction centers of purple bacteria. J Mol Evol 45: 131–136 Nogi T, Fathir I, Kobayashi M, Nozawa T and Miki K (2000) Crystal structures of photosynthetic reaction center and highpotential iron-sulfur protein from Thermochromatium tepidum: Thermostability and electron transfer. Proc Natl Acad Sci USA 97: 13561–13566 Nozawa T and Madigan MT (1991) Temperature and solvent effects on reaction centers from Chloroflexus aurantiacus and Chromatium tepidum. J Biochem 110: 588–594 Nozawa T, Fukada T, Hatano M and Madigan MT (1986) Organization of intracytoplasmic membranes in a novel thermophilic purple photosynthetic bacterium as revealed from absorption, circular dichroism, and emission spectra. Biochim Biophys Acta 852: 191–197 Overmann J and Schubert K (2002) Phototrophic consortia: Model systems for symbiotic interrelations between prokaryotes. Arch Microbiol 177: 201–208 Overmann J, Beatty JT and Hall KJ (1994) Photosynthetic activity and population dynamics of Amoebobacter purpureus in a meromictic saline lake. FEMS Microbiol Ecol 15: 309–320 Overmann J, Beatty JT and Hall KJ (1996) Purple sulfur bacteria control the growth of aerobic heterotrophic bacterioplankton in a meromictic salt lake. Appl Environ Microbiol 62: 3251–3258 Overmann J, Hall KJ, Northcote TG and Beatty JT (1999) Grazing of the copepod Diaptomus connexus on purple sulphur bacteria
Michael T. Madigan and Deborah O. Jung in a meromictic salt lake. Environ Microbiol 1: 213–221 Pattaragulwanit K, Brune DC, Trüper HG and Dahl C (1998) Molecular genetic evidence for extracytoplasmic localization of sulfur globules in Chromatium vinosum. Arch Microbiol 169: 434–444 Permentier HP, Neerken S, Overmann J and Amesz J (2001) A bacteriochlorophyll a antenna complex from purple bacteria absorbing at 963 nm. Biochemistry 40: 5573–5578 Pfennig N (1967) Photosynthetic bacteria. Ann Rev Microbiol 21: 285–324 Pfennig N (1969) Rhodopseudomonas acidophila, sp. n., a new species of the budding purple nonsulfur bacteria. J Bacteriol 99: 597–602 Pfennig N (1974) Rhodopseudomonas globiformis, sp. n., a new species of the Rhodospirillaceae. Arch Microbiol 100: 197–206 Pfennig N (1975) The phototrophic bacteria and their role in the sulfur cycle. Plant Soil 43: 1–16 Pfennig N (1978a) General physiology and ecology of photosynthetic bacteria. In: Clayton RK and Sistrom WR (eds) The Photosynthetic Bacteria, pp 3–18. Plenum Press, New York Pfennig N (1978b) Rhodocyclus purpureus gen. nov. and sp. nov., a ring-shape, vitamin B12-requiring member of the family Rhodospirillaceae. Int J Syst Bacteriol 28: 283–288 Pfennig N (1989) Ecology of phototrophic purple and green sulfur bacteria. In: Schlegel HG and Bowien B (eds) Autotrophic Bacteria, pp 97–116. Springer-Verlag, Heidelberg Raymond J, Zhaxybayeva O, Gogarten JP and Blankenship RE (2003) Evolution of photosynthetic prokaryotes: A maximumlikelihood mapping approach. Phil Tran Roy Soc Lond B Biol Sci 358: 223–230 Resnick SM and Madigan MT (1989) Isolation and characterization of a mildly thermophilic nonsulfur purple bacterium containing bacteriochlorophyll b. FEMS Microbiol Lett 65: 165–170 Satoh T, Hoshino Y and Kitamura H (1976) Rhodopseudomonas sphaeroides forma sp. denitrificans, a denitrifying strain as a subspecies of Rhodopseudomonas sphaeroides. Arch Microbiol 108: 265–269 Sattley WM and Madigan MT (2006) Isolation, characterization and ecology of cold-active, chemolithotrophic sulfur-oxidizing bacteria from perennially ice-covered Lake Fryxell, Antarctica. Appl Environ Microbiol 72: 5562–5568 Schultz JE and Weaver PF (1982) Fermentation and anaerobic respiration by Rhodospirillum rubrum and Rhodopseudomonas capsulata. J Bacteriol 149: 181–190 Siefert E and Koppenhagen VB (1982) Studies on the vitamin B12 auxotrophy of Rhodocyclus purpureus and two other vitamin B12-requiring purple nonsulfur bacteria. Arch Microbiol 132: 173–178 Siefert E, Irgens RL and Pfennig N (1978) Phototrophic purple and green bacteria in a sewage treatment plant. Appl Environ Microbiol 35: 38–44 Sojka GA (1978) Metabolism of nonaromatic organic compounds. In: Clayton RK and Sistrom WR (eds) The Photosynthetic Bacteria, pp 707–718. Plenum Press, New York Stadtwald-Demchick R, Turner FR and Gest H (1990) Rhodopseudomonas cryptolactis, sp. nov., a new thermotolerant species of budding phototrophic purple bacteria. FEMS Microbiol Lett 71: 117–121 Tabita FR (1995) The biochemistry and metabolic regulation of
Chapter 1
Overview of Purple Bacteria
carbon metabolism and CO2 fixation in purple bacteria. In: Blankenship RE, Madigan MT and Bauer CE (eds) Anoxygenic Photosynthetic Bacteria (Advances in Photosynthesis and Respiration, Vol 2), pp 885–914. Kluwer Academic Publishers, Dordrecht Takahashi M and Ichimura S (1968) Vertical distribution and organic matter production of photosynthetic sulfur bacteria in Japanese lakes. Limnol Oceanog 13: 644–655 Takaichi S, Jung DO and Madigan MT (2001) Accumulation of unusual carotenoids in the spheroidene pathway, demethylspheroidene and demethylspheroidenone, in an alkaliphilic purple nonsulfur bacterium Rhodobaca bogoriensis. Photosynth Res 67:207–214 Trüper HG (1978) Sulfur metabolism. In: Clayton RK and Sistrom WR (eds) The Photosynthetic Bacteria, pp 677–690. Plenum Press, New York Trüper HG (1981) Versatility of carbon metabolism in phototrophic bacteria. In: Dalton H (ed) Microbial Growth on C1 Compounds, pp 116–121. Heyden, London Trüper HG and Fischer U (1982) Anaerobic oxidation of sulphur compounds as electron donors for bacterial photosynthesis. Phil Trans Roy Soc Lond B 298: 529–542 Trüper HG and Pfennig N (1981) Characterization and identification of the anoxygenic phototrophic bacteria. In: Starr MP, Stolp H, Trüper HG, Balows A and Schlegel HG (eds.) The Prokaryotes, a Handbook on Habitatss, Isolation, and Identification of
15 Bacteria, pp 299–312. Springer-Verlag, New York Uffen RL and Wolfe RS (1970) Anaerobic growth of purple nonsulfur bacteria under dark conditions. J Bacteriol 104: 462–472 van Gemerden H (1968) On the ATP generation by Chromatium in darkness. Arch Mikrobiol 64: 118–124 van Gemerden H and Mas J (1995) Ecology of phototrophic sulfur bacteria. In: Blankenship RE, Madigan MT and Bauer CE (eds) Anoxygenic Photosynthetic Bacteria (Advances in Photosynthesis and Respiration, Vol 2), pp 50–85. Kluwer Academic Publishers, Dordrecht van Niel CB (1932) On the morphology and physiology of the purple and green sulphur bacteria. Arch Mikrobiol 3: 1–112 van Niel CB (1944) The culture, general physiology, morphology, and classification of the non-sulfur purple and brown bacteria. Bacteriol Rev 8: 1–118 Ward DM, Weller R, Shiea J, Castenholz RW and Cohen Y (1989) Hot spring microbial mats: anoxygenic and oxygenic mats of possible evolutionary significance. In: Cohen Y and Rosenberg E (eds) Microbial Mats: Physiological Ecology of Benthic Microbial Communities, pp 3–15. American Society for Microbiology, Washington, DC Zengler K, Heider J, Rossello-Mora R and Widdel F (1999) Phototrophic utilization of toluene under anoxic conditions by a new strain of Blastochloris sulfoviridis. Arch Microbiol 172: 204–212
Chapter 2 Evolutionary Relationships Among Purple Photosynthetic Bacteria and the Origin of Proteobacterial Photosynthetic Systems Wesley D. Swingley Institute of Low Temperature Science, Hokkaido University, N19W8, Sapporo 060-0819, Japan
Robert E. Blankenship Departments of Biology and Chemistry, Washington University in St. Louis, Campus Box 1337, St. Louis, MO 63130, U.S.A.
Jason Raymond* School of Natural Sciences, University of California-Merced, Merced, CA 95344, U.S.A.
Summary ................................................................................................................................................................. 17 I. Introduction....................................................................................................................................................... 18 II. The Alphaproteobacteria .................................................................................................................................. 18 III. Aerobic Purple Bacteria.................................................................................................................................... 19 IV. The Photosynthesis Gene Cluster and its Role in Evolution ............................................................................ 20 V. Proteobacterial Comparative Genomics: Photosynthetic versus non-Photosynthetic Proteins ....................... 21 VI. Origin and Evolution of Proteobacterial Phototrophy ....................................................................................... 22 VII. Origin and Evolution of Proteobacterial Carbon-fixation .................................................................................. 24 VIII. Future Directions: High-Throughput Sequencing and Metagenomics.............................................................. 27 Acknowledgments ................................................................................................................................................... 28 References .............................................................................................................................................................. 28
Summary The purple bacteria occupy a unique position among photosynthetic bacteria. Nested within the various proteobacterial lineages, the origin and evolution of purple bacterial photosynthesis has been the topic of innumerable debates. Attempts to reconstruct the evolutionary history of individual photosynthetic protein families have further fueled debate over lateral vs. vertical transfer of genetic elements. In this era of high-throughput sequencing we can begin to distance ourselves from this dependency on single-gene and single-protein phylogenies. Here we present automated comparative genomics-based methods useful for reconstructing the genomic history of not only the purple bacterial lineage, but the proteobacterial lineage as a whole. These reconstructions integrate phylogenetic data inferred from 200 to more than 1000 protein families common to all or part of the proteobacterial lineage. This framework allows us to reconstruct the evolutionary history of *Author for correspondence, email:
[email protected] C. Neil Hunter, Fevzi Daldal, Marion C. Thurnauer and J. Thomas Beatty (eds): The Purple Phototrophic Bacteria, pp. 17–29. © 2009 Springer Science + Business Media B.V.
18
Wesley D. Swingley, Robert E. Blankenship and Jason Raymond
each proteobacterial class and parse out the finer relationships among photosynthetic species. By telescoping inward on protein families of interest, we can delve deeper than ever before into the convoluted evolutionary origin of the primary photosynthetic traits, phototrophy and autotrophy. While these full-genome comparisons clarify the nature of many poorly understood phylogenetic relationships, they do not yet serve to resolve the entire mystery surrounding the history of proteobacterial phototrophy. I. Introduction Among six phyla of photosynthetic prokaryotes, the purple proteobacteria are the most metabolically diverse. Unlike some other groups of phototrophs such as cyanobacteria or green sulfur bacteria, in which all members are capable of photosynthesis, the proteobacteria have a remarkable range of metabolic lifestyles. Within this group are organisms that grow aerobically and anaerobically, photoautotrophically and photoheterotrophically, and many that utilize a diverse number of metabolic pathways for energy generation, carbon assimilation, as well as nitrogen, sulfur, and phosphorous metabolism (Imhoff et al., 2005; Chapter 1, Madigan and Jung). This diversity reflects the widespread nature of these organisms throughout the proteobacterial lineage. In many cases, phototrophic proteobacteria share genus-level relationships with non-phototrophs. This fact could be explained as either a loss of ancestral phototrophy or a lateral gain of the genes necessary to produce photosynthetic pigments and proteins. Although the proteobacteria are the most sequenced bacterial phylum to date, with more than 500 genomes complete or in progress (according to NCBI), the purple bacteria have been far less sampled, with fewer than 30 genome projects as of this writing, fewer than half of which are completed. This disparity is, in part, due to the overwhelming medical interest in proteobacteria, but it speaks volumes that the first purple bacterial sequence, Rhodopseudomonas palustris CGA009, was not published until 2004 (Larimer et al., 2004), nine years after the first complete bacterial genome sequence (Fleischmann et al., 1995). Notwithstanding, preliminary sequence data from Rhodobacter capsulatus and Rhodobacter sphaeroides 2.4.1 have been available since 2001 (http://www. integrated genomics.com/genomereleases.html; Mackenzie et al., 2001).
Early purple bacteria sequencing projects focused on model organisms, a common trend throughout the genome sequencing arena, completing the heavily studied species Rhodobacter sphaeroides (Choudhary et al., 2007), Rhodopseudomonas palustris (Larimer et al., 2004), Rhodospirillum rubrum (NCBI), and both phototrophic and nonphototrophic Bradyrhizobia (Kaneko et al., 2002; Giraud et al., 2007). Recently, the focus has begun to shift to those purple bacteria with a significant involvement in oceanic carbon and nutrient cycling, with the completed sequence of Roseobacter denitrificans Och 114 (Swingley et al., 2007) leading the way to a tremendous number of marine organismal genome sequences in progress or recently completed . While all of the above sequences are from the alpha class of proteobacteria, few other phototrophic proteobacteria have been targeted for sequencing. Three of these belong to the gammaproteobacteria, Halorhodospira halophila SL1, Congregibacter litoralis KT71, and the undescribed marine strain HTCC2080 (NCBI). Of these, Halorhodospira and Congregibacter have been characterized as phototrophic (Imhoff and Süling, 1996; Fuchs et al., 2007), but HTCC2080, which carries the genetic capability for phototrophy, has not yet been shown to produce photosynthetic pigments. Two major clades of phototrophic betaproteobacteria, genera Rubrivivax and Rhodoferax, have yet to be sequenced, as the only sequenced members, the former Rubrivivax gelatinosus PM1 (now Methylibium petroleiphilum PM1) and Rhodoferax ferrireducens T118, are non-phototrophic strains. M. petroleiphilum was only recently disambiguated from a cluster of closely-related species (both phototrophic and nonphototrophic) lumped under a relatively small number of genera (Nakatsu et al., 2006). II. The Alphaproteobacteria
Abbreviations: AAP – aerobic anoxygenic phototrophs; BChl – bacteriochlorophyll; GOS – Global Ocean Survey; kb – kilobase(s); PGC – photosynthesis gene cluster; rubisco– ribulose 1,5-bisphosphate carboxylase
The vast majority of phototrophic proteobacteria are spread throughout the alpha lineage. However, phototrophy is by no means ubiquitous in alphapro-
Chapter 2
Evolution of Purple Bacteria
teobacteria. Less than 22% of all alphaproteobacteria currently sequenced bear signatures of phototrophy (presence of chlorophyll biosynthesis or reaction center genes). While the number of sequenced phototrophic taxa is low, they are widespread throughout the alpha cluster (Fig. 1), which raises important questions on the origin of their phototrophy. Furthermore, a sequencing bias for the large number of agriculturally-important alphaproteobacteria could skew the aforementioned ratio. In fact, 18 of 39 currently available ‘in progress’ proteobacteria genomes at NCBI are potentially phototrophic. Unlike phototrophy, carbon-fixation is not common among alphaproteobacteria. The typical method of autotrophic carbon-fixation in all proteobacteria is through the Calvin Cycle using the key enzyme ribulose 1,5-bisphosphate carboxylase (rubisco). Variations of this enzyme are found throughout the tree of life (Tabita, 1999), but it is most famously associated with chlorophyll and photoautotrophy in photosynthetic organisms. However, a unique class of alphaproteobacteria (discussed below) grows photoheterotrophically rather than photoautotrophically. As a result, a relatively low proportion of alphaproteobacteria are photoautotrophic when compared to the beta- and gammaproteobacteria, where phototrophy is more rare. III. Aerobic Purple Bacteria Canonically speaking, purple bacteria grow photoautotrophically only at low oxygen levels, while they down-regulate their photosynthetic apparatus under higher oxygen levels and instead grow chemotrophically using organic carbon (Harashima et al., 1982; Yurkov and van Gemerden, 1993; Bauer et al., 2003; Yurkov and Csotonyi, 2003). Recent oceanic metagenomic surveys have created a burgeoning interest in a related class of purple bacteria that grow primarily under aerobic conditions. Dubbed the aerobic anoxygenic phototrophs (AAP or AAnP) or aerobic phototrophic bacteria (APB), these purple bacteria produce photosynthetic pigments and carry out phototrophic metabolism only under aerobic conditions (Shiba et al., 1979; Shiba and Harashima, 1986; Yurkov and van Gemerden, 1993;Yurkov and Beatty, 1998; Chapter 3, Yurkov and Csotonyi). However, they produce bacteriochlorophyll (BChl) a only in the dark, possibly to avoid the potential for oxidative damage caused by the excitation of chlorophyll
19
Fig. 1. 16S rRNA tree of the Proteobacteria, based on sequences from the Ribosomal Database [http://rdp.cme.msu.edu]. To facilitate visualization, 16S rRNA gene data are filtered so that only lineages at 97% or less 16S rRNA gene distance are shown (i.e. filtered at roughly the species level). Bold lineages are those proteobacterial species with one or more genome-sequenced members. Bold, numbered lineages indicate the positions of sequenced purple/phototrophic proteobacteria in the tree. Other lineages have 16S rRNA data available from isolates or environmental samples, but do not have genome projects in progress as of this writing. Proteobacterial classes are shown in Greek letters. Numbered purple proteobacteria are: 1. Rhodospirillum rubrum, 2. Rhodospirillum centenum, 3. Erythrobacter NAP1, 4. Roseobacter denitrificans (multiple strains), 5. Roseovarius sp. (multiple strains), 6. Loktanella, 7. Jannaschia sp. CCS1, 8. Rhodobacter sphaeroides (incl. R. capsulatus), 9. Fulvimarina, 10. Methylobacterium, 11. Bradyrhizobium (multiple species), 12. Rhodopseudomonas palustris (multiple strains), 13. Thermochromatium tepidum (proprietary sequence), 14. Halorhodospira sp. SL1, 15. Rubrivivas gelatinosus (photosynthetic gene cluster only, no genome)
intermediates (Beatty, 2002). While these AAP have long been overlooked in oceanic studies, recent data shows that they are widely distributed in the oceans and, consequently, their contribution to the global carbon cycle could be significant (Kolber et al., 2001; Buchan et al., 2005; Yutin et al., 2007). Historically, the origin of aerobiosis in the AAP has been a puzzling problem, especially in regards to the regulation of photosynthetic gene expression. The largest cluster of AAP, the Roseobacter clade, fall within the order Rhodobacterales and these species are closely related to the model species Rhodobacter sphaeroides and Rhodobacter capsulatus (Yurkov and
20
Wesley D. Swingley, Robert E. Blankenship and Jason Raymond
Fig. 2. Schematic comparison of photosynthesis gene clusters (PGC) from various purple bacteria. Lines represent transpositions of syntenous gene regions between two PGC. Numbers annotated above the Rhodospirillum rubrum PGC represent the location on the genome of each separate PGC segment. The lines with arrows represent inversion of the genes contained between the lines and the other lines represent shifts in position without inversions. See also Color Plate 1, Fig. 1, for a color version with additional information.
Beatty, 1998). Even within this group of organisms the common proteobacterial trend holds true in that not all members are phototrophic. Thus far, nearly all sequenced species in this cluster lack rubisco and thus the ability to fix carbon autotrophically (Swingley et al., 2007). The reason for the seemingly detrimental absence of autotrophic carbon-fixation is unclear. Distantly related families of AAP, such as some Erythrobacter and Congregibacter species, also lack rubisco (Swingley et al., 2007; Yutin et al., 2007). One possibility is that the presence of rubisco with a poor specificity for CO2 vs. O2 is a liability in aerobic conditions, which would lead to a large amount of photorespiration — which has led to multiple rubisco gene losses throughout evolutionary history. The only AAP shown to exhibit photoautotrophic carbon-fixation are several members of the plant endosymbiont genus Bradyrhizobia (Giraud et al., 2007). IV. The Photosynthesis Gene Cluster and its Role in Evolution In all known cases, the majority of the genetic information needed to build the photosynthetic apparatus
in purple bacteria is clustered in large groups of genes known as the Photosynthesis Gene Cluster (PGC). In most organisms, a single cluster of 40–50 kb contains all the structural genes for the photosystem, as well as genes for the latter stages of BChl and carotenoid biosynthesis (Fig. 2). Exceptions to this pattern are Rhodospirillum rubrum and Methylobacterium species, which contain three separate PGC clusters that are widely dispersed in the chromosome. Figure 2 shows gene neighborhoods for the PGC in several purple bacterial species. It is apparent that while the genes are organized into one or a few large clusters, the precise gene organization within the cluster is highly variable. It is possible to produce the various gene synteny arrangements found in PGCs by inversions and rearrangements, as shown in Fig. 2. The dramatic clustering of nearly all genes involved in phototrophy has provided a perfect framework for the lateral spread of phototrophy. It is intriguing that there is not a similar clustering of photosynthesisrelated genes in other groups of phototrophs, with the exception of the heliobacteria (Xiong et al., 1998). The gene organization of the PGC in heliobacteria is not generally similar to that found in the purple bacteria, with the exception of some of the operons
Chapter 2
Evolution of Purple Bacteria
for multisubunit enzyme complexes involved in BChl biosynthesis. This suggests that the assembly of genes into the PGC in the purple bacteria and heliobacteria may have been independent evolutionary events. The heliobacteria seem to have little in common with the purple bacteria in that they have a type I reaction center, are strict anaerobes and are incapable of photoautotrophic metabolism. Why these two very different groups of phototrophs have both assembled the genetic information needed to do photosynthesis into large gene clusters while other groups of phototrophs have not done so is a question for which no answer is yet available. The possible functional role of such extreme clustering of genes that code for functionally related proteins has a long history in biology, starting with the lac operon of Escherichia coli (Jacob and Monod, 1961). The utility of clustering and co-expression of genes that code for parts of a multiprotein complex is easy to appreciate. A single mRNA encoding each component of a complex, which is then translated into proteins that then associate with each other, permits precise control of the expression levels and spatial proximity of the components. However, what is not so clear is the benefit of a large gene cluster such as the PGC, which consists of more than thirty genes. This cluster has genes that are divergently transcribed, so they cannot be part of a single operon, which is confirmed by transcriptional analysis (Kiley and Kaplan, 1988; Wellington and Beatty, 1991). Furthermore, the dosage requirements of the various photosynthetic complexes and biosynthetic enzymes are vastly different. Regulatory elements can easily coordinate gene expression in genes that are distant on the chromosome, so there seems to be little functional need for physical clustering. The ‘selfish operon’ theory as formulated by Lawrence and Roth (Lawrence and Roth, 1996; Lawrence, 2003) proposes that operons assemble to facilitate horizontal transfer of an operon and its easy replacement if it happens to be lost. This theory has been questioned and some predictions of the selfish operon theory have not been observed (Pál and Hurst, 2004; Price et al., 2005). Nagashima et al. (1997) analyzed the gene sequences of the L and M reaction center proteins and concluded that significant amounts of horizontal gene transfer has taken place among purple bacteria. This same conclusion was extended to the PGC in a later study (Igarashi et al., 2001). However, this issue has not been addressed in detail since the advent of large-scale genome sequencing
21 of photosynthetic bacteria and it represents an area that needs to be investigated in more detail. V. Proteobacterial Comparative Genomics: Photosynthetic versus Non-Photosynthetic Proteins The foremost challenge in using a wealth of genomic data to reconstruct evolutionary events is how to integrate thousands of genes worth of (often discordant) phylogenies. One approach to this end involves concatenating multiple aligned gene or protein sequences and inferring a single phylogeny for this concatenated dataset. The underlying assumption for this approach is that the incongruence in single phylogenies, caused by data that are poorly fit by evolutionary models or by horizontal gene transfer, will be overtaken by the reinforcing signal of vertical evolution. Simulation studies have illustrated the robustness of concatenated datasets while raising the important caveat that, under some circumstances, concatenation only serves to reinforce the wrong tree (Gadagkar et al., 2005). No doubt the approach will continue to improve with more rigorous methods for determining which genes are suitable for concatenation. Concatenated phylogenies represent an averaged evolutionary overview of a set of genes or proteins, providing a potentially useful broad brush history but also masking potentially interesting details. It is important that this, and other approaches, be considered along with — not in lieu of — rigorous, single gene or protein phylogenies. Keeping these assumptions and ideas in mind, it is useful to examine the photosynthesis gene phylogeny for a large set of sequenced proteobacteria and contrast this with the 16S rRNA tree. This method involves the concatenation of specific PGC proteins to compare the evolutionary history of proteobacterial photosynthesis with the history of the phylum. Figure 3 (left) shows the phylogeny inferred from concatenating 208 orthologous proteins found in 29 proteobacterial genomes and several non-proteobacterial phototrophic bacteria, yielding an alignment of nearly 69,000 positions. Figure 3 (right) shows the concatenation of 38 proteins from the PGC of Rhodobacter sphaeroides with their homologs in other bacterial phototrophs, which resulted in an alignment of nearly 16,400 positions. The 208-protein (all-protein) concatenation (Fig. 3, left) is generally consistent with the 16S rRNA tree in the distinct
22
Wesley D. Swingley, Robert E. Blankenship and Jason Raymond
Fig. 3. A comparison of purple bacterial phylogeny based on whole genome proteins versus only photosynthesis-related proteins. On the left is the phylogeny based on concatenation of 208 proteins found in all, or all but one, genome from representative proteobacteria and several other phototrophic phyla. The tree on the right shows the phylogeny based on the concatenation of 38 proteins from the photosynthetic gene cluster (PGC) in purple proteobacteria and their homologs in other phototrophic phyla. Taxa in gray are present in only one of the two trees, whereas taxa in black are present in both trees. The lineage line-width is proportional to topological (branching order) differences between the two trees. Proteobacterial classes are indicated by Greek letters and surrounded by dashed lines. PGC protein homologs included in the concatenation are: LhaA, Lhb1, PpaA, PpsR, PucC, PufL, PufM, PufQ, PufX, PuhH, AcsF, BchB, BchC, BchD, BchE, BchF, BchG, BchH, BchI, BchJ, BchL, BchM, BchN, BchO, BchP, BchX, BchY, BchZ, CrtA, CrtB, CrtC, CrtD, CrtE, CrtF, CrtI, CrtK, Idi, and Lha1.
separation of the proteobacterial classes as well as with the branching order of species within these classes. In contrast, the 38-protein PGC concatenation shows no congruence at the class level with either the 16S rRNA tree or the all-protein concatenation. Identical branching is observed at lower taxonomic levels, as seen for instance in the topology of order Rhodobacteriales (Rhodobacter sphaeroides through Roseobacter denitrificans), and also in conservation of the Rhodopseudomonas/Bradyrhizobium clade. At face value, the PGC tree is consistent with the idea of an alpha origin of proteobacterial phototrophy, followed by subsequent horizontal gene transfer into members of the beta- and gammaproteobacterial classes. Note that the position of the non-proteobacterial phototrophs as outgroups on both trees is based on midpoint rooting on the longest branch in the tree; in principle, one could use additional lines of evidence to root the trees along other lineages.
