The Molecular Biology of Chloroplasts and Mitochondria in (Advances in Photosynthesis and Respiration)

The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas

Advances in Photosynthesis

VOLUME 7

Series Editor: GOVINDJEE University of Illinois, Urbana, Illinois, U.S.A.

Consulting Editors: Jan AMESZ, Leiden, The Netherlands Eva-Mari ARO, Turku, Finland James BARBER, London, United Kingdom Robert E. BLANKENSHIP, Tempe, Arizona, U.S.A. Norio MURATA, Okki, Japan Donald R. ORT, Urbana, Illinois, U.S.A.

Advances in Photosynthesis is an ambitious book series seeking to provide a comprehensive and state-of-the-art account of photosynthesis research. Pho­ tosynthesis is the process by which higher plants, algae and certain species of bacteria transform and store solar energy in the form of energy-rich organ­ ic molecules. These compounds are in turn used as the energy source for all growth and reproduction in these organisms. As such, virtually all life on the planet ultimately depends on photosynthetic energy conversion. This series of multiauthored books spans topics from physics to agronomy, from femtosecond reactions to season long production, from the photophysics of reaction centers to the physiology of whole organisms, and from X-ray crys­ tallography of proteins to the morphology of intact plants. The intent of this series of publications is to offer beginning researchers, graduate students, and even research specialists a comprehensive current picture of the remark­ able advances across the full scope of photosynthesis research.

The titles to be published in this series are listed on the backcover of this volume.

The Molecular Biology of

Chloroplasts and

Mitochondria

in Chlamydomonas

Edited by

J.-D. Rochaix M. Goldschmidt-Clermont Departments of Molecular Biology and Plant Biology,

University of Geneva,

Geneva, Switzerland

and

S. Merchant Department of Chemistry and Biochemistry

and Molecular Biology Institute,

University of California-Los Angeles,

Los Angeles, U.S.A.

KLUWER ACADEMIC PUBLISHERS NEW YORK, BOSTON, DORDRECHT, LONDON, MOSCOW

eBook ISBN: Print ISBN:

0-306-48204-5 0-7923-5174-6

©2004 Kluwer Academic Publishers New York, Boston, Dordrecht, London, Moscow Print ©1998 Kluwer Academic Publishers Dordrecht All rights reserved No part of this eBook may be reproduced or transmitted in any form or by any means, electronic, mechanical, recording, or otherwise, without written consent from the Publisher Created in the United States of America Visit Kluwer Online at: and Kluwer's eBookstore at:

http://kluweronline.com http://ebooks.kluweronline.com

This book is dedicated to Paul Levine for his pioneering studies on the genetics of photosynthesis in Chlamydomonas and to the memory of Ruth Sager for her seminal contributions to organellar genetics.

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Contents

Preface

xvii

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1–11

Introduction to Chlamydomonas Elizabeth H. Harris

Summary I. Why Chlamydomonas? II. CellArchitecture III. Life Cycle IV. Laboratory strains of Chlamydomonas reinhardtii V. Genetic Analysis VI. Molecular Biology VII. Resources Acknowledgment References

2

Perspectives on Early Research on Photosynthesis in Chlamydomonas Robert K. Togasaki and Stefan J. Surzycki

1

1

3

3

4

6

7

7

8

8

13–23

Summary I. General Background II. The Levine Laboratory in the Early 1960s III. Establishment of Chlamydomonas reinhardtii as a Legitimate Model Organism IV. Development of New Techniques V. Emergence of New Research Targets VI. Old Experiments Becoming Reality Acknowledgment References

3

Organization of the Nuclear Genome Carolyn D. Silflow

Summary I. Introduction and Scope II. General Characteristics of the Nuclear Genome III. Organization of the Genome IV. Characteristics of Chlamydomonas Genes Transcribed by Polymerase II IV. Physical Mapping of the Chlamydomonas Genome VI. Future Prospects Acknowledgments References

4

Nuclear Transformation: Technology and Applications Karen L. Kindle

Summary I. Introduction II. A Brief History of C. reinhardtii Nuclear Transformation

13

13

14

15

18

19

21

21

22

25–40 25

26

26

26

30

36

37

37

37

41–61 42

42

42

III. Selectable Markers IV. Methods for Introducing DNA into the Nuclear Genome of C. reinhardtii V. Reporters and Promoters VI. Characteristics of Transformation Events VII. Insertional Mutagenesis and Gene Tagging VIII. Gene Isolation by Complementation of a Mutant Phenotype IX. Homologous Recombination and Gene Targeting X. The Use of Nuclear Transformation to Study Promoter Function XI. Conclusion Acknowledgments References

5

43

45

46

48

51

54

54

56

58

58

58

Modes and Tempos of Mitochondrial and Chloroplast Genome Evolution in Chlamydomonas: A Comparative Analysis 63–91 Aurora M. Nedelcu and Robert W. Lee

Summary I. Introduction II. Phylogenetic Position of Chlamydomonas III. Monophyletic versus Polyphyletic Origin of Mitochondria and Plastids:

The Chlamydomonas Case IV. Evolution of Mitochondrial and Chloroplast Genome Size in Chlamydomonas V. Evolution of Mitochondrial and Chloroplast Genome Organization in

Chlamydomonas VI. Evolution of Mitochondril and Chloroplast Gene Structure and Organization

in Chlamydomonas VII. Evolution of Mitochondrial and Chloroplast DNA Sequences in

Chlamydomonas VIII. Conclusions Acknowledgments References

6

Uniparental Inheritance of Chloroplast Genomes E. Virginia Armbrust Summary I. Introduction II. Historical Overview of the Uniparental Inheritance of Chloroplast DNA III. Mating-Type Control of Life Cycle Events IV. Protection of Plus Chloroplast DNA V. Zygote Specific Elimination of Minus Chloroplast DNA VI. Regulation of Chloroplast DNA Inheritance VI. Evolution of the Uniparental Inheritance of Organelle Genomes Acknowledgments References

7

Replication, Recombination, and Repair in the Chloroplast Genetic System of Chlamydomonas Barbara B. Sears

Summary I. Introduction II. Replication III. Recombination IV. Repair

63

64

64

65

69

79

82

85

87

87

87

93–113 93

94

95

98

101

103

108

110

110

110

115–138 115

116

116

123

130

viii

V. Perspectives and Conclusions Acknowledgments References

8

Chloroplast Transformation and Reverse Genetics Michel Goldschmidt-Clermont

133

133

133

139–149

Summary I. Introduction II. Delivery of DNA to the Chloroplast III. Selectable Markers and Reporters IV. Fate of Transforming DNA V. Reverse Genetics VI. Conclusion and Perspective Acknowledgments References

9

Chloroplast RNA Stability Jörg Nickelsen

139

140

140

140

142

147

147

148

148

151–163

Summary I. Introduction II. Cell Cycle Dependent Regulation of Chloroplast RNA Stability III. Nuclear Mutants Affected in Chloroplast RNA Stability IV. Towards a Molecular Model of Chloroplast RNA Stabilization/Degradation V. Conclusions and Perspectives Acknowledgments References

10 Chloroplast RNA Synthesis and Processing David B. Stern and Robert G. Drager

Summary I. Transcription of Chloroplast Genes II. Processing of Chloroplast mRNAs Acknowledgments References

151

152

152

154

154

161

161

161

165–183 165

166

171

177

177

11 RNA Splicing in the Chloroplast 183–195 David L. Herrin, Tai-Chih Kuo and Michel Goldschmidt-Clermont

Summary I. Introduction II. Group I Introns III. Group II Introns and Trans-Splicing IV. Perspective Acknowledgments References

183

184

184

190

193

193

193

197–217 12 Regulation of Chloroplast Translation Charles R. Hauser, Nicholas W. Gillham and John E. Boynton

197

Summary 198

I. Introduction II. The Role of Physiological and Environmental Factors in Translational Control 200

ix

III. Current Biochemical and Genetic Approaches to Dissect Mechanisms of

Translational Regulation IV. Cis-acting Sequences Involved in Translation Initiation V. Translational Regulation Involves Interactions between cis-Acting

Sequences and trans-Acting Factors VI. Ribosomes, Membranes and Tethers VII. Translational Regulation of Complex Assembly VIII. How are the Regulatory Proteins Regulated? IX. Is there Hierarchical Control of Chloroplast mRNA Translation? Acknowledgments References

13 Chloroplast Protein Translocation Mireille C. Perret, Karen K. Bernd and Bruce D. Kohorn

219

220

220

222

223

226

228

229

229

233–254

Summary I. Introduction II. Cell and Chloroplast Morphology III. Ultrastructural Organization of Thylakoid Membranes IV. Dynamic Aspects of Thylakoid Membrane Organization V. Biogenesis VI. Conclusion Acknowledgment References

15 Assembly of Photosystem II Jeanne Marie Erickson

205

209

211

212

213

214

214

219–231

Summary I. Introduction II. Chloroplast Import III. Sorting of Proteins Within the Chloroplast IV. Thylakoid Translocation V. Mutations Affecting Translocation VI. Perspectives Acknowledgments References

14 Supramolecular Organization of the Chloroplast and of the Thylakoid Membranes Jacqueline Olive and Francis-André Wollman

202

203

234

234

235

239

246

248

250

251

251

255–285

Summary I. Introduction II. Developmental Biogenesis of Photosystem II III. Assembly of Photosystem II Complexes IV. Assembly of the Extrinsic Membrane Polypeptides of the PS II

Oxygen-Evolving Complex V. Assembly of Manganese: The Catalytic Center of the Oxygen-Evolving

Complex Acknowledgments References

x

255

256

257

260

270

273

277

277

16 Functional Analysis of Photosystem II Stuart V. Ruffle and Richard T. Sayre Summary I. Introduction II. The Photosystem II Complex III. The Chloroplast DNA Encoded Small Polypeptides of Photosystem II IV. The Nucleus Encoded Polypeptides of the Photosystem II Complex V. Perspectives Acknowledgments References

17 Structure and Function of Photosystem I Andrew N. Webber and Scott E. Bingham

Summary I. Introduction II. Structure of Photosystem I III. Nature and Function of Electron Transfer Cofactors IV. Antenna Structure and Function V. Function of Photosystem I Subunits VI. Biogenesis of Photosystem I Acknowledgments References

18 Reexamining the Validity of the Z-Scheme: Is Photosystem I

Required for Oxygenic Photosynthesis in Chlamydomonas? Kevin Redding and Gilles Peltier

Summary I. The Z-Scheme of Oxygenic Photosynthesis and Alternative Schemes II. Electron Transport in the Absence of PS II III. Photosynthesis in the Absence of PS I IV. Putative Electron Transport Pathways Outside of the Z-Scheme V. Thermodynamic Considerations VI. Evolutionary Considerations VII. Conclusions Acknowledgments References

19 Assembly of Light-Harvesting Systems J. Kenneth Hoober, Hyoungshin Park, Gregory R. Wolfe, Yutaka Komine and Laura L. Eggink Summary I. Thylakoid Biogenesis in Chlamydomonas II. Analysis of LHCII Assembly III. Site of Assembly of LHCII During Initial Greening IV. Conclusions Acknowledgments References

xi

287–322

287

288

289

308

311

315

315

315

323–348 324

324

325

328

332

333

341

344

344

349–362

349

350

351

354

357

358

358

359

359

360

363–376 363

364

366

368

371

373

373

20 Pigment Biosynthesis: Chlorophylls, Heme, and Carotenoids Michael P. Timko

377–414

Summary I. Introduction II. Tetrapyrroles and Their Derivatives—An Overview III. Formation of ALA IV. The Pathway from ALA to Protoporphyrin IX V. The Magnesium Branch—Chlorophyll Formation VI. The Iron Branch—Formation of Heme VII. Light and Metabolic Regulation of Chlorophyll Formation VIII. Carotenoids Acknowledgments References