VI. Origin and Evolution of Proteobacterial Phototrophy The scattered distribution of phototrophic organisms throughout the proteobacteria has presented a longstanding puzzle. On one hand, the sheer number of lateral transfer events necessary to accommodate such a distribution seems overwhelming. Conversely, the number of deletion events required seems to rule out a phototrophic proteobacterial ancestor. However, by breaking down the problem into manageable pieces, some conclusions can be drawn. Because the alphaproteobacteria represent the largest repository for phototrophy, they are the best group for studying the elusive origin of this trait. One thing made clear by the growing library of sequence data is that alphaproteobacterial phototrophy is evenly dispersed across evolutionary space (Fig. 1), indicating that vertical transfer coupled with gene loss may be more important than lateral gene transfer in this group. However, it is clear that the earliest
Chapter 2
Evolution of Purple Bacteria
phototrophic ancestor would likely occur after the divergence of the Rickettsiales — none of which are phototrophic — although the highly reduced genomes of these intracellular parasites could have selectively lost such an extraneous metabolic activity. In fact, the next earliest-branching clade includes the genus Acidiphilium, which contains phototrophic species (Hiraishi and Shimada, 2001). One of the largest and best studied alphaproteobacterial lineages is the Rhodobacteraceae. Over half of the in progress or complete phototrophic alpha genomes belong to this clade. However, even though the Rhodobacteraceae is a monophyletic, terminal clade, it contains a large number of non-phototrophic species. The conservation of phototrophy in this cluster suggests that the loss of phototrophy (and indeed photosynthesis as a whole) in these species may be quite common. One possible reason for such a phenomenon stems from the highly diverse metabolic library of Rhodobacteraceae species (Buchan et al., 2005). It is possible that such species modulate their metabolism based on availability of nutrients and/or light availability in their ecological niche. Given that bacterial genomes are generally streamlined, the loss of photosynthesis is likely in environments where photosynthesis is not a practical metabolic option. Most of the other sequenced phototrophic proteobacteria are found in the alphaproteobacterial family Bradyrhizobiaceae. This family is composed of three major genera, Bradyrhizobium, Rhodopseudomonas, and Nitrobacter, of which the former two both contain phototrophic species. Internal membranes in Nitrobacter species that are reminiscent of purple bacterial photosynthetic membranes suggest that this genus may have originated from a phototrophic ancestor. While all known Rhodopseudomonas species are phototrophic, there is only one monophyletic cluster of phototrophic Bradyrhizobium species, typified by the first discovered phototrophic strain Bradyrhizobium sp. BTAi1. Because Bradyrhizobium strains have typically been isolated from legume root nodules, it has been speculated that selective pressure has caused the loss of photosynthetic activity in many species (Giraud et al., 2007). This hypothesis is supported by the fact that all photosynthetic strains have been isolated from light-accessible stem nodules rather than light-inaccessible root nodules (Willems et al., 2003). Naturally, a ‘chicken or egg’ argument can be posed for any case of photosynthesis gene loss vs. gain. In cases such as the Rhodobacteraceae where a vast
23 majority of polyphyletic species are phototrophic, it is likely that phototrophy is a vertically inherited trait. However, in the case of Bradyrhizobium, the monophyletic phototrophic cluster could be the result of a single lateral transfer. While the presence of the closely related phototrophic Rhodopseudomonas and remnant ‘photosynthetic’ membranes in Nitrobacter support a phototrophic ancestry, they are not conclusive proof. A wealth of sequenced Bradyrhizobia, both phototrophic (strains BTAi1 and ORS278) and nonphototrophic (strain USDA110), and other Bradyrhyzobiaceae (five Rhodopseudomonas and three Nitrobacter species) allow for thorough analysis of this problem. One striking case is the similarity of gene orientation (synteny) between the phototrophic and non-phototrophic Bradyrhizobia (Fig. 4). While the number of homologous genes (E-value greater than 10–50) is greater between BTAi1 and ORS278, the overall gene order is more conserved between either of the two phototrophic strains and USDA110. This suggests two possible conclusions: a) genome rearrangement is a common occurrence and syntenic homology between distant strains occurs by coincidence or lower rates of recombination; or b) phototrophic and non-phototrophic strains are more closely related to each other than previous work has shown (Willems et al., 2003). It is very difficult to support the second case, which would contradict multiple phylogenetic reconstructions from 16S rRNA and rRNA ITS sequences as well as that presented in this chapter. The immense library of literature discussing genome rearrangement strongly supports the first conclusion. A high rate of genome rearrangement also leads to an increase in the rate of gene duplication and loss. This could have greater implications on the scattered nature of phototrophy throughout the proteobacterial clade, where a high rate of gene loss could account for numerous closely-related strains with and without phototrophic ability. Recent work by Haffa et al. (A.L. Haffa, personal communication) suggests that viral invasion has had a great influence on both random and directed gene losses in proteobacterial species, a fact that further convolutes evolutionary history in these species. Expanding beyond the phototrophic alphaproteobacteria, we see only a few select phototrophic clades throughout the rest of the proteobacterial tree. Methylibium petroleiphilum strain PM1and Rhodoferax ferrireducens T118 are the only sequenced members from phototrophic betaproteobacterial
24
Wesley D. Swingley, Robert E. Blankenship and Jason Raymond
Fig. 4. Dot-plot comparison representing a genome alignment between Bradyrhizobium species. Dots represent aligned nucleotide regions with higher than 75% identity. Only alignments with a BLAST score better than E=10-50 are shown. Axes represent sequence location in megabases. Dots in sequence along the 1:1 axis represent laterally homologous gene regions. Dots in sequence along the -1:1 axis represent contiguous translocations.
clades. Unfortunately, neither is a phototrophic species. However, Nagashima et al. (1993) sequenced the entire PGC from a phototrophic member of the same family, Rubrivivax gelatinosus, providing the most detailed glimpse so far into betaproteobacterial phototrophy. Halorhodospira halophilum SL1 is the only completely sequenced phototrophic gam-
maproteobacterium. Sequence data for two new phototrophic gammas, Congregibacter litoralis KT71 and unnamed marine strain HTCC2080, are currently available, but as yet incomplete. The sparse distribution of phototrophic beta- and gammaproteobacteria suggests that phototrophy did not originate in these clades. This idea is supported by the evolutionary history of their photosynthetic proteins (Fig. 3, right). Rubrivivax phototrophy appears to originate early on the Bradyrhizobiaceae branch, before the divergence of Rhodopseudomonas and Bradyrhizobium. The lack of conservation in the gene order between Rubrivivax and either alpha PGC suggests that this represents a relatively old lateral transfer event. On the other hand, the PGC gene order in Congregibacter and gamma HTCC2080 shares a great deal of similarity to their apparent origin from a close relative of Fulvimarina pelagi HTCC2506. The lesser homology between Halorhodospira and Fulvimarina illustrates the longer phylogenetic distance between the PGC in these two species; however, the origin is still quite near the Fulvimarina cluster. The precise origin of phototrophy in individual families of beta and gamma proteobacteria will not be resolved until further taxa are sequenced (if not full genomes, then at least PGC sequences). However, this chapter shows that only a few lateral transfer events could account for all non-alpha proteobacterial phototrophy. Less clear is the origin of the few chlorophyll biosynthesis genes that are not always contained within the PGC, such as the aerobic and anaerobic monomethyl ester cyclases (acsF and bchE, respectively) and divinyl reductase (gene not yet identified for all purple bacteria). These genes are not always present in the PGC of beta and gamma phototrophs, implying that they were either transferred independently by a later event or were originally in the PGC and have since translocated away. Such an outward translocation seems in direct opposition to our current understanding of bacterial gene clusters/operons (as discussed above). VII. Origin and Evolution of Proteobacterial Carbon-fixation Carbon-fixation is an inextricable component of photosynthesis. However, when studying purple bacterial photosynthesis we must be careful to extract and analyze the two portions independently. As discussed in the preceding section, phototrophy is
Chapter 2
Evolution of Purple Bacteria
unevenly distributed throughout the proteobacteria and the same is true for carbon-fixation. Nevertheless, these two traits do not appear to be linked in any clear fashion. Certainly there are many species that grow photoautotrophically using Calvin Cycle carbon-fixation, but just as many grow photoheterotrophically. On the other hand, many proteobacterial species fix CO2 without the use of light energy. Unlike photosynthetic proteins, the subunits of rubisco, RbcL/CbbL and RbcS/CbbS, are evolutionarily split into several classes (Watson and Tabita, 1997; Tabita, 1999; Ashida et al., 2005; Tabita et al., 2007). Therefore, while carbon-fixation may appear conserved in any given monophyletic cluster, the actual genes may have very different origins. This
25 is most striking in the Rhodobacteraceae, where some AAP species contain a rubisco that shares a close phylogenetic origin with Type IV rubisco-like proteins (Swingley et al., 2007) (Fig. 5 and Table 1). The role of the Type IV enzymes has not been clearly described; however, they have not been shown to fix CO2 (Hanson and Tabita, 2001; Imker et al., 2007; Tabita et al., 2007). There are also many cases where members of a single genus carry copies of rubisco from different family-types. Most notable among these are the Rhodopseudomonas species — which contain up to four different types of rubisco — and some Rhodobacter species that code for a number of rubisco variations (Table 1). Even with the likelihood of widespread rubisco
Fig. 5. Phylogenetic representation of purple bacterial ribulose 1,5-bisphosphate (rubisco) sequences. Roman numeral labels identify the recognized distinct ‘Form’ family of rubisco or rubisco-like protein. This analysis revealed three divergent groups of Form IV proteins that are further split into types A, B, and C, which are analogous to those described by Tabita (2007): A = IV-DeepYkr, B = IV-Photo, C = IV-NonPhoto.
Wesley D. Swingley, Robert E. Blankenship and Jason Raymond
26
Table 1. Ribulose 1,5-bisphosphate (rubisco) content of the purple bacteria and non-phototrophic members of genera that contain known phototrophs. rubisco Form-types are labeled as shown in Fig. 5 Family
α
Organism Bradyrhizobium japonicum USDA 110 Bradyrhizobium sp. BTAi1 Bradyrhizobium sp. ORS278 Dinoroseobacter shibae DFL 12 Erythrobacter litoralis HTCC2594 Erythrobacter sp. NAP1 Erythrobacter sp. SD-21 Fulvimarina pelagi HTCC2506 Jannaschia sp. CCS1 Loktanella vestfoldensis SKA53 Magnetospirillum magneticum AMB-1 Magnetospirillum magnetotacticum MS-1 Methylobacterium chloromethanicum CM4 Methylobacterium extorquens PA1 Methylobacterium sp. 4-46 Oceanicola batsensis HTCC2597 Oceanicola granulosus HTCC2516 Rhodobacter capsulatus Rhodobacter sphaeroides 2.4.1 Rhodobacter sphaeroides ATCC 17025 Rhodobacter sphaeroides ATCC 17029 Rhodocista centenaria Rhodopseudomonas palustris BisA53 Rhodopseudomonas palustris BisB18 Rhodopseudomonas palustris BisB5 Rhodopseudomonas palustris CGA009 Rhodopseudomonas palustris HaA2 Rhodospirillum rubrum ATCC 11170 Roseobacter denitrificans OCh 114 Roseobacter sp. AzwK-3b Roseobacter sp. CCS2 Roseobacter sp. MED193 Roseobacter sp. SK209-2-6 Roseovarius nubinhibens ISM Roseovarius sp. 217 Roseovarius sp. HTCC2601 Roseovarius sp. TM1035 Sulfitobacter sp. EE-36 Sulfitobacter sp. NAS-14.1
β
Methylibium petroleiphilum PM1 Rhodoferax ferrireducens T118
Form I ID IA, ID (x2) IA, ID ––––-
Form II
Form IV
IVC IVC –II II (×2) ––IVC –IVA IA IC IA, IC IC IA, ID ID ID IA, ID ID ID
II II II II II II II II II II
IVA, IVB IVA, IVB IVA, IVB IVA, IVB IVA, IVB IVA
–––IVC –––ID –––-
IVC (×2)
IC (x2) II
Chapter 2
Evolution of Purple Bacteria
27
Table 1. Continued Family
Organism
Form I
γ
Congregibacter litoralis KT71 Halorhodospira halophila SL1 marine gamma proteobacterium HTCC2080
–IA –-
Chlorobium tepidum TLS Chloroflexus aurantiacus J-10-fl Clostridium acetobutylicum ATCC 824 Desulfitobacterium hafniense Y51 Pelodictyon luteolum DSM 273 Prochlorococcus marinus MIT 9313 Roseiflexus sp. RS-1 Synechococcus sp. JA-2-3B´a(2-13)
lateral gene transfer, some phylogenetic information can be drawn from the proteobacterial genome tree. The aforementioned Rhodopseudomonas and Bradyrhizobium species likely share a Form ID rubiscocontaining ancestor with the related non-phototrophic genus Nitrobacter. However, the presence of many proteins from unrelated species in this phylogenetic cluster indicates that lateral transfer has likely occurred on multiple occasions. The presence of Form II in all Rhodobacter species also indicates that it is ancestral to this genus. Given the lack of autotrophic CO2-fixation in the AAP, Form II rubisco may not be ancestral to the Rhodobacteraceae in general. Nevertheless, the presence of Form IV-like proteins in AAPs suggests its possible ancestry to this group. Although Form IB rubisco, representing the traditional plant and cyanobacterial class of enzymes, is not present in any purple bacteria, other Form I enzymes are common throughout the proteobacteria. Like phototrophy, the origin and evolution of carbon-fixation in purple bacteria is difficult to clarify. The apparently rampant lateral transfer of Calvin Cycle enzymes in proteobacteria (and bacteria/Archaea as a whole) creates a high level of background noise that is difficult to separate from phylogenetic signals. Even with such a confusing situation, carbon-fixation is clearly an ancestral trait for the Bradyrhizobiaceae and likely many other lineages. However, given the current level of sequence coverage, we cannot make any further conclusions on the origin of purple bacterial — or proteobacterial for that matter — carbon-fixation.
Form II
Form IV IVA IVB
–––IVB IA –IB
VIII. Future Directions: High-Throughput Sequencing and Metagenomics One of the exciting recent advances in microbiology is the application of genomics to complex microbial communities taken directly from their natural environments. Environmental genomics, or metagenomics, provides an opportunity to sidestep the so-called cultivation bias: the fact that only a few percent of microbes from natural communities have been grown and studied in a laboratory. Indeed the recent discovery of phototrophy among the phylum Acidobacteria (Bryant et al., 2007) was founded on analysis of a Yellowstone metagenome sequencing project. Environmental sequencing was also used to demonstrate the abundance of rhodopsin-based phototrophy in the oceans (Béjà et al., 2000). This has been borne out in large-scale metagenomics of plankton in the Sargasso Sea (Venter et al., 2004) and other marine samples taken as part of the ongoing Global Ocean Survey (GOS) (Yooseph et al., 2007). These data underscore the remarkable phenotypic diversity that has yet to be characterized in the laboratory. For instance, the GOS dataset contains several hundred homologs of genes encoding the purple reaction center subunits. Homologs to known marine strains are particularly abundant, but the reaction center proteins of nearly every phototrophic proteobacterium discussed in this chapter has multiple close homologs in the GOS data. Beyond the enormous repository of marine metagenomes, a growing number of projects should provide additional insight into the genetic diversity of
28
Wesley D. Swingley, Robert E. Blankenship and Jason Raymond
phototrophy. These include samples from hot springs, wastewater treatment ponds, hypersaline microbial mats, even soil and sediment samples that receive some solar input. While exciting, these approaches also demand new ways of thinking about experimental validation and follow-through. In many cases, clone libraries are built from small inserts of DNA, typically 3–10 kb in length. If and when truly novel discoveries are sifted out of these enormous datasets, tracing the origin of sequences in these data will be a serious challenge. Small insert libraries will provide, at most, a handful of contextual genes that provide almost no insight into the organism’s physiology. Isolating the genome sequence of a single, specific organism from a complex community is incredibly difficult — though it is amenable with genetic ‘clues’ that would come from higher levels of targeted sequencing. This underscores the importance of either sequencing, or at the very least archiving, large insert (>100 kb) libraries that can provide a more substantial glimpse into the neighborhoods surrounding genes of interest. Ultimately, extrapolating useful information from enormous repositories of ‘letters on a page’ will require unprecedented synergism of genomics with clever new microbiological techniques. Acknowledgments WDS is supported by a Postdoctoral Fellowship for Foreign Researchers from the Japan Society for the Promotion of Science. Work on the evolution of photosynthesis by REB has been supported by a grant from the Exobiology program from NASA. References Ashida H, Danchin A and Yokota A (2005) Was photosynthetic rubisco recruited by acquisitive evolution from rubisco-like proteins involved in sulfur metabolism? Res Microbiol 156: 611–618 Bauer CE, Elsen S, Swem LR, Swem DL and Masuda S (2003) Redox and light regulation of gene expression in photosynthetic prokaryotes. Philos Trans R Soc Lond B Biol Sci 358: 147–154 Beatty JT (2002) On the natural selection and evolution of the aerobic phototrophic bacteria. Photosynth Res 73: 109–114 Béjà O, Aravind L, Koonin EV, Suzuki MT, Hadd A, Nguyen LP, Jovanovich SB, Gates CM, Feldman RA, Spudich JL, Spudich EN and DeLong EF (2000) Bacterial rhodopsin: Evidence for a new type of phototrophy in the sea. Science 289: 1902–1906 Bryant DA, Garcia Costas AM, Maresca JA, Chew AGM,
Klatt CG, Bateson MM, Tallon LJ, Hostetler J, Nelson WC, Heidelberg JF and Ward DM (2007) Candidatus Chloracidobacterium thermophilum: An aerobic phototrophic acidobacterium. Science 317: 523–526 Buchan A, Gonzalez J and Moran MA (2005) Overview of the marine Roseobacter lineage. Appl Environ Microbiol 71: 5665–5677 Choudhary M, Zanhua X, Fu YX and Kaplan S (2007) Genome analyses of three strains of Rhodobacter sphaeroides: Evidence of rapid evolution of chromosome II. J Bacteriol 189: 1914–1921 Fleischmann RD, Adams MD, White O, Clayton RA, Kirkness EF, Kerlavage AR, Bult CJ, Tomb JF, Dougherty BA, Merrick JM, McKenney K, Sutton G, Fitzhugh W, Fields C, Gocayne JD, Scott J, Shirley R, Liu LI, Glodek A, Kelley JM, Weidman JF, Phillips CA, Spriggs T, Hedblom E, Cotton MD, Utterback TR, Hanna MC, Nguyen DT, Saudek DM, Brandon RC, Fine LD, Fritchman JL, Fuhrmann JL, Geoghagen NSM, Gnehm CL, McDonald LA, Small KV, Fraser CM, Smith HO and Venter JC (1995) Whole-genome random sequencing and assembly of Haemophilus influenzae Rd. Science 269: 496–512 Fuchs BM, Spring S, Teeling H, Quast C, Wulf J, Schattenhofer M, Yan S, Ferriera S, Johnson J, Glockner FO and Amann R (2007) Characterization of a marine gammaproteobacterium capable of aerobic anoxygenic photosynthesis. Proc Natl Acad Sci USA 104: 2891–2896 Gadagkar SR, Rosenberg MS and Kumar S (2005) Inferring species phylogenies from multiple genes: Concatenated sequence tree versus consensus gene tree. J Exp Zoolog B Mol Dev Evol 304B: 64–74 Giraud E, Moulin L, Vallenet D, Barbe V, Cytryn E, Avarre J-C, Jaubert M, Simon D, Cartieaux F, Prin Y, Bena G, Hannibal L, Fardoux J, Kojadinovic M, Vuillet L, Lajus A, Cruveiller S, Rouy Z, Mangenot S, Segurens B, Dossat C, Franck WL, Chang W-S, Saunders E, Bruce D, Richardson P, Normand P, Dreyfus B, Pignol D, Stacey G, Emerich D, Verméglio A, Medigue C and Sadowsky M (2007) Legumes symbioses: Absence of nod genes in photosynthetic Bradyrhizobia. Science 316: 1307–1312 Hanson TE and Tabita FR (2001) A ribulose-1,5-bisphosphate carboxylase/oxygenase (RubisCO)-like protein from Chlorobium tepidum that is involved with sulfur metabolism and the response to oxidative stress. Proc Natl Acad Sci USA 98: 4397–4402 Harashima K, Nakagawa M and Murata N (1982) Photochemical activities of bacteriochlorophyll in aerobically grown cells of aerobic heterotrophs, Erythrobacter species (Och 114) and Erythrobacter longus (Och 101). Plant Cell Physiol 23: 185–193 Hiraishi A and Shimada K (2001) Aerobic anoxygenic photosynthetic bacteria with zinc-bacteriochlorophyll. J Gen Appl Microbiol 47: 161–180 Igarashi N, Harada J, Nagashima S, Matsuura K, Shimada K and Nagashima KVP (2001) Horizontal transfer of the photosynthesis gene cluster and operon rearrangement in purple bacteria. J Mol Evol 52: 333–341 Imhoff JF, Hiraishi A and Suling J (2005) Anoxygenic phototrophic purple bacteria. In: Brenner DJ, Krieg NR and Staley JT (eds) Bergey’s Manual of Systematic Bacteriology, pp 119–132. Springer, New York Imhoff JF and Süling J (1996) The phylogenetic relationship
Chapter 2
Evolution of Purple Bacteria
among Ectothiorhodospiraceae: A reevaluation of their taxonomy on the basis of 16S rDNA analyses. Arch Microbiol 165: 106–113 Imker HJ, Fedorov AA, Fedorov EV, Almo SC and Gerlt JA (2007) Mechanistic diversity in the rubisco superfamily: The ‘enolase’ in the methionine salvage pathway in Geobacillus kaustophilus. Biochemistry 46: 4077–4089 Jacob F and Monod J (1961) On regulation of gene activity. Cold Spring Harb Symp Quant Biol 26: 193 Kaneko T, Nakamura Y, Sato S, Minamisawa K, Uchiumi T, Sasamoto S, Watanabe A, Idesawa K, Iriguchi M, Kawashima K, Kohara M, Matsumoto M, Shimpo S, Tsuruoka H, Wada T, Yamada M and Tabata S (2002) Complete genomic sequence of nitrogen-fixing symbiotic bacterium Bradyrhizobium japonicum USDA110. DNA Res 9: 189–197 Kiley PJ and Kaplan S (1988) Molecular genetics of photosynthetic membrane biosynthesis in Rhodobacter sphaeroides. Microbiol Mol Biol Rev 52: 50–69 Kolber ZS, Plumley FG, Lang AS, Beatty JT, Blankenship RE, VanDover CL, Vetriani C, Koblizek M, Rathgeber C and Falkowski PG (2001) Contribution of aerobic photoheterotrophic bacteria to the carbon cycle in the ocean. Science 292: 2492–2495 Larimer FW, Chain P, Hauser L, Lamerdin J, Malfatti S, Do L, Land ML, Pelletier DA, Beatty JT, Lang AS, Tabita FR, Gibson JL, Hanson TE, Bobst C, Torres J, Peres C, Harrison FH, Gibson J and Harwood CS (2004) Complete genome sequence of the metabolically versatile photosynthetic bacterium Rhodopseudomonas palustris. Nat Biotechnol 22: 55–61 Lawrence JG (2003) Gene organization: Selection, selfishness, and serendipity. Annu Rev Microbiol 57: 419–440 Lawrence JG and Roth JR (1996) Selfish operons: Horizontal transfer may drive the evolution of gene clusters. Genetics 143: 1843–1860 Mackenzie C, Choudhary M, Larimer F, Predki P, Stilwagen S, Armitage J, Barber R, Donohue T, Hosler J, Newman J, Shapleigh J, Sockett R, Zeilstra-Ryalls J and Kaplan S (2001) The home stretch, a first analysis of the nearly completed genome of Rhodobacter sphaeroides 2.4.1. Photosynth Res 70: 19–41 Nagashima KVP, Shimada K and Matsuura K (1993) Phylogenetic analysis of photosynthetic genes of Rhodocyclus gelatinosus — possibility of horizontal gene-transfer in purple bacteria. Photosynth Res 36: 185–191 Nagashima KVP, Hiraishi A, Shimada K and Matsuura K (1997) Horizontal transfer of genes coding for the photosynthetic reaction centers of purple bacteria. J Mol Evol 45: 131–136 Nakatsu CH, Hristova K, Hanada S, Meng X-Y, Hanson JR, Scow KM and Kamagata Y (2006) Methylibium petroleiphilum gen. nov., sp. nov., a novel methyl tert-butyl ether-degrading methylotroph of the Betaproteobacteria. Int J Syst Evol Microbiol 56: 983–989 Pál C and Hurst LD (2004) Evidence against the selfish operon theory. Trends Genet 20: 232–234 Price MN, Huang KH, Arkin AP and Alm EJ (2005) Operon formation is driven by co-regulation and not by horizontal gene transfer. Genome Res 15: 809–819 Shiba T and Harashima K (1986) Aerobic photosynthetic bacteria. Microbiol Sci 3: 376–378 Shiba T, Simidu U and Taga N (1979) Distribution of aerobic bacteria which contain bacteriochlorophyll a. Appl Environ Microbiol 38: 43–45
29 Swingley WD, Sadekar S, Mastrian SD, Matthies HJ, Hao J, Ramos H, Acharya CR, Conrad AL, Taylor HL, Dejesa LC, Shah MK, O’Huallachain ME, Lince MT, Blankenship RE, Beatty JT and Touchman JW (2007) The complete genome sequence of Roseobacter denitrificans reveals a mixotrophic rather than photosynthetic metabolism. J Bacteriol 189: 683–690 Tabita FR (1999) Microbial ribulose 1,5-bisphosphate carboxylase/oxygenase: A different perspective. Photosynth Res 60: 1–28 Tabita FR, Hanson TE, Li H, Satagopan S, Singh J and Chan S (2007) Function, structure, and evolution of the RubisCO-like proteins and their RubisCO homologs. Microbiol Mol Biol Rev 71: 576–599 Venter JC, Remington K, Heidelberg JF, Halpern AL, Rusch D, Eisen JA, Wu D, Paulsen I, Nelson KE, Nelson W, Fouts DE, Levy S, Knap AH, Lomas MW, Nealson K, White O, Peterson J, Hoffman J, Parsons R, Baden-Tillson H, Pfannkoch C, Rogers Y-H and Smith HO (2004) Environmental genome shotgun sequencing of the Sargasso Sea. Science 304: 66–74 Watson GMF and Tabita FR (1997) Microbial ribulose 1,5bisphosphate carboxylase/oxygenase: A molecule for phylogenetic and enzymological investigation. FEMS Microbiol Lett 146: 13–22 Wellington CL and Beatty JT (1991) Overlapping mRNA transcripts of photosynthesis gene operons in Rhodobacter capsulatus. J Bacteriol 173: 1432–1443 Willems A, Munive A, de Lajudie P and Gillis M (2003) In most Bradyrhizobium groups sequence comparison of 16S-23S rDNA internal transcribed spacer regions corroborates DNADNA hybridizations. Syst Appl Microbiol 26: 203–210 Xiong J, Inoue K and Bauer CE (1998) Tracking molecular evolution of photosynthesis by characterization of a major photosynthesis gene cluster from Heliobacillus mobilis. Proc Natl Acad Sci USA 95: 14851–14856 Yooseph S, Sutton G, Rusch DB, Halpern AL, Williamson SJ, Remington K, Eisen JA, Heidelberg KB, Manning G, Li W, Jaroszewski L, Cieplak P, Miller CS, Li H, Mashiyama ST, Joachimiak MP, van Belle C, Chandonia J-M, Soergel DA, Zhai Y, Natarajan K, Lee S, Raphael BJ, Bafna V, Friedman R, Brenner SE, Godzik A, Eisenberg D, Dixon JE, Taylor SS, Strausberg RL, Frazier M and Venter JC (2007) The Sorcerer II Global Ocean Sampling expedition: Expanding the universe of protein families. PLoS Biol 5: e16 Yurkov V and van Gemerden H (1993) Impact of light/dark regime on growth rate, biomass formation and bacteriochlorophyll synthesis in Erythromicrobium hydrolyticum. Arch Microbiol 159: 84–89 Yurkov VV and Beatty JT (1998) Aerobic anoxygenic phototrophic bacteria. Microbiol Mol Biol Rev 62: 695–724 Yurkov VV and Csotonyi JT (2003) Aerobic anoxygenic phototrophs and heavy metal reducers from extreme environments. In: Pandalai SG (ed) Recent Research Developments in Bacteriology, pp 247–300. Transworld Research Network, Trivandrum, India Yutin N, Suzuki MT, Teeling H, Weber M, Venter JC, Rusch DB and Béjà O (2007) Assessing diversity and biogeography of aerobic anoxygenic phototrophic bacteria in surface waters of the Atlantic and Pacific Oceans using the Global Ocean Sampling expedition metagenomes. Environ Microbiol 9: 1464–1475
Chapter 3 New Light on Aerobic Anoxygenic Phototrophs Vladimir Yurkov* and Julius T. Csotonyi Department of Microbiology, University of Manitoba, Winnipeg, Manitoba, Canada R3T 2N2
Summary ................................................................................................................................................................. 31 I. Introduction....................................................................................................................................................... 32 II. Morphological Diversity, Taxonomic Nuances, Phylogeny and Evolution ........................................................ 34 A. Morphological Richness ..................................................................................................................... 34 B. Taxonomic Diversity ........................................................................................................................... 34 C. Persistent Taxonomic-Phylogenetic Tangles ..................................................................................... 37 D. Evolution ............................................................................................................................................ 38 III. Nutritional Versatility and Peculiarities of Carbon Metabolism ......................................................................... 40 A. Carbon Sources ................................................................................................................................. 40 B. Alternative Electron Donors and Acceptors........................................................................................ 41 IV. Photosynthetic Pigment Composition and Synthesis Reveal Surprises........................................................... 41 A. Carotenoids ........................................................................................................................................ 41 B. Bacteriochlorophyll ............................................................................................................................. 42 C. Highly Evolved Regulation of Pigment Synthesis .............................................................................. 42 V. The Mysterious Photosynthetic Apparatus of Aerobic Anoxygenic Phototrophs.............................................. 44 A. The Reaction Center of Aerobic Anoxygenic Phototrophs Has Purple Bacterial Roots ..................... 44 B. Light-Harvesting Complexes: Ordinary and Extraordinary ................................................................. 45 C. The Riddle of Aerobic Anoxygenic Phototrophy................................................................................. 46 VI. Speculation on Ecological Roles ...................................................................................................................... 47 A. Technical Challenges of Enumerating Aerobic Anoxygenic Phototrophs .......................................... 47 B. Ocean Surface ................................................................................................................................... 47 C. Vertical Distribution of Deep Ocean Aerobic Anoxygenic Phototrophs .............................................. 50 D. Aerobic Anoxygenic Phototrophs Thrive in Extreme Environments ................................................... 51 E. Soil and Freshwater ........................................................................................................................... 51 VII. Concluding Remarks and Perspectives ........................................................................................................... 51 Acknowledgments ................................................................................................................................................... 52 References .............................................................................................................................................................. 52
Summary Discovered 30 years ago, aerobic anoxygenic phototrophs (AAP) represent an entirely new bacterial functional group that was surprisingly found to constitute nearly 10% of microbial cells in the world’s biggest surface ecosystem, the ocean. These intriguing and colorful descendents of anaerobic anoxygenic phototrophs possess a fully functional photosynthetic apparatus that is paradoxically operative only under oxic conditions. An obviously ancient group, the AAP display numerous extensive evolutionary modifications to their photosynthetic machinery from that of their ancestors, such as different suites of light-harvesting 2 complexes and, in some *Author for correspondence, email:
[email protected] C. Neil Hunter, Fevzi Daldal, Marion C. Thurnauer and J. Thomas Beatty (eds): The Purple Phototrophic Bacteria, pp. 31–55. © 2009 Springer Science + Business Media B.V.