21 Glycerolipids: Composition, Biosynthesis and Function in Chlamydomonas Antoine Trémolières

378

378

378

379

383

388

397

398

401

406

406

415–431

Summary Introduction II. Glycerolipid and Fatty Acid Composition of Chlamydomonas III. Lipid Metabolic Pathway in Chlamydomonas spp. IV. In vivo Modifications of Lipid Composition in Chlamydomonas V. Mutants Affected in Lipid Composition VI. Involvement of Lipids in the Functional Organization and the Biogenesis

of the Photosynthetic Apparatus Acknowledgments References

22 In vivo Measurements of Photosynthetic Activity: Methods Pierre Joliot, Daniel Béal and René Delosme Summary I. Introduction II. Kinetic Analysis of the Fluorescence Yield III. Fluorescence Emission Spectra at Low Temperature IV. Delayed Fluorescence Measurements V. Oxygen Measurements VI. Absorption Spectroscopy VII. Photoacoustic Measurements VIII. Conclusion and Perspectives Appendix A: Estimation of the Signal-to-Noise Ratio in Fluorescence

Measurements Appendix B: Flash Spectrophotometer Acknowledgment References

23 New Digital Imaging Instrument For Measuring Fluorescence and Delayed Luminescence Pierre Bennoun and Daniel Béal

Summary I. Introduction II. Setup for Fluorescence and Delayed Luminescence Video Imaging

xii

415

416

417

422

425

426

428

429

429

433–449 433

434

436

439

439

439

440

443

445

446

446

448

448

451–458 451

452

452

III. Digital Fluorescence Imaging Related to Photosynthetic Electron Transfer IV. Digital Fluorescence Imaging Related to the Permanent Thylakoid

Electrochemical Gradient V. Digital Delayed Luminescence Imaging Related to Light-Induced and

Permanent Thylakoid Electrochemical Gradient Acknowledgments References

24 The Structure, Function and Biogenesis Of Cytochrome Complexes Francis-André Wollman

Summary I. General Traits II. Biochemical and Structural Studies III. Functional Studies IV. The pet Genes V. Biogenesis and Assembly VI. Concluding Remarks Acknowledgments References

25 Assembly and Function of the Chloroplast ATP Synthase Heinrich Strotmann, Noun Shavit and Stefan Leu

Summary I. Introduction II. Structure of

III. Molecular Genetics of IV. Mechanism of V. Regulation of VI. Conclusions References

452

453

455

457

457

459–476 460

460

461

463

466

467

472

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477–500 477

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478 482

487

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26 Molecular Aspects of Components of the Ferredoxin/Thioredoxin 501–514 Systems Jean-Pierre Jacquot, Mariana Stein, Stéphane Lemaire, Paulette Decottignies, Pierre Le Maréchal and Jean-Mark Lancelin Summary I. Introduction II. Ferredoxin Dependent Systems III. Thioredoxin Dependent Systems IV. Conclusion Acknowledgment References

501

502

505

508

511

512

512

515–527

27 Genetic Engineering of Rubisco Robert J. Spreitzer

Summary I. Introduction II. Chloroplast Genetic Screening and Selection

xiii

515

516

518

III. Directed Mutagenesis and Chloroplast Transformation IV. Rubisco Nuclear Mutants V. Conclusion and Perspective Acknowledgments References

28

Acquisition. Acclimation to Changing Carbon Availability

Martin H. Spalding

Summary

I. Introduction II. Photosynthetic Carbon Assimilation III. Induction of the CCM and Related Adaptations to Limiting Acknowledgments References

521

523

524

524

524

529–547 529

530

530

539

544

544

549–567

29 Regulation of Starch Biosynthesis Steven G. Ball

Summary

I. Starch: Structure and Function II. The Starch Pathway III. The Genetics of Starch Biosynthesis IV. A Model Explaining the Biogenesis of the Plant Starch Granule V. Future Prospects Acknowledgments References

30 State Transition and Photoinhibition Nir Keren and Itzhak Ohad

Summary

I. Introduction II. State Transition: The Phenomenon III. Light Stress: Photoinhibition and Recovery IV. Concluding Remarks and Perspectives Acknowledgments References

31 Synthesis of Metalloproteins Involved in Photosynthesis: Plastocyanin and Cytochromes Sabeeha Merchant

Summary

I. Introduction II. Copper-Responsive Synthesis of Plastocyanin and Cytochrome III. Genetic Analysis of Chloroplast Metalloprotein Assembly IV. Conclusions Acknowledgments References

32 Responses to Deficiencies in Macronutrients John P. Davies and Arthur R. Grossman Summary I. Introduction

549

550

554

559

563

564

565

565

569–596 569

570

574

578

590

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597–611 598

598

600

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608

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613–635 614

614

xiv

II. Nutrients in the Environment III. Specific Responses IV. Common Responses V. Model Integrating the Responses to Nutrient Deprivation VI. Regulation of the Responses to Nutrient Deprivation VII. Identification of Mutants Deficient in the Acclimation to Nutrient Deprivation Acknowledgments References

33 Nitrogen Assimilation and its Regulation Emilio Fernández, Aurora Galván and Alberto Quesada

Summary I. Introduction. Pathways for Nitrogen Assimilation in Chlamydomonas II. Assimilation of Ammonium III. Assimilation of Amino Acids IV. Assimilation of Purines V. Assimilation of Nitrate and Nitrite VI. Concluding Remarks Acknowledgments References

34 Mitochondrial Genetics Claire Remacle and René F. Matagne

Summary I. Introduction II. Mitochondrial Genome III. Mitochondria and the Electron Transport Chain IV. Mutations Affecting the Mitochondrial Genome V. Transmission of Mitochondrial Genes in Meiotic Zygotes VI. Transmission of Mitochondrial Genes in Vegetative Zygotes and

Mapping of Mitochondrial Mutations by Recombinational Analysis VII. Mitochondrial Transformation Acknowledgments References

35 Chlororespiration, Sixteen Years Later Pierre Bennoun

Summary I. Introduction II. The Thylakoid Electrochemical Gradient Present in the Dark III. Reduction of Plastoquinone in the Dark IV. Oxidation of Plastoquinol in the Dark V. Conclusion Cautionary Note Acknowledgments References

36 Perspectives Lauren J. Mets and Jean-David Rochaix

Summary I. Introduction II. The Niche of Chlamydomonas in Photosynthesis Research

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615

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629

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637–659 638

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661–674 661

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662

664

665

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669

671

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675–683 675

676

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680

680

682

682

682

685–703 685

686

687

III. Forefront Problems in Photosynthesis and Organelle Research Acknowledgments References

Index

696 700 700

705

xvi

Preface

The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas is the seventh volume to be published in the series Advances in Photosynthesis of Kluwer Academic Publishers (Series Editor: Govindjee). Volume 1 dealt with The Molecular Biology of Cyanobacteria; Volume 2 with Anoxygenic Photosynthetic Bacteria; Volume 3 with Biophysical Techniques in Photosynthesis; Volume 4 with Photosynthesis and the Environment; and Volume 6 with Lipids in Photosynthesis: Structure, Function and Genetics. The main goal of this book is to provide a comprehensive overview of current research with the green alga Chlamydomonas on chloroplast and mitochondrial biogenesis and function, with special emphasis on the assembly and structure-function relationships of the constituents ofthe photosynthetic apparatus. The editors have encouraged the contributors of this volume to emphasize the particular features of Chlamydomonas that make this unicellular organism uniquely suited for study­ ing photosynthesis and its multiple regulatory mechanisms operating under various environmental and stress conditions. A second, but equally important aim is to show that current research in photosyn­ thesis is multidisciplinary and combines molecular genetics, biochemical, biophysical and physiological approaches. Although Chlamydomonas has also proven to be a powerful system for understanding the structure, function and assembly of flagella, this topic is not covered in the book. Chlamydomonas research would not have reached its present status without the pioneering studies of the late Ruth Sager and of Paul Levine. Organellar genetic analysis of this alga started over 40 years ago when Ruth Sager discovered that during crosses certain traits were transmitted uniparentally to the progeny from the mating-type plus parent, but not from the mating-type minus parent. These uniparental traits were shown later to be specified by the chloroplast genome. Sager also found that, in rare cases, the uniparental traits of both parents could be inherited and that the analysis of their segregation pattern during crosses could be used to construct a genetic map. The potential of using C. reinhardtii for

a genetic dissection of photosynthesis was first recognized by Paul Levine. Together with his coworkers, he initiated a long-range genetic approach which proved to be highly successful. It provided genetic support for the linear Z scheme of photosynthesis and led to the identification of new components of the photosynthetic electron transfer chain such as the Rieske protein ofthe cytochrome complex. During the past 20 years, the powerful techniques of molecular biology and genetics, and the development of methods for efficient nuclear and chloroplast transformation of C. reinhardtii have greatly enhanced the potential of this organism as an experimental system for studying chloroplast biogenesis. This has led to impressive advances in our understanding of the regulation of chloroplast gene expression and it has provided important new insights into the complex cooperative interplay between the chloroplast and nuclear compartments in the assembly of the photosynthetic apparatus. At the same time, the ability to manipulate the chloroplast genome with surgical precision has opened the door for a detailed structure-function analysis of photosynthetic complexes in vitro, and thanks to the refinements and new developments in spectroscopic and fluorescence techniques, also in vivo. We feel strongly that a book on these recent exciting advances in research on photosynthesis in Chlamydomonas is timely and important. The first part of the book provides a general introduction to Chlamydomonas (Chapter 1, Harris), a historical chapter on early research on photo­ synthesis in this organism (Chapter 2, Togasaki and Surzycki) and chapters on nuclear genome organi­ zation (Chapter 3, Silflow), nuclear transformation (Chapter 4, Kindle), mitochondrial and chloroplast genome evolution (Chapter 5, Nedelcu and Lee), chloroplast uniparental inheritance (Chapter 6, Armbrust), chloroplast DNA metabolism (Chapter 7, Sears) and chloroplast transformation and reverse genetics (Chapter 8, Goldschmidt-Clermont). The second part includes several chapters on chloroplast gene expression: RNA stability (Chapter 9, Nickelsen), RNA processing (Chapter 10, Stern and

xvii

Drager), splicing (Chapter 11, Herrin et al.) and translation (Chapter 12, Hauser et al.). Protein targeting in the chloroplast is discussed in Chapter 13 (Perret et al.). The third part includes articles on the biosynthesis and function of thylakoid membranes (Chapter 14, Olive and Wollman), Photosystem II (Chapter 15, Erickson; Chapter 16, Ruffle and Sayre), Photosystem I (Chapter 17, Webber and Bingham; Chapter 18, Redding and Peltier), LHCII (Chapter 19, Hoober et al.), pigments (Chapter 20, Timko), glycerolipids (Chapter 21, Trémolières), the complex (Chapter 24, Wollman), the cytochrome ATP synthase (Chapter 25, Strotmann et al.), ferredoxin and thioredoxin (Chapter 26, Jacquot et al.) and of ribulose 1,5 bisphosphate carboxylaseoxygenase (Chapter 27, Spreitzer). In addition, Chapters 22 (Joliot et al.) and 23 (Bennoun and Beal) describe new and powerful techniques used for measurements of photosynthetic activity in vivo. These techniques are particularly suited for Chlamydomonas. The fourth part includes chapters uptake (Chapter 28, Spalding) and starch on biosynthesis (Chapter 29, Ball). Several articles are devoted to the responses of Chlamydomonas to various stress conditions, such as high light (Chapter 30, Keren and Ohad), copper deficiency (Chapter 31, Merchant) and macronutrient depri­ vation (Chapter 32, Davies and Grossman). Nitrogen assimilation and its regulation is discussed in Chapter 33 (Fernández et al.). Chapter 34 (Remacle and Matagne) describes mitochondrial genetics and

Chapter 35 (Bennoun) discusses the current models of chlororespiration. The last Chapter (36, Mets and Rochaix) offers a perspective on research on photosynthesis with Chlamydomonas. We thank the authors for their invaluable contributions which we hope will make this book very useful for researchers and students interested in photosynthesis and organellar biology in Chlamy­ domonas. The book is also intended for a wide audience, but is specifically designed for advanced undergraduate and graduate students and researchers in the fields of biochemistry, molecular biology, physiology, biophysics, plant biology, phycology and biotechnology. We also hope that this book will stimulate scientists outside the Chlamydomonas research community to use this organism for their studies. We wish to express our gratitude to Larry Orr for his patience with inexperienced editors, generous help and remarkable efficiency, to Govindjee for his continued interest and for his many helpful suggestions, and to Nicolas Roggli and Michael Hippler for their help in designing the cover graphics. Finally, we hope that by showing the extraordinary power and uniqueness of Chlamydomonas as a research tool in photosynthesis and by documenting the fast pace of progress achieved in the past years with this unicellular organism, this book will help promote Chlamydomonas as the ‘green yeast’ among the plant research community. Jean-David Rochaix Michel Goldschmidt-Clermont Sabeeha Merchant

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Chapter 1

Introduction to Chlamydomonas Elizabeth H. Harris

DCMB Group, Department of Botany, Box 91000,

Duke University, Durham, NC 27708-1000, U.S.A.

Summary I. Why Chlamydomonas? II. Cell Architecture III. Life Cycle IV. Laboratory strains of Chlamydomonas reinhardtii V. Genetic Analysis VI. Molecular Biology VII. Resources Acknowledgment References

1 1 3 3 4 6 7 7 8 8

Summary The unicellular green alga Chlamydomonas has found widespread use as a model experimental system for diverse studies in cell and molecular biology. The ability of C. reinhardtii to grow heterotrophically with acetate as its sole carbon source has made this species especially useful for investigation of chloroplast biogenesis and function, since mutants unable to carry out photosynthesis are viable. The simple vegetative and sexual cycles are easily manipulated in the laboratory, making this organism a powerful tool for genetic analysis of photosynthesis as well as many other cellular functions. The usual laboratory strain of C. reinhardtii is the descendant ofan isolate made in Massachusetts in 1945. Several additional strains interfertile with this one have been isolated from nature, all from North America, and are providing a useful source of molecular diversity. More than 300 genetic and molecular loci have now been identified in seventeen linkage groups in the nuclear genome. Maps, references, cultures and other resources for Chlamydomonas research are available from the Chlamydomonas Genetics Center and other collections.

I. Why Chlamydomonas? Sometimes called the ‘green yeast’ (Goodenough, 1992; Rochaix, 1995), this unicellular chlorophyte alga has achieved recognition as a model system for the study of photosynthesis, organelle biogenesis, and many other aspects of cell biology. A simple life cycle that is easily manipulated in the laboratory, minimal nutritional requirements, and rapid growth have all favored selection of this alga as an

experimental organism. Although mutants of a Chlamydomonas species were isolated and crossed as early as 1918 by A. Pascher (see Harris, 1989 for review), the modern age of Chlamydomonas research can be dated from the 1940s, with isolation of the principal laboratory strains of C. reinhardtii and C. moewusii. (Smith, 1946; Lewin, 1949). Conditions were established for laboratory culture and for manipulation of the sexual cycle (Smith and Regnery, 1950; Lewin, 1951; Sager and Granick, 1953, 1954),

J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 1–11. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

2 and the first mutants were selected (Lewin, 1952, 1953, 1954; Sager, 1954, 1955). Nearly fifty years and roughly 5,000 research papers later, we know many intimate details of the life of our favorite subject, but there are secrets still to be revealed. R. K. Togasaki and S. J. Surzycki (Chapter 2) have traced the history of research on photosynthesis in Chlamydomonas. The present chapter introduces the organism and provides suggestions for further reading on other aspects of its biology. Species of the genus Chlamydomonas are distinguished from other unicellular Volvocales by the presence of a cell wall, a pair of apical flagella, and a basal chloroplast surrounding one or more pyrenoids (Fig. 1). H. Ettl (1976) recognized 459 species based on morphological criteria, but the trend in recent years has been toward consolidation into a smaller number (Ettl and Schlösser, 1992; Schlösser, 1994). Molecular phylogenies now suggest that the genus comprises a diverse collection of organisms, some only distantly related (Buchheim et al., 1990, 1996; Chapter 5, Nedelcu and Lee). Some Chlamydomonas species have close affinity to certain colonial Volvocales (Buchheim et al., 1994; Liss et al., 1997). Although the Volvocales are regarded as a side branch from the evolutionary tree leading to land plants (Chapman and Buchheim, 1992), components of the photosynthetic process are highly conserved, and the relevance of research with Chlamydomonas to our understanding of chloroplast function in vascular plants is undisputed. Studies of photosynthesis and chloroplast biogenesis have been greatly facilitated by the ability of some species, notably C. reinhardtii, to grow with acetate as their sole carbon source (Harris, 1989). Mutants unable to carry out photosynthesis are thus viable when supplied with acetate. In nature this metabolism is usually found in organisms living in environments rich in organic compounds, and these so-called ‘acetate flagellates’ can tolerate the low oxygen tension and relatively high levels of associated with these conditions (Pringsheim, 1937; Hutner and Provasoli, 1951). Most laboratory isolates of C. reinhardtii have in fact come originally from nutrient-rich soil in cultivated fields or gardens (Sack et al., 1994). C. moewusii and its close relative, the strain usually known in laboratory research as C. eugametos (Gowans, 1963), are also widely used as experimental subjects, especially for studies of the sexual cycle and to some extent for flagellar biogenesis and

Elizabeth H. Harris

nutritional investigation. Because these species, which are only distantly related to C. reinhardtii (Lemieux et al., 1985; Boudreau and Turmel, 1996; Buchheim et al., 1996), are obligate phototrophs, they have not been employed in many photosynthesis studies. C. monoica, a homothallic species, has been developed by VanWinkle-Swift and colleagues (VanWinkle-Swift and Hahn, 1986; van den Ende, 1995) as a genetic system for study of processes involved in the sexual cycle, including investigations on mechanism of uniparental transmission of the chloroplast genome in crosses (VanWinkle-Swift and Aubert, 1983), but few physiological and molecular studies have been done with this species. Thus most research on chloroplast biogenesis and function has been carried out with C. reinhardtii, and in the present book, ‘Chlamydomonas’ can be

Chapter 1

Introduction to Chlamydomonas

assumed to refer to C. reinhardtii unless otherwise specified.

II. Cell Architecture Cells of C. reinhardtii are spherical to ovate in shape, varying from approximately 8 to as much as in length over the course of the vegetative cell cycle (see Ettl, 1976). A pair of apical flagella extend through a specialized collar region in the cell wall, and are typically 1.5 to 2× the length of the cell body. Within the cell, the flagella terminate in a pair of basal bodies connected by a striated fiber (see Jarvik and Suhan, 1991; Johnson and Rosenbaum, 1992). The basal bodies are connected to the nucleus by centrin fibers (Taillon et al., 1992; Salisbury, 1995), and to a cruciate system of four sets of microtubules, the flagellar roots (Lechtreck and Melkonian, 1991). Two contractile vacuoles are usually seen between the basal bodies and the nucleus (Luykx et al., 1997). For concise reviews of flagellar architecture and biogenesis in Chlamydomonas, see Johnson and Rosenbaum (1993), Johnson (1995), Dutcher (1995a), and Smith and Lefebvre (1997). Bernstein (1995) has reviewed flagellar kinesins, and Porter (1996; also Porter et al., 1996) has discussed dynein structure and genetics. Experimental procedures and reviews relating to the motility system are presented in a series of articles in the volume edited by Dentler and Witman (1995). The nucleus is partially surrounded by a cupshaped chloroplast in C. reinhardtii (Fig. 1), whereas in some other species the chloroplast may be displaced laterally (Ettl, 1976). A prominent pyrenoid is situated within the chloroplast distal to the nucleus. More information on chloroplast structure and the functions of the pyrenoid can be found in Chapters 14 (Olive and Wollman), 28 (Spalding) and 29 (Ball). The eyespot or stigma lies at the anterior of the chloroplast, just within the outer chloroplast envelope and close to the overlying cytoplasmic membrane, and contains carotenoid pigments (Crescitelli et al., 1992; Lawson and Satir, 1994). A unique rhodopsin has been identified as the photoreceptor for both phototaxis and photophobic responses (Derguini et al., 1991; Kroeger and Hegemann, 1994; Deininger et al., 1995), and the nature of the rhodopsin-mediated photoresponse is becoming understood (Pazour et al. 1995; Holland et al., 1996, 1997; Nonnengaesser et al., 1996). Witman (1993) has summarized earlier

3 investigations of phototaxis in Chlamydomonas. For general reviews of the cell biology of algal photoresponses, see Kreimer (1994) and Hegemann (1997). The evolution of algal visual proteins is reviewed by Walne and Gualtieri (1994). Mitochondria may appear in sections as small oval or elongate bodies (Fig. 1), or under some growth conditions can form an interconnecting, branching network throughout the cell (see Harris, 1989). Mitochondrial DNA and genetics are covered in this volume in Chapter 5 (Nedelcu and Lee) and Chapter 34 (Matagne), respectively. Respiratory physiology, including the cytochrome and alternative oxidase pathways, has been discussed by Weger and colleagues (Weger et al., 1990; Derzaph and Weger, 1996; Weger, 1996), and Eriksson et al. (1996) have published an improved method for isolation of physiologically active mitochondria. Geraghty and Spalding (1996) have discussed mitochondrial responses to changing concentrations. Cells of C. reinhardtii are surrounded by a complex cell wall composed of glycoproteins with high hydroxyproline content (Goodenough and Heuser, 1985; Matsuda, 1988; Adair and Snell, 1990; Woessner and Goodenough, 1994). Genes encoding some of the constituent proteins have been cloned and sequenced (Waffenschmidt et al., 1993; Kurvari, 1997). Mutants with absent or defective cell walls have been isolated (Harris, 1989; Voigt et al., 1997), and have proved useful as recipients for trans­ formation and in other experimental applications where the wall would otherwise present a barrier to manipulation.

III. Life Cycle Vegetative cells of C. reinhardtii are haploid. Growing logarithmically on a 12:12 light-dark cycle, they divide synchronously during the dark period in (usually) two mitotic divisions in rapid succession, releasing four daughter cells from a single sporangial wall after secretion of a specific lytic enzyme (Schlösser et al., 1976; Spessert and Waffenschmidt, 1990). Under optimal laboratory conditions in sufficient light, stationary phase is reached at about 1 cells/ml. Gene regulation and expression to through the cell cycle have been investigated by several laboratories (Savard et al., 1996; Voigt et al., 1996; Lechtreck and Silflow, 1997), and mutants blocked at specific points in the cycle have been

4 isolated (Howell and Naliboff, 1973; Harper et al., 1995; Wu et al., 1997). A number of biological processes in Chlamydomonas exhibit circadian rhythms (e.g. Fujiwara et al., 1996; Hwang et al., 1996; Jacobshagen et al., 1996), and Goto and Johnson (1995) have proposed that the cell division cycle itself is controlled by a circadian clock. In nature, the sexual cycle is a response to adverse conditions. In the laboratory, gametogenesis is controlled by nitrogen deprivation and a blue lightresponsive signal transduction pathway (Matsuda et al., 1992; Beck and Haring, 1996; Gloeckner and Beck, 1997; Pan et al., 1997). The light requirement has been obviated by mutation to constitutive expression of one or more regulatory genes in some laboratory strains (Gloeckner and Beck, 1995). is genetically determined Mating type by a complex locus on linkage group VI (Ferris and Goodenough, 1994, 1997) and gametes express mating type-specific glycoproteins (agglutinins) on their flagellar surfaces (Goodenough et al., 1995). Flagellar pairing initiates a cascade of events, including lysis of the gametic cell walls by a specific enzyme distinct from the vegetative cell lysin (Matsuda, 1988; Buchanan et al., 1989), contact of mating type-specific structures at the cell apices (Ferris et al., 1996; Wilson et al., 1997), and fusion of the gametic cytoplasms. For review of the extensive literature on signal transduction in mating, see Musgrave (1993) and Quarmby (1994). A transient quadriflagellate stage is followed by resorption of all flagella, and nuclear and chloroplast fusion follow within a few hours. Within the first 24 hours after mating, a hard, impermeable wall is secreted (Woessner and Goodenough, 1989, 1992), creating a durable zygospore that is resistant to desiccation or other environmental insults. After a maturation period of four to seven days, or longer, restoration of nitrogen, moisture and light induce the germination process. Over the next 18 to 24 hours, meiosis takes place, sometimes followed by a mitotic division, and the zygospore wall bursts to release the four or eight haploid progeny. Separation of these meiotic products to allow each to form a clone is the basis of Chlamydomonas genetics by tetrad analysis (Harris, 1989; Dutcher, 1995b). Nuclear genes are inherited in a Mendelian fashion and segregate 2:2 among the tetrad products. Chloroplast genes are inherited uniparentally from parent in most zygotes (>95% under the usual the laboratory conditions), whereas mitochondrial DNA

Elizabeth H. Harris in C. reinhardtii is transmitted uniparentally from parent. The mechanism by which the the inheritance oforganelle DNA is controlled was hotly debated for years, and is still not fully understood (Chapter 6, Armbrust). This book covers many aspects of metabolism in Chlamydomonas, including synthesis of lipids (Chapter 21, Trémolières), pigments (Chapter 20, Timko), and starch (Chapter 29, Ball). Nitrogen metabolism is discussed by Fernández et al. (Chapter 33), and micronutrient metabolism by Davies and Grossman (Chapter 32). For additional background on Chlamydomonas physiology, see Harris (1989).