32
Vladimir Yurkov and Julius T. Csotonyi
species, the only zinc-based chlorophyll pigments found anywhere in nature. Whereas AAP are incapable of photoautotrophy and rely on heterotrophy for 80% or more of their cellular energetics, sunlight can double organic carbon assimilatory efficiency over that of strict heterotrophs, making AAP key players in the marine carbon cycle. The AAP inhabit not just soil, rivers and oceans, but also hypersaline waters, thermal springs and even the dark realm of deep ocean hydrothermal vents. Ubiquity and atypical photosynthetic nature has inspired an ever-increasing scientific interest in the AAP, for which there are more exceptions than rules. I. Introduction It is astonishing that a bacterial functional group constituting nearly a tenth of microorganisms in the earth’s largest surface ecosystem, the ocean, could have been overlooked until 1978. Yet, this unlikely tale describes obligately aerobic anoxygenic phototrophs (AAP), which have long lain hidden beneath the shortsighted assumption that anoxygenic photosynthesis, originally evolving to generate metabolic energy from light in reducing Proterozoic environments, has for the past 3.5 billion years remained an exclusively anoxic process. Oxygen does generally repress synthesis of the bacteriochlorophyll (BChl)-containing photosynthetic apparatus in conventional anoxygenic phototrophs, relegating purple, green and heliobacterial light-harvesting to illuminated zones of anaerobic habitats such as stratified lakes and sulfide springs (Pfennig, 1978). However, O2 release by cyanobacteria ~2.5 GYa exerted sufficient selection pressure for an aerobic counterpart to evolve (Beatty, 2002). Indeed, Japanese scientists’ intuitive leap to search for such organisms was met with immediate success (Shiba et al., 1979). Since then, description of new species has increased in pace (Fig. 1A). AAP have turned up in practically every environment probed for them, including rivers, soil, acidic mine drainage, hypersaline springs, nutrient-rich microbial mats, oligotrophic ocean surface waters, and even near marine hydrothermal vents (Yurkov and Csotonyi, Abbreviations: AAP – aerobic anoxygenic phototrophs; BChl – bacteriochlorophyll; Cmi. – Citromicrobium; Cyt – cytochrome; D – Dinoroseobacter; DMSP – dimethylsulfoniopropionate; E. – Erythromicrobium; Erb. – Erythrobacter; GYa – 109 years ago; H. – Hoeflea; IREM – infrared epifluorescence microscopy; IRFRR – infrared fast repetition rate fluorometry; LH – lightharvesting; PEP – phosphoenol pyruvate; PSU – photosynthetic unit; QPCR – quantitative polymerase chain reaction; Qy – electronic transition of BChl a from ground state to lowest excited singlet state; R. – Roseicyclus; Rba. – Rhodobacter; RC – reaction center; Rps. – Rhodopseudomonas; Rsc. – Roseococcus; Rst. – Roseateles; Rubisco – ribulose-1,5-bisphosphate carboxylase/oxygenase; Rva – Roseovarius; S. – Stappia; Srb. – Sandaracinobacter; Stl. – Staleya; TMAO – trimethylamine N-oxide
2003). Comprehensive reviews have already set the groundwork for an introduction to the AAP (Yurkov and Beatty, 1998; Hiraishi and Shimada, 2001; Yurkov, 2006; Yurkov and Csotonyi, 2003; Rathgeber et al., 2004). However, the veritable explosion of research on this group during the last four years justifies both a synthesis of exciting new findings and suggestion of new investigative directions. Two other branches of bacteria produce BChl exclusively aerobically: photosynthetic methylotrophs such as Methylobacterium extorquens, and photosynthetic rhizobia like Bradyrhizobium denitrificans (van Berkum et al., 2006). Distinct metabolic strategies or specialized plant symbioses impart distinct functional group status on these organisms, and they will not be treated further in this chapter. Interested readers are directed to reviews by Komagata (1989) and Giraud and Fleischman (2004). The AAP are primarily chemoheterotrophic Proteobacteria that nonetheless produce a functional purple bacterial photosynthetic reaction center (RC) and one or more peripheral light-harvesting (LH) complexes that can amend heterotrophic energy generation by up to 20% (Kolber et al., 2001; Yurkov and van Gemerden, 1993). As their name implies, they evolve no O2 photosynthetically, because they do not use water for reducing power. Five overarching traits distinguish AAP from classical anoxygenic phototrophs: (1) requirement of O2 for photosynthesis, granting escape from restrictive anaerobic illuminated environments; (2) inhibition, paradoxically, of BChl synthesis by light; (3) absence of the Calvin cycle and inability to subsist on inorganic carbon; (4) much lower number of photosynthetic units (PSU) per cell; but (5) great abundance of carotenoids (Yurkov and Csotonyi, 2003; Rathgeber et al., 2004). With such a peculiar mixture of characteristics, where do the mysterious yet functionally and numerically significant AAP fit into our conception of the microbial world? On one hand, they are typical proteobacterial cells, constructed from the same building blocks as their relatives, expressing no earth-shatteringly novel metabolic modes, possess-
Chapter 3
Aerobic Anoxygenic Phototrophs
33
Fig. 1. Taxonomic richness and phylogeny of AAP. A. Cumulative growth in number of described genera over 24 years. B. Neighborjoining phylogenetic tree of AAP and select nonphototrophs (marked with asterisks) based on 16S rDNA sequences. Scale bar, 10% substitution rate. Bootstrap values at branch points are based on 500 resamplings. Phylogenetic tree was prepared by E. Stackebrandt and J. Swiderski.
34 ing typical cell walls, synthesizing mostly trivial pigments and electron carriers, etc. On the other hand, evolutionary forces have combined some of these common components in chimeric ways, permitting anoxygenic phototrophy to function under conditions contrary to those for which it originally evolved, thus opening for exploitation huge niches not available to their ancestors. This principal evolutionary development necessarily caused cascades of corollary adaptations, such as major changes to the regulation of photosynthetic pigment synthesis, and proliferation of photoprotective carotenoids. In the end, the peculiar AAP display more exceptions than rules in form and function. II. Morphological Diversity, Taxonomic Nuances, Phylogeny and Evolution A. Morphological Richness The AAP are morphologically as well rounded as are their anaerobic relatives, even expressing properties unique among bacteria (Fig. 2). Pleomorphism is common in the Erythrobacter clade. Erythromicrobium (E.) ramosum was named for its propensity to produce branches (Fig. 2A), while Citromicrobium (Cmi.) bathyomarinum forms unusual Y-cells during trinary division (Yurkov and Csotonyi, 2003) (Fig. 2B). Rathgeber et al. (2005) reported the first AAP that can form cyclical cells, Roseicyclus (R.) mahoneyensis (Fig. 2C), reminiscent of Rhodocyclus purpureus (Pfennig, 1978), to which it is only distantly related. Uncultured spirilloid cells ascribed to AAP have turned up in the Sargasso Sea (Sieracki et al., 2006). Cell accessories and intercellular connective structures provoke further interest, especially ecologically. Pleomorphic Porphyrobacter neustonensis and Porphyrobacter meromictius produce appendages, perhaps enhancing intake of nutrients by maximizing surface-to-volume ratio (Fig. 2D) (Yurkova et al., 2002; Rathgeber et al., 2007). More interestingly, cell chains are connected via bubble-like formations with a central tubular structure, an arrangement heretofore unobserved in the domain Bacteria (Fig. 2E) (Yurkova et al., 2002; Rathgeber et al., 2007). Bubble-like membranous connective structures are also seen between cells of Cmi. bathyomarinum (Yurkov and Beatty, 1998). The relevance of these structures is unknown. Perhaps they facilitate mate-
Vladimir Yurkov and Julius T. Csotonyi rial exchange between cells. Several members of the α-2, α-3 and α-4 Proteobacteria, including Staleya (Stl.) guttiformis, Stappia (S.) marina, Roseovarius (Rva.) mucosus, Roseisalinus antarcticus and deep ocean strain C8 (Fig. 2F) form star-shaped rosettes of cells, joined by unknown means (Biebl et al., 2005a; Labrenz et al., 2005; Kim et al., 2006). The taxonomically unassigned meromictic lake strains BL7 and BL14, however, form the most impressive rosettes, 10 µm or more in diameter and resembling dandelion heads (Fig. 2G) or even brain coral (Fig. 2H), in which cells are linked by a polar hook-like structure (Fig. 2I) (Rathgeber et al., 2004). Interestingly, all of these rosette-forming species except S. marina and C8 are meromictic lake isolates, suggesting that aggregation may bestow an ecological advantage, such as buoyancy regulation, a useful feature in lake habitats where salinity varies with depth. Alternately, the convoluted surface of rosettes may assist in trapping particulate nutrients. B. Taxonomic Diversity AAP are dispersed throughout the Alphaproteobacteria, with one known betaproteobacterial representative, Roseateles (Rst.) depolymerans (Fig. 1B) (Yurkov and Csotonyi, 2003) and a recently described gammaproteobacterial species, Congregibacter litoralis (Fuchs et al., 2007). Thus far, alphaproteobacterial species occur in five cohesive clusters, in all four subclasses. Soil, hot spring and acidophilic isolates define the α-1 cluster, consisting of Acidiphilium, Acidisphaera, Craurococcus, Geminicoccus, Paracraurococcus, Roseococcus and Rubritepida. The exclusively marine α-2 species form two unrelated branches, the first containing Labrenzia, Roseibium and S. marina, the second Hoeflea (H.) phototrophica. The oceanic or halophilic α-3 Roseobacter clade, allied closely with purple nonsulfur Rhodobacter and Rhodovulum, includes Dinoroseobacter, Roseibacterium, Roseicyclus, Roseinatronobacter, Roseisalinus, Roseivivax, Roseobacter, Roseovarius, Rubrimonas, Staleya and Thalassobacter. Finally, Blastomonas natatoria, Citromicrobium, Erythrobacter, Erythromicrobium, Erythromonas, Sandaracinobacter and Sandarakinorhabdus represent the α-4 AAP, hailing from diverse freshwater and marine habitats. Citromicrobium, Erythrobacter, Erythromicrobium and Porphyrobacter form a clade so cohesive that 16S rDNA sequence analysis is often insufficient to place species into proper genera, generic assignment rely-
Chapter 3
Aerobic Anoxygenic Phototrophs
35
Fig. 2. Morphological diversity of AAP. A. Branching in Erythromicrobium ramosum. B. Y-cell of Citromicrobium bathyomarinum. C. Tightly curved vibroid cells of Roseicyclus mahoneyensis. D. Appendage (arrow) of Porphyrobacter meromictius. E. Tube-in-bubble type of intercellular connections of Porphyrobacter meromictius. F. Rosettes of cells in strain C8. G. Dandelion head-like rosettes of strain BL7. H. Brain coral-like rosettes of strain BL14. I. Hooked appendage of strain BL14. C, F, G, H. Phase contrast micrographs. A, B, C (inset), D, E, I. Electron micrographs. Scale bars 0.25 µm (C inset); 0.5 µm (D, I); 1 µm (A, B, E); 2.5 µm (G, H); 5 µm (C, F).
ing heavily on phenotypic characteristics. Rainey et al. (2003) suggested unification of this cluster under a single genus. Instead of taxonomic abridgement, it is advisable to utilize the phenotypic diversity of this group (e.g., presence/absence of the unique B798-832 LH2 in Erythromicrobium and appendage formation in Porphyrobacter) to assign generic and specific status. Perhaps Porphyrobacter dokdonensis, possessing an 835-nm LH2 absorption peak and lacking appendages (Yoon et al., 2006), should have been placed within Erythromicrobium. Additional powerful phylogenetic analysis techniques, such as 23S rRNA sequence analysis (Peplies et al., 2004) or internal transcribed spacer analysis (Brown and Fuhrman, 2005), may be more phyletically infor-
mative for tight clusters than traditional 16S rDNA sequence analysis. The number of known AAP (currently 52 species in 33 genera) is waxing rapidly (Fig. 1A), with 18 new species described in the last four years (Table 1). H. phototrophica represents the first AAP from the α-2 proteobacterial Phyllobacteriaceae, which also contains rhizobia (Biebl et al., 2006). Meanwhile, R. mahoneyensis is the first inland Roseobacter-clade representative (Rathgeber et al., 2005). Description of the nearly completely genome-sequenced C. litoralis is extraordinary because not since 1999 has a new proteobacterial subclass been added to the short list that contains AAP (Fuchs et al., 2007). Gathering interest in this group, use of new high-throughput
36
Table 1. Determinative characteristics of novel species of aerobic anoxygenic phototrophs described since 2003 1. Habitat, site of isolation
Phylogenetic affiliation 2
Color
Cell shape
Congregibacter litoralis 3 (Fuchs et al., 2007)
sea water, North Sea
NOR5/OM60 clade (γ)
pink
pleomorphic
In vivo BChl peaks (nm) N.D.4
Dinoroseobacter shibae (Biebl et al., 2005b)
dinoflagellate culture, North Sea Rhodobacteraceae (α-3) Jannaschia helgolandensis, 94.1%
beige to light red
cocci or ovoid
804, 868
Geminicoccus roseus (Foesel et al., 2007)
marine aquaculture biofilter, Israel
diplococci
N.D.
Hoeflea phototrophica (Biebl et al., 2006)
dinoflagellate culture, North Sea Phyllobacteriaceae (α-2) Hoeflea marina, 98.5%
colorless to pink
short rod with capsule
N.D.
Labrenzia alexandrii (Biebl et al., 2007)
dinoflagellate culture, North Sea Rhodobacteraceae (α-2) Stappia alba and Stappia marina, 98.0%
faint pink
rod, uneven ends
801, 865
Porphyrobacter cryptus (Rainey et al., 2003)
hot spring water, Portugal
Sphingomonadaceae (α-4) Porphyrobacter tepidarius, 98.3%
reddish-orange
short rod
800, 870
Porphyrobacter dokdonensis (Yoon et al., 2006)
sea water, Dokdo, Korea
Sphingomonadaceae (α-4) Porphyrobacter cryptus, 98.7%
reddish-orange
pleomorphic: cocci, ovoid 800, 835, 862 or rod
Porphyrobacter donghaensis (Yoon et al., 2004b)
sea water, East Sea, Korea
Sphingomonadaceae (α-4) Erythromicrobium ramosum, 99.0%
reddish-orange
pleomorphic: cocci, ovoid 808, 867 or rod
Porphyrobacter meromictius (Rathgeber et al., 2007)
meromictic lake water, Canada
Sphingomonadaceae (α-4) Erythrobacter aquamaris, 98.1%
reddish-brown
short rod or pleomorphic
806-808, 866-867
Roseibacterium elongatum (Suzuki et al., 2006)
sand, Shark Bay, Australia
Rhodobacteraceae (α-3) Marinosulfonomonas methylotropha, 93%,
pink
rod, variable length
800, 879
Roseicyclus mahoneyensis (Rathgeber et al., 2005)
meromictic lake water, Canada
Rhodobacteraceae (α-3) Ketogulonicigenium vulgare, 92.6%
pinkish-purple to purple
pleomorphic: ovoid, long rod, vibrioid, cyclical rod
805-806, 870-871
Roseinatronobacter monicus (Boldareva et al., 2007)
hypersaline lake, USA
Rhodobacteraceae (α-3) Roseinatronobacter thiooxidans, 98.1%
pink
ovoid
804, 870
Roseisalinus antarcticus (Labrenz et al., 2005)
meromictic lake water, Antarctica
Rhodobacteraceae (α-3) Jannaschia helgolandensis, 94%
red
rod
800-801, 870
Novel clade (α−1) whitish-grey to pink Candidatus “Alysiosphaera europaea,” 90.8%
Vladimir Yurkov and Julius T. Csotonyi
Species (described by)
1 New taxa described before 2003 are covered by Yurkov and Beatty (1998), Yurkov and Csotonyi (2003) and Rathgeber et al., (2004). 2 family (nomenclature after Garrity et al., 2005), proteobacterial subclass (in parentheses), nearest 16S rDNA relative, phylogenetic distance to nearest relative; 3 names in bold indicate novel genera. 4 N.D., not determined.
N.D. ovoid to irregular rod salmon-pink Rhodobacteraceae (α-3) Jannaschia helgolandensis, 95.7% Thalassobacter stenotrophicus (Macián et al., 2005)
sea water, Mediterranean Sea
N.D. N.D. rod Rhodobacteraceae (α-2) Stappia aggregata, 98.5% Stappia marina (Kim et al., 2006)
tidal flat, Korea
N.D. rod orange-red soil, South Korea Sphingomonas kaistensis (Kim et al., 2007)
Sphingomonadaceae (α-4) Sphingomonas oligophenolica, 95.8%
800, 837, 865 rod Sandarakinorhabdus limnophila freshwater lake water, Germany Sphingomonadaceae (α-4) Sandaracinobacter sibiricus, 92.8% (Gich and Overmann, 2006)
orange-red
ovoid to rod whitish to faint pink dinoflagellate culture, North Sea Rhodobacteraceae (α-3) Roseovarius tolerans, 96.4% Roseovarius mucosus (Biebl et al., 2005a)
In vivo BChl peaks (nm) N.D. Cell shape Phylogenetic affiliation 2 Habitat, site of isolation Species (described by)
Table 1. Continued
Aerobic Anoxygenic Phototrophs
Color
Chapter 3
37 isolation methods, and the application of phylogenetic techniques to detect photosynthesis genes are responsible for this taxonomical flurry (Gich et al., 2005). Our current delineation of AAP diversity is by no means the end of the taxonomic tale. Sequencing marine photosynthesis genes or their mRNA, Béjà et al. (2002) and Allgaier et al. (2003) detected much higher AAP diversity than suggested by culture-dependent studies. Description of novel lotic β-proteobacterial species is soon anticipated (Waidner and Kirchman, 2005), while procedures facilitating recent isolation of previously uncultivable SAR11 marine bacterioplankton (Rappé et al., 2002) may yield additional elusive γ-proteobacterial AAP. C. Persistent Taxonomic-Phylogenetic Tangles Embedding a thorn in taxonomists’ sides, 16S ribosomal genes of AAP often ally more closely with non-phototrophs than with purple nonsulfur bacteria. This phyletic interspersion still baffles enough researchers that, regrettably, an old argument must be revisited regarding the confusion of taxonomy with phylogeny in the classification of AAP. Taxonomic convention recognizes major metabolic capabilities such as phototrophy as markers to distinguish genera. After all, in spite of the deceptive visual homogeneity of bacteria harboring divergent metabolic strategies, the difference made by possession of a photosynthetic apparatus is as significant as would be the discovery of a fish breathing nitrate rather than oxygen. Yet, since 2002, no less than nine species of BChl-lacking bacteria have been proposed as members of phototrophic genera (Erythrobacter aquamaris, Erythrobacter citreus, Erythrobacter flavus, Erythrobacter gaetbuli, Erythrobacter luteolus, Erythrobacter vulgaris, Roseovarius crassostreae, Roseovarius nubinhibens) (Denner et al., 2002; Yoon et al., 2003, 2004a, 2005a,b; González et al., 2003; Boettcher et al., 2005; Ivanova et al., 2005). Conversely, and despite repeated appeals to the contrary since 1998 (Yurkov and Beatty, 1998), BChl-producing species such as H. phototrophica and S. marina are included in definitively non-phototrophic genera. Ironically, the very phylogenetic analyses sparking original misclassifications, upon closer inspection, support genus-level distinction of BChl-free isolates from true AAP. New non-phototrophic erythrobacters were sufficiently unrelated to phototrophic congener-
38 ics (16S rDNA sequence similarities as low as 94.1% and DNA-DNA reassociation levels often below 10%) to justify erection of new genera (Yoon et al., 2003, 2004a, 2005a). Similarly, non-phototrophic Rva. nubinhibens and Rva. crassostreae can clearly be differentiated from the phototrophic Roseovarius cluster by specific signature nucleotides at several positions of the secondary structure of the 16S rRNA gene (Biebl et al., 2005a). Judicious phylogenetic analysis supports the repeated insistence by the International Committee on Systematic Bacteriology that phenotypic traits should remain important criteria in delineation of genera (Wayne et al., 1987; Murray et al., 1990; Stackebrandt et al., 2002). Phylogenetic analysis is not sufficiently advanced for consensus on the use of a standard suite of purely genetic techniques for resolution of both macro- and microdiversity, and thus it remains inappropriate to use phylogenetic relatedness as a sole or overriding determinative factor in assigning taxonomic status. Errors are occasionally corrected by reclassifying miscategorized species — see Yurkov and Csotonyi (2003) for examples — but lamentably, some new papers still contain the original misnomers and cite erroneous papers as major references. For instance, Boettcher et al. (2005) alluded to ‘Roseobacter gallaciensis,’ which was long ago reclassified as Ruegeria gallaciensis (Uchino et al., 1998). Clearly, reviewers and editors of premier bacterial systematics journals must exercise greater discrimination when considering proposals of taxonomic placement to avoid subsequent confusion from rearrangements necessary to correct taxonomic faux pas. D. Evolution We must distinguish evolution of aerobic anoxygenic phototrophy as a physiological process from the evolutionary history of its phylogenetic distribution among AAP clades. In hindsight, it is not surprising that aerobic anoxygenic phototrophy arose from anaerobic phototrophy after global oxygenation; given life’s rapid niche-occupying tendency, such a transition ought to occur over 2 billion years after the earth’s atmosphere became oxygenated (Rye and Holland, 1998). This question of how such evolution played out still deserves much research effort, but speculation is fruitful. Similarity of the AAP photosynthetic apparatus to that of purple nonsulfur bacteria implicates the latter as ancestors. Probably
Vladimir Yurkov and Julius T. Csotonyi the most serious hurdle for evolution to clear was aerobic phototoxicity (Beatty, 2002). Exposure of BChl to light in the presence of O2 generates highly reactive chemical species called triplet BChl and singlet oxygen, which can cause considerable oxidative damage to cells (Krinsky, 1971; Beatty, 2002). Hence, repression of aerobic BChl synthesis by anaerobic phototrophs in response to global O2 pollution served as much a protective as a resource conservation role. Interestingly, a logical solution to phototoxicity conveniently explains three definitive features of AAP. First, low PSU abundance minimizes generation of triplet BChl and singlet O2 (Beatty, 2002). Second, BChl is only generated during dark periods, when it is safe to do so (Beatty, 2002). This paradoxical trait seems counterintuitive only until photochemistry is considered. Third, synthesis of light-harvesting photoactive carotenoids by anaerobic phototrophs also preadapted their descendents for an aerobic existence. Not only do carotenoids with more than nine conjugated double bonds filter out high-energy radiation, but also their low lying first excited triplet state enables them to efficiently quench the destructive energy of singlet O2 and triplet BChl (Krinsky, 1971; Fraser et al., 2001). AAP needed only to upregulate carotenoid synthesis to achieve better protection from the dangerous combination of light, O2 and BChl — which is exactly what we see, especially for the particularly photoprotective carotenoid sulfates (Krinsky, 1971; Yurkov and Beatty, 1998). In light of the inherent self-limitation of aerobic phototrophic energy generation, AAP have understandably lost the energetically expensive Calvin cycle for carbon fixation, relying instead on heterotrophy. Beyond explaining their appearance as a functional group, however, phylogenetic interspersion of AAP with non-phototrophs seriously challenges evolutionary microbiologists wishing to trace the phylogenetic roots of each clade. Unfortunately, genetic research has found that many of the pieces of this evolutionary puzzle do not fit snugly. There is little doubt that AAP are polyphyletic at the level of the 16S rRNA gene. However a battle ensues between two opposing views: (1) that phototrophy is ancestral in Proteobacteria, with non-phototrophs such as Paracoccus or Ketogulonicigenium arising through loss of photosynthesis, and (2) that aerobic photosynthetic genes were transferred laterally to unrelated non-phototrophs (Yurkov and Beatty, 1998). The argument hinges on the relative topologies of ribosomal and photosyn-
Aerobic Anoxygenic Phototrophs C
B Absorbance
A
39
Absorbance
Chapter 3
869
857
804
799
400
500
600
700
800
900
1000
400
500
D
E
600
700
800
900
1000
Wavelength (nm)
Wavelength (nm)
P870*
F
Absorbance
BChl BPh
+
QA
E0'
868 836
cyt bc1
799
cyt c2 400
500
600
700
800
900
1000
P870
Wavelength (nm)
G
H
I
Fig. 3. Photosynthetic and habitat diversity of aerobic anoxygenic phototrophs (AAP). A–E. In vivo absorbance spectra of cells grown under different physiological conditions, with numerals above peaks denoting LH and RC wavelengths. Insets, photographs of liquid cultures. A. Roseicyclus mahoneyensis, illuminated (dashed line) and dark (solid line). B,C. Dashed line, rich organic medium (3 g·l–1 organics); solid line, minimal acetate (1 g·l–1) medium (B. Roseococcus thiosulfatophilus, C. Erythrobacter litoralis). D. Erythromicrobium ramosum, rich organic medium, dark (dashed line) and oligotrophic medium, light:dark regimen of 12h:12h, illuminated with diffuse ambient sunlight (solid line). E. Citromicrobium bathyomarinum strain JF1, rich organic medium (dashed line), minimal glucose medium (dotted line), compared with Erythrobacter litoralis (solid line). F. Hypothetical electron transfer system of aerobic anoxygenic photosynthesis, showing electron flow through major carriers: P870, special pair of BChl in RC (photoexcited state indicated by ‘*’); BChl-BPh, accessory BChl and bacteriopheophytin in RC; QA, quinone primary electron acceptor; cyt bc1, cytochrome bc1 complex; cyt c2, cytochrome c2. The symbol ‘+’ indicates that the midpoint potential of QA in all tested AAP is positive and higher than in anaerobic phototrophs. G–I. Extreme environment habitats of AAP. G. Hydrothermal vent field in Eastern Pacific Ocean, showing smoker chimney. H. Hypersaline spring system (East German Creek) in Manitoba, Canada: spring pool in foreground and playa in background, with white patches of salt precipitates. I. Cyanobacterial-Thiothrix mat development in thermal springs of Banff National Park, Alberta, Canada. See also Color Plate 2, Fig. 2.
thetic phylogenetic trees, and on the diversity of the aerobic photosynthetic apparatus. The answer probably lies between opposing schools of thought: loss of phototrophy may account for close non-phototrophic relatives, and lateral gene transfer for the rise of a few
AAP clades. Ribosomal and pufL (the gene encoding the L subunit of the photosynthetic reaction center) cladistic trees agree topologically at the highest level of similarity (such as the Erythrobacter clade), but a few deep incongruities exist (Allgaier et al., 2003).