IV. Laboratory strains of Chlamydomonas reinhardtii The standard laboratory strains of C. reinhardtii derive from a zygospore isolated by G. M. Smith from a soil sample collected near Amherst, Massachusetts, in 1945 (Hoshaw, 1965). Smith gave his strains to several of the early investigators, with the result that three principal lineages exist today, all apparently tracing their ancestry to the same sample (Table 1; see Harris, 1989, for further discussion). Most of the early nonphotosynthetic mutants were isolated in the strain used by R. P. Levine, W. T. Ebersold and coworkers, usually known as 137C (Smith’s original designation for his isolate, now usually applied specifically to the Levine line). These strains lack nitrate reductase activity and thus require a reduced nitrogen source for growth. Comparison of the strains used by R. A. Lewin, Levine and others suggests that this deficiency was present in this stock as early as 1949. Ruth Sager also obtained strains from Smith in the early 1950s. Her wild type isolate, 21 gr, was selected as a clone that remained green in the dark, in contrast to other strains, collectively designated y mutants, that fail to accumulate chlorophyll in the dark and maintain only a rudimentary chloroplast structure under these conditions (Sager and Palade, 1954). Beginning in the late 1960s, I. Ohad, J. K. Hoober, and others carried out extensive studies of rediffer­ entiation of chloroplast structure on transfer of y mutants into light (Chapter 2, Togasaki and Surzycki; Chapter 19, Hoober et al.). In contrast to the Levine 137C strain, Sager’s 21 gr and the original y1 strains used by her and by Ohad are able to grow on nitrate

Chapter 1

Introduction to Chlamydomonas 5

Elizabeth H. Harris

6 as their sole nitrogen source. The C8 and C9 strains ofthe Japanese culture collection (IAM) are probably strains from Sager, deposited by Y. Tsubo. This fundamental physiological difference between Sager’s and Levine’s strains produced speculation that these strains might have independent origins (Harris, 1989). Molecular analysis, particularly of insertion of the Gulliver transposon (Ferris, 1989) and ofpolymorphisms in chloroplast DNA (Harris et al., 1991), argues, however, for a common origin, followed by loss early on ofnitrate reductase activity in the Levine line, through mutations in the structural gene for nitrate reductase (Nit1) and in an unlinked regulatory gene (Nit2) (Chapter 33, Fernández et al.). Spreitzer and Mets (1981) crossed the Sager 21 gr strain with a Levine 13 7C isolate to create the hybrid strain 2137. Many nonphotosynthetic mutants have been isolated in this background. A third line of C. reinhardtii deriving from Smith’s original isolate made its way to the British (CCAP), German (SAG) and American (Indiana University, subsequently UTEX) algal collections (Table 1; Harris, 1989). These strains have been less frequently used than the aforementioned strains for studies of chloroplast function. There are several strains extant in the major culture collections that are listed as independent isolates of C. reinhardtii, but, based on molecular criteria, they appear to be incorrectly labeled as cultures of one of the Smith strains (Table 1). Ferris (1989) found that insertion patterns for the transposon Gulliver were remarkably similar when four cultures from the SAG collection, ostensibly of independent origin, were compared with the usual laboratory isolates, and chloroplast DNA from these strains shows restriction digest patterns identical to the Smith strains (Harris et al., 1991). In contrast, several strains of undisputed inde­ pendent origin (Table 1) are cross-fertile with laboratory strains of C. reinhardtii, and in recent years have been exploited as sources of molecular diversity. The oldest ofthese strains is the one isolated from South Deerfield, Massachusetts, and identified by Smith as C. reinhardtii but subsequently redescribed by Hoshaw and Ettl (1966) as the type specimen of the new species C. smithii. Hoshaw and Ettl identified it as mating type minus, based on tests against strains 89 and 90 of the Indiana collection. These strains were subsequently found to have been labeled with their mating types reversed (Harris, 1989; Starr and Zeikus, 1993), and the C. smithii

strain from Massachusetts is in fact The strain identified by Hoshaw and Ettl as the opposite mating type of C. smithii was isolated from Santa Cruz CA, and appears to be only slightly interfertile, if at all, with C. reinhardtii. Based on sequence analysis of the ITS-2 region of the nuclear ribosomal repeats, Coleman and Mai (1997) have suggested that this strain bears closer affinity to C. culleus, a species also found on the U.S. west coast, than to C. reinhardtii. Several other more recent ‘wild’ Chlamydomonas strains (Gross et al., 1988; Spanier et al., 1992; Sack et al., 1994; E. H. Harris, unpublished) are fully interfertile with laboratory strains of C. reinhardtii (Table 1). These strains show molecular polymorphisms in both nuclear and chloroplast DNA, and variations in incidence and insertion sites of the Gulliver and TOC transposons (Ferris, 1989; Harris et al., 1991 and unpublished; Sack et al., 1994; Chapter 3, Silflow). It is perhaps worth noting that all the authentic interfertile strains of C. reinhardtii isolated to date have been found in North America east of the Rocky Mountains.

V. Genetic Analysis The first genetic maps published for the nuclear genome of C. reinhardtii (Levine and Goodenough, 1970) comprised non-motile (pf, ‘paralyzed flagella’), non-photosynthetic (ac, ‘acetate-requiring’), anti­ biotic resistant (e.g. streptomycin, cycloheximide), and auxotrophic mutants (requiring arginine, nicotinamide, thiamine or p-aminobenzoic acid). Although the specific mutations are now understood in much greater detail, and some additional mutant types have been identified, these remain the major broad categories. More than 200 genetic loci have been defined based on mutation, and an additional 170 molecular loci have been identified (Ranum et al., 1988; Chapter 3, Silflow,). Levine and his colleagues used mutants to dissect the path of photosynthetic electron transport (Levine, 1969; Chapter2,Togasaki and Surzycki). Flagellarstructure and biogenesis have likewise been elucidated primarily through mutant studies (Curry and Rosenbaum, 1993; Dutcher, 1995a), as have the pathways of nitrogen assimilation (Chapter 33, Fernández et al.). Specific mutations affecting chloroplast structure and function are discussed in detail throughout the present book. Seventeen nuclear linkage groups are currently

Chapter 1

Introduction to Chlamydomonas

recognized (Harris, 1993; Chapter 3, Silflow). Linkage groups XII and XIII as defined by Levine and Goodenough (1970) have been consolidated (Dutcher et al., 1991), as have XVI and XVII (Harris, 1989; Dutcher et al., 1991). Chromosome cytology in C. reinhardtii is poor (see Harris, 1989, for review), and electrophoretic separation of chromosomes has not been fully satisfactory (Hails et al., 1993). Most of the chromosomes co-migrate and are resolved into four bands on pulsed-field gels. Sixteen linkage groups have now been localized to these bands by blot analysis, permitting unmapped DNA probes to be rapidly assigned to a subset of co-migrating chromosomes (A. Day, personal communictation). Linkage group XIX, or UNI (for the uniflagellate mutation uni1), was defined by Ramanis and Luck (1986), who reported genetic evidence suggesting that it might be circular rather than linear, as well as novel effects of temperature during zygospore maturation on recombination frequency. As first described, this linkage group contained only markers relating to motility, but subsequently mutations with no apparent flagellar association have been found linked to other loci on this group (Dutcher et al., 1992). Hall et al. (1989) reported that molecular probes for the UNI linkage group hybridized to basal bodies and concluded that the UNI linkage group corresponds to basal body DNA. This report was challenged by K. A. Johnson and Rosenbaum (1990), who did not find immunologically detectable DNA associated with basal bodies; by D. E. Johnson and Dutcher (1991), who found that DNA sequences from this linkage group were present in the same copy number per cell as other nuclear linkage groups in both haploid and diploid strains, and in mutants lacking basal bodies; and by Holmes et al. (1993) who obtained a linear genetic map in multifactor crosses. Hall and Luck (1995) subsequently confirmed the linearity of the genetic map and the stoichiometry reported by Johnson and Dutcher, but based on in situ hybridization data, they believe that DNA of this linkage group does occupy a unique position in interphase cells specifically at the anterior edge of the nucleus proximal to the basal body complex.

VI. Molecular Biology The past ten years have seen a virtual revolution in the style of Chlamydomonas research, and the types

7 of studies that are feasible, entirely because of advances in techniques for molecular analysis and genetic engineering. Successful introduction of exogenous DNA into Chlamydomonas chloroplasts by biolistic bombardment (Boynton et al., 1988) was soon followed by application of this procedure to nuclear genes (Debuchy et al., 1989; Kindle et al., 1989), and then by a simple procedure based on agitation of cells in the presence of DNA and glass beads (Kindle, 1990) that made transformation feasible in any laboratory. High efficiency trans­ formation by electroporation has recently been reported by Shimogawara et al. (1998). Kindle (Chapter 4) and Goldschmidt-Clermont (Chapter 8) have reviewed transformation technology for nuclear and chloroplast genes, respectively in this volume. Whereas DNA introduced into the chloroplast genome usually integrates at homologous sites, in the nuclear genome integration is more often by non­ homologous recombination. Tam and Lefebvre (1993) turned this seeming difficulty into advantage by using transformation as a means of generating tagged mutants by insertional mutagenesis, and their technique has now been used by many laboratories to identify a diverse collection of genes (see Kindle, Chapter 4, for details). Expression of foreign genes in Chlamydomonas has also been a stumbling block, owing perhaps to a combination of codon bias, gene silencing, and perhaps other factors, but this obstacle too now is being surmounted (Hall et al., 1993; Sizova et al., 1996; Stevens et al., 1996;Cerutti et al., 1997a,b). With solution ofthese problems specific to Chlamydomonas, application of general techniques of molecular biology to algal research becomes straightforward. The results will be apparent throughout this volume.

VII. Resources Historical background, descriptions of mutants, and many methods are available in The Chlamydomonas Sourcebook (Harris, 1989). The Chlamydomonas Genetics Center maintains a collection of wild type and mutant strains of C. reinhardtii, C. moewusii and C. eugametos, as well as bacterial cultures carrying plasmids with inserts of Chlamydomonas DNA (Chlamydomonas Genetics Center, c/o Dr. Elizabeth H. Harris, DCMB Box 91000, Duke University, Durham NC 27708-1000, U.S.A.; e-mail chlamy@ duke.edu; phone 919-613-8164; fax 919-613-8177).

8 The Center also maintains a web page (http:// www.botany.duke.edu/DCMB/chlamy.htm) with methods files, announcements, and links to pages of many laboratories engaged in Chlamydomonas research, and a database that is part of the Plant Genome project of the U.S. National Agricultural Library (http://probe.nalusda.gov:8300/cgi-bin/ browse/chlamydb). Cultures of other species of Chlamydomonas can be obtained from several major algal collections, i n c l u d i n g UTEX (University of Texas Algal Collection, Department of Botany, Austin TX 78713­ 7640, U.S.A.; see also Starr and Zeikus, 1993), CCAP (Culture Centre of Algae and Protozoa, Freshwater Biology Association, The Ferry House, Ambleside, Cumbria LA22 OLP, U.K.), SAG (Sammlung von Algenkulturen, Pflanzenphysio­ logisches Institut, Universität Göttingen, Nikolaus­ berger Weg 18, D-3400 Göttingen, Germany; see also Schlösser, 1994), and IAM (Institute of Applied Microbiology, The University of Tokyo, 1-1-1 Yayoi, Bunkyou-ku, Tokyo 113, Japan). A few Chlamy­ domonas cultures are also maintained by the American Type Culture Collection (12301 Parklawn Drive, Rockville MD 20852, U.S.A.). Several specialized collections also have significant numbers of Chlamydomonas accessions: Peterhof Chlamy­ domonas Collection (Genetics Department, St. Petersburg State University, St. Petersburg V 164, 199164, Russia); Culture Collection of Autotrophic Organisms (Institute of Botany, Czech Academy of Sciences, Dukelska 145, CS-379 82 Trebon, Czech Republic); Culture Collection of Microalgae (K. A. Timiryazev Institute of Plant Physiology, Russian Academy of Sciences, 35 Botanicheskaya Street, Moscow, 127276, Russia), and the University of Toronto Culture Collection (Department of Botany, University of Toronto, Toronto, Ontario M5S 3B2, Canada).