40 Both α-3 Staleya and β-proteobacterial Roseateles possess pufL genes characteristic of α-4 Erythrobacter (Allgaier et al., 2003). Even more surprising are three marine strains, HTCC2080, HTCC2148 and HTCC2246, which, despite forming a tight 16S rDNA gammaproteobacterial cluster in the NOR5/OM60 clade, possess pufL genes of gammaproteobacterial, α-1 and α-2 proteobacterial affiliation, respectively (Cho et al., 2007). Other studies (Béjà et al., 2002; Oz et al., 2005) have underscored the observed interspersion of photosynthesis genes among different clades, but the emerging picture is complicated, the pattern of interspersion depending on the gene that is sequenced. For example, unlike pufL, the pufM gene of Rst. depolymerans clusters more closely with those of other Betaproteobacteria rather than with those of Erythrobacter (Béjà et al., 2002). It remains to be explained how such a linked gene pair as pufL and pufM can exhibit such phylogenetic topological variety. Furthermore, although the plasmid pBLM2 was used to mobilize the entire photosynthesis gene cluster of Rhodobacter capsulatus, pBLM2 did not induce functional pigment expression in Escherichia coli (Marrs, 1981). Thus, lateral photosynthetic gene transfer might occur successfully only between bacteria that are already phototrophic, possibly because specific membrane configurations are prerequisite for functional photosynthetic machinery (Marrs, 1981). Adding a further twist, the apparent α-proteobacterial origin of the entire 37-kb photosynthetic gene cluster of the β-proteobacterial anoxygenic phototroph Rubrivivax gelatinosus (an anaerobic 16S rDNA close relative of Rst. depolymerans) (Igarashi et al., 2001), implies multiple independent origins of aerobic phototrophy, as Rst. depolymerans probably evolved from a Rubrivivax gelatinosus-like ancestor. Moreover, the difference of LH2 complexes of AAP both from each other and from those of anaerobic phototrophs (see Section V.B.) also implicates multiple origins of aerobic phototrophy even within Alphaproteobacteria. As an attempt to synthesize these arguments into perhaps the most accurate picture of AAP evolution, we suggest that each major AAP clade evolved separately from a different purple nonsulfur bacterial branch. Moderate gene shuffling then occurred only among these phototrophs, and non-phototrophic Proteobacteria arose through loss of phototrophy, as proposed by Woese (1987).
Vladimir Yurkov and Julius T. Csotonyi III. Nutritional Versatility and Peculiarities of Carbon Metabolism A. Carbon Sources Metabolic diversity in AAP is extremely great, as expected for a functional group relying exclusively on heterotrophy for carbon needs. Erythrobacter, Erythromicrobium, Porphyrobacter, Roseicyclus, Roseobacter and Sandaracinobacter are able to grow on most carbon sources tested, including organic acids, carbohydrates, alcohols and complex organics (Yurkov and Csotonyi, 2003). However, some species are more specialized, with a second cluster (deep ocean Citromicrobium; meromictic Mahoney Lake strains ML15, ML17, ML35; Manitoba hypersaline spring strains EG4, EG6 and EG15) growing best on complex organics and on only a few defined carbon sources (Yurkov and Beatty, 1998; Yurkova et al., 2002; Csotonyi and Yurkov, unpublished). Most intriguingly, a third cluster (e.g., Mahoney Lake strains ML37, ML47; Manitoba hypersaline spring strain EG12) requires yeast extract or casamino acids, and subsists on no known defined medium (Yurkova et al., 2002; Csotonyi and Yurkov, unpublished). These species may be adapted to utilizing specific carbon sources produced by organisms or geological processes exclusive to their native environment, and they likely evolved locally by sympatric speciation based on metabolic niche differentiation. Endemism may explain the diversity of AAP in isolated habitats such as meromictic lakes or hypersaline springs. Certain carbon sources exploited by AAP bear ecological significance. Much recent attention focuses on dimethylsulfoniopropionate (DMSP), an osmolyte produced by algae and an important marine store of sulfur. The prominent role of Roseobacter relatives in DMSP consumption by cleavage or demethylation and dethiolation, and the resulting liberation of sulfide or climate-influencing dimethyl sulfide, implicates them as key modulators of the marine sulfur cycle (Wagner-Döbler and Biebl, 2006). The next significant step will be discrimination of BChl-containing obligately aerobic representatives from strict heterotrophs, because phylogenetic and [35S]DMSP-incorporation studies alone cannot distinguish phototrophs from non-phototrophs. Photoautotrophic CO2 fixation is lacking in all physiologically tested AAP, with both enzymological and full genome sequence studies (for Roseobacter
Chapter 3
Aerobic Anoxygenic Phototrophs
(Rsb.) denitrificans and C. litoralis) confirming the absence of the key Calvin cycle enzyme ribulose1,5-bisphosphate carboxylase/oxygenase (Rubisco) (Yurkov and Beatty, 1998; Fuchs et al., 2007; Swingley et al., 2007). However, nonexistence of autotrophy in AAP need not contradict the finding of a low level of CO2 fixation in Acidiphilium rubrum, Erythrobacter longus, Rsb. denitrificans, Sandaracinobacter sibiricus and other AAP (Kolber et al., 2001; Rathgeber et al., 2004). All heterotrophs anaplerotically fix CO2 (at efficiencies of 1–4% of that expressed by autotrophs) via phosphoenolpyruvate (PEP) carboxylase, a key respiratory enzyme that replaces organic acids sequestered from the citric acid cycle by other biochemical pathways. Light-stimulated citric acid cycle reversal may also account for some carbon fixation (Rathgeber et al., 2004). B. Alternative Electron Donors and Acceptors Although most AAP appear to rely on reduced carbon compounds as electron donors, a few species reflect anaerobic phototrophic ancestry by oxidizing reduced sulfur sources. Interestingly, most S-oxidizers belong to the α-1 proteobacterial lineage (e.g., Acidiphillium acidophilum, Roseococcus (Rsc.) thiosulfatophilus and Rubritepida flocculens). The latter two species, as well as Roseinatronobacter thiooxidans and Roseinatronobacter monicus, oxidize thiosulfate (Yurkov and Beatty, 1998; Yurkov and Csotonyi, 2003; Boldareva et al., 2007), whereas Acidiphilium acidophilum oxidizes elemental S via glutathione at a rate nine to fourteen times as high as that of the lithoautotroph, Acidithiobacillus thiooxidans (Rohwerder and Sand, 2003). Most AAP are described as obligately aerobic, but some species are capable of anaerobic chemoheterotrophic growth, substituting O2 with alternate electron acceptors. Dinoroseobacter (D.) shibae can dissimilatorily reduce nitrate, and Rsb. denitrificans reduces nitrate, nitrite and trimethylamine N-oxide (TMAO), a common odoriferous byproduct of decomposing marine organisms (Yurkov and Csotonyi, 2003; Biebl et al., 2005b). Even more interesting, and of bioremediative value, is the pH-elevating effect of dissimilatory Fe3+-reduction by A. acidophilum, perhaps key to facilitating transition of mine drainage systems from acidic to neutral conditions (Marchand and Silverstein, 2003).
41 IV. Photosynthetic Pigment Composition and Synthesis Reveal Surprises A. Carotenoids A ubiquitous feature of AAP is their diverse range of intense colors, conferred by abundant carotenoids. Colors span the spectrum from watermelon red to brown, orange, yellow, pink, purple and intermediates (Fig. 3A-E and Color Plate 2). Most interestingly, in contrast to the light-harvesting function of carotenoids in anaerobic purple phototrophs, the majority of AAP carotenoids are disengaged from energy transduction, and are distributed evenly throughout the cytoplasmic membrane, cytoplasm and cell wall (Yurkov and Beatty, 1998; Koblížek et al., 2003). The primary purpose of this non-photosynthetic carotenoid pool is uncertain, but preventative or remediative amelioration of phototoxicity is a leading suggestion (Yurkov and Beatty, 1998; Fraser et al., 2001; Beatty, 2002). The photoprotection to which AAP appear to have enlisted the bulk of their carotenoids is reflected not only in up-regulation, but also in extensive structural and compositional modification. Whereas AAP LHbound carotenoids resemble those of anaerobic phototrophs, the LH-dissociated pool displays extreme variety (Fig. 3A–E and Color Plate 2), with E. ramosum possessing about 20 types (Yurkov and Beatty, 1998). Moreover, despite predominance of purple nonsulfur-type spheroidenone and spirilloxanthin in some AAP (e.g., Roseobacter and Acidiphilium), the majority in most species are carotenes, which are more widely distributed among oxygenic than anoxygenic phototrophs and give AAP decidedly more orange hues than in purple nonsulfur bacteria (Yurkov and Beatty, 1998) (Fig. 3A–E and Color Plate 2). Some of these carotenoids are structurally unique among phototrophs (Chapter 6, Takaichi). AAP are especially well-endowed with polar carotenoids, such as the short-chain C30 molecule, 4,4´-diapocarotene-4,4´dioate in Rsc. thiosulfatophilus (Fig. 3B and Color Plate 2), or erythroxanthin sulfate in Erythrobacter and Erythromicrobium (Yurkov and Csotonyi, 2003) (Fig. 3C, D and Color Plate 2). Acquisition of the latter carotenoid, which is particularly antioxidative (Krinsky, 1971), is a logical adaptation to illuminated oxic environments.
42 B. Bacteriochlorophyll One of the namesake features of AAP is their possession of BChl, the remarkable bacterial pigment capable of absorbing the electromagnetic energy of light. All known AAP synthesize BChl a esterified to phytol like many purple nonsulfur bacteria (Yurkov and Beatty, 1998). However, members of the genus Acidiphilium are the only organisms known to naturally chelate the BChl tetrapyrrole ring to Zn instead of to Mg, producing a 7-nm blue-shift in the absorbance spectrum of the aberrant BChl a (Hiraishi and Shimada, 2001). Striking similarity in the electrochemistries of Zn and Mg permits this novel pigment to function, while metals such as Ni would produce ineffective bacteriochlorins, exhibiting too short an excited state lifetime to facilitate photosynthetic electron flow (Cogdell et al., 2002). However, a key difference in the acid chemistries of Zn and Mg explains the ecological significance of this pigment in Acidiphilium. Zn-BChl is 106 times more resistant to loss of the metal ligand (pheophytinization) by acids than is Mg-BChl, granting exceptional acid-stability in habitats with a pH as low as 2. Supporting this idea, the ratio of Zn-BChl to Mg-BChl synthesized by A. rubrum increases with acidity (Hiraishi and Shimada, 2001). Regardless of its chemical form, low cellular BChl content is an important determinative trait of the predominantly heterotrophic AAP (Fig. 3A–E and Color Plate 2), which produce at least ten times less BChl than the ~20 nmol/mg dry weight typical of purple nonsulfur bacteria: 2.0 nmol/mg cells in Erb. longus, 1.0 nmol/mg protein in Srb. sibiricus and too low to reliably measure in Rva. mucosus (Yurkov and Csotonyi, 2003; Biebl et al., 2005a). Consequently, absorbance peaks of BChl integrated into RC and LH complexes are about ten-fold lower than those of carotenoids. However, BChl:carotenoid ratios of 1.33:1 (Manitoba hypersaline spring strain EG14; Csotonyi and Yurkov, unpublished) and 1.2:1 (R. mahoneyensis; Rathgeber et al., 2005) (Fig. 3A–E and Color Plate 2) were also found. These resemble anaerobic phototroph ratios, begging two substantial questions. First, how do these AAP species circumvent phototoxicity incurred by aerobic operation of their high levels of BChl? Perhaps unique membrane environments or particularly antioxidant suites of carotenoids nullify this oxidative hazard. Second, does high BChl content indicate recent divergence from anaerobic phototrophy, or does it bespeak an
Vladimir Yurkov and Julius T. Csotonyi even more advanced condition than exists in other AAP, representing the next stage in evolution toward efficient exploitation of aerobic illuminated environments? Whatever the answers to these questions, the elevated BChl content in such strains makes them ideal candidates for investigation of regulatory factors of photosynthetic pigment expression. C. Highly Evolved Regulation of Pigment Synthesis If, as we propose, there was a transition from an anaerobic to an aerobic mode of existence, this necessitated modifications in the regulation of pigment expression. BChl synthesis in AAP is repressed by light (Fig. 3A and Color Plate 2) more extremely than in anaerobic phototrophs (Yurkov and Csotonyi, 2003). However, AAP also exhibit regulatory systems involving nutrients (Fig. 3B–E and Color Plate 2), temperature, pH and salinity (Alarico et al., 2002, Koblížek et al., 2003; Rathgeber et al., 2004; Macián et al., 2005; Biebl et al., 2006). Ongoing research is elucidating the mechanisms driving these regulatory regimes and delineating the range of responses. Ouchane et al. (2004) recently investigated AcsF, a strictly aerobically-active functional equivalent to the anaerobic Mg-protoporphyrin monomethyl ester cyclase, BchE, both of which catalyze formation of a key intermediate in BChl and Chl synthesis (Chapter 4, Willows and Kriegel). AcsF is thought to require O2, whereas BchE is thought to use H2O for the O atom needed for this step in (B)Chl synthesis (Ouchane et al., 2004). Sequencing studies revealed that whereas the acsF gene was absent in strict anaerobes such as green sulfur bacteria, homologs existed in facultative aerobes (purple and green nonsulfur bacteria) and in strict aerobes (cyanobacteria, green algae and plants), with the latter group lacking bchE (Ouchane et al., 2004). Although AAP would be expected to possess acsF and lack bchE, full genome sequencing of Rsb. denitrificans has turned up both genes (Swingley et al., 2007), making it unclear which genetic elements are responsible for restricting BChl synthesis to aerobic conditions in AAP. The final picture of O2 regulation of BChl synthesis is clearly more complex than indicated simply by presence or absence of two complementary genes. Maximal BChl synthesis in obligately aerobic Rst. depolymerans occurs at an atmospheric O2 content of only 2% (Suyama et al., 2002), and R. thiooxidans best expresses BChl microaerophilically (Yurkov and
Chapter 3
Aerobic Anoxygenic Phototrophs
Csotonyi, 2003) and C. litoralis exhibits chemotactic preference for about 10% O2 concentration (Fuchs et al., 2007). What governs species-specific fine-tuning of optimum O2 concentration for pigment synthesis is currently unknown. When complemented with Rsb. denitrificans homologs of the anaerobic phototrophic two-component O2 regulatory system (RegA and RegB) of pigment synthesis, Rhodobacter capsulatus still only expressed O2-deprived pigment synthesis (Masuda et al., 1999), indicating that the AAP system functions as does the Rhodobacter counterpart (Chapter 35, Bauer et al.). The mechanism by which light regulates pigment synthesis in AAP is even cloudier than is the story for O2. Possibly, light repression of pigmentation is an overexpression of a similar but less potent process that purple nonsulfur bacteria employ in response to dangerously intense radiation. In most AAP, such as Erythromicrobium hydrolyticum, even dim light is strongly repressive to BChl synthesis (Yurkov and van Gemerden, 1993). However, Suyama et al. (2002) demonstrated 25% maximal BChl content in Rst. depolymerans at 10 W/m2 light intensity, while D. shibae produced BChl at 15% of its dark rate in the light (Biebl and Wagner-Döbler, 2006). Most interesting is a recent investigation of pigment synthesis by D. shibae under light-dark cycles of varying periodicity (Biebl and Wagner-Döbler, 2006), an elaboration on similar work by Yurkov and van Gemerden (1993). When the light:dark regimen was changed from 8h:16h (simulating winter conditions) to 16h:8h (summer conditions), the nightly rate of BChl synthesis increased nearly four-fold, indicating not only sensitivity to instantaneous level of illumination, but also a response to the duration of the previous illuminated interval (Biebl and Wagner-Döbler, 2006). Such regulation would be advantageous in the organism’s native marine environment, preparing more photosynthetic pigment on summer nights in anticipation of longer, warmer days, when higher doubling rates more extensively dilute the cell’s PSU complement. The authors suggest that signal transduction systems such as the recently discovered bacteriophytochrome (Evans et al., 2005; Chapter 40, Evans et al.) might mediate the effects of light in BChl synthesis in AAP (Biebl and Wagner-Döbler, 2006). Such speculation sets the stage for exciting future biochemical research. AAP hold additional fascinating surprises. Pigment expression in Thalassobacter stenotrophicus, H. phototrophica and Cmi. bathyomarinum is salt-sensitive;
43 T. stenotrophicus exhibits no pigmentation at 7% salinity, H. phototrophicum produces maximal BChl at 0.6% salinity and no BChl at 3.5% salinity, and Cmi. bathyomarinum manufactures the most BChl in the absence of NaCl, despite the native marine environment (~3.5% salinity) of all three species (Rathgeber et al., 2004; Macián et al., 2005; Biebl et al., 2006). Indeed, numerous studies have demonstrated that AAP tend to upregulate pigment synthesis under suboptimal growth conditions, implying that photosynthesis proves especially advantageous under stress (Rathgeber et al., 2004). Supporting this idea, thermophilic Rubritepida flocculens synthesizes BChl at 30 °C, but not at its preferred growth temperature of 50 °C (Alarico et al., 2002), and whereas marine Erythrobacter relative NAP1 grew optimally at 32.5 °C, BChl production maximized at 22.5 °C and halted at temperatures above 30 °C (Koblížek et al., 2003). Given the prominence of heterotrophic metabolism in AAP, nutritional status is a paramount regulatory factor of photosynthetic pigment synthesis (Beatty, 2002). Sudden carbon source dilution caused Rst. depolymerans to transcriptionally induce BChl synthesis (Suyama et al., 2002). This response makes sense in light of competition for electron carriers by heterotrophy and phototrophy in AAP: redox agents are maximally available for phototrophy when exogenous carbon sources are limiting (Beatty, 2002). Such is the case for E. ramosum, which produces over twice as much BChl when starved as when amended richly with nutrients (Fig. 3D and Color Plate 2). Most unusually, however, BChl synthesis occurred only in stationary phase cultures of Rst. depolymerans, and illumination only extended the viability of mature cultures instead of enhancing growth rate as in E. hydrolyticum and D. shibae (Yurkov and van Gemerden, 1993; Biebl and Wagner-Döbler, 2006). These observations inspired suggestion of an innovative use for the photosynthetic apparatus in some AAP: maintenance of the electrochemical proton gradient by light-induced cyclic electron flow, a condition that is essential for cellular integrity during starvation conditions (Suyama et al., 2002). In other species, the role of nutrients in regulation of pigment synthesis is more obscure. Erythrobacter litoralis produces more carotenoids but the same amount of BChl when organic carbon is limiting (Fig. 3C and Color Plate 2), and production of BChl and carotenoids in Rsc. thiosulfatophilus are both stimulated by a rich organic medium (Fig. 3B and
44 Color Plate 2), necessitating more research to unravel such mysteries. Even more interestingly, regulatory effects of two factors can interact nonlinearly in unexpected ways, best illustrated by exposure of H. phototrophica and strain DFL-11 to 8-hour-long pulses of light and starvation (Biebl and Wagner-Döbler, 2006). Whereas pulses of light or starvation alone had little or no effect on BChl levels, simultaneous exposure to both factors caused an order-of-magnitude increase in BChl, especially significant in these otherwise poorly pigmented species (Biebl and Wagner-Döbler, 2006). Thus, transient illumination and nutrient deprivation may facilitate the detection of BChl in strains in which its presence is uncertain and therefore aid in AAP identification. In light of this discovery, the tirade about incorrect taxonomic assignment of AAP to non-phototrophic genera in Section II.C deserves the balance of an important counterpoint. Several recent isolates, including Rva. mucosus and H. phototrophica, initially tested spectrophotometrically negative for BChl production and were recognized as phototrophs solely by their pufLM genes (Allgaier et al., 2003). The repeated insistence that photosynthetic apparatus expression is a useful taxonomic marker for AAP because BChl is easily detected (e.g., Yurkov and Csotonyi, 2003) must be tempered with caution. Indeed, Rva. mucosus failed to produce measurable BChl even under the pulse of starvation and illumination that induced such a marked increase in pigmentation in H. phototrophica and strain DFL-11 (Biebl and Wagner-Döbler, 2006). Whereas a negative conclusion of BChl production should only follow a reasonably wide application of culture conditions, it is nonetheless conceivable that some isolates will evade all reasonable attempts to induce BChl synthesis, and would therefore understandably be misclassified as non-phototrophs in the absence of genetic sequencing. Aside from applications in taxonomic hair-splitting, this point inspires a potentially informative investigation. A PCR-based screening of described proteobacterial species for pufLM, acsF and bchE genes could delineate the true phylogenetic distribution of aerobic phototrophy in this group. The pufLM genes would establish phototrophic capability, while presence of acsF and absence of bchE could indicate an obligately aerobic nature. Physiological work on species possessing pufLM should then be performed as a precaution, because Swingley et al. (2007) and Fuchs et al. (2007) have reported bchE in the completely or nearly completely
Vladimir Yurkov and Julius T. Csotonyi sequenced genomes of the AAP R. denitrificans and C. litoralis, repspectively. V. The Mysterious Photosynthetic Apparatus of Aerobic Anoxygenic Phototrophs Demonstration that AAP incorporate BChl into a functional photosynthetic apparatus certainly raised eyebrows in a skeptical research community struggling to accept the idea of an aerobic variant of anoxygenic photosynthesis. Nevertheless, AAP proved to possess a properly transcribed, bonafide purple bacterial-type photosynthetic superoperon, encoding apoproteins of RC, LH1 and often LH2 complexes (Yurkov, 2006). Exactly how a sophisticated set of anaerobically evolved pigment-protein complexes was enlisted to operate only aerobically is still debated, and is grievously neglected in recently published research. The striking genetic and structural similarities of AAP photosynthetic machinery to that of purple nonsulfur bacteria may have stifled some interest in this area. Increasing accessibility of relevant tools and techniques should cause this field to burgeon. A. The Reaction Center of Aerobic Anoxygenic Phototrophs Has Purple Bacterial Roots As in anaerobic purple phototrophs, the heart of the AAP photosynthetic apparatus consists of a type II (quinone-type) RC with three structural subunits (L, M and H), four BChls, two bacteriopheophytins, two ubiquinones, a nonheme high-spin Fe2+, and a carotenoid (Yurkov and Beatty, 1998). In purple bacteria, components are encoded on the ~45 kb photosynthesis gene cluster, and superoperons and polycistronic mRNAs contribute to stoichiometric consistency (Klug, 1993). The structural cornerstone of the RC makes the genes encoding the L and M subunits (pufLM) most frequently utilized in genetic surveys for phototrophs (Yutin et al., 2005). RC gene sequences in AAP are similar to genes from anaerobic anoxygenic phototrophs, illustrating that only a few modifications were required to enable anoxygenic phototrophy to function in the presence of O2. As a prime example, a single key replacement of His L168 with glutamic acid in the RC of Acidiphilium may be largely responsible for stabilization and function
Chapter 3
Aerobic Anoxygenic Phototrophs
of Zn-BChl (Nagashima et al., 1997). However, the flip-side of high structural and functional dependence on fine-scale gene sequence variation is that coarse genetic analysis techniques may fail to resolve much diversity. Indeed, introducing greater variation in two new pufM primers returned 14 times as many phototrophic sequences from Eastern Mediterranean Sea water samples as did probes constructed to amplify the single pufM sequence (Yutin et al., 2005). Some reports of diversity in photosynthetic gene organization are controversial. Exemplifying this, the recent claim by Pradella et al. (2004) that the pufLM genes of Roseobacter litoralis and Stl. guttiformis are not chromosomal but reside on linear plasmids of 91 and 120 kb size, respectively, could provide another ‘smoking gun’ for the idea of lateral photosynthetic gene transfer among AAP. However, these conclusions must be weighed hesitantly, because both partial and complete genome sequencing have established a chromosomal location for the puf operon of Rsb. denitrificans (Liebetanz et al., 1991; Swingley, et al., 2007). It seems unlikely that congeneric organisms (two species of Roseobacter) could possess such vastly different genomic arrangements. If further research rules out the possibility of an artifact, then such a discovery would bestow on the AAP uniquely flexible photosynthesis gene location. Great molecular diversity in AAP exists in Cyt electron carriers that close the photosynthetic electron transfer circuit by returning electrons to photo-oxidized BChl in the RC. In Erythrobacter, Erythromicrobium and Mahoney Lake strain ML31, the RC is reduced by a soluble periplasmic Cyt c2, as in Rhodobacter (Rba.) sphaeroides (Rathgeber et al., 2004). In addition to a soluble Cyt, Cmi. bathyomarinum, Erythromonas ursincola, R. mahoneyensis, Rsb. denitrificans, Rsc. thiosulfatophilus and Srb. sibiricus also possess a tetraheme Cyt c (encoded by pufC in Rsb. denitrificans) tightly bound to the RC, a characteristic of the anaerobic anoxygenic phototroph Blastochloris viridis (Rathgeber et al., 2004; Yutin and Béjà, 2005). Interestingly, Rsb. denitrificans lacks the posttranslational N-terminal cleavage of PufC that anchors the resulting tetraheme Cyt c to the membrane via a diacylglycerol molecule in purple bacterial species such as Blastochloris viridis (Hucke et al., 2003). Instead, the elongated N-terminus fastens Cyt c and contains a transmembrane helix motif resembling the core of another polypeptide, PufX. The Rhodobacter PufX protein affects the orientation and dimerization of the RC-LH1-PufX complex (Francia et al., 1999;
45 Siebert et al., 2004; Chapter 9, Bullough et al.), and the pufX gene is located where pufC sits in other species (Yutin and Béjà, 2005). Roseobacter thus opens a unique window on the evolutionary history of the PufX protein, which may have evolved by incomplete deletion of pufC in ancestral phototrophs switching from a RC-bound to a soluble Cyt c that donates electrons to the RC (Hucke et al., 2003). Although AAP have not been shown to contain pufX, bacterial artificial chromosome (BAC) cloning vectors constructed from marine samples have yielded putative AAP sequences (such as eBAC 60D04) from uncultured Gammaproteobacteria containing a gene strongly resembling pufX (Yutin and Béjà, 2005). However, ascribing uncultured anoxygenic photosynthetic gene sequences from aerobic environments to the AAP is risky, as purple nonsulfur bacterial sequences may masquerade as AAP to unwary eyes. B. Light-Harvesting Complexes: Ordinary and Extraordinary Surrounding the RC, AAP possess LH antenna complexes that sequester additional photonic energy, both by increasing light-harvesting cross-sectional area and by broadening the range of captured wavelengths. The core LH1 complex, existing in all AAP, shows little variation, but accessory LH2 manifests unexpected diversity. Reflecting the conserved nature of LH1 is its close genetic association with the RC: LH1 subunits β and α are encoded by genes (pufBA) cotranscribed with and immediately upstream of pufLMC (Yutin and Béjà, 2005). Carotenoid-rich LH1 forms an intimate ring of about 30 BChls around the RC (Yurkov and Beatty, 1998), facilitating highly efficient transfer of excitons to the RC primary electron donor, P870. In this respect, AAP strongly resemble purple nonsulfur bacteria such as Rba. sphaeroides, sharing similar RC-LH1 near-IR absorption spectra, with maxima at around 800 nm (RC) and 870 nm (LH1) (Yurkov and Csotonyi, 2003). With the exception of Zn-BChl of Acidiphilium, however, a few AAP display further unexplained spectral deviation of LH1, such as blue-shifting (856 nm maximum) in Rsc. thiosulfatophilus (Fig. 3B and Color Plate 2) or red-shifting (879 nm maximum) in Roseovarius tolerans (Yurkov and Beatty, 1998; Yurkov and Csotonyi, 2003). The structural factors responsible for this spectral fine-tuning remain to be elucidated. In contrast to LH1, the peripheral LH2 complex of AAP varies widely in both expression and structure.