Acknowledgment The Chlamydomonas Genetics Center is supported by NSF Grant DBI-9319941.

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9 Goto K and Johnson CH (1995) Is the cell division cycle gated by a circadian clock? The case of Chlamydomonas reinhardtii. J Cell Biol 129: 1061–1069. Gowans CS (1963) The conspccificity of Chlamydomonas eugametos and Chlamydomonas moewusii: An experimental approach. Phycologia 3: 37–44 Gross CH, Ranum LPW and Lefebvre PA (1988) Extensive restriction fragment length polymorphisms in a new isolate of Chlamydomonas reinhardtii. Curr Genet 13: 503–508 Hails T. Jobling M and Day A (1993) Large arrays of tandemly repeated DNA sequences in the green alga Chlamydomonas reinhardtii. Chromosoma 102: 500–507 Hall JL and Luck D (1995) Basal body-associated DNA: In situ studies in Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 92: 5129–5133 Hall JL, Ramanis Z and Luck DJL (1989) Basal body/centriolar DNA: Molecular genetic studies in Chlamydomonas. Cell 59: 121–132 Hall LM, Taylor KB and Jones DD (1993) Expression of a foreign gene in Chlamydomonas reinhardtii. Gene 124: 75–81 Harper J, Wu L, Sakuanrungsirikul S and John P (1995) Isolation and partial characterization of conditional cell division cycle mutants in Chlamydomonas. Protoplasma 186: 149–162 Harris EH (1989) The Chlamydomonas Sourcebook. A Comprehensive Guide to Biology and Laboratory Use. Academic Press, San Diego Harris EH (1993) Chlamydomonas reinhardtii. In: O’Brien SJ (ed) Genetic Maps. Locus Maps of Complex Genomes, 6th edition, pp 2.157–2.169. Cold Spring Harbor Laboratory, Cold Spring Harbor Harris EH, Boynton JE, Gillham NW, Burkhart BD and Newman SM (1991) Chloroplast genome organization in Chlamy­ domonas. Arch Protistenk 139: 183–192 Hegemann P (1997) Vision in microalgae. Planta 203:265–274 Holland EM, Braun FJ, Nonnengaesser C, Harz H and Hegemann P (1996) Nature of rhodopsin-triggered photocurrents in Chlamydomonas .1. Kinetics and influence of divalent ions. Biophys J 70: 924–931 Holland EM, Harz H, Uhl R and Hegemann P (1997) Control of phobic behavioral responses by rhodopsin-induced photo­ currents in Chlamydomonas. Biophys J 73: 1395–1401. Holmes JA, Johnson DE and Dutcher SK (1993) Linkage group XIX of Chlamydomonas reinhardtii has a linear map. Genetics 133: 865–874 Hoshaw RW (1965) Mating types of Chlamydomonas from the collection of Gilbert M. Smith. J Phycol 1: 194–196 Hoshaw RW and Ettl H (1966) Chlamydomonas smithii sp. nov.—a Chlamydomonad interfertile with Chlamydomonas reinhardtii. J Phycol 2: 93–96 Howell SH and Naliboff JA (1973) Conditional mutants in Chlamydomonas reinhardtii blocked in the vegetative cell cycle. I. An analysis of cell cycle block points. J Cell Biol 57: 760–772 Hutner SH and Provasoli L (1951) The phytoflagellates. In: Lwoff A (ed) Biochemistry and Physiology of Protozoa, Vol 1, pp 27–128. Academic Press, New York Hwang S, Kawazoe R and Herrin DL (1996) Transcription of tufA and other chloroplast-encoded genes is controlled by a circadian clock in Chlamydomonas. Proc Natl Acad Sci USA 93: 996–1000 Jacobshagen S, Kindle KL and Johnson CH (1996) Transcription

10 of CABII is regulated by the biological clock in Chlamydomonas reinhardtii. Plant Mol Biol 31: 1173–1184 J a r v i k JW and Suhan JP (1991) The role of the flagellar transition region: Inferences from the analysis of a Chlamydomonas mutant with defective transition region structures. J Cell Sci 99: 731–740 Johnson DE and Dutcher SK (1991) Molecular studies of linkage group XIX of Chlamydomonas reinhardtii: Evidence against a basal body location. J Cell Biol 113: 339–346 Johnson KA (1995) Keeping the beat: Form meets function in the Chlamydomonas flagellum. BioEssays 17: 847–854 Johnson KA and Rosenbaum JL (1990) The basal bodies of Chlamydomonas reinhardtii do not contain immunologically detectable DNA. Cell 62: 339–346 Johnson KA and Rosenbaum JL (1992) Replication of basal bodies and centrioles. Curr Opin Cell Biol 4: 80–85 Johnson KA and Rosenbaum JL(1993) Flagellar regeneration in Chlamydomonas: A model system for studying organelle assembly. Trends Cell Biol 3: 156–161 Kindle KL (1990) High-frequency nuclear transformation of Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 87: 1228–1232 Kindle KL, Schnell RA, Fernández E and Lefebvre PA (1989) Stable nuclear transformation of Chlamydomonas using the Chlamydomonas gene for nitrate reductase. J Cell Biol 109: 2589–2601 Kreimer G (1994) Cell biology of phototaxis in flagellate algae. Int Rev Cytol 148: 229–310 Kroeger P and Hegemann P (1994) Photophobic responses and phototaxis in Chlamydomonas arc triggered by a single rhodopsin photoreceptor. FEBS Lett 341: 5–9 Kurvari V (1997) Cell wall biogenesis in Chlamydomonas: Molecular characterization ofa novel protein whose expression is up-regulated during matrix formation. Mol Gen Genet 256: 572–580 Lawson MA and Satir P (1994) Characterization of the eyespot regions of ‘blind’ Chlamydomonas mutants after restoration of photophobic responses. J Euk Microbiol 41: 593–601 Lechtreck KF and Melkonian M (1991) An update on fibrous flagellar roots in green algae. Protoplasma 164: 38–44 Lechtreck KF and Silflow CD (1997) SF-assemblin in Chlamydomonas: Sequence conservation and localization during the cell cycle. Cell Motil Cytoskeleton 36: 190–201 Lemieux B, Tunnel M and Lemieux C (1985) Chloroplast DNA variation in Chlamydomonas and its potential application to the systematics of this genus. BioSystems 18: 293–298 Levine RP (1969) The analysis of photosynthesis using mutant strains of algae and higher plants. Annu Rev Plant Physiol 20: 523–540 L e v i n e RP and Goodenough UW (1970) The genetics of p h o t o s y n t h e s i s and the chloroplast in Chlamydomonas reinhardi. Annu Rev Genet 4: 397–408 Lewin RA (1949) Genetics of Chlamydomonas—paving the way. Biol Bull 97: 243–244 Lewin RA (1951) Isolation of sexual strains of Chlamydomonas. J Gen Microbiol 5: 926–929 Lewin RA (1952) Ultraviolet induced mutations in Chlamy­ domonas moewusii Gerloff. J Gen Microbiol 6: 233–248 L e w i n RA (1953) The genetics of Chlamydomonas moewusii Gerloff. J Genet 51: 543–560 Lewin RA (1954) Mutants of Chlamydomonas moewusii with

Elizabeth H. Harris impaired motility. J Gen Microbiol 11: 358–363 Liss M, Kirk DL, Beyser K and Fabry S (1997) Intron sequences provide a tool for high-resolution phylogenetic analysis of volvocine algae. Curr Genet 31: 214–227 Luykx P, Hoppenrath M, Robinson DG (1997) Structure and behavior of contractile vacuoles in Chlamydomonas reinhardtii. Protoplasma 198: 73–84. Matsuda Y (1988) The Chlamydomonas cell walls and their degrading enzymes. Japan J Phycol 36: 246–264 Matsuda Y, Shimada T and Sakamoto Y (1992) Ammonium ions control gametic differentiation and dedifferentiation in Chlamydomonas reinhardtii. Plant Cell Physiol 33: 909–914 Musgrave A (1993) Mating in Chlamydomonas. Prog Phycol Res 9: 193–237 Nonnengaesser C, Holland EM, Harz H and Hegemann P (1996) The nature of rhodopsin-triggered photocurrents in Chlamy­ domonas .2. Influence of m o n o v a l e n t i o n s . Biophys J 70: 932– 938 Pan JM, Having MA and Beck CF (1997) Characterization of blue light signal transduction chains that control development and maintenance of sexual competence in Chlamydomonas reinhardtii. Plant Physiol 1 1 5 : 1241–1249 Pazour GJ, Sineshchekov OA and Witman GB( 1995) Mutational analysis of the phototransduction pathway of Chlamydomonas reinhardtii. J Cell Biol 131: 427–440. Porter ME (1996) Axonemal dyneins: Assembly, organization, and regulation. Curr Opin Cell Biol 8: 10–17 Porter ME, Knott JA, Myster SH and Farlow SJ (1996) The dynein gene family in Chlamydomonas reinhardtii. Genetics 144: 569–585 Pringsheim EG (1937) Beiträge zur Physiologie saprotropher Algen und Flagellaten. 3. Mitteilung: Die Stellung der Azetatflagellaten in einem physiologischen Ernährungssystem. Planta 27: 61–72 Quarmby LM (1994) Signal transduction in the sexual life of Chlamydomonas. Plant Mol Biol 26: 1271–1287 Ramanis Z and Luck DJL( 1986) Loci affecting flagellar assembly and function map to an unusual linkage group in Chlamy­ domonas reinhardtii. Proc Natl Acad Sci USA 83: 423–426 Ranum LPW, Thompson MD, Schloss JA, Lefebvre PA and Silflow CD (1988) Mapping flagellar genes in Chlamydomonas using restriction fragment length polymorphisms. Genetics 120: 109–122 Rochaix J - D ( 1 9 9 5 ) Chlamydomonas reinhardtii as the photosynthetic yeast. Annu Rev Genet 29: 209–230 Sack L, Zeyl C, Bell G, Sharbel T, Reboud X, Bernhardt T and K o e l e w y n H ( 1 9 9 4 ) I s o l a t i o n of four new strains of Chlamydomonas reinhardtii (Chlorophyta) from soil samples. J Phycol 30: 770–773 Sager R (1954) Mendelian and non-Mendelian inheritance of streptomycin resistance in Chlamydomonas reinhardi. Proc Natl Acad Sci USA 40: 356–363 Sager R (1955) Inheritance in the green alga Chlamydomonas reinhardi. Genetics 40: 476–489 Sager R and Granick S (1953) N u t r i t i o n a l studies with Chlamydomonas reinhardi. Ann New York Acad Sci 56: 831– 838 Sager R and Granick S (1954) Nutritional control of sexuality in Chlamydomonas reinhardi. J Gen Physiol 37: 729–742 Sager R and Palade GE (1954) Chloroplast structure in green and yellow strains of Chlamydomonas. Exp Cell Res 7: 584–588