46 In Rhodobacter and Rhodopseudomonas anaerobic phototrophic species, the LH2 genes reside in one or more puc operons, presumably enabling more independent transcriptional regulation (Klug, 1993; Larimer et al., 2004). Most tantalizing is the striking spectral differences of LH2 antennas in AAP from those of purple bacteria. When present, the LH2 complexes of anaerobic purple phototrophic bacteria exist in several spectral forms, their nomenclature denoting primary near-IR absorption maxima in nm: e.g., B800-850 (typical Rhodobacter-type), B800-820 (in Rhodopseudomonas (Rps.) acidophila) and the BChl b-containing B800-1020 (in Ectothiorhodospira) (Steiner and Scheer, 1985; Cogdell et al., 2002). The unusual purple nonsulfur bacterium Rps. palustris possesses four different peripheral LH species, including a monomodal LH2 complex with a single absorption peak (Hartigan et al., 2002). Interestingly, and for reasons unknown, the LH2 spectral classes of AAP often differ from those of their anaerobic relatives: B798-832 (in α-4 Erythromicrobium, Sandarakinorhabdus and Porphyrobacter dokdonensis) (Fig. 3D and Color Plate 2), B800-814 (in α-4 Porphyrobacter donghaensis) and B806 (in the α-3 Roseobacter, Roseicyclus and Rubrimonas) (Fig. 3A and Color Plate 2) (Yurkov and Beatty, 1998; Rathgeber et al., 2004; Yoon et al., 2004b, 2006; Gich and Overmann, 2006). The latter is a rare monomodal LH complex, similar to that found in only one anaerobic species, Rps. palustris (Hartigan et al., 2002). However, a 4 nm difference in its absorption maximum calls into question how structurally similar B806 is to the Rps. palustris LH2, which is an octamer with four BChl per αβ peptide instead of the usual three, each arranged radially rather than tangentially (Hartigan et al., 2002). Cogdell et al. (2002) propose that controlling the rotation of the BChl acetyl group relative to the bacteriochlorin ring plane by adjusting the content of hydrogen bonding amino acid residues at key positions in the LH2 apoprotein permits effective modulation of the Qy absorption band shift. Because loss of hydrogen bonding robustly explains the development of B800-820 from B800-850 (Fowler et al., 1992, 1994), a similar transition may have generated B798-832. Curiously, however, Raman spectroscopy indicates the presence of H bonds to the acetyl substituents of BChl in E. ramosum (Yurkov and Beatty, 1998), meaning that the case is definitely not closed on the source of LH2 Qy band shifts in AAP.
Vladimir Yurkov and Julius T. Csotonyi C. The Riddle of Aerobic Anoxygenic Phototrophy Exploring PSU biophysics, biochemistry and molecular genetics should resolve the question addressing the definitive quality of AAP, i.e., what allows their PSU to function aerobically. Unfortunately, similarity in RC architecture between AAP and anaerobic phototrophs (Section V.A.) complicates matters. Further smearing the distinction between aerobic and anaerobic phototrophy is the recent discovery of obligately microaerophilic photosynthesis in R. mahoneyensis (Rathgeber et al., 2004). Still, several studies provide a smattering of reasons that may explain why AAP photosynthesis is obligately aerobic. Energy-conserving photosynthetic electron flow in AAP appears to trace a circuit typical of anaerobic anoxygenic phototrophic bacteria, from the photooxidized special pair of BChl (P870+) to auxiliary BChl, bacteriopheophytin, the ubiquinone primary e– acceptor QA, then exiting the RC to the membrane quinone pool, a membrane bound Cyt bc1 complex, Cyt c2, and back to the RC (Fig. 3F and Color Plate 2). AAP differ subtly from purple nonsulfur bacteria at three principal points, the first involving the redox poise of the primary acceptor, QA. In most AAP, QA has a much higher midpoint potential (+5 to +150 mV) than in anaerobic purple bacteria (usually negative potentials), causing it to become and remain overreduced anoxically and halting electron flow (Fig. 3F and Color Plate 2) (Rathgeber et al., 2004). The structural basis of this functional discrepancy remains a mystery, as homology modeling of quinone binding pockets in the RC of Rsb. denitrificans demonstrates remarkably similar interactions between protein and quinone as compared with Rba. sphaeroides (Kohler et al., 2005). The second distinction of AAP is the lack of an alternative quinol oxidase pathway, thought to maintain the quinone pool at a proper redox state for efficient electron transport under photosynthetic conditions (Yurkov and Beatty, 1998). Finally, failure of soluble Cyts to transfer electrons anoxically from the Cyt bc1 complex to the RC-bound Cyt c in Rsb. denitrificans also implicates these components in arresting anaerobic photosynthetic electron flow in AAP (Schwarze et al., 2000). Hence, properties of one or more electron carriers found in the RC, membrane lipid phase and periplasm may prohibit anaerobic photosynthesis in the AAP, but the relative contribution of each element is presently unknown.
Chapter 3
Aerobic Anoxygenic Phototrophs
Whether scarcity of recent publications is a cause or an effect of the recalcitrance of the central biophysical question encompassing obligately aerobic photosynthesis is uncertain, but hopefully this lull will soon end. VI. Speculation on Ecological Roles Despite strides over the last four years in both culture-based and culture-independent studies, AAP ecological understanding is still in its adolescence. Researchers have discovered experimental tools with great potential, but proper combination of techniques to guarantee maximal analytical power still requires refinement. At the crux of the issue, comprehension of ecological significance requires qualitative description of interactions to go hand in hand with both assessment of biogeochemical cycling and reliable enumeration. Whereas some interesting associations have been described, work on elemental cycling is scant. Also, in spite of its seeming triviality, measuring the abundance of AAP in nature has proven exceptionally difficult. A. Technical Challenges of Enumerating Aerobic Anoxygenic Phototrophs AAP were traditionally enumerated by culturing on rich organic media (Yurkov and Beatty, 1998). This technique is very powerful for assessing the abundance of a physiological subset of the community and offers the added benefit that all enumerated organisms can subsequently be examined in physiological and phylogenetic detail. However, realization of the miniscule proportion of microorganisms amenable to cultivation spurs a growing interest in census techniques that circumvent the need to culture organisms. Regrettably, most culture-independent methods possess significant shortfalls. Phylogenetic surveys based on 16S rRNA genes alone, constituting much of marine microbial research, fail to discriminate between closely related phototrophic and non-phototrophic organisms. Efforts to screen for photosynthesis genes, such as quantitative or real-time PCR (Schwalbach and Fuhrman, 2005; Du et al., 2006), overcome this phylogenetic interspersion issue, but using primers constructed from described organisms inherently limits the diversity of detectable phototrophs (Yutin et al., 2005). Even bulk pigment analysis of ocean water by HPLC, or of individual cells by infrared
47 epifluorescence microscopy (IREM) or its variants (Schwalbach and Fuhrman, 2005; Jiao et al., 2006; Sieracki et al., 2006) are faulty as they measure a pool of BChl produced by both AAP and purple nonsulfur bacteria. IREM can even misidentify cyanobacteria as anoxygenic phototrophs and underestimate Prochlorococcus populations (oxygenic phototrophs particularly numerous in oligotrophic ocean waters) by orders of magnitude (Jiao et al., 2006), although newer studies attempt to correct for this effect (Lami et al., 2007). Finally, the sensitivity of BChl synthesis to a slew of environmental factors (Section IV.C.) generates great variation in pigment content from cell to cell, invalidating estimates of overall AAP abundance based on average cellular BChl concentration (e.g., Kolber et al., 2001; Koblížek et al., 2005). Obviously, measuring the abundance of AAP in nature is no trivial task. Thus far, three preferable alternatives exist: (1) infrared fast repetition rate fluorometry (IRFRR), which unfortunately requires state-of-the-art instruments, but can measure and distinguish between aerobic and anaerobic photosynthetic electron transport; (2) QPCR of pufLM (and of genes that can be used to discriminate between aerobic/anaerobic photosynthesis, such as acsF and bchE); and (3) combination of culture-independent ribosomal and photosynthetic gene surveys with cultivation, followed by 16S rDNA genetic analysis of cultured organisms. To date, only a few studies have applied either the first (Kolber et al., 2000, 2001; Koblížek et al., 2003, 2006) or the third (e.g., Gich et al., 2005; Salka et al., 2006) strategy to AAP, and no studies have yet employed the second. The majority of reported marine AAP enumeration must therefore be judged inconclusive, because abundance estimates represent only an upper limit, pooling AAP with nonphototrophs or anaerobic phototrophs, depending on methods employed. B. Ocean Surface Between 2003 and 2006, nearly 75% of approximately 22 ecologically relevant publications focused on marine environments, from which AAP were originally isolated. Three reports likely instigated this return to the sea. First, AAP were surprisingly recovered from deep ocean hydrothermal vent plumes (Yurkov et al., 1999). Second, although AAP have long been known to constitute up to 10 to 30% of the cultivable bacterial community in hypersaline and hydrothermal environments (Yurkov and Csotonyi, 2003) (Fig. 3G, I and
A, B ND NW Atlantic: Sargasso Sea (11–15 m) Sieracki et al. (2006)
Mar 02
1.3–2.5%; (0.71–1.49 × 104 ml–1); 2.4–3.3% of biomass
IREM
A, B ND NW Atlantic: USA coast to Gulf Stream (11–15 m) Sieracki et al. (2006)
Mar 02
0.8–2%; (0.76–2.1 × 104 ml–1); 1.8–2.5% of biomass
IREM
A, B ND NW Atlantic: Sargasso Sea (11–15 m) Sieracki et al. (2006)
Oct 01
0.8–2.6%; (0.82–1.26 × 104 ml–1); 3.4–5% of biomass
IREM
A, B
A, B NW Atlantic: USA E coast to Gulf Stream (11–15 m) Sieracki et al. (2006)
Oct 01
ND
IREM
A, B
2.3–9.4%; (2.16–9.84 × 104 ml–1); 7–12% of biomass
IREM ND 25 ps was calculated (Hsu et al., 2001), also in agreement with the very low efficiency obtained experimentally (Macpherson, 2001; Polívka et al., 2002; Wohlleben, 2003). Although the trend of decreasing efficiency of the S1-mediated energy transfer was confirmed by experiments using LH2 complexes from Rba. sphaeroides having the different carotenoids, neurosporene, spheroidene, and spheroidenone, the drop in efficiency when going from spheroidene to spheroidenone was less than expected on the basis of a change in spectral overlap. The S1 lifetimes of carotenoids in LH2 complexes of 1.4, 1.5 and 1.4 ps for neurosporene, spheroidene, and spheroidenone were compared with their lifetimes in solution (24, 8.5 and 6 ps), resulting in energy transfer efficiencies of 94, 82 and 76% (Polívka et al., 2004). Knowing that the S1 energy of spheroidenone is around 13000 cm–1 (Zigmantas et al., 2004), the efficiency of energy transfer from spheroidenone does not match that expected from calculations by Hsu et al. (2001). Also, comparison of 76% with the nearly zero efficiency of the S1-mediated energy transfer obtained for rhodopin glucoside that has a LH2 S1 energy of 12550 ± 150 cm–1 (Polívka et al., 2002) would imply that ~500 cm–1 decrease in energy is enough to decrease efficiency from 76 to 5%. Taking into account that S1 emission of carotenoids is quite broad (Fujii et al., 1998; Frank et al., 2000a), such a drop can be hardly explained as due solely to change in the spectral overlap (Ritz et al., 2000). Thus, it indicates that other factors besides conjugation length may play a role. One possibility is that the high efficiency of the S1mediated energy transfer of spheroidenone is related to the fact that it belongs to the family of carbonyl carotenoids, which possess an excited state with chargetransfer character (Frank et al., 2000b; Zigmantas et al., 2004) that may enhance the spheroidenone-BChl coupling. Another explanation was offered by Ritz et al. (2000) who noted that the conjugated systems of neurosporene and spheroidene, which systematically exhibit highly-efficient energy transfer via the S1 state, have a non C2h-symmetrical arrangement of their methyl side groups. This is, however, not
Harry Frank and Tomáš Polívka the case for lycopene and rhodopin glucoside, both exhibiting very low efficiencies of the S1-mediated energy transfer. Consequently, Ritz et al. (2000) suggested that symmetry breaking of neurosporene and spheroidene may be an important factor in explaining why energy transfer via the S1 state is much more efficient for these carotenoids compared to lycopene and rhodopin glucoside. Since the conjugated system of spheroidenone has also asymmetric arrangement of methyl groups (Fig. 2), the observation of efficient S1-mediated energy transfer, despite its low S1 energy, provides further support for the conjecture proposed by Ritz et al. (2000). In addition to the S1-mediated energy transfer occurring from a thermalized S1 state discussed above, it has been proposed that a portion of the pathway may involve a vibrationally hot S1 state. Using global analysis, Papagiannakis et al. (2002) reported the decay of a species associated spectrum corresponding to the vibrationally hot S1 state, suggesting the presence of an energy transfer channel via this route. However, the contribution of this pathway to the total energy transfer efficiency was only 5%. Indirect evidence for the presence of this channel also could be found in experiments carried out by Krueger et al. (1999) who measured the spectral profile of the S0-S1 transition in the LH2 complex by two-photon excitation techniques that detect emission from BChl a. The 0-0 band of the S0-S1 transition was very weak but the intensity of higher vibrational bands was high which may be due to energy transfer occurring from the hot vibrational states. In any case, this energy transfer channel plays only a minor role in the overall carotenoid-BChl energy transfer in LH2. An upper limit of 3% was found in the LH2 complex from Rps. acidophila for this channel (Wohlleben et al., 2003). Moreover, as no evidence for energy transfer via a hot S1 state was provided by a number of other experiments using various LH2 complexes (Papagiannakis et al., 2003a; Koyama et al., 2004; Rondonuwu et al., 2004; Polli et al., 2006), further experiments using both advanced experimental approaches and sophisticated data analysis will be needed to verify the presence of this energy transfer channel. C. The S* State Reports of other carotenoid excited states located between the S2 and S1 states (see Polívka and Sundström, 2004, for a review) initiated lively debates regarding whether these states act as energy donors
Chapter 12
Carotenoid-to-BChl Energy Transfer
in carotenoid-BChl energy transfer. New ways of data analysis allowed for a more rigorous assignment of various excited state species (van Stokkum et al., 2004), and it was shown in a number of cases that the two-state (S2 and S1) model is not sufficient to describe all features revealed in experimental data. The so-called S* state is the best studied in this respect; it was first reported in the excited state manifold of the carotenoid spirilloxanthin in both solution and the LH1 complex of Rsp. rubrum (Gradinaru et al., 2001). Using global analysis of data in the 470–720 nm spectral region, it was shown that the S1-Sn band of spirilloxanthin in solution (peaking at 590 nm) possessed a distinct shoulder at ~540 nm. While the 590 nm band decayed with 1.4 ps corresponding to the S1 lifetime of spirilloxanthin, the 540 nm shoulder exhibited a much longer decay time of ~6 ps (Gradinaru et al., 2001). This result was explained in terms of two parallel pathways of S2 depopulation; a major part (70%) decaying to form the S1 state, while a minor pathway (30%) leads to population of the S* state which then decays to the ground state with a 6 ps lifetime. Interestingly, as shown in subsequent studies (Papagiannakis et al., 2002; Papagiannakis et al., 2003a; Wohlleben et al., 2003), the S* state is formed with much higher yield when carotenoids are incorporated into purple bacterial LH complexes. Moreover, in LH1 and LH2 complexes the S* state was found to be a precursor of fast carotenoid triplet state formation. A relatively high triplet yield of 25–30% was explained to be a result of a conformational distortion of spirilloxanthin in the LH1 complex, promoting triplet formation via singlet homofission from the S* state (Gradinaru et al., 2001). Further studies confirmed the triplet state formation via the S* state for other complexes containing different carotenoids. The yields of triplet formation varied from nearly 40% for rhodopin glucoside in LH2 from Rps. acidophila (Wohlleben et al., 2003) to less than 10% for spheroidene in both native LH2 from Rba. sphaeroides (Papagiannakis et al., 2002) and incorporated into the Rb. sphaeroides R26.1 mutant (Papagiannakis et al., 2003a). The fact that no triplet formation is observed in solution suggests that the protein environment is a crucial factor governing the formation of the triplet via singlet homofission, and it strongly supports the conclusion that the deviation from planar conformation in antenna complexes of purple bacteria is a necessary condition for efficient triplet formation (Papagiannakis et al., 2003a).
223 However, it turns out that the decay to the S0 state and triplet formation are not the only possible fates of the S* state in LH2 complexes. In LH2 complexes containing either spheroidene or rhodopin glucoside, the S* state also contributes to carotenoid-BChl energy transfer. For spheroidene in the LH2 of Rba. sphaeroides, S*-mediated energy transfer contributes 10–15% to the total spheroidene-BChl energy transfer (Papagiannakis et al., 2002). For rhodopin glucoside in Rps. acidophila LH2, the S*-mediated channel was found to be ~10% efficient; i.e., even higher than the energy transfer efficiency from the S1 state (Wohlleben et al., 2003). The possibility of carotenoid-BChl energy transfer via the S* state provided important information for determining the origin of the S* state, which is still a matter of debate. The presence of the S*-mediated energy transfer puts the S* energy above the Qy bands of BChl a, eliminating one of the proposed origins, a vibrationally hot ground state (Wohlleben et al., 2004). Instead, it seems that S* is indeed a separate excited state as originally suggested by Gradinaru et al. (2001), but its symmetry and relation to other states remains unknown. It also must be noted that although only a few carotenoids were subject to studies focusing on the S* state so far, it seems obvious that the lifetime of the S* state does not follow any clear dependence on conjugation length. While the S1 lifetime is changed systematically from ~9 ps (spheroidene) to 4.1 ps (rhodopin glucoside) and 1.5 ps (spirilloxanthin) as a result of increased conjugation length from 10 to 13 (Polívka and Sundström, 2004), the intrinsic S* lifetimes (in the absence of energy transfer) in LH2 and LH1 complexes are scattered in the 6 to 30 ps range without any obvious relation to the conjugation length (Gradinaru et al., 2001; Papagiannakis et al., 2002, 2003a, 2006a; Wohlleben et al., 2003). The lack of correlation between lifetime and conjugation length also casts doubts on assignment of the S* state to the 1Bu– state proposed by some authors (Papagiannakis et al., 2002; Wohlleben et al., 2003). A recent thorough study of the S* state in LH2 complexes from Rps. acidophila and Rba. sphaeroides revealed another aspect of the spectroscopic properties of the S* state in LH2 complexes. Population of the S* state exhibits a dependence on intensity of excitation pulses that differs from that observed for the S1 state (Papagiannakis et al., 2006a), indicating that the S* and S1 state cannot have a common precursor, the S2 state, as previously thought. These authors proposed
224 two models to explain their data. One assumes two different ground-state populations each leading to population of either S1 or S* state, the other involves higher excited states. These upper excited states are populated via excited state absorption from the S2 state that is resonant with the excitation pulse, thus the S2 population created by the front of the excitation pulse can be re-excited into higher excited states by photons arriving in the tail of the pulse. These higher excited states exhibit a relaxation pattern that favors population of the S* state. Because increasing excitation intensity increases the probability of the re-excitation, the S* state gets more populated with higher excitation intensities (Papagiannakis et al., 2006a). This model involving higher excited states has gained support from another study showing that direct excitation of high-lying excited states enhances population of the S* state (Billsten et al., 2005). These authors also suggested that the S* state in solution may be related to a conformational change of the carotenoid molecule. This possibility was later supported by a study on a series of carotenoids with different structures, indicating that the S* state may be a minimum in the S1 potential surface corresponding to a conformational change (Niedzwiedzki et al., 2006). This conclusion also explains why the S* state is preferentially populated in LH2 complexes where such distortions have been confirmed by X-ray crystallography. D. Other Pathways Another excited state widely discussed as a potential energy donor in carotenoid-BChl energy transfer in LH2 complexes is the 1Bu– state. Its presence in the excited state manifold was predicted two decades ago by calculations on polyenes carried out by Tavan and Schulten (1987), and it was shown that for conjugation lengths corresponding to naturally-occurring carotenoids (N = 9-13) it may be located below the strongly absorbing S2 state. Due to the forbidden nature of the 1Bu– state (it is forbidden for both oneand two-photon transitions from the ground state), experimental verification of the presence of the 1Bu– state between the S2 and S1 states is very difficult. In the late nineties Sashima et al. (1999) detected this state using measurements of resonance Raman profiles. Later, the 1Bu– state was also proposed to be active in energy transfer between carotenoids and BChl in LH2 complexes (Rondonuwu et al., 2004; Koyama et al., 2004). Based on previous assignments
Harry Frank and Tomáš Polívka of spectral signatures thought to be associated with the 1Bu– state in transient absorption spectra recorded in the near-infrared and visible ranges (Koyama et al., 2004), Rondonuwu et al. (2004) used global analysis of data taken on a few LH2 complexes containing different carotenoids and concluded that the 1Bu– state transfers energy to BChl with ~20% efficiency. This corresponds to an energy transfer time of ~0.6 ps for neurosporene and spheroidene in the LH2 complexes from Rba. sphaeroides. This pathway was proposed to be inactive for LH2 complexes accommodating the longer carotenoids lycopene and rhodopin glucoside. The absence of this channel in these complexes was explained by the 1Bu– state lying below the Qx state of BChl a, which was assumed to be the energy acceptor. For these carotenoids, however, it was hypothesized that another dark excited state, the 3Ag– state, may be active in carotenoid-BChl energy transfer because the expected 3Ag– energies may be favorable for this state to act as an energy donor. However, no experimental evidence for such a pathway was given (Rondonuwu et al., 2004). It must be noted that both 1Bu– lifetimes and spectral signatures in the visible region obtained by Rondonuwu et al. (2003) are essentially identical to those assigned earlier to the hot S1 state (Billsten et al., 2002b; de Weerd et al., 2002). Consequently, the data by Rondonuwu et al. (2004) may also be interpreted in terms of the hot S1 state being the energy donor instead of the 1Bu– state. Another issue that awaits further clarification is the relationship between the 1Bu– and S* states. It has been proposed that the 1Bu– state is identical with the S* state (Papagiannakis et al., 2002, Wohlleben et al., 2003; Rondonuwu et al., 2003). This assignment may seem correct as the symmetry and origin of the S* state are still unknown, and the 1Bu– state was predicted to be a precursor of ultrafast triplet formation (Rondonuwu et al., 2004; Koyama et al., 2004), the same process reported for the S* state (Gradinaru et al., 2001; Papagiannakis et al., 2002). On the other hand, for the S* state it was shown that there is no S1 ↔ S* conversion (Gradinaru et al., 2001; Papagiannakis et al., 2002; Wohlleben et al., 2003), which is in contradiction with the 1Bu– state being an intermediate state in S2-S1 internal conversion (Koyama et al., 2004). Similarly, the 1Bu– lifetimes (see Koyama et al., 2004, for a review) are about an order of magnitude shorter than those measured for the S* state, which argues that the 1Bu– and S* states are not equivalent.
Chapter 12
Carotenoid-to-BChl Energy Transfer
E. The Role of B800 For the LH2 and LH3 complexes, B850 (B820 in LH3) and B800 BChl a molecules may be acceptors in carotenoid-BChl energy transfer. The question of partitioning between these two possible acceptors has been addressed in a few studies. Macpherson et al. (2001) investigated S2-mediated energy transfer by fluorescence up-conversion in B800-B850 and B850only LH2 complexes from Rps. acidophila. Upon combining the results for these two LH2 complexes, they concluded that the S2 state of rhodopin glucoside transfers energy with 20% efficiency to B800 and with 31% efficiency to B850, leading to the total efficiency of 51% (Macpherson et al., 2001). Similar analysis was carried out by Papagiannakis et al. (2003a) who used transient absorption spectroscopy to study LH2 from a carotenoidless Rba. sphaeroides R26.1 mutant lacking the B800 BChl but incorporated with spheroidene. It was reported that the S2 pathway operates with 25% efficiency, which, when compared with the 57% efficiency of this pathway known for the wild type B800-B850 complex, allowed them to conclude that approximately half of the S2 population transfers energy to B800 (Papagiannakis et al., 2003a). It should be noted that although this work clearly showed that B800 accepts up to 50% of energy from the S2 state, calculations based on the X-ray structure of the LH2 complex from Rps. acidophila gave appreciable couplings only for the S2-B850 Qx channel (Krueger et al., 1998). On the other hand, similar calculations using full Coulombic couplings performed on the basis of the LH2 structure of Rsp. molischianum, yielded appreciable couplings of the S0-S2 transition with both the B850 and B800 BChls, resulting in close to a 1:1 branching ratio between the B800 and B850 acceptors (Damjanovic et al., 1999). Essentially the same results were obtained by calculations of the lycopene-BChl couplings by means of the collective electronic oscillators algorithm (Tretiak et al., 2000). Regarding energy acceptors in the S1-mediated energy transfer route, both B800 and B850 BChls are capable of accepting energy from carotenoids, but the S1-B800 channel seems to dominate. In experiments employing samples having spheroidene incorporated into LH2 from the carotenoidless Rba. sphaeroides R26.1 mutant lacking B800 BChl, the efficiency of energy transfer via the S1 pathway reached only 35% (Papagiannakis et al., 2003a). This is significantly less than the ~80% observed for the LH2 containing both
225 B800 and B850 (Walla et al., 2000; Zhang et al., 2000; Polívka et al., 2002), signaling that the main pathway involves B800 as an acceptor. The same conclusion was reached for LH2 from Rps. acidophila. Although the S1 efficiency is only 4–5% in the wild type complex, selective removal of the B800 BChls led to a complete absence of S1 energy transfer (Macpherson et al., 2001). Polívka et al. (2007) used lithium dodecyl sulfate (LDS) which selectively perturbs the B800 site (Chadwick et al., 1987). The LDS-treated complexes lack the B800 band and can be therefore used to investigate the role of B800 in energy transfer (Fig. 6). By measuring the S1 lifetimes of neurosporene, spheroidene and spheroidenone in LDS-treated LH2 complexes and comparing them with data taken on untreated complexes (Fig. 6), Polívka et al. (2007) showed that the carotenoid S1 lifetime has values in the range 1.4–1.8 ps for the untreated LH2 complexes, but it is prolonged to 2.7–3.5 ps for LDS-treated complexes. This is consistent with slower energy transfer via the S1 state. The S1 state branching ratios for energy transfer to the B850 and B800 BChls were calculated using the S1 lifetimes of the carotenoids in solution and in untreated and LDS-treated LH2 complexes. The calculations showed that the B800: B850 branching ratio remained essentially the same regardless of the conjugation length of the carotenoid. The values were 65:35 for LH2 with spheroidene and 60:40 for the complexes containing neurosporene
Fig. 6. Kinetics recorded at the maximum of the S1-Sn band for LH2 complexes from Rba. sphaeroides containing spheroidene. Kinetics are shown for spheroidene in solution (open triangles), LDS-treated LH2 complexes (open circles) and untreated LH2 complexes (full squares). The LH2 complexes were excited in the 0-0 band of the carotenoid S2 state at 515 nm. All kinetics are normalized to maximum. Solid lines represent fits.
226 or spheroidenone. This confirmed further that the B800 BChl a represents the dominant acceptor in carotenoid-BChl energy transfer via the S1 state. Interestingly, the B800:B850 branching ratio was not affected by conjugation length, indicating that even for spheroidenone having an S1 energy of ~13000 cm–1, the S1-B800 pathway remains dominant. The significance of the B800 BChl a molecule is further underlined by reports that it is likely the only acceptor in the minor energy transfer routes via the hot S1 state or the S* state (Papagiannakis et al., 2003a). IV. Energy Transfer in Light-Harvesting 1 Complexes and Reaction Centers Carotenoid-BChl energy transfer in LH1 complexes has been much less studied, and most of our current knowledge is limited to information provided by time-resolved studies of the LH1 complex from Rsp. rubrum. First experiments, however, were carried out by Ricci et al. (1996) who demonstrated efficient energy transfer via the S2 state of spheroidene in the LH1 complex from Rba. sphaeroides. Based on the comparison between fluorescence up-conversion data on spheroidene in solution and in LH1 they concluded that S2-mediated energy transfer operates with 65% efficiency (energy transfer time of 90 fs), which is even more efficient than for the spheroidenecontaining LH2 complex (Ricci et al., 1996). Later experiments on LH1 complexes from Rsp. rubrum containing the long (N=13) carotenoid, spirilloxanthin, revealed only 35% efficient S2-mediated energy transfer (Gradinaru et al., 2001). Moreover, it was shown that no energy transfer proceeds via the S1 state of spirilloxanthin. Thus, 35% is the total efficiency of the spirilloxanthin-BChl energy transfer, in agreement with earlier results based on measurements of fluorescence excitation spectra (Rademaker et al., 1980). The absence of the S1-mediated energy transfer pathway for spirilloxanthin likely results from its very long conjugated system which pushes its S1 energy too low to transfer energy to BChl a. Measurements of the S1-S2 spectra of spirilloxanthin in the LH1 complex determined the S1 energy of the carotenoid to be 11500 cm–1, which is below the energy of both B800 and B850 Qy transitions (Papagiannakis et al., 2003b). An investigation of LH1 complexes reconstituted with carotenoids with conjugation lengths in the range 9–13 showed that carotenoid-BChl energy transfer
Harry Frank and Tomáš Polívka in LH1 obeys the same trend as shown earlier for LH2. The overall carotenoid-BChl energy transfer decreases with increasing conjugation length, dropping from 78% for LH1 with neurosporene (N=9) to 36% for the spirilloxanthin-containing LH1 complex (Akahane et al., 2004). These are slightly less than values obtained for LH2. These authors found efficiencies for the S1-mediated energy transfer of 20 and 19% for LH1 complexes reconstituted with neurosporene or spheroidene, respectively. No transfer via the S1 state (efficiency 5M) the excitation is almost localized (1 < Ndel < 1.1). The energy transfer has the form of hopping between the two molecules. In this case the Förster and Redfield theories give approximately the same rate. Decreasing the energy gap (E1–E2 < 5M) results in formation of delocalized states (with Ndel increasing from 1.1 up to 2). In this case the excitonic interactions create a coherent mixing of the two sites. Instead of an excitation jumping from one site to the other, we now have a relaxation between two delocalized states. Increase of the spatial overlap between the
Fig. 1. The energy transfer rate as a function of the energy gap between two molecules calculated with the Förster and modified Redfield theories (only downhill rates are shown). The delocalization length Ndel is calculated as the inverse participation ratio of the exciton wavefunctions. The Ndel = 1 line is shown to highlight the deviations from the localized limit. The two frames correspond to Mnm = 255 and 55 cm−1. We use the exciton-phonon spectral density measured for the plant light-harvesting complex (Peterman et al., 1997). The relaxation rates have been calculated for 77 K. The specific non-monotonous dependence of the rates on the energy gap is determined by the shape of the phonon wing.
delocalized wavefunctions corresponds to having much faster transfer. The deviation of the Redfield rates compared with those predicted by the Förster equation increases in proportion to the deviation of delocalization length from the localized limit. This example suggests that interband transfers in purple bacteria (for example the energy transfer from B800 to B850, which corresponds to a large energy gap and small coupling) can be calculated using the generalized Förster approach. On the contrary, intraband dynamics (for instance the B850-B850 equilibration corresponding to a small energy gap and large exciton couplings) should be modeled by the Redfield theory.
Chapter 13
Spectroscopy and Dynamics of Excitation Transfer
II. Structure and Exciton Spectra of LightHarvesting 1 and 2 Bacterial Antenna Complexes A. Structure and the Excited-state Properties Since the determination of the crystal structure of the peripheral B800-B850 LH2 antenna of purple bacteria Rhodopseudomonas (Rps.) acidophila (McDermott et al., 1995) and Rhodospirillum (Rsp.) molischianum (Koepke et al., 1996) a direct calculation of the exciton Hamiltonian (Eq. 1) from first principles has become possible (Alden et al., 1997; Scholes et al., 1999; Tretiak et al., 2000a,b; Damjanović et al., 2002; Hu et al., 2002). These calculations revealed strong exciton couplings of the Qy transitions within the tightly packed B850 ring. The corresponding values of nearest-neighbor interaction energy vary around 250–300 cm−1, depending on the method of computation; see Tretiak et al. (2000b) for a review. Such strong interactions produce a broad exciton band with the lowest states near 850 nm and higher exciton states spreading up to the 800 nm region where they are superimposed with the degenerate levels of the B800 ring (Dracheva et al., 1996; Sauer et al., 1996; Alden et al., 1997; Hu et al., 1997; Koolhaas et al., 1997b; Wu et al., 1997b; Georgakopoulou et al., 2002; Novoderezhkin et al., 2003). The B800 ring is characterized by weak pigment-pigment couplings of about 30 cm−1, corresponding to localized excited states. Nevertheless, the excitonic interactions should be taken into account to correctly reproduce the B800 line shapes (Cheng and Silbey, 2006) and the excitation dynamics within the B800 ring (Novoderezhkin et al., 2003; Zigmantas et al., 2006). Interaction of the exciton states with the phonons induces a homogeneous line broadening (given by the function gkkkk), a red-shift of the pure electronic (zero-phonon) transitions (due to reorganization effects determined by λkkkk), and the transfer of excitation energy in the complex (relaxation or hopping between different states or sites). All these features are determined by the exciton-phonon spectral density Cn(ω) that can be extracted from spectroscopic data (for example, from the low-temperature FL spectra (Kühn et al., 2002)), or obtained numerically using molecular dynamics and quantum chemical calculations (Damjanović et al., 2002).
235
B. Disordered Exciton Model Besides fast nuclear motion (phonons and vibrations) the excitonic transitions are affected by slow conformational motion of the proteins. Such motion induces random shifts (quasi-static disorder) in the transition energies En and couplings Mnm. The exciton energies and wavefunctions become perturbed correspondingly. In conventional bulk spectroscopies (with averaging over many complexes) the conformational disorder produces inhomogeneous line broadening. In single molecule techniques a slow conformational motion can be viewed as fluctuations of the spectral shapes on a time scale from milliseconds to minutes. In the homogeneous limit (without disorder) all the dipole strength of the circular aggregate (made up of N molecules having almost in-plane orientation of the Qy transition dipoles) is concentrated in a degenerate pair of levels (k = ±1). The participation ratio is PR = 1/N and 3/(2N) (i.e., 0.056, and 0.083 for N = 18) for the lowest and higher states, respectively. This homogenous exciton structure is changed dramatically in the presence of disorder depending on the ratio between the disorder width σ and the nearest neighbor interaction energy M (Monshouwer et al., 1997; Alden et al., 1997; Novoderezhkin et al., 1999a). An example of how weak, moderate, and strong disorder (σ/M = 0.52, 1.29, and 2.23) affect the absorption spectrum of the B850 antenna (with N = 18) is shown in Fig. 2. In the case of weak disorder (σ/M = 0.52) the disorder value is less than the nearest neighbor interaction energy and the exciton structure is close to that of the homogeneous limit. The k = ±1 levels are almost degenerate and each have a dipole strength of about 8.5 (in the units of monomeric dipole strength). The lowest k = 0 state becomes weakly allowed borrowing a dipole strength of about 1 from the k = ±1 levels. The PR values are 0.6–1.2 and 0.9–1.1 for k = 0 and higher levels, respectively. In the second case the disorder value exceeds the exciton coupling (σ/M = 1.29), producing significant mixing of the exciton wavefunctions. The splitting between the k = ±1 states increases to 70 cm−1. The lowest state becomes superradiant (with dipole strength of 2) and more localized (with PR values increasing to 0.4–0.5 for the red-shifted realizations). Higher states become allowed and more localized (PR is spread between 0.09 and 0.2).
236
Rienk van Grondelle and Vladimir I. Novoderezhkin
Fig. 2. Top frames: Measured (open circles) and calculated (solid lines) absorption spectra for the LH2-B850 antenna of Rps. acidophila at room temperature. The calculated absorption is shown together with the individual exciton components (thin lines). Bars show dipole strengths of the exciton components (in units of monomeric dipole strength) averaged over disorder. Line shapes are obtained with the spectral density modeled by an overdamped Brownian oscillator with coupling value of λ0 = 200 cm−1 and width (inverse bath relaxation time) of γ0 = 100 cm−1. The energies of the 1α1β, 1β2α, 1α2α, 1β2β, and 1α2β interactions were taken to be 291, 273, −50, −36, and 12 cm−1.The absorption spectrum is averaged over disorder, which was modeled by uncorrelated shifts of the site energies (diagonal disorder) taken from a Gaussian distribution with FWHM of σ = 150, 375, and 650 cm−1. Bottom frames: Participation ratio PR(k) = ∑n(cnk)4 for the 7 lowest exciton levels (i.e., k = 0, −1, 1, −2, 2, −3 and 3) calculated for 2000 realizations of the disorder. Each point shows the PR value for one exciton state for one realization as a function of the wavelength corresponding to the zero-phonon line (ZPL) of this state.
A further increase of the disorder (σ/M = 2.23 in the third case) produces an even more uniform distribution of the dipole strength. The lowest state has (on average) become more superradiant, with an average dipole strength of 3. On the other hand, there are red-shifted realizations with an almost localized lowest state (with PR value up to 0.6–0.8). The higher states are more localized as well (with PR values distributed between 0.09 and 0.35). The LH1 antenna has a structure similar to that of the B850 ring of LH2. The larger size (N = 30–32 instead of 16–18 in LH2) results in less spacing between the zero-order exciton levels, which in turn produces stronger disorder-induced mixing of the
exciton states. In particular, the lowest k = 0 state becomes more superradiant and the higher k = ±2 states (which in LH2 stay almost forbidden even with moderate disorder) now give a significant contribution to the absorption profile (Novoderezhkin et al., 1999a; 2000; 2002). C. Steady-state Spectra The steady-state spectroscopic properties of LH1/ LH2 were studied with polarized light spectroscopy (Van Mourik et al., 1992; Visschers et al., 1995), nonlinear polarized absorption (Leupold et al., 1996), hole-burning (Reddy et al., 1991, 1992, 1993; Wu
Chapter 13
Spectroscopy and Dynamics of Excitation Transfer
et al., 1997a), low-temperature FL (Freiberg et al., 2003), superradiance (Monshouwer et al., 1997), circular dichroism (CD) (Koolhaas et al., 1997b, 1998; Georgakopoulou et al., 2002), Stark spectroscopy (Beekman et al., 1997), and single-molecule techniques (Bopp et al., 1997, 1999; Van Oijen et al., 1998, 1999a,b; Tietz et al., 1999; Ketelaars et al., 2001; Gerken et al., 2003a,b; Hoffman et al., 2003; Rutkauskas et al., 2004, 2005, 2006).The data revealed important details of the exciton structure such as the exciton splitting value, disorder values (for different types of disorder), dipole strengths of the exciton transitions, and the delocalization length. The majority of the steady-state spectroscopic results can be explained within the framework of the exciton theory taking into account the influence of the static disorder. Low temperature polarized FL (Van Mourik et al., 1992) and hole-burning (Reddy et al., 1992) data revealed the presence of the allowed lowest k = 0 level together with intense higher levels broadened due to relaxation. A specific shape of the CD spectra with a red-shift of the zero-crossing (Koolhaas et al., 1997b) reflects a collective (coherent) contribution of the pigments within a large part of a complete ring. The disorder manifests itself through a mixing of the k = ±1 exciton states with the k = 0 state producing a strongly allowed superradiant lowest state observed in FL experiments (Monshouwer et al., 1997). On the other hand, it appeared that including uncorrelated disorder into the exciton model is not sufficient to explain the large splitting between the k = 0 and k = ±1 states observed in hole-burning spectra and consequently correlated disorder was introduced (Wu et al., 1997b). Also, the large splitting between the orthogonally polarized transitions observed in single molecule FL excitation spectra was explained in terms of correlated disorder produced by an elliptical deformation of the ring (Van Oijen et al., 1999b; Ketelaars et al., 2001). The simple exciton model with static disorder (uncorrelated or correlated) also fails to explain the anomalously broad FL profiles observed for LH1/LH2 at low temperature (Freiberg et al., 2003; Timpmann et al., 2004). The data suggested the presence of strong exciton-phonon coupling producing polaron features in the low temperature emission spectra (Freiberg et al., 2003; Timpmann et al., 2004). Including strong exciton-phonon coupling into the modified Redfield theory (see Eqs. 3, 4, and 7) yields more realistic line shapes with reorganization shifts
237
and phonon-induced broadening of the exciton levels in addition to the exciton splitting. These phonon-induced effects, proportional to the PR value according to Eq. (4), are more pronounced for the lowest k = 0 state (especially for strongly disordered realizations giving rise to a localized and red-shifted k = 0 state as seen in Fig. 2). This produces a much broader low-temperature FL profile as compared to the simple exciton model with just static disorder. On the other hand, the disorder of the reorganization shift values produces additional splitting between the exciton states and additional inhomogeneous broadening. Application of the modified Redfield approach allowed a quantitative explanation of the shapes of FL profiles and fluctuations of the FL peak positions observed in a single LH2 complex (Novoderezhkin et al., 2006; Rutkauskas et al., 2006; see Section VI). D. Multiple Exciton Delocalization Sizes The precise character of the excitation within the circular aggregates of the B850-LH2 and LH1 antennae, such as the degree of delocalization, multiple coherence sizes, dynamic localization, etc., has been the subject of intense debate (Reddy et al., 1992; Novoderezhkin and Razjivin, 1993, 1994, 1995a; Bradforth et al., 1995; Dracheva et al., 1995, 1997; Pullerits et al., 1996; Alden et al., 1997; Monshouwer et al., 1997; Nagarajan et al., 1999; Freiberg et al., 2003) and was critically reviewed and summarized in Novoderezhkin et al. (1999b); Meier et al. (1997c); Dahlbom et al. (2001). It should be noted that there exist many different definitions of the degree of delocalization which implies that even for the same system one may find different sizes of an exciton. Delocalization of individual exciton states can be characterized by the inverse participation ratio PR(k), which is different for different states (as illustrated in Fig.2). The effective (averaged over k) delocalization of the exciton wavefunctions Neff can be defined as PR−1(k) weighed with the populations of the exciton states at thermal equilibrium (Meier et al., 1997a). Typically the excited state in the antenna is given by a superposition of exciton states, i.e., an exciton wavepacket is formed whose evolution in time is described by the timedependent density matrix (Mukamel, 1995; Kühn and Sundström, 1997b; Meier et al., 1997a,b,c). The coherence length (or delocalization length) of the wavepacket Ncoh can be defined as the width of antidiagonal distribution of the density matrix in the site
238 representation (Kühn and Sundström, 1997). Modeling of the LH1 and LH2 antenna proteins in the presence of static disorder gave the values Neff = 5–11 (Jimenez et al., 1996; Alden et al., 1997; Meier et al., 1997c; Novoderezhkin et al., 1999) and Ncoh = 4–6 (Kühn and Sundström, 1997b; Novoderezhkin et al., 1999a,b) at room temperature. The dynamic disorder induced by exciton-phonon coupling may give rise to a further reduction of these exciton sizes due to polaron formation (Meier et al., 1997a). Notice that both Neff and Ncoh values reflect some kind of nonuniformity of the shape of the exciton wavefunction, but not a true ‘physical size.’ Generally, spectral responses of a strongly coupled aggregate reflect the cooperative behavior of some number of pigments which can be very different from the effective localization sizes, Neff , Ncoh (Koolhaas et al., 1997b; Meier et al., 1997c; Koolhaas et al., 2000). The number of molecules that contribute to the spectral response can also be different for different spectroscopic techniques. For example, the shape of the CD spectra of the B850 band of the LH2 antenna is determined by cooperativity within at least 10–12 molecules (Koolhaas et al., 1997b; 2000). The shape and amplitude of the transient absorption measured for B850 can only be reproduced by taking into account the exciton coupling between more than 12 molecules of the antenna (Novoderezhkin et al., 1999b). One could define the spectroscopic subunit as the minimal fragment of the ring which is big enough to reproduce all spectral features of the whole antenna (Koolhaas et al., 2000). Due to disorder, the size of such a subunit is less than the aggregate size N, but larger than both effective localization sizes, Neff and Ncoh. E. Superradiance and Anomalously High Nonlinear Responses One of the manifestations of the collective character of excitations in the antenna is an increase in dipole strength of the lowest exciton states (i.e., k = 0 and k = ±1 states as shown in Fig. 2). An anomalously high dipole strength of the main k = ±1 transitions gives rise to an increase of the nonlinear response amplitudes in proportion to N (Novoderezhkin and Razjivin, 1993; 1995b; Leupold et al., 1996; Novoderezhkin et al., 1999b). Pump-probe studies on the core RC-LH1 complex revealed bleaching amplitudes in the antenna that were several times bigger than those corresponding
Rienk van Grondelle and Vladimir I. Novoderezhkin to the bleaching of the special pair due to oxidation of the RC (Novoderezhkin and Razjivin, 1993, 1995b; Kennis et al., 1994; Xiao et al., 1994). In the LH2 complex the bleaching amplitude of the B850 ring was found to be much higher than for the B800 monomeric band (Kennis et al., 1996). Quantitative modeling of the shapes and relative amplitudes of the TA spectra for the LH2 antenna and isolated B820 dimeric subunit (Novoderezhkin et al., 1999b) provided direct evidence for a delocalization of the exciton over many BChls in the B850 ring; corresponding to effective localization sizes for the B850 antenna of Neff = 7.87 and Ncoh = 5.0. The dipole strength of the lowest exciton state (sensitive to the amount of the static disorder) can be determined from the low-temperature transient absorption (TA) (Kennis et al., 1997b) and superradiance (Monshouwer et al., 1997) data. Thus, the integrated intensity of the stimulated emission component in B850 corresponds to the dipole strength of the lowest exciton level of 2.3–3.4 (Kennis et al., 1997b). From the low-temperature superradiance data this value was estimated as 2.8 and 3.8 for the LH2 and LH1 antenna, respectively (Monshouwer et al., 1997). Taken together, these nonlinear spectroscopic results have lead to the disordered exciton model for the LH2-B850 and LH1 antennae (Novoderezhkin et al., 1999a). F. Exciton Transitions Viewed by Polarized Single Molecule Spectroscopy One of the most prominent manifestations of excitonic effects in the LH2-B850 and LH1 antennae is the polarization of the two main transitions (k = ±1) of a circular aggregate. Even in the presence of disorder the largest part of the dipole strength is still concentrated in the original two k = ±1 transitions that have perpendicular polarization in the plane of the ring (Novoderezhkin et al., 1999a). In contrast, localization of the excitation at one site (or on one dimeric BChl subunit) would produce N (or N/2) transitions with different (almost in-plane) polarizations having angles uniformly distributed between 0 and 2π. Such localized features were indeed observed in a single molecule study of the monomeric B800 band of LH2 (Van Oijen et al., 1999b). On the other hand, FL excitation spectra in the 850 nm region (Bopp et al., 1999; van Oijen et al., 1999a) demonstrated the existence of just two perpendicularly polarized transitions in a single LH2 complex, thus giving
Chapter 13
Spectroscopy and Dynamics of Excitation Transfer
direct evidence for pronounced excitonic features in the B850 antenna. Such a confirmation of one of the main predictions of the exciton model was not evident a priori, since in a real pigment-protein complex the ideal exciton picture may easily be destroyed by slow conformational dynamics of the complex inducing quasi-static energetic disorder, and fast phonons producing polaron self-trapping. The anomalously large splitting of the two major orthogonal excitonic transitions observed in the low temperature FL spectra of LH2 was attributed to a modulation of the coupling strength in the B850 ring that was asserted to be associated with an elliptical deformation of the LH2 ring (Bopp et al., 1999; van Oijen et al., 1999a; Mostovoy and Knoester, 2000; Ketelaars et al., 2001; Matsushita et al., 2001) (see also Section VI). III. Equilibration Dynamics The energy transfer dynamics within LH2 includes migration of localized excitations around the BChl 800 outer ring, superimposed on excitation transfer to the exciton states of the BChl 850 inner ring with subsequent equilibration in the B850 exciton manifold and the motion of the quasi-steady-state wavepacket (delocalized over 4–6 BChls) around the B850 inner ring (see Sundström et al., 1999; Renger et al., 2001; Van Grondelle and Novoderezhkin, 2001, 2006 for reviews.). The dynamics within the B850 band was studied by fs FL up-conversion (Jimenez et al., 1996), relative measurements of the induced absorption changes (Novoderezhkin et al., 1999b, Kennis et al., 1996), polarized pump-probe (Nagarajan et al., 1996, 1999; Chachisvilis et al., 1997; Vulto et al., 1999), and photon-echo (Joo et al., 1996; Jimenez et al., 1997). Modeling has included the quantitative fitting of linear and nonlinear responses (Kühn and Sundström, 1997b; Novoderezhkin et al., 1999b, 2003; Brüggemann and May, 2004). The dynamics within the LH1 antenna are very similar to the equilibration dynamics within the B850 band of LH2. But due to its larger size the LH1 antenna is characterized by a more pronounced contribution to the absorption from higher exciton levels (k = ±2 and ±3) and this feature is responsible for the increased amplitude of the dynamic Stokes shift observed in LH1 as compared with LH2 (Monshouwer et al., 1998; Novoderezhkin and Van Gron-
239
delle, 2002). Another specific property of LH1 is the stronger coupling to some low-frequency vibrations, manifested as more pronounced oscillatory features in nonlinear responses due to vibrational coherences (Chachisvilis and Sundström, 1996; Monshouwer et al., 1998; Novoderezhkin et al., 2000). A. Exciton Relaxation and Dynamic Red-Shift The direct observation of excitation dynamics in the tightly packed ring-like aggregates of the LH1 and B850-LH2 antennae (including relaxation of the exciton states and migration of the quasi-steady-state exciton wavepacket) is possible by transient absorption studies via the ps/fs pump-probe technique. Such measurements have revealed the initial ultrafast (sub100 fs) relaxation in LH1/LH2 complexes followed by slower (ps) dynamics (Xiao et al., 1994; Visser et al., 1995, 1996; Nagarajan et al., 1996, 1999; Savikhin and Struve, 1996; Chachisvilis et al., 1997; Kennis et al., 1997a,b; Monshouwer et al., 1998; Vulto et al., 1999; Polívka et al., 2000; Book et al., 2001). Fast relaxation in such a system gives rise to specific TA dynamics typical of assemblies of excitonically coupled antenna pigments, but absent from isolated dimeric subunits and monomers. Thus, Visser et al. (1995) found pronounced TA dynamics for LH1 of Rsp. rubrum with an 8–12 nm red-shift reflecting fast exciton equilibration, whereas no dynamic red-shift was found in the lowest-state absorption band of either the B820 dimeric subunit or for monomeric BChl. Short-pulse (35–50 fs) pump-probe studies of LH1 and LH2 of Rhodobacter (Rba.) sphaeroides (Nagarajan et al., 1996, 1999; Chachisvilis et al., 1997; Vulto et al., 1999) showed relaxation components in the 10–100 fs range; short-lived koff = 103 s–1. In this case, electron transfer occurs upon each binding event. In the diffusion limited case the rate limiting step is not the electron transfer rate but is determined by the formation of the bound state. A different behavior occurs when the dissociation rate is faster than the electron transfer rate, the fast exchange limit (i.e., koff >> ke). In this case electron transfer does not occur upon each binding event and depends upon the electron transfer rate and the occupancy of the bound state. In the fast exchange limit, the second order rate constant becomes k2 ≅ ke/KD, and monophasic rather than biphasic kinetics is observed in flash kinetic experiments (i.e., no fast first order rate is observed). The different features of the kinetics for electron transfer following a single saturating laser flash in the diffusion-limited and fast exchange regimes are shown in Fig. 3. For native RCs for which the reaction with Cyt is diffusion-limited, the reduction of the oxidized primary donor monitored at 865 nm is biphasic (Fig. 3A). The fast phase is due to electron transfer from bound Cyt c2. The time constant for this reaction (unresolved in this trace) is τe = 1µs. The slow phase has decays with a rate that is dependent on the Cyt concentration: k = k2 [Cyt]. The relative amplitudes of the fast and slow phases and the rate of the slow phase depend on the cytochrome concentration. For some mutant RCs having changes to hydrophobic residues the reaction rates and binding
affinity lead to reaction dynamics in the fast exchange limit (Fig. 3B). In the fast exchange limit the donor reduction is monophasic and the rate is dependent on the Cyt c2 concentration, reaching a limit at high Cyt concentration (Gong et al., 2003). A critical parameter in the electron transfer process is the dissociation rate koff. Graige (1998) measured the dissociation rate for Cyt c2 from the RC (Rba. sphaeroides) and found koff = 1.7 × 103 s–1 by using two laser flashes in the presence of Cyt c2 where reduction of the oxidized donor after the second flash was limited by the rate of dissociation of oxidized Cyt. Gerencser et al. (1999) found a dissociation rate for horse heart Cyt c from Rba. sphaeroides RC to be koff =103 s–1 and showed that at low ionic strength the dissociation rate of horse heart Cyt became the rate limiting step for turnover at high light intensities. The dissociation rate of electron transfer found in these studies is fast enough not to be rate limiting in photosynthetic membranes under physiological light conditions (Crofts and Wraight, 1983). To a rough approximation the dissociation rate may be estimated from the dissociation constant and the association rate. At low ionic strength this gives a value of approximately koff = kon × KD = 109 × 10–6 = 103 s–1. However, this is a rough estimate since kon is measured for the association of reduced Cyt, and koff is measured for the dissociation of oxidized Cyt and the dissociation and association rates, and thus the binding constant, can depend on the redox state of the Cyt and D. However, several studies indicate that the differences of binding constants and rates between
Chapter 17
Cytochrome c2 Interactions
oxidized and reduced forms are relatively small. In early studies Rosen (1980) measured the KD values for reduced and oxidized Rba. sphaeroides Cyt c2 to the Rba. sphaeroides RC (10 mM Tris pH 8) to be 1.0 and 1.5 µM, respectively. Larson and Wraight (2000) found a preferential binding of oxidized horse Cyt to Rba. sphaeroides RC compared to the reduced form by a factor of 3. However, experiments by Devanathan et al. (2004) on membrane reconstituted RCs using plasmon-waveguide resonance spectroscopy have found larger differences in binding of oxidized and reduced forms. They find two binding sites, a high affinity and low affinity site, and preferential binding of the reduced Cyt to the high affinity site. In some kinetic studies involving Cyt c2 and RCs, a slow first order component (τ~500 µs) was observed in addition to the slow second order phase (Overfield et al., 1979; Moser and Dutton, 1988). This slow phase has been proposed to be due to the transition from a configuration in which the Cyt is bound in an inactive state to an active configuration. However, the presence of the slow first order phase is variable and not observed in all studies. The amount of the slow phase varies with different preparations (Tiede and Dutton, 1993). Two other explanations for the 500 µs phase have been advanced: 1) dissociation of a fraction of oxidized Cyt, and 2) heterogeneity of the RC preparation due to aggregation. Larson and Wraight (2000) reported a problem of double-hits to RCs excited with xenon flash lamps leading to multiphasic kinetics which may explain results from earlier experiments. The longer illumination times result in the excitation of RCs containing oxidized Cyt giving rise to an additional slow phase where donor reduction is rate limited by the dissociation of oxidized Cyt. Tiede (2000) used neutron diffraction and found that different preparations of isolated RCs are in different states of aggregation. The presence of the slow first order phase was found in aggregated RCs but not in monomeric RCs. He suggested that the slow first order phase is due to aggregation of RCs. The presence of aggregation effects agrees with recent studies which indicate that the apparent KD for Cyt c2 binding increases at higher RC concentrations which could be due to an aggregation of RC that blocks Cyt binding (E. Abresch, unpublished). More recent measurements of the dissociation constant and second order rate constant have been routinely performed at low RC concentrations (~0.1 µM) in order to reduce the effect of aggregation (Tetreault et al., 2001; Gong et al., 2003).