Chapter 1

Introduction to Chlamydomonas

Salisbury JL (1995) Centrin, centrosomes, and mitotic spindle poles. Curr Opin Cell Biol 7: 39–45 Savard F, Richard C and Guertin M (1996) The Chlamydomonas reinhardtii LI818 gene represents a distant relative of the cabI/ II genes that is regulated during the cell cycle and in response to i l l u m i n a t i o n . Plant Mol Biol 32: 461–473 Schlösser UG (1994) SAG—Sammlung von Algenkulturen at the University of Göttingen. Bot Acta 107: 111–186 Schlösser UG, Sachs H and Robinson DG (1976) Isolation of protoplasts by means of a ‘species-specific’ autolysine in Chlamydomonas. Protoplasma 88: 51–64. Shimogawara K, Fujiwara S, Grossman A and Usuda H (1998) High efficiency transformation of Chlamydomonas reinhardtii by electroporation. Genetics, in press Sizova I A , Lapina TV, Frolova ON, Alexandrova NN, Akopiants KE and Danilenko VN (1996) Stable nuclear transformation of Chlamydomonas reinhardtii with a Streptomyces rimosus gene as the selective marker. Gene 181: 13–18 Smith GM (1946) The nature of sexuality in Chlamydomonas. Amer J Bot 33: 625–630 Smith EF and Lefebvre PA (1997) The role of central apparatus components in flagellar motility and microtubule assembly. Cell Motil Cytoskeletonj 38: 1–8 Smith GM and Regnery DC (1950) Inheritance of sexuality in Chlamydomonas reinhardii. Proc Natl Acad Sci USA 36: 246– 248 Spanier JG, Graham JE and Jarvik JW (1992) Isolation and preliminary characterisation of three Chlamydomonas strains interfertile with Chlamydomonas reinhardtii (Chlorophyta). J Phycol 28: 822–828 Spessert R and Waffenschmidt S (1990) Studies on the vegetative autolysin during the vegetative life cycle in Chlamydomonas reinhardtii. Eur J Cell Biol 51: 17–22 Spreitzer RJ and Mets L (1981) Photosynthesis-deficient mutants of Chlamydomonas reinhardii with associated light-sensitive phenotypes. Plant Physiol 67: 565–569 Starr RC and Zeikus JA (1993) UTEX—the culture collection of algae at the University of Texas at Austin. J Phycol 29 suppl: 1–106 Stevens DR, Rochaix JD and Purton S (1996) The bacterial phleomycin resistance gene ble as a dominant selectable marker in Chlamydomonas. Mol Gen Genet 251: 23–30 Taillon BE, Adler SA, Suhan JP and Jarvik JW(1992) Mutational analysis of centrin: An EF-hand protein associated with three distinct contractile fibers in the basal body apparatus of Chlamydomonas. J Cell Biol 119: 1613–1624 Tam LW and Lefebvre PA (1995) Insertional mutagenesis and isolation of tagged genes in Chlamydomonas. Methods Cell Biol 47: 519–523 van den Ende H (1994) Vegetative and gametic development in the green alga Chlamydomonas. Adv Bot Res 20: 125–161 van den Ende H (1995) Sexual development in the homothallic

11 green alga Chlamydomonas monoica Strehlow. Sexual Plant Reprod 8: 139–142 Van W i n k l e - S w i f t KP and Aubert B ( 1 9 8 3 ) Uniparental inheritance in a homothallic alga. Nature 303: 167–169 VanWinkle-Swift KP and Hahn JH (1986) The search for matingtype-limited genes in the homothallic alga Chlamydomonas monoica. Genetics 1 1 3 : 601–619 Voigt J, Hinkelmann B, Liebich I and Mix M (1996) Alteration of the cell surface during the vegetative cell cycle of the unicellular green alga Chlamydomonas reinhardtii. Plant Cell Physiol 37: 726–733 Voigt J, Hinkelmann B and Harris EH (1997) Production of cell wall polypeptides by different cell wall mutants of the unicellular green alga Chlamydomonas reinhardtii. Microbiol Res 152: 189–198 Waffenschmidt S, Woessner JP, Beer K and Goodenough UW (1993) Isodityrosine cross-linking mediates insolubilization of cell walls in Chlamydomonas. Plant Cell 5: 809–820 Walne PL and Gualtieri P (1994) Algal visual proteins: An evolutionary point of view. Crit Rev Plant Sci 13: 185–197 Weger HG (1996) Interactions between respiration and inorganic phosphate uptake in phosphate-limited cells of Chlamydomonas reinhardtii. Physiol Plant 97: 635–642 Weger HG, Chadderton AR, Lin M, Guy RD and Turpin DH (1990) Cytochrome and alternative pathway respiration during transient ammonium assimilation by N-limited Chlamy­ domonas reinhardtii. Plant Physiol 94: 1131–1136 Wilson NF, Foglesong MJ, Snell WJ (1997) The Chlamydomonas mating type plus fertilization tubule, a prototypic cell fusion organelle: Isolation, characterization, and in vitro adhesion to mating type minus gametes. J Cell Biol 137: 1537–1553 Witman GB (1993) Chlamydomonas phototaxis. Trends Cell Biol 3: 403–408 Woessner JP and Goodenough UW (1989) Molecular charac­ terization of a zygote wall protein: An extensin-like molecule in Chlamydomonas reinhardtii. Plant Cell 1: 901–911 Woessner JP and Goodenough UW (1992) Zygote and vegetative cell wall proteins in Chlamydomonas reinhardtii share a common epitope, (SerPro)x. Plant Sci 83: 65–76. Woessner JP and Goodenough UW (1994) Volvocine cell walls and t h e i r c o n s t i t u e n t g l y c o p r o t e i n s : An e v o l u t i o n a r y perspective. Protoplasma 181: 245–258 Woessner JP, Molendijk AJ, Van Egmond P, Klis FM, Goodenough UW and Haring MA (1994) Domain conservation in several volvocalean cell wall proteins. Plant Mol Biol 26: 947–960 Wu LP, Hepler PK, John PCL (1997) The met 1 mutation in Chlamydomonas reinhardtii causes arrest at mitotic metaphase with persisting p34cdc2-like H1 histone kinase activity that can promote mitosis when injected into higher-plant cells. Protoplasma 199: 135–150

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Chapter 2

Perspectives on Early Research on Photosynthesis in Chlamydomonas Robert K. Togasaki and Stefan J. Surzycki Department of Biology, Indiana University, Bloomington, Indiana 47405, U.S.A.

Summary I. General Background II. The Levine Laboratory in the Early 1960s III. Establishment of Chlamydomonas reinhardtii as a Legitimate Model Organism IV. Development of New Techniques V. Emergence of New Research Targets VI. Old Experiments Becoming Reality Acknowledgment References

13 13 14 15 18 19 21 21 22

Summary Personal perspectives of the authors on research conducted in the laboratory of R. P. Levine at the Biological Laboratories, Harvard University in the latter half of the 1960s to the early 1970s is described. The chapter summarizes the state ofresearch in photosynthesis and chloroplast biology in the early 1960s. The authors recall experiments and events that led to the establishment of Chlamydomonas reinhardtii as a model organism for the study of photosynthesis and the molecular biology of chloroplasts. These reminiscences include personal anecdotes that try to convey the excitement, and elation and disappointments, that were experienced during these pioneering times in Chlamydomonas research. I. General Background As of 1964, the field of photosynthesis had several dominant topics. Most important among them were photosynthetic electron transport, photophos­ phorylation, and photosynthetic carbon assimilation. The precise measurement of photosynthetic activities in micro algae, pioneered by Warburg, led to the discovery of the red drop phenomenon by Emerson and Lewis (Emerson and Lewis, 1943). This was followed by the discovery of the enhancement phenomenon by Emerson’s group (Emerson et al., 1957), and Hill and Bendall’s proposal of two photosystems, the Z scheme, for photoelectron transport (Hill and Bendall, 1960). Subsequently the

Z scheme was elegantly corroborated by Duysen’s work on red algae (Duysens et al., 1961). Thus, the foundation for today’s generally accepted model of photosynthetic electron transport in photosynthetic eukaryotes and cyanobacteria was well established by the mid 1960’s. Furthermore, electron transport, using water as the electron donor and as the electron acceptor, was demonstrated in vitro by work in San Pietro’s laboratory at Johns Hopkins (San Pietro and Lang, 1957). Photophosphorylation in isolated chloroplast fragments was achieved by Arno’s group at Berkeley (Arnon et al., 1954). In addition, in the 1960s, work on the chemiosmotic basis of ATP synthesis by thylakoid membranes was initiated by

J. -D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 13–23. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

14 Jagendorf and coworkers in their pioneering paper ( H i n d and Jagendorf, 1963). In the area of photosynthetic carbon metabolism, the reductive photosynthetic carbon cycle was well established by mid-1950 (Bassham, 1957), but analysis of photosynthesis was just beginning with Kortschak’s paper (Kortschak et al., 1965). This was to be followed later by the elucidation ofthe carbon assimilation pathway by Hatch’s and Slack’s group (see Edwards and Walker, 1983). At this time, research with intact organisms was conducted mainly with cells of the unicellular green algae Chlorella, Scenedesmus, the cyanobacterium Anacystis and Euglena. The measurements made were either physical measurements, such as or gas exchange, or the fluorescence and biochemical analysis of fixation products. At the subcellular level, thylakoid membranes were isolated from plant leaves, mainly spinach and peas, for the analysis of photosynthetic electron transport and photophosphorylation. Chloroplasts with intact envelopes were also isolated successfully from these leaves and used extensively in studies of photo­ synthetic carbon metabolism (for review, see Edwards and Walker, 1983). Many plants and unicellular organisms were analyzed by the application of both biophysical and biochemical techniques, with a focus on the components of the photosynthetic apparatus and their function. The discovery of one component of the photosynthetic machinery in one organism was quickly extended to other systems in a quest to find all common components of the photosynthetic apparatus. For example, the discovery of plastocyanin in Chlorella was extended rapidly to other plant systems (Katoh et al., 1961) and its position in the electron transport chain was incorporated into most models for eukaryotic and cyanobacterial photo­ synthesis. In general, the emphasis was on refinement and exploitation of biophysical and biochemical research tools, better spectrophotometers, fluor­ imeters, and more specific chemical inhibitors. The idea that the experimental organism itself can offer a powerful research advantage remained dormant. Among these early studies, however, Duysens’ study of photosynthetic electron transport in Abbreviations: DCMU – 3-(3,4-dichlorophenyl) 1,1 -dimethyl­ urea; DC IP –2,6-dichloroindophenol; and – components of chloroplast ATP synthase; PMS – phenazine methosulfate; PS I – Photosystem I; PS II – Photosystem II; Rubisco – ribulose1,5-biphosphate carboxylase/oxygenase; TAP – trisacetate-phosphate

Robert K. Togasaki and Stefan J. Surzycki Porphyridium cruentum stands out as an example of superb exploitation of a biological trait, unique to a given experimental material (Duysens et al., 1961). In this red alga, light absorbed by an accessory pigment, phycoerythrin (563 nm), drives PS II while light absorbed by chlorophyll a (680 nm) drives PS I activities exclusively. Thus, Duysens could clearly demonstrate photoreduction and photooxidation of cytochrome f by PS II and PS I, respectively. This was an exceptional case; the more common approach was to use ever more powerful biochemical and biophysical tools on the usual experimental organisms. In an alternative approach, the alteration of the genetic constitution and hence the phenotype of a photosynthetic organism became a major research tool. This approach, pioneered by R. P. Levine with Chlamydomonas reinhardtii and N. Bishop with Scenedesmus obliquus (see Bishop, 1973), used the paradigm ofTatum and Beadle’s work of elucidating the order of enzymatic reactions in biochemical pathways through the study of mutants. It was reasoned that the best way to discover all of the components of the photosynthetic apparatus, as well as the mode and place of their action, would be to create mutants that affectphotosynthesis. One should marvel on the farsightedness of this approach at that time, realizing that it is still in use thirty years after its introduction into photosynthetic research. The analysis of photosynthetic mutants, using established experimental procedures began to yield new information by the early 1960s. At that time, the S. obliquus system was already yielding much new information whereas research on photosynthesis with C. reinhardtii was only beginning. However, the potential of C. reinhardtii as a model organism to study photosynthesis appeared much greater than for S. obliquus because of the possibility of genetic analysis. This possibility led one of us (R. Togasaki) to choose Levine’s laboratory over Bishop’s for postdoctoral studies.