329 In summary, the high second order rate of electron transfer, the rapid first order rate of electron transfer, and the relatively low binding affinity (giving a dissociation rate that is not rate limiting for turnover) are well optimized for the function of the electron transfer cycle through the RC. The electron transfer rates and binding will be discussed in more detail below in relation to the structure of the Cyt:RC complex. V. Effects of Mutation A. Effects on Binding, KD Residues in the interface region between the Cyt and the RC were mutated to determine how changing the interactions between the two proteins affects binding. Tetreault et al. (2001) found that mutation of negatively charged acid residues to neutral or positively charged residues on the RC increased KD. The largest change (~1000 fold) was obtained for the mutation of Asp M184 → Lys. Asp M184 is in the center of a cluster of acid residues and close to Tyr L162 in the center of the interface region. Gong et al. (2003) found that mutation of RC hydrophobic residues Tyr L162 and Leu M191 to Ala and other residues increased KD. The largest increase in KD (~100-fold) due to Ala substitution was observed for the mutation of Tyr L162. This residue is located in the center of the short-range interaction domain (see Fig. 2C). This large change in binding shows that Tyr L162 is a ‘hot spot’ for binding. Similar, ‘hot spots’ have been observed in studies of Ala replacement mutagenesis on protein-protein association (Bogan and Thorn, 1998). Interestingly, the mutation of Val M192 to Ala resulted in an increase in binding affinity. The changes in binding affinity due to Ala mutation were found to correlate with the solvent accessibility of the mutated residues. Tyr L162, whose mutation produced a large decrease in binding affinity, was inaccessible to solvent in the Cyt:RC complex, while Val M192 whose mutation resulted in increased binding affinity was solvated in the Cyt:RC complex. These results support the idea that the hydrophobic binding interaction is due to a closely fitting association between protein residues with the exclusion of solvent. The function of the cation-S interaction between Lys C32 and Tyr M295 in the Cyt:RC complex was studied by mutation of both residues (Paddock et al., 2005). The mutation of Tyr M295 to non-aromatic
Herbert Axelrod, Osamu Miyashita and Melvin Okamura
330
A) Electrostatic
8
B) Hydrophobic
6
10
5
7
ke (s )
10
-1
-1
ke (s )
10
10
4
10
6
10
3
3
10
4
10
5
10
6
7
10
10
8
10
-1
KA=1/KD (M )
10
3
10
4
10
5
10
6
10
7
10
-1
KA=1/KD (M )
Fig. 4. Electron transfer rate ke with Cyt c2 in the bound state vs. the association constant KA for different mutant RCs. A) RCs modified by changes to charged residues (Tetreault et al., 2001). B) RCs modified by changes to hydrophobic residues (Gong et al., 2003).
residues had only a relatively small (3-fold ) effect on KD in contrast to theoretical estimates which predict a large binding interaction for the complex (Gallivan and Dougherty, 1999). The small effect on binding due to the mutation probably results from the energy needed for desolvation of the positively charged Arg C32 in order to form the cation-π complex. This desolvation energy reduces the binding energy for formation of the cation-π bond. Mutation of residues forming hydrogen bonds across the Cyt:RC interface resulted in changes in KD (up to six-fold) indicating that hydrogen bonding interactions stabilize the bound state. (Abresch et al., 2006) As with the case of hydrophobic interactions the magnitude of the change in KD was correlated with solvent accessibility. The residues exhibiting the largest changes in KD showed the lowest solvent accessibility in the Cyt:RC complex. B. Effects on Electron Transfer in the Bound State, ke Residues in the interface region were mutated to determine how structural changes affect electron transfer. The changes to ke were different for mutations of charged residues or mutations of residues having short-range inter-protein contacts. Mutation of charged residues changed the binding affinity KA=1/KD but did not change the electron transfer rate, ke, in the bound state (Fig. 4A) (Tetreault et al., 2001; Tetreault et al., 2002). In contrast, the mutation of hydrophobic residues changed both the binding affinity and electron transfer rate (Fig. 4B) (Gong et al., 2003). These results can be understood
in terms of changes in the electronic coupling interaction between donor and acceptor groups, which is responsible for electron transfer. The electronic coupling interaction decreases exponentially with increasing distance between donor and acceptor and depends on the pathway for electron transfer between donor and acceptor. The mutational results indicate that the pathway for electron transfer is through the short-range interaction domain around Tyr L162 in contact with the heme edge (Fig. 2C). The mutations of charged residues change the binding energy, but do not change the structure of the tunneling contact region in the bound state. Thus, ke is not changed by mutation of charged residues. On the other hand, mutations of residues in the short-range interaction region change the short-range contacts between the electron donor (heme) and acceptor (BChl2). Modifications of these contacts change the electronic coupling responsible for electron transfer. Thus, ke is changed by mutations in the short-range interaction region. An interesting feature of the results of mutations is the strong correlation between the changes in binding affinity KA and in the change in electron transfer rate ke. The correlation suggests that there is a relationship between the interactions responsible for electron transfer and the interactions responsible for binding (Gong et al., 2003). C. Effects on the Second Order Rate Constant, k2 The effects of mutations on the second order rate constant k2 for electron transfer between Cyt c2 and the RC were studied to determine the mechanism of
Cytochrome c2 Interactions
Chapter 17
331
10
10
9
10
k2 (s M )
9
1
1
10
1
1
k2 (s M )
A) Electrostatic
8
10
B) Hydrophobic
8
10
7
10 3
10
4
10
5
6
10
7
10
10
8
10
- -11
K =1/K (M ) A
D
3
10
4
5
10
6
10
10
K =1/K A
D
7
10
-1
(M )
Fig. 5. Second order electron transfer rate constant, k2, vs. association constant KA for reaction between Cyt c2 and mutant RCs. A) RCs modified by changes to charged residues (Tetreault et al., 2001). B) RCs modified by changes to hydrophobic residues (Gong et al., 2003). The triangles indicate reactions displaying single exponential (fast exchange) kinetics.
the reaction. Lin et al. (1994) showed that the driving force dependence of k2 is independent of the driving force for electron transfer. This finding indicates that electron transfer is not the rate limiting step for thesecond order rate. Instead, the association process for the formation of the bound state is the rate limiting step expected for a diffusion limited reaction. (i.e., k2 = kon ). The mutation of charged residues on the RC (Tetreault et al., 2001, 2002) and on the Cyt (Caffrey et al., 1992) changed the second order rate constant, indicating the role of electrostatic interactions in the association process. The changes in k2 due to mutations that modify charge were generally smaller than the changes in KD but were correlated (Fig. 5A). The effects of charge changes on k2 can be understood in terms of transition state theory for a chemical reaction. The transition state is the configuration in which the Cyt is in position to form the bound state of the Cyt: RC complex. Electrostatic interactions stabilize the Cyt in the transition state. However, since the distance between the Cyt and the RC is greater in the transition state than in the bound state, the electrostatic effect due to the Coulomb potential energy (which falls as 1/r ) is smaller for the transition state (affecting k2) than for the bound state (affecting KD). The relative changes in k2 and KD can be accounted for if the Cyt in the transition state is about 10 Å farther away from the RC than in the bound state (Tetreault et al., 2001; Miyashita et al., 2003b). The changes in k2 due to mutations of hydrophobic residues (Gong et al., 2003) showed complex behavior (Fig. 5B). Hydrophobic mutations resulting in small
decreases in KA resulted in only small changes in k2 (i.e., the slope is low on the right side of Fig. 5B). These small changes in k2 indicate that kon is almost unaffected by short-range interactions. The hydrophobic mutations change KD by changing koff. Mutations that produced large decreases in KA resulted in large decreases in k2. (i.e., the slope is higher on the left side of Fig. 5B). The onset of the large decreases in k2 was coincident with the appearance of monophasic rather than biphasic kinetics (triangles in Fig. 5B). This change in kinetics indicates a change from the diffusion limit (ke >> koff) to the fast exchange limit ke koff ), thus allowing the reaction to proceed in the diffusion-limited regime. In addition, the binding interactions are weak enough to allow a dissociation rate (koff ~ 103 s–1) fast enough not to be rate limiting in turnover. This mechanism for electron transfer between Cyt and RC explains the highly optimized diffusion-limited rate constant that is important for the rapid turnover. The role of a specific complex between electron transfer partners may vary for different systems. Several stable protein complexes have been studied in which the cofactors are in position for fast electron transfer as in the case for the Cyt:RC complex. These include complexes between cytochrome c and cytochrome c peroxidase (Pelletier and Kraut, 1992) and cytochrome c and cytochrome bc1 (Hunte et al., 2002).
333 On the other hand, in other systems a stable complex between the two proteins has not been observed. For these systems the reactions most likely occur in the fast-exchange regime by a dynamic docking mechanism (Liang et al., 2004). When a permanent bound state is not observed, the electron transfer can occur from transient formation of a specific complex in which rapid electron transfer occurs. In this case the rapid electron transfer (k2 = ke/KD) would be enhanced by close tunneling contacts between donor and acceptor that increase ke to compensate for a large KD. Such a specific complex may explain the reaction between the Cyt c2 and RC from Rsp. rubrum where the kinetics are strictly monophasic (van der Wal et al., 1987). Although there is no direct evidence for complex formation, a computational study of the binding between Cyt c2 and the RC from Rsp. rubrum shows that a complex similar to that found in Rba. sphaeroides can be formed but with lower binding affinity (Pogorelov et al., 2007). Alternatively, electron transfer may occur from an ensemble of states such as in a non-specific electrostatic encounter complex. In this case, the complex between the two proteins may not be very specific or well optimized for electron transfer as suggested in the ‘Velcro model’ of McClendon (1991) and the electron transfer may proceed through longer distances through water. Studies that indicate that long distance electron transfer through water can occur (Lin et al., 2005; Miyashita et al., 2005). The actual electron transfer mechanism for these systems depends upon the relative rates and occupancies at different docking positions. Further studies of the electron transfer are needed to establish the mechanisms that form the molecular basis for these critical biological processes. Acknowledgments We wish to thank the many colleagues who contributed to the work from our group on the cytochrome: RC complex reported here. Special thanks to George Feher, Mark Paddock, Ed Abresch and Charlene Chang for contributions to ongoing research and to José Onuchic for theoretical discussions. Work supported by NIH grant GM 41637. References Abresch EC, Villalobos M, Paddock ML, Chang C and Okamura MY (2006) The importance of buried H-bonds on binding and
334
Herbert Axelrod, Osamu Miyashita and Melvin Okamura
electron transfer in the cytochrome c2:reaction center complex. Biophysical Society Meeting Abstracts, Biophys J, Supplement, 503a, Abstract, 2400 Adir N, Axelrod H, Beroza P, Isaacson R, Rongey S, Okamura M and Feher G (1996) Co-crystallization and characterization of the photosynthetic reaction center-cytochrome c2 complex from Rhodobacter sphaeroides. Biochemistry 35: 2535–2547 Allen JP, Feher G, Yeates TO, Rees DC, Deisenhofer J, Michel H and Huber R (1986). Structural homology of reaction centers from Rhodopseudomonas sphaeroides and Rhodopseudomonas viridis as determined by x-ray diffraction. Proc Natl Acad Sci USA 83: 8589–8593 Allen JP, Feher G, Yeates TO, Komiya H and Rees DC (1987) Structure of the reaction center from Rhodobacter sphaeroides R-26: The protein subunits. Proc Natl Acad Sci USA 84: 6162–6166 Aquino A, Beroza P, Reagan J and Onuchic J (1997) Estimating the effect of protein dynamics on electron transfer to the special pair in the photosynthetic reaction center. Chem Phys Lett 275: 181–187 Autenrieth F, Tajkhorshid E, Schulten K and Luthey-Schulten Z (2004) Role of water in transient cytochrome c2 docking. J Phys Chem B 108: 20376–20387 Axelrod H and Okamura M (2005) The structure and function of the cytochrome c2:reaction center electron transfer complex from Rhodobacter sphaeroides. Photosynth Res 85: 101–114 Axelrod H, Feher G, Allen J, Chirino A, Day M, Hsu B and Rees D (1994) Crystallization and X-ray structure determination of cytochrome c2 from Rhodobacter sphaeroides in three crystal forms. Acta Cryst D: Biol Crystallogr 50: 596–602 Axelrod HL, Abresch EC, Okamura MY, Yeh AP, Rees DC and Feher G (2002) X-ray structure determination of the cytochrome c2:reaction center electron transfer complex from Rhodobacter sphaeroides. J Mol Biol 319: 501–515 Bendall D (1996) Interprotein electron transfer. In: Bendall D (ed) Protein Electron Transfer, pp 43–68. Bios Scientific Publishers Ltd, Oxford Beratan D, Betts J and Onuchic J (1991) Protein electron transfer rates set by the bridging secondary and tertiary structure. Science 252: 1285–1288 Blankenship RE (2002) Molecular Mechanisms of Photosynthesis. Blackwell Science, London Bogan A and Thorn K (1998) Anatomy of hot spots in protein interfaces. J Mol Biol 280: 1–9 Caffrey MS, Bartsch RG and Cusanovich MA (1992) Study of the cytochrome c2-reaction center interaction by site-directed mutagenesis. J Biol Chem 267: 6317–6321 Chance B and Smith L (1955) Respiratory pigments of Rhodospirillum rubrum. Nature 175: 803–806 Chang C-H, Tiede D, Tang J, Smith U, Norris J and Schiffer M (1986) Structure of Rhodopseudomonas sphaeroides R-26 reaction center. FEBS Lett 205: 82–86 Crofts AR and Wraight CA (1983) The electrochemical domain of photosynthesis. Biochim Biophys Acta 726: 149–185 Deisenhofer J and Michel H (1989) The photosynthetic reaction center from the purple bacterium Rhodopseudomonas viridis. EMBO J 8: 2149–2170 Devanathan S, Salamon Z, Tollin G, Fitch J, Meyer T and Cusanovich M (2004) Binding of oxidized and reduced cytochrome c2 to photosynthetic reaction centers: Plasmon-waveguide resonance
spectroscopy. Biochemistry 43: 16405–16415 Devault D (1980) Quantum Mechanical Tunneling in Biological Systems. Q Rev Biophys 13: 387–564 Drepper F, Dorlet P and Mathis P (1997) Cross-linked electron transfer complex between cytochrome c2 and the photosynthetic reaction center of Rhodobacter sphaeroides. Biochemistry 36: 1418–1427 Dutton PL and Prince RC (1978) Reaction center driven cytochrome interactions. In: Clayton RK and Sistrom WR (eds) The Photosynthetic Bacteria, pp 525–570. Plenum Press, New York Duysens L (1952) Transfer of excitation energy in photosynthesis. Doctoral dissertation. Utrecht University, Utrecht, The Netherlands Duysens L (1954) Reversible photo-oxidation of a cytochrome pigment in photosynthesizing Rhodospirillum rubrum. Nature 173: 692–693 Ermler U, Fritsch G, Buchanan S and Michel H (1994) Structure of the photosynthetic reaction center from Rhodobacter sphaeroides at 2.65 Å resolution — Cofactors and protein cofactor interactions. Structure 2: 925–936 Hopfield JJ (1974) Electron transfer between biological molecules by thermally activated tunneling. Proc Natl Acad Sci USA 71: 3640–3645 Gallivan JP and Dougherty DA (1999) Cation-π interactions in structural biology. Proc Natl Acad Sci USA 96: 9459–9464 Gerencsér L, Laczkó G and Maróti P (1999) Unbinding of oxidized cytochrome c from photosynthetic reaction center of Rhodobacter sphaeroides is the bottleneck of fast turnover. Biochemistry 38: 16866–16875 Gong X, Paddock M and Okamura M (2003) Interactions between cytochrome c2 and photosynthetic reaction center from Rhodobacter sphaeroides: Changes in binding affinity and electron transfer rate due to mutation of interfacial hydrophobic residues are strongly correlated. Biochemistry 42: 14492–14500 Graige M, Feher G and Okamura M (1998) Conformational gating of the electron transfer reaction QA–.QB → QAQB–. in bacterial reaction centers of Rhodobacter sphaeroides determined by a driving force assay. Proc Natl Acad Sci USA 95: 11679–11684 Hunte C, Solmaz S and Lange C (2002) Electron transfer between yeast cytochrome bc1 complex and cytochrome c: A structural analysis. Biochim Biophys Acta 1555: 21–28 Ke B, Chaney TH and Reed DW (1970) The electrostatic interaction between the reaction center bacteriochlorophyll derived from Rhodopseudomonas sphaeroides and mammalian cytochrome c and its effects on light activated electron transport. Biochim Biophys Acta 216: 373–383 Kurlancheek W and Cave RJ (2006) Tunneling through weak interactions: A comparison of through-space, H-bond, and throughbond mediated tunneling. J Phys Chem 110: 14018–14028 Larson J and Wraight C (2000) Preferential binding of equine ferricytochrome c to the bacterial photosynthetic reaction center from Rhodobacter sphaeroides. Biochemistry 39: 14822–14830 Liang ZX, Kurnikov IV, Nocek JM, Mauk AG, Beratan DN and Hoffman BM (2004) Dynamic docking and electron-transfer between cytochrome b5 and a suite of myoglobin surface-charge mutants. Introduction of a functional-docking algorithm for protein-protein complexes. J Am Chem Soc 126: 2785–2798
Chapter 17
Cytochrome c2 Interactions
Lin J, Balabin I and Beratan D (2005) The nature of aqueous tunneling pathways between electron-transfer proteins. Science 310: 1311–1313 Lin X, Williams JC, Allen J and Mathis P (1994) Relationship between rate and free energy difference for electron transfer from cytochrome c2 to the reaction center in Rhodobacter sphaeroides. Biochemistry 33: 13517–13523 Long J, Durham B, Okamura M and Millett F (1989) Role of specific lysine residues in binding cytochrome c2 to the Rhodobacter sphaeroides reaction center in optimal orientation for rapid electron transfer. Biochemistry 28: 6970–6974 Marcus RA (1964) Chemical and electrochemical electron transfer theory. Ann Rev Phys Chem 15: 155–196 McLendon G (1991) Control of biological electron transport via molecular recognition and binding: The ‘Velcro’ model. Struct Bond 75: 160–174 Meyer T and Donohue T (1995) Cytochromes, Iron-sulfur, and copper proteins mediating electron transfer from the Cyt bc1 complex to photosynthetic reaction center complexes. In: Blankenship RE, Madigan MT and Bauer CE (eds) Anoxygenic Photosynthetic Bacteria (Advances in Photosynthesis and Respiration, Vol 2), pp 725–745. Kluwer Academic Publishers, Dordrecht Miyashita O, Okamura MY and Onuchic JN (2003a) Theoretical understanding of the interprotein electron transfer between cytochrome c2 and the photosynthetic reaction center. J Phys Chem B 107: 1230–1241 Miyashita O, Onuchic JN and Okamura MY (2003b) Continuum electrostatic model for the binding of cytochrome c2 to the photosynthetic reaction center from Rhodobacter sphaeroides. Biochemistry 42: 11651–11660 Miyashita O, Onuchic J and Okamura M (2004) Transition state and encounter complex for fast association of cytochrome c2with bacterial reaction center. Proc Natl Acad Sci USA 101: 16174–16179 Miyashita O, Okamura and Onuchic J (2005) Interprotein electron transfer from cytochrome c2 to photosynthetic reaction center: Tunneling across an aqueous interface. Proc Natl Acad Sci USA 102: 3558–3563 Moser C and Dutton PL (1988). Cytochrome c and c2 binding dynamics and electron transfer with photosynthetic reaction center protein and other integral membrane redox proteins. Biochemistry 27: 2450–2461 Moser C and Dutton P (1992) Engineering protein structure for electron transfer in photosynthetic reaction centers. Biochim Biophys Acta 1101: 171–176 Nitschke W and Dracheva SM (1995) Reaction center associated cytochromes. In: Blankenship RE, Madigan MT and Bauer CE (eds) Anoxygenic Photosynthetic Bacteria (Advances in Photosynthesis and Respiration), pp 775–805. Kluwer Academic Publishers, Dordrecht Nogi T, Hirano Y and Miki K (2005) Structural and functional studies on the tetraheme cytochrome subunit and its electron donor proteins: The possible docking mechanisms during the electron transfer reaction. Photosynth Res. 85: 87–99 Overfield RE, Wraight CA and Devault DC (1979) Microsecond photooxidation kinetics of cytochrome c2 from Rhodopseudomonas sphaeroides: In vivo and solution studies. FEBS Lett. 105: 137–142 Paddock M, Weber K, Chang C and Okamura M (2005) Interac-
335 tions between cytochrome c2 and the photosynthetic reaction center from Rhodobacter sphaeroides: The cation-π interaction. Biochemistry 44: 9619–9625 Parson W (1968) The role of P870 in bacterial photosynthesis. Biochim Biophys Acta 153: 248–259 Pelletier H and Kraut J (1992) Crystal structure of a complex between electron transfer partners, cytochrome c peroxidase and cytochrome c. Science 258: 1748–1755 Pogorelov T, Autenrieth F, Roberts E and Luthey-Schulten Z (2007) Cytochrome c2 exit strategy: Dissociation studies and evolutionary implications. J Phys Chem B 111: 618–634 Prince RC, Cogdell RJ and Crofts AR (1974) The photo-oxidation of horse heart cytochrome c and native cytochrome c2 by reaction centres from Rhodopseudomonas spheroides R-26. Biochim Biophys Acta 347: 1–13 Prytkova TR, Kurnikov IV and Beratan DN (2005) Ab initio based calculations of electron-transfer rates in metalloproteins. J Phys Chem B 109: 1618–1625 Rosen D, Okamura MY and G Feher (1980) Interaction of cytochrome c with reaction centers of Rhodopseudomonas sphaeroides R-26: Determination of number of binding sites and dissociation constants by equilibrium dialysis. Biochemistry 19: 5687–5692 Rosen D, Okamura MY, Abresch EC, Valkirs GE and Feher G (1983) Interaction of cytochrome c with reaction centers of Rhodopseudomonas sphaeroides R-26: Localization of the binding site by chemical cross-linking and immunochemical studies. Biochemistry 22: 335–341 Tetreault M, Rongey SH, Feher G and Okamura M (2001) Interaction between cytochrome c2 and the photosynthetic reaction center from Rhodobacter sphaeroides: Effects of charge-modifying mutations on binding and electron transfer. Biochemistry 40: 8452–8462 Tetreault M, Cusanovich M, Meyer T, Axelrod H and Okamura M (2002) Double mutant studies identify electrostatic interactions that are important for docking cytochrome c2 onto the bacterial reaction center. Biochemistry 41: 5807–5815 Tiede D (1987) Cytochrome c orientation in electron transfer complexes with photosynthetic reaction centers of Rhodobacter sphaeroides and when bound to the surface of negatively charged membranes: Characterization by optical linear dichroism. Biochemistry 26: 397–410 Tiede DM and Chang CH (1988) The cytochrome-c binding surface of reaction centers from Rhodobacter sphaeroides. Isr J Chem 28: 183–191 Tiede D and Dutton P (1993) Electron transfer between bacterial reaction centers and mobile c-type cytochromes. In: Deisenhofer J and Norris J (eds) The Photosynthetic Reaction Center, Vol 1, pp 258–288. Academic Press, San Diego Tiede D, Vashishta A and Gunner M (1993) Electron-transfer kinetics and electrostatic properties of the Rhodobacter sphaeroides reaction center and soluble c-cytochromes. Biochemistry 32: 4515–4531 Tiede DM, Littrell K, Marone PA, Zhang R and Thiyagarajan P (2000) Solution structure of a biological bimolecular electron transfer complex: characterization of the photosynthetic reaction center-cytochrome c2 protein complex by small angle neutron scattering. J Appl Cryst 33: 560–564 van der Wal H, van Grondelle R, Millett F and Knaff D (1987) Oxidation of cytochrome c2 and of cytochrome c by reaction cen-
336
Herbert Axelrod, Osamu Miyashita and Melvin Okamura
ters of Rhodospirillum rubrum and Rhodobacter sphaeroides. The effect of ionic strength and of lysine modification on oxidation rates. Biochim Biophys Acta 893: 490–498 Vernon L and Kamen M (1953) Studies on the metabolism of
photosynthetic bacteria. XV. Photoautoxidation of ferrocytochrome c in extracts of Rhodospirillum rubrum. Arch Biochem Biophys 44: 298–311
Chapter 18 Directed Modification of Reaction Centers from Purple Bacteria JoAnn C. Williams and James P. Allen* Department of Chemistry and Biochemistry and Center for Bioenergy & Photosynthesis, Arizona State University, Tempe AZ 85287-1604, U.S.A.