II.The Levine Laboratory in the Early 1960s R. P. Levine began to use Chlamydomonas for his research by focusing on the isolation and charac­ terization of amino acid auxotrophs, as well as by studying genetic recombination mechanisms (Levine and Ebersold, 1958). By the early 1960s, the main focus of the laboratory shifted to the genetics and

Chapter 2 Perspective on Early Research biochemistry of photosynthesis. It is our under­ standing that the very low yield of amino acid auxotrophs and the high yield of acetate auxotrophs during early mutant screening was one factor for this change. The other was Levine’s association with Bob Smillie at Brookhaven National Laboratory and their collaborative work on photosynthesis (Levine and Smillie, 1962). In 1962, while Levine was away in France, N. Gillham, G. Hudock and G. Russell in Levine’s laboratory carried out a large scale isolation of acetate auxotrophs (Hudock, personal commun­ ication). They characterized some of these mutants as being deficient in photosynthesis by using an autoradiography screen developed earlier by Levine (Levine, 1960). Thus, by the fall of 1964, several papers on the photosynthetic electron transport system and photophosphorylation in Chlamydomonas had been published (for review, see Harris, 1988). In the fall of 1964, after Levine returned from France, three new postdoctoral fellows joined his laboratory, R. K. Togasaki, S.J. Surzycki and J.P. Hastings. Robert Togasaki arrived from the laboratory ofM. Gibbs where he hadjust finished his dissertation work on photosynthetic carbon metabolism in micro algae. He intended to study potential photosynthetic carbon cycle mutants in C. reinhardtii. Stefan Surzycki came from Gajewski’s laboratory in Warsaw where he was studying the mechanism of meiotic gene conversion in Ascobolus immersus. He, in turn, wanted to study the molecular basis of recombination using Chlamydomonas as an experimental system. Philip Hastings came from the Whitehouse laboratory at Cambridge University, England where he worked on the formulation of the molecular theory ofgenetic recombination. During their stay, they witnessed and participated in the evolution and development of approaches to study the function (photosynthesis) and structure (molecular biology) of the chloroplast in Chlamydomonas.

III. Establishment of Chlamydomonas reinhardtii as a Legitimate Model Organism During the 1960s, C. reinhardtii had to earn its citizenship in the community of photosynthetic research model organisms. The green algae Chlorella and Scenedesmus, the cyanobactenum Anacystis, as well as Euglena were dominant model organisms for biochemical, biophysical and physiological analysis of photosynthesis at the level of intact cells. Spinach

15 and peas were, in turn, favored material for in vitro studies using isolated, intact chloroplasts or thylakoid membranes. A researcher using C. reinhardtii had to face the dual task of establishing reliable experi­ mental procedures for this organism, and of demonstrating the unique advantage ofthis organism for the study of photosynthetic and molecular processes. By 1964, much of the biochemical and biophysical characterization of the photosynthetic apparatus had been accomplished, including the determination of the sequence ofevents from the photophysics of light absorption to the biochemical events of carbon assimilation. The notable exception was the oxygen evolving mechanism in PS II. Thus, much of early photosynthesis research activity in Chlamydomonas involved the isolation and characterization ofmutants deficient in photosynthesis. The aim of this research was to localize the site of the biochemical lesion in the photosynthetic apparatus caused by these mutations. Mutants lacking PS II and PS I activities, or photophosphorylation activity, and those lacking a component of photosynthetic electron transport, such as plastocyanin or cytochrome f, were discovered and characterized (see Chapter 31, Merchant). Figure 1 presents a diagram of the electron transport system and shows the positions of components affected in mutants isolated in Levine’s laboratory. Table 1 lists properties ofthese mutants. The study of these Chlamydomonas mutants confirmed, at first, the findings of published biochemical research. In addition, their genetic mapping provided the genetic basis for understanding the biochemical data and the organization of genes encoding photosynthetic components (Hastings et al., 1965). However, very soon, the mutant approach yielded new and unexpected results. The first major demonstration of an advantage of the Chlamydomonas system came with the unequiv­ ocal demonstration of the positions of cytochrome f and plastocyanin in the electron transport chain (Gorman and Levine, 1966b). At that time, the copperchelating agent, salicylaldoxime, was shown to inhibit photoreduction of cytochrome f. Since plastocyanin is a copper-containing protein, the results implied that plastocyanin is located before cytochrome f in the electron transport chain (Fork and Urbach, 1965). Gorman and Levine used two mutants in their study, one lacking cytochrome f activity (ac-206), and the other, missing plastocyanin (ac-208). In the absence of plastocyanin (ac-208), cytochrome f was

16

photoreduced by PS II but not photooxidized by PS I. The absence of cytochrome f in ac-206 did not prevent photooxidation of the exogenously added reductant (ascorbate and DCIP) by PS I, while the absence of plastocyanin did. These data showed that cytochrome f preceded plastocyanin in the electron transport chain. The Gorman and Levine experiments demonstrated the usefulness of mutants in photo­

Robert K. Togasaki and Stefan J. Surzycki

synthetic research and illustrated the danger of artifacts that can arise from the use of chemical inhibitors in in vitro experiments. Gorman and Levine’s conclusions were later confirmed for spinach chloroplasts (Kimimura and Katoh, 1972). Moreover, this research was also an early example of data from Chlamydomonas being confirmed by later bio­ chemical work and not the other way around. Robert

Chapter 2 Perspective on Early Research Togasaki recalls the following episode. A very prominent researcher in the photosynthetic electron transport field was visiting the Levine laboratory during the time when work with mutants ac-208 and ac-206 was in progress. He was shown the latest data on the relative positions of cytochrome f and plastocyanin in the electron transport chain. His comment was, ‘Well, is that how you want to order them? But, most other people think otherwise!’ This was symptomatic of the time when the power of the genetic approach was not fully appreciated by established researchers in the field. Another important mutant isolated during this time was ac-21. This mutation identified a new essential component, called M, in the photosynthetic electron transport chain located between PS II and cytochrome f (Levine and Smillie, 1962; Gorman and Levine 1966a; Levine 1968). Many years later it was found that M is the Rieske iron-sulfur protein (Bendall et al., 1986). One advantage ofworking in a new field is thatjust about anything you do or find is novel. Eventually many of these findings pioneered new fields of research. Some of the early doctoral dissertations from Levine’s laboratory centered on the isolation and characterization of a single mutant affecting a key component ofthe photosynthetic apparatus. These were: The isolation and characterization of the first mutant defective in PS I function, ac-80 (Givan and Levine, 1967). A. Givan showed that ac-80 did not display the absorbance change associated with P700. The mutant could not reduce with electrons from either water or an artificial donor (the dye DCIP and ascorbate), but it had considerable Hill activity. The cells could not carry out cyclic photophosphorylation but were capable still ofnoncyclic photophosphorylation coupled to the reduction of ferricyanide. These experiments allowed them to conclude that one coupling site is situated between the two photosystems. Analysis of a 520 nm light-induced absorbance change (Chua and Levine, 1969). N. -H. Chua studied the 520 nm light-induced absorbance change in wild type and in mutants deficient in electron transport to analyze the function of quinones in the electron transport system.

17 Effects ofmanganese on photosynthetic activities and chloroplast structure of C. reinhardtii (Teichler-Zallen and Levine, 1969). D. TeichlerZallen found that Mn deficiency led to severe reduction of PS II activity but did not affect PS I activity. The extent of thylakoid membrane stacking decreased in parallel to the reduction of PS II activity. This established a significant role for Mn in the structure and function ofthylakoid membranes. Isolation and characterization ofthe first mutant lacking phosphoribulokinase activity with near wild type levels of several other Calvin cycle enzymes, F-60 (Moll and Levine, 1970). B. Moll provided the first clear genetic evidence to support the Calvin cycle. Interestingly, in spite of extensive searching, this mutant remained the only photosynthetic carbon cycle mutant until the isolation of Rubisco mutants (Spreitzer and Mets, 1980; Chapter 27, Spreitzer). The phosphoribulokinase mutation eventually was used in many research projects including studies of fermentation and photohydrogen evolution and the hydrogenase system by Gibbs and coworkers (for review, see Togasaki and Whitmarsh, 1983). Isolation and characterization of the first mutant deficient in photophosphorylation that lacked coupling factor, F-54 (Sato et al., 1971). V. Sato provided the first genetic evidence, linking the absence of thylakoid membrane surface particles, the lack of ATPase, and photophosphorylation activity to a single mutation. Analysis of the structure of chloroplast DNA using electron microscopy (Rochaix, 1972). This work was one of the first attempts to create a physical map of organelle DNA. Excitation energy transfer and chlorophyll orientation was studied in intact cells of Chlamydomonas (Whitmarsh and Levine, 1974). Fluorescence polarization for intact cells was measured with horizontally polarized exciting light, in the presence and absence of DCMU to test the validity of the Förster mechanism of excitation energy transfer.

18 The pioneering work on chloroplast genetics and biology of Chlamydomonas was not restricted to the Levine’s laboratory at this time. Ruth Sager and Nicholas Gillham performed independently an extensive genetic analysis of uniparental inheritance in Chlamydomonas (Sager, 1960; Gillham, 1965). Laurie Mets and Laurie Bogorad began the analysis of the chloroplast translational apparatus, studying erythromycin binding by plastid ribosomes (Mets and Bogorad, 1971).

IV. Development of New Techniques Since C. reinhardtii was a new model organism, most of the experimental techniques had to be modified or developed de novo. A few examples given below will demonstrate the effort ofthis task in the early period of working with this alga in Levine’s laboratory. In 1960, R. P. Levine developed an effective screening method to identify mutants deficient in photosynthesis (Levine, 1960a). Mutagenized cells were plated on acetate supplemented agar plates, and allowed to develop colonies that, in turn, were replicated to minimal media plates to screen for acetate auxotrophs. Acetate auxotrophic colonies were replica plated and allowed to carry out assimilation in the light. The incorporation was stopped by exposure to acid, and colonies were incorporation by autoradiography. analyzed for This double screen, involving both acetate auxotrophy and autoradiography, was labor intensive and lacked a positive selection for mutants. However, this test was the least biased mutant isolation procedure developed to date. It is no wonder, therefore, that this test yielded many different photosynthetic mutants affecting electron transport, photophosphorylation and carbon metabolism. All of the early acetate requiring mutants isolated in Levine’s laboratory were obtained using this procedure. In the summer of 1966, P. Bennoun visited the Levine laboratory and developed a rapid screen for the isolation of photosynthetic mutants. This procedure was based on the fluorescent light output ofindividual colonies (Bennoun, 1967). Mutagenized cells were plated on acetate containing plates (TAP), allowed to form colonies, irradiated with blue light and photographed through a red filter using a red and infrared sensitive Polaroid film. Colonies of the mutants with defects localized after PS II gave high

Robert K. Togasaki and Stefan J. Surzycki fluorescence and could be easily differentiated from wild type colonies with normal fluorescence. While a positive selection for mutants deficient in photosynthesis was still absent in this procedure, the simplicity and rapidity ofthis detection method more than made up for it. The development of this elegant procedure literally began with a big bang! In the very first experiment, the absence of a heat filter between the Zeiss microscope lamp and the primary blue glass filter caused the expensive glass filter to overheat and shatter to pieces. The new method permitted Bennoun, Chua, and Togasaki, to isolate and partially characterize a large number of photosynthesis negative mutants in short time. A similar method was published by Garnier’s group (Garnier, 1967). This non-invasive screening method for photosynthetic mutants was eventually adopted for plants, in particular maize, by D. Miles (Miles and Daniel, 1973). The Bennoun screening procedure is an elegant example of a hybrid between the genetic and microbiological advantage of the Chlamydomonas system with the powerful biophysical tool of fluorescence analysis (Chapter 23, Bennoun and Béal). It was extended later to a method capable of analyzing fluorescence kinetics ofindividual colonies that became a powerful tool in the isolation of photosynthetic electron transport and photophos­ phorylation mutants, and is still used today to isolate mutants and study the phenotypes of strains carrying site-directed mutations. While a well established procedure for assaying photophosphorylation by spinach thylakoid mem­ branes existed by then, the spinach procedure did not work with Chlamydomonas cells. Trying to adapt the method developed for spinach to Chlamydomonas, D. Gorman found that grinding cells in mortar with sand was the only effective means of cell disruption for obtaining chloroplast fragments capable of noncyclic phosphorylation (Gorman and Levine, 1965). This method made it possible to isolate and characterize the first photophosphorylation negative mutant (Sato et al., 1971). It is said that a noted expert in the photophosphorylation field was visiting the Levine laboratory when this work was in progress. He saw how thylakoid fragments with photophos­ phorylation activity were isolated from Chlamy­ domonas by grinding the cells with sand. He was bemused, and set out to demonstrate how it should really be done. After a period of frustration, he also began using sand grinding for his experiments. This procedure was in use for more than 10 years until