Summary ................................................................................................................................................................... 1 I. Introduction......................................................................................................................................................... 2 II. Properties of the Cofactors................................................................................................................................. 2 A. Identity, Substitution, and Removal ...................................................................................................... 2 B. Optical Spectra ..................................................................................................................................... 3 C. Oxidation/Reduction Midpoint Potentials ............................................................................................. 4 D. Modeling the Electronic Structure of the Bacteriochlorophyll Dimer .................................................... 6 III. Electron Transfer Concepts................................................................................................................................ 7 A. Energetics ............................................................................................................................................ 7 B. Coupling ............................................................................................................................................... 9 C. Dynamics ............................................................................................................................................. 9 IV. Pathways of Electron Transfer ......................................................................................................................... 10 A. B-side Electron Transfer..................................................................................................................... 10 B. New Electron Transfer Reactions....................................................................................................... 12 V. Conclusions ...................................................................................................................................................... 13 Acknowledgments ................................................................................................................................................... 13 References .............................................................................................................................................................. 13
Summary Reaction centers from purple bacteria form a superb test system for the manipulation of electron transfer parameters. The wealth of cofactors and electron transfer reactions provides opportunities for directed modification of specific properties. In particular, the energies of each cofactor can be selectively changed by mutations of neighboring amino acid residues. The starting point for the initial electron transfer, the bacteriochlorophyll dimer, has proven to be exceptionally malleable, allowing large changes in energetics and rates. Most of the other cofactors can be exchanged or eliminated entirely, permitting considerable alteration of pathways. By orchestrating multiple changes in the reaction center, the light-initiated electron transfer pathway can be directed towards alternate ends, for example down the B branch of cofactors rather than the naturally preferred A branch. Extensive modeling of features of electron transfer such as the energetics, the coupling, and the protein dynamics has been corroborated by observed changes in the characteristics of the reactions after modification of the cofactor properties. For example, the maximum rates for several electron transfer reactions, determined by application of Marcus theory to the rates of reactions in a range of mutants, show a correlation with the
*Author for correspondence, email:
[email protected] C. Neil Hunter, Fevzi Daldal, Marion C. Thurnauer and J. Thomas Beatty (eds): The Purple Phototrophic Bacteria, pp. 337–353. © 2009 Springer Science + Business Media B.V.
338
JoAnn C. Williams and James P. Allen
distance between the cofactors. Other measurements revealing the intimate interaction of the protein and cofactors show that protein motion controls the rate of the initial electron transfer. Thus the reaction center provides a natural and modifiable template for understanding the factors governing electron transfer. I. Introduction What happens when the reaction center is excited by light? The light energy is converted into chemical energy through a series of electron and proton transfer reactions involving the cofactors of the reaction center. These reactions are able to proceed with essentially every light photon producing useful reactions, corresponding to a quantum efficiency of nearly 100%. The balancing act of capturing light energy while not destroying the molecules involved or producing unfavorable side reactions is achieved by fine tuning the properties of the cofactors through interactions with the protein in which they are embedded. The protein can influence several aspects of electron transfer identified by theoretical treatments, including the energetics, the coupling, and the protein dynamics. Although it is difficult to separate the contributions of each of these components, experiments have probed their effects, notably by altering the light-induced reactions in specific ways. This chapter will review the partnership between the cofactors and the protein scaffold as it relates to the parameters of electron transfer models, focusing on examples where changes in the properties have measurable effects on electron transfer. The examples will be primarily of reaction centers from Rhodobacter (Rba.) sphaeroides and Rba. capsulatus, the two commonly studied reaction center systems of purple bacteria. II. Properties of the Cofactors A. Identity, Substitution, and Removal The reaction center from Rba. sphaeroides and Rba. capsulatus is an integral membrane protein complex composed of three proteins: the L, M, and H subunits. The L and M subunits form the core of the protein and are largely composed of five transmembrane helices Abbreviations: BA – bacteriochlorophyll monomer on A branch of cofactors; BB – bacteriochlorophyll monomer on B branch of cofactors; HA – bacteriopheophytin on A branch of cofactors; HB – bacteriopheophytin on B branch of cofactors; P – bacteriochlorophyll dimer; QA – quinone on A branch of cofactors; QB – quinone on B branch of cofactors; Rba. – Rhodobacter
that are structurally related to each other by an approximate two-fold symmetry axis. The H subunit is more peripheral, containing one transmembrane helix and a large cytoplasmic domain. Embedded in the middle of the L and M subunits are ten cofactors, all of which can participate in some manner in energy or electron transfer. The ten cofactors in the reaction center are: two bacteriochlorophyll a molecules that form a dimer (P), two bacteriochlorophyll a monomers (BA and BB), two bacteriopheophytin a molecules (HA and HB), two ubiquinone molecules (QA and QB), a carotenoid molecule, and an iron (Fig. 1). These cofactors are arranged into two branches, identified as the A and B branches, which are related by the same two-fold symmetry as found for the L and M subunits. Although the cofactors are normally expressed with a well-defined composition, some of these cofactors can be substituted by molecules in the same class, for example bacteriochlorophylls for bacteriopheophytins. The monomer bacteriochlorophylls and bacteriopheophytins, the quinones, the carotenoid, and the iron can be biochemically removed and replaced. Mutagenesis can also result in biosynthetic substitutions, primarily by replacement of the amino acid residues coordinating the cofactors. For example, when the ligand to one of the central Mg atoms of P is changed, a bacteriopheophytin is incorporated rather than bacteriochlorophyll in a mutant that has been termed a heterodimer (His L173 to Leu and His M202 to Leu in Rba. sphaeroides, L173 and M200 in Rba. capsulatus) (Bylina and Youvan, 1988; Kirmaier et al., 1988; McDowell et al., 1991; Allen et al., 1996; van Brederode et al., 1999; King et al., 2001). Both halves of the dimer can be individually replaced this way, although the double bacteriopheophytin dimer appears to be unstable. Similarly the binding site for the B-side bacteriochlorophyll monomer is found to contain a bacteriopheophytin when the residue forming its Mg ligand is changed (φ mutant, His M182 to Leu in Rba. sphaeroides), although analogous mutations on the A side do not appear to have the same substitution effect (Katilius et al., 1999, 2004). Conversely, the bacteriopheophytin on the A side can be converted to bacteriochlorophyll by introduction of a residue to act as a ligand (β mutant, Leu M214
Chapter 18
Reaction Center Alteration
Fig. 1. Three-dimensional structure of the cofactors of the reaction center from Rba. sphaeroides R-26. Shown are the bacteriochlorophyll a dimer (P) (shaded dark), the two bacteriochlorophyll a monomers (BA and BB), the two bacteriopheophytin a molecules (HA and HB), the two ubiquinone molecules (QA and QB), and the iron (Fe). Although the carotenoid is present in wild type, it is not present in the R-26 strain. The view is perpendicular to the approximate two-fold symmetry axis that passes from P to Fe in the plane of the paper.
to His in Rba. sphaeroides, M212 in Rba. capsulatus) (Kirmaier et al., 1991). The addition of a histidine near the bacteriopheophytin on the B branch also results in incorporation of a bacteriochlorophyll (Leu L185 to His in Rba. sphaeroides) (Watson et al., 2005). Changing iron ligands results in a loss of metal specificity, and in one case a significant amount of zinc is incorporated (His M266 to Cys in Rba. sphaeroides) (Williams et al., 2007). In addition to altering the cofactor composition, some of the cofactors can be removed or their incorporation can be blocked. Many studies of reaction centers have been performed on a carotenoid-less strain of Rba. sphaeroides, identified as R-26, which shows properties essentially identical to the carotenoid-containing wild type except for the loss of the ability to trap excess energy. The quinones can be taken out by exposing the reaction centers to a detergent treatment, which initially results in a decrease in QB followed by loss of QA. Biosynthetic incorporation of QA can be blocked by substitution of amino acid residues forming the binding pocket,
339 for example by removing the tryptophan in van der Waals contact with QA (M252 in Rba. sphaeroides and M250 in Rba. capsulatus), or adding a tryptophan in place of a smaller residue (Ala M260 to Trp in Rba. sphaeroides) (Breton et al., 2004). Certain mutations near P result in reaction centers that lack a functional P (Val L157 to Arg, His L153 to Glu, Leu, Gln, or Tyr, His L173 to Gly, and His M202 to Gly in Rba. sphaeroides) (Jackson et al., 1997; Moore and Boxer, 1998; Katilius et al., 2004). The loss of HA is one outcome of the large-scale alterations of the DLL mutant, in which the D transmembrane sequence of the M subunit is replaced with the symmetry-related segment of the L subunit (M192 to M217 replaced with L165 to L190 in Rba. capsulatus) (Robles et al., 1990). Similarly, the B-branch bacteriopheophytin is not required for assembly of the reaction center as shown by a mutant with the change of an alanine that is adjacent to HB to tryptophan (M149 in Rba. sphaeroides)(Watson et al., 2005). A loss of bacteriochlorophyll (presumably in the dimer) was also reported to be due to structural and electrostatic changes in a residue located between BB and P (Ile L177 to His in Rba. sphaeroides) (Khatypov et al., 2005). See Chapter 16, Jones, for a summary of the effects of exclusion and replacement of reaction center cofactors. The ability to alter the cofactor composition provides the opportunity to manipulate the electron transfer reactions as discussed below. B. Optical Spectra One of the most accessible properties of the cofactors is the absorption spectrum (Fig. 2). The tetrapyrrole pigments (P, BA, BB, HA, and HB) have absorption peaks in the near-infrared region, the visible region and the UV region, and the quinones have an unresolved band in the visible region. The bacteriopheophytins, monomer bacteriochlorophylls, and dimer bacteriochlorophylls can be distinguished from each other in the near-infrared peaks at 760 nm, 800 nm, and 865 nm, respectively. In the visible region, the 540 nm peak arises from the bacteriopheophytins, and the 590 nm peak is from all four bacteriochlorophylls. The A and B branch pigments of the same type overlap, except at low temperature where the broad peak in the 540 nm region of the spectrum is resolved into two peaks at 533 nm and 546 nm associated with HB and HA, respectively. The contributions of the tetrapyrrole pigments in the Soret region have been delineated, with H contributing primarily on the
340
Fig. 2. Absorption spectrum of reaction centers from wild-type Rba. sphaeroides. The primary contributions of the bacteriochlorophyll dimer (P), bacteriochlorophyll monomers (B), and bacteriopheophytin monomers (H) to each of the absorption bands are identified. The Soret band arises from all the tetrapyrrole cofactors.
blue side and P absorbing on the red side (Wang et al., 2006). The carotenoid has a major but poorly resolved contribution to the absorption near 500 nm, where absorption of the other cofactors is weak, and can transfer excitation energy to the bacteriochlorophylls, performing a light-harvesting function in addition to photoprotection (Lin et al., 2003). The absorption peaks in the near-infrared region are shifted from the solution spectra, notably in the shift of the bacteriochlorophyll bands to longer wavelengths, presumably because of their protein environment. However, specific alteration of the spectrum generally has not been amenable to mutagenesis. Modeling of the excited states has also proven difficult (Dahlbom and Reimers, 2005). However in certain instances, shifts in the spectrum are attributed to changing particular residues. A notable example is the change in the visible region peak of the bacteriopheophytins due to changing a hydrogen bond to the keto group of HA, which established the assignment of these optical bands to the individual bacteriopheophytins (Bylina et al., 1988). However, changing a hydrogen bond is not typically correlated with a shift in the peak of the tetrapyrrole pigments. Another relatively malleable absorption peak is that of the dimer. It can shift up to approximately 15 nm to shorter wavelengths as a result of mutations, mostly mutations in which a hydrogen bond to the acetyl group is changed, precipitating a rotation of this side group. Major changes in the absorption spectra also occur when substitutions
JoAnn C. Williams and James P. Allen of the cofactors are introduced. For example, the nearinfrared absorption peak of the dimer is significantly different in the heterodimer mutant. Likewise, shifts in the both the visible and near-infrared regions of the spectra are observed in mutants with alterations of ligands to the monomer bacteriochlorophylls that result in pigment changes (Katilius et al., 1999, 2004). Thus, the optical spectrum is a sensitive indicator of the effects of certain types of modifications to the reaction center. The absorption bands in the near-infrared region arise from transitions from the ground state to the first excited state and so are markers of the excited state energy, indicating the maximum amount of energy that can be captured. For the primary donor of Rba. sphaeroides, the absorption peak at 865 nm corresponds to an energy difference of 1.4 eV. The properties of the reaction center can also be characterized from measurement of the spontaneous and stimulated emission of the excited state of the dimer, centered near 915 nm. Because the absorption and emission bands change as the cofactors undergo excitation, oxidation, and reduction, transient optical spectroscopy is one of the major techniques utilized to follow the light-induced transfer of electrons in the reaction center. C. Oxidation/Reduction Midpoint Potentials The oxidation/reduction midpoint potentials of the reaction center cofactors are critical properties for their function as electron transfer components. The midpoint potential of the dimer, at approximately 500 mV, is the only one easily measured directly (Fig. 3). The midpoint potentials of the other cofactors can only be inferred. A change in the chemical nature of a cofactor has a direct effect on its midpoint potential. For example, incorporation of a bacteriopheophytin in the heterodimer mutant increases the potential by approximately 130 mV due to the intrinsically higher potential of bacteriopheophytin (Allen et al., 1996). In addition, the energies of the electronic states are sensitive to the environment so protein interactions with the dimer, including hydrogen bonds and electrostatic forces from charged residues, can affect the dimer midpoint potential and be modulated by mutagenesis. The midpoint potentials of other tetrapyrroles in the reaction center can presumably also be changed by similar modifications, although the evidence is based upon alterations of the electron transfer rates rather than direct measurements.
Chapter 18
Reaction Center Alteration
341
Fig. 3. (Left) The near-infrared region of the absorption spectrum showing a systematic decrease in the band associated with the bacteriochlorophyll dimer as the ambient potential is increased. The absorbance of this band is used to determine the fraction of P+ at any given potential compared to the total amount of P. (Right) Fits of the dependence of the fraction of P+ on the ambient potential are used to determine the oxidation-reduction midpoint potential, Em. A shift in the midpoint potential is observed for a number of different mutants (Allen and Williams, 1995).
Whereas some modifications lead to large changes, analysis of various mutations indicates that most result in small (4.4 Å) and point away from His M219. This represents a striking departure from the overall
Chapter 20
Acceptor Quinones of Purple Photosynthetic Bacteria
389
Fig. 5. View down the two-fold rotational symmetry axis, showing that the methoxy groups (C2M and C3M) of QA (left) and QB (right) violate the symmetry that is followed by the isoprene tail and the major features of the reaction center organization (cofactor positions and architecture of the L and M subunits).
C2 symmetry relating the two sites (Fig. 5). C. Hydrogen Bonding in the Quinone Binding Sites There is a variety of evidence for the importance of hydrogen bonding of the two quinones of the RC. The carbonyls and the methoxy oxygens are both nominally capable of functioning as hydrogen bond acceptors, and hydrogen bonds to the carbonyls, in particular, are expected to affect the affinity and specificity of QA and QB binding and to modify their physico-chemical properties to be sufficient for the reduction of quinone to quinol. In general, however, the resolution of the X-ray structures is too low to allow unequivocal assignment of the interactions between the quinones and their binding sites, and the energetics and specific properties of, e.g., hydrogen bonds, must be determined from spectroscopy and computation. The crystal structures, measurements of affinity of modified quinones, electron paramagnetic resonance (EPR), electron nuclear double resonance (ENDOR), solid state nuclear magnetic resonance (NMR) and Fourier transfer infra-red (FTIR) spectroscopy all offer complementary pictures of the quinone interactions. Fruitful comparisons might also be expected with an increasing selection of approximately 30 structurally defined quinone and 10 inhibitor binding sites in other proteins (Fisher and Rich, 2000), including cytochromes bc1 and b6 f, succinate dehydrogenase and fumarate reductase, as well as the binding of inhibitors as quinone or quinol analogs such as stigmatellin in RCs and cytochromes bc1 and b6 f complexes. However, the crystal structures residing in the protein databank in March 2007 reveal a very wide variation of hydrogen bond partners to quinone carbonyls, with no evident motifs defining a canonical quinone binding site. Thus, the quinone site architecture found in RCs is not replicated in
any other known quinone binding sites and useful correlations will require considerable sophistication, if they exist at all. IV. Spectroscopy of the Acceptor Quinones Vibrational spectroscopy provides an independent source of data on the quinone binding interactions, and FTIR spectroscopy in particular has provided significant insights, while magnetic resonance methods have been applied with great effect to the semiquinones. An advantage of IR absorbance and Raman over magnetic resonance spectroscopies is that the former can see all redox states of the quinones, not just the radical species. The use of light-induced (or redox potential-induced) difference spectra has allowed excellent resolution of the difference between quinone and semiquinone states in RCs, against the much greater background of mostly unchanging protein vibrations (for reviews, see Breton and Nabedryk, 1996; Nabedryk and Breton, 2008). The theoretical and experimental bases for the IR spectra of ubiquinones, in solution, have been systematically explored by Breton and Nonella and coworkers (Nonella and Brändli, 1996; Burie et al., 1997; Nonella, 1997, 1998; Nonella et al., 2003), and others (Boesch and Wheeler, 1997; O’Malley, 2001; Wheeler, 2001). The IR absorbance spectrum of isoprenyl ubiquinone (UQ-n) in solution, at room temperature, is characterized by a C=C stretch at 1610 cm–1 and two C=O stretches at 1665 and 1650 cm–1. The C=C and C=O modes are substantially coupled to each other but those of the two carbonyls are not. The splitting indicates asymmetry of substitution, which, for ubiquinone, arises from the conformation of the two methoxy groups, since an in-plane methoxy donates electrons into the ring while those out of plane are electron withdrawing (Burie et al.,
390 1997; Meyerson, 1985). The lone pair electrons of an in-plane methoxy oxygen are conjugated to the quinone C=C—C=O π electron system, decreasing the bond order, downshifting the distal C=O stretch frequency and upshifting the proximal C=O (Nonella and Brändli, 1996; Burie et al., 1997; Boullais et al., 1998; Nonella et al., 2003). The in-plane conformation is favored by the stabilizing electron resonance but is countered by unfavorable van der Waals interactions with any adjacent substituents.3 The steric clash is diminished by the larger bond angle arising from increased sp2 character with hydridization of the inplane methoxy oxygen. There is only room for one of the adjacent methoxy groups in UQ to be in-plane at a time, so the two carbonyls are not equivalent. However, both carbonyls contribute equally to both C=O stretch frequencies since neither methoxy has precedence in solution (Burie et al., 1997; Boullais et al., 1998; Remy et al., 2003). The quinone C=O and C=C bands are generally not very sensitive to changing from a protic (hydrogen bonding) to an aprotic solvent (Bauscher and Mäntele, 1992; Burie et al., 1995). This reflects the relatively weak hydrogen bonding propensity of β-unsaturated carbonyls (Rasmussen et al., 1949; Bellamy, 1968; 1975). Extensive IR studies have been made of the quinones in bacterial RCs (reviewed in Breton and Nabedryk, 1996; Boullais et al., 1998). The two C=O vibrations are very weakly coupled, if at all, indicating at least as much conformational and environmental asymmetry as is found in solution. Also as in solution, both carbonyls are substantially mixed with the C=C stretch. Assignment of the dominant C1=O, C4=O and C2=C3 and C5=C6 stretch bands has been made through the use of site-specific isotopic labels (Breton et al., 1994a, 1995; Brudler et al., 1994, 1995). A. The Primary Quinone, QA For QA, the quinone mode dominated by C1=O, associated with a hydrogen bond to the backbone of Ala M260, is found at 1660 cm–1, close to the average of the two solution values. A band at 1628 cm–1 is identified as having major C=C character, signifi3 Both the methyl and the non-conjugated lone pair electrons of the methoxy oxygen interact with adjacent groups — the proximal carbonyl oxygen and the other methoxy group. In the absence of a neighboring substituent, the preferred in-plane conformation is with the methoxy methyl group pointing away from the carbonyl. However, even a methyl in the adjacent position hinders this conformer significantly more than the other in-plane alignment, which is therefore preferred.
Colin A. Wraight and Marilyn R. Gunner cantly up-shifted from solution (1610 cm–1) but still recognizable. However, the C4=O stretch, hydrogen bonded to His M219, has been convincingly assigned to a band at 1601 cm–1, which is downshiftedfrom the solution value by 50–65 cm–1 (Breton et al., 1994a; Brudler et al., 1994). The relative positions of the C1=O and C4=O stretch frequencies are consistent with the C2- and C3-methoxy groups being roughly in-plane and outof-plane, respectively, but the magnitude of the C4=O downshift is far larger than can be accounted for by this effect. Furthermore, a substantially in-plane orientation for either group is not well supported by a consensus of the more recent X-ray structures, nor by an FTIR analysis of the methoxy marker bands at 1260-1290 cm–1 (Remy et al., 2003). However, FTIR spectra of RCs reconstituted with monomethoxy-ubiquinone-4 analogs (with tetra-isoprenyl sidechains) show that absence of the 2-methoxy group does result in an upshift of the C4=O stretch, although only by about 10 cm–1 (A. Vakkasoglu, B. Lipshutz and C. A. Wraight, unpublished). Consistent with such a role, tetramethylbenzoquinone (duroquinone) does not exhibit a substantially downshifted C=O stretch, although small, tailless quinones also appear to be distinct from isoprenylated analogs in other ways (Breton et al., 1994b). Strong hydrogen bonding has been suggested as the origin of the C4=O stretch frequency, but it is a very large shift, especially since hydrogen bonding by protic solvents has little influence on the C=O stretch frequency (Bauscher and Mäntele, 1992). Linear relationships between hydrogen bond strength and relative frequency shift (∆νC=O /νC=O), have been shown; a specific dependence of 4 x 10–3 (kcal/mol)–1, reported by Zadorozhnyi and Ishchenko (1965) for both intra- and intermolecular hydrogen bonds, has been widely cited. Using the relationship of Zadorozhnyi and Ishchenko, the 50–65 cm–1 frequency shift for C4=O of QA (depending on whether 1665 or 1650 cm–1 is used as a reference, see above) would correspond to a hydrogen bond strength (enthalpy) of about 7.5–10 kcal/mol. This is a large value, but IR and Raman band shifts have suggested hydrogen bond strengths of this magnitude in other enzyme systems (Deng and Callender, 1999). However, other solution studies have shown that, unlike the classic Badger-Bauer relationship for νOH, different carbonyl species do not fall on a unique line but exhibit distinct linear dependencies (Bellamy and Pace, 1970; Thijs and Zeegers-Huyskens, 1984; Meyerson, 1985).
Chapter 20
Acceptor Quinones of Purple Photosynthetic Bacteria
Thus, caution is needed in applying the Zadorozhnyi and Ishchenko linear relationship, as it has not been validated for quinones Some support for one strong hydrogen bond to QA comes from measurements of the binding affinity of the QA site of Rba. sphaeroides RCs for a series of compounds with different numbers of carbonyls (Gunner et al., 1985; Warncke and Dutton, 1992; 1993a). Thus, anthrone (one C=O) is found to bind 1.2 kcal/mol weaker than 9,10-anthraquinone, while 9,10-dimethyl-anthracene (no C=O) binds more than 6.5 kcal/mol weaker (Gunner et al., 1985). This is consistent with quinone binding utilizing one strong and one weak hydrogen bond but does not prove it, since the removal of one carbonyl could strengthen the remaining interaction. Despite the evidence of the affinity measurements, assigning the C=O band shift in the protein to hydrogen bonding is not well founded for quinones, which show only weak interactions with hydrogen bonding solvents (Bauscher and Mäntele, 1992). Large downshifts of the carbonyl bands are seen in quinones capable of intramolecular hydrogen bonding, such as anthraquinones with OH-substituted in the α position adjacent to the carbonyl (Flett, 1948). However, this is thought not to be primarily due to the hydrogen bonding, per se, but to the electronic redistribution made possible through resonance in a dimer that is stabilized by H-bonding (Rasmussen et al., 1949; Bloom et al., 1959). In contrast, the O-H band for these compounds is greatly shifted by hydrogen bonding, in a typical Badger-Bauer relationship (Badger and Bauer, 1937; Gordy, 1940; Rao et al., 1975). Similar conclusions have been drawn from studies on quinhydrones (dimers of quinone and hydroquinone in a charge transfer complex), which suggest that π electronic interactions make the major contribution to the C=O downshift rather than a strong H-bonding between the quinone carbonyl and adjacent hydroquinone hydroxyl (Bloom et al., 1959; Slifkin and Walmsley, 1969; Kruk et al., 1993). Other evidence that the downshift of the C4=O stretch is not due to strong hydrogen bonding is that incubation with D2O has no effect on the 1601 cm–1 QA band (Breton and Nabedryk, 1995; Wells et al., 2003). It was suggested that no (or very slow) H/D exchange occurred at His M219. Slow exchange is supported by ENDOR data, which showed very slow D2O exchange of two proton hyperfine interactions with identical half times of ≈3 hrs (Okamura and Feher, 1986). More recent ENDOR studies found
391
one of the two hyperfine interactions (assigned to the peptide NH of Ala M260) to exchange in about 1 hour, significantly more quickly than the other (assigned to NδH of His M219), which takes several hours at room temperature (Flores et al., 2006).4 However, H/D shifts are seen in the semiquinone IR spectrum (Breton and Nabedryk, 1995; Wells et al., 2003), suggesting that the light-dark cycling used in IR difference spectroscopy may accelerate the exchange. Thus, the lack of a significant D/H-shift in the quinone C=O stretch at 1600 cm–1 may not be due to lack of exchange, but to insensitivity to the isotopic substitution. This would not be supportive of an unusually strong H-bond. Considering X-ray structures with a resolution ≤2.8 Å, the heavy atom distances from O4 to Nδ of His M219 and O1 to the carbonyl of Ala M260 are both about 2.8 Å — neither bond is particularly short and both are of similar length. In addition neither Ala nor His donor N is in plane with the carbonyl, which is not optimal. The hydrogen bonding strength has been estimated using a standard molecular mechanics force field, which considers the Lennard-Jones attractive and repulsive terms and the electrostatic interactions that the quinone makes with the protein. In 1aig the interactions with the His sidechain are estimated at only –1.6 kcal/mol, and –0.4 kcal/mol for the amide backbone, neither of which are especially strong (Zhu and Gunner, 2005). Solid state NMR studies on 13C-labeled quinones in RCs also provide little support for the strong hydrogen bond to C4=O (van Liemt et al., 1995), although de Groot (1995) has made a brave effort to show that the expected chemical shift could be masked by other effects. Thus, although the FTIR and NMR results may not be fundamentally inconsistent, it seems fair to say that the origin of the C4=O frequency shift for 4 A mutant RC with His M266 changed to cysteine (mutant M266HC) has been widely used for ENDOR studies of the semiquinones, because it readily incorporates Zn in place of Fe (Williams et al., 1991). Although no substantial perturbations are seen in the X-ray structure (E. C. Abresch, unpublished, reported in Lubitz and Feher, 1999), Paddock et al. (1999) have reported much faster H/D exchange (