Chapter 2

Perspective on Early Research

Brand, in San Pietro’s laboratory, developed a more convenient and rapid preparation procedure (Brand et al, 1975). When Togasaki joined the Levine laboratory, fixation was measured in 25 ml flasks using 2 ml of cell suspension. One flask was used for one time point to measure the rate of photosynthetic carbon fixation after stopping the reaction by injecting acid into the flask. A number of flasks were required for a single kinetic experiment. Togasaki modified this method by using the newly available automatic pipettor to withdraw samples. This reduced sample size to 0.1 ml or less, and made the kinetic analysis of fixation more rapid, more reliable and less expensive. In the process, he also developed a convenient method to control the gas composition fixation (Togasaki and Botos, 1972). during The Levine laboratory also developed a new liquid growth medium for culturing photosynthetic mutants that remains in use presently. Sueoka’s high salt medium, commonly used at this time, formed a precipitate after autoclaving and the liquid medium appeared cloudy. To analyze the growth rate of the cells, the change in optical density at 750 nm was used usually as an index of cell growth. The precipitate in the liquid medium interfered with these measure­ ments. Furthermore, to maintain the pH of the medium at pH 7.0, Sueoka’s medium used phosphate buffer, the pK of which is 6.83. As acetate was consumed by growing Chlamydomonas cells, the pH of the medium increased to the point where the phosphate buffer ceased to have buffering capacity. Gorman and Levine devised Tris Acetate Phosphate (TAP) medium that solved both problems. The TAP medium does not form a precipitate during autoclaving and the correct pH is maintained. In the course ofstudies ofchloroplast development and biogenesis, synchronously grown cultures of Chlamydomonas were required. Existing methods for synchronizing algae were time consuming and resulted in very poor synchrony of cell division. To remedy this, Surzycki developed a new method of synchronizing cell division in Chlamydomonas using a specific light and temperature regime (Surzycki, 1967).

V. Emergence of New Research Targets During the 1960s, the focus of the entire laboratory was largely on the isolation of new mutants and the

19 analysis of components of the photosynthetic apparatus affected in these mutants. Thus, the study of one of these photosynthetic mutants, ac-20, yielded quite unexpected and surprising results that led some of us to the study of chloroplast biogenesis. At the beginning of 1965, Levine asked R. Togasaki to analyze a number of acetate requiring mutants that still retained the ability to fix in the light at wildtype levels. The basic idea was that these mutants were normal in their photocarboxylation capacities but were defective at some later step in the photosynthetic carbon cycle. Determining these steps was a very interesting problem. This kind of study could possibly uncover unknown steps in photo­ synthesis. The photo-assimilation of in such mutants should result in the accumulation of radioactive intermediates that would indicate the site of the genetic lesion leading to their unusual phenotype. To tackle this problem, R. Togasaki developed a high resolution (for that time) column chroma­ tography method for the analysis of radioactive products of short term photosynthesis. Using these columns he compared the product profile of a short term labeling of the mutant cells to that of wild-type cells. R. Togasaki analyzed seven mutants which were provided by Levine from his mutant collection. All ofthese mutants, except ac-20, yielded a product profile indistinguishable from wild-type cells. In contrast, ac-20 produced the same level of radioactive products of light-independent fixation as wild type cells, but much reduced products fixation. Moreover, as of light-dependent R. Togasaki learned later, this mutant was the only mutant in the group that fixed at a much lower rate than wild type. Upon further analysis, he found that the ac-20 cells had greatly reduced ribulose-1,5bisphosphate carboxylase (Rubisco) activity. At first, it was presumed, that the genetic lesion in these cells resulted from defective Rubisco function (Levine and Togasaki, 1965). However, ac-20 cells did not die on minimal medium, and much to our surprise, had a much higher Rubisco activity when grown on minimal medium than when grown on acetate supplemented medium. With background training in pure biochemical analysis of photo­ synthetic carbon metabolism, R. Togasaki was not ready to handle this new phenomena, namely the regulation of enzyme biosynthesis. To learn more about regulation of enzyme biosynthesis he attended a Gordon conference on the regulation of enzyme

20 synthesis in the summer of 1965 and discovered that 90% of the meeting was focused on the regulation of the lac operon. Armed with a new perspective, he began to analyze the effect of growth conditions on Rubisco activity in ac-20 cells. The conclusion of this research was, that in ac-20 cells, cultured in acetate in either the light or dark, there is a block in the biosynthesis of Rubisco. This block is removed during the incubation of the cells in the absence of acetate in either light or dark conditions. Because Rubisco activity in this mutant was light-dependent, the removal of acetate, when cells were grown in the dark, would not result in synthesis of the enzyme. The appearance of Rubisco activity had a distinct time lag suggesting that de novo enzyme synthesis occurred at this time rather than induction of activity (Togasaki and Levine, 1970). Clearly a new approach was needed to solve the problem of induction of Rubisco activity in the mutant cells of ac-20. Around this time, Ursula Goodenough began to examine the structure of photosynthetic Chlamy­ domonas mutants by electron microscopy (Goode­ nough and Levine, 1969). Analyzing the structure of the chloroplast in light- or dark-grown cells of ac-20 she discovered that, in the presence of acetate, in the light or dark, this mutant had a very small number of chloroplast ribosomes which increased upon removal of acetate. Thus, she identified the chloroplast ribosomes as the rate limiting block in the biosynthesis of Rubisco in these cells. She further demonstrated the participation of chloroplast ribosomes in Rubisco biosynthesis and went on to characterize this regulatory phenomena in more detail (Goodenough and Levine, 1970). This research added a cell biology perspective to the laboratory’s research. This was one of the early examples of regulation of biosynthesis of the photosynthetic apparatus. Subsequent work in Levine’s laboratory extended this approach to the analysis of the site of coding and synthesis of many components of the chloroplast photosynthetic, transcriptional andtranslational apparatus (Surzycki, 1969; Surzycki et al., 1970; Armstrong et al. 1971; Beck and Levine, 1974). It is intriguing looking back today, that some ofthe ‘anomalies’ we found at the time, became the starting point for very fruitful later investigations. For example, in their analysis of the ac-208 mutant, Gorman and Levine found that these cells easily acquired some suppressor mutations that could restore much ofthe photosynthetic activity. This observation

Robert K. Togasaki and Stefan J. Surzycki eventually led to the work of Wood on regulation of plastocyanin and cytochrome (now called biosynthesis (Wood, 1978). His work was followed by an elegant study of copper involvement in the regulation of plastocyanin (Merchant and Bogorad, 1986). Another observation was that in order to have ac-208 cells with a ‘non-suppressed’ phenotype, the cells ofthis mutant must always be maintained in the logarithmic growth phase. We had no idea what caused this phenomenon. In hindsight, the organism was trying to tell us that there is an interesting and complex regulatory problem awaiting investigation. Goodenough provided the combined analysis of thylakoid membrane structure and photosynthetic electron transport activities on ac-31,a mutant devoid of thylakoid membrane stacking (Goodenough et al., 1969). This paper, on the effect of membrane architecture on photosynthetic efficiency, was another seminal work for the field of excitation energy distribution. When S. Surzycki and P. Hastingsjoinedthe Levine laboratory in 1964, genetics and molecular biology of the organelle, in this case the chloroplast, was in its infancy. A quote from a 1970 paper reflects the new direction in the Levine laboratory that began to emerge at this time. ‘In the early 1960s, the presence of DNA and RNA in the chloroplast was beginning to be documented. At this time, several models were published that depicted a highly autonomous chloroplast carrying the synthesis of most of its components, and largely independent of the cytoplasm, except for the supply of small metabolites. Such models were of some puzzlement to students of Chlamydomonas, for during the same period many genes controlling chloroplast functions were being mapped to what appeared to be classical nuclear linkage groups.’ (Surzycki et al., 1970). Genetic data and specific chemical tools were combined to study chloroplast structure and function at a molecular level. Characterized mutants and antibiotics that specifically inhibit chloroplastic transcription (rifampicin) ortranslation(spectinomycin), were used to ask more molecular questions, namely the sites of transcription and translation for some chloroplast components. The work of Surzycki and Amstrong (Surzycki et. al., 1970; Amstrong et al., 1971) established the similarity between the effect of spectinomycin and the ac-20 mutation which reduced the level of 70S ribosomes in the presence of acetate. Using an

Chapter 2 Perspective on Early Research inhibitor of chloroplast translation together with rifampicin, an inhibitor of transcription in chloro­ plasts, they determined the site of synthesis and transcription of many components of the photo­ synthetic and genetic apparatus ofthe chloroplast. In a variation of this approach they used transcriptional and translational inhibitors in synchronized cultures of wild-type cells. This permitted the analysis of transcription and translation in normal, photo­ synthetically active cells, that were in the same physiological state at a given time. This refinement permitted the analysis of the chloroplastic trans­ cription and translation requirements, as well as cytoplasmic translational requirements for several chloroplast components. The work of P. Hastings, J.-D. Rochaix and Surzycki on the regulation of expression and on the structure of the ribosomal gene cluster was also carried out in the Levine laboratory (Surzycki and Hastings, 1968; Surzycki and Rochaix, 1971). This work was one of the first examples of the application of molecular biology methods to the field of chloroplast biogenesis in Chlamydomonas. Thus, the molecular analysis of chloroplast biogenesis was well on its way.

VI. Old Experiments Becoming Reality During the 1960s, the investigation of chloroplasts and mitochondria took place separately in cell free preparations. However, in intact organisms, these organelles coexist and function in close proximity within the same cell. Since chloroplast or mitochon­ drial functions can be genetically perturbed in Chlamydomonas, this alga should provide a powerful tool for the analysis of interactions between these organelles. A glimpse of this possibility emerged in 1967, and recent findings appear to support it. In 1967, Robert Togasaki attempted to duplicate a fixation protocol designed for spinach chloroplasts using intact cells of Chlamydomonas. He reasoned that if DCIP/ascorbate can supply electrons to PS I and cyclic photophosphorylation can supply ATP, the cell should be able to photoassimilate in the absence of PS II contribution (namely in the presence of 0.1 mM fixationby spinach chloroplasts DCMU). Since was favored in a nitrogen atmosphere, his experiments were also carried out under nitrogen. The results showed a small but significant light-dependent

21 fixation by the cells in the presence of 0.1 mM DCMU. However, in the absence of ascorbate and even of DCIP, control cells fixed Further experiments showed that DCMU inhibition of fixation was complete only for cells kept in aerobic conditions. There was always a significant rate of fixation in the presence of DCMU when the nitrogen atmosphere was maintained. Furthermore, addition of DCIP and especially PMS increased this anaerobic, DCMU-independent, fixation substantially (Levine, 1969). Togasaki concluded that there must be an electron donor pool within Chlamydomonas cells, and both atmospheric oxygen, and PS I are competing for it. In air, oxygen out competes PS I due to its more positive redox potential. However, in the absence of oxygen, PS I serves as an alternate electron acceptor. This PS II independent fixation under nitrogen suggests the diversion of electrons from mitochondria to the chloroplast. Thus, we can imagine a flow of electrons between chloroplasts and mitochondria. If this prediction is true then mutants defective in oxidative electron transport should fix even under aerobic conditions in the presence of DCMU. Recent data of G. Peltier, K. Redding and P. Bennoun reveal that a small amount ofPS II-dependent electron flow occurs in the absence of PS I, but that the reducing power is somehow transported and fed into the mitochondrial electron transport chain thus revealing the existence ofelectron traffic between the two organelles (Chapter 18, Redding and Peltier; Chapter 35, Bennoun). In summary, during the latter half of the 1960s to the early 1970s, we were exploring the research potential of Chlamydomonas and witnessed the emergence of many areas of research. It was a very productive period, and one of the major driving forces was R. P. Levine’s abundant curiosity and drive, coupled with his ability to assimilate new thinking and data, from both within and outside of his laboratory. He provided us with freedom to pursue our ideas and with an environment which stimulated productive interactions. It was an exciting time.

Acknowledgment The authors wish to express their thanks to Judith A. Surzycki for reading, discussion and editorial advice in the preparation of this manuscript.

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References Armstrong JJ, Surzycki SJ, Moll, B and Levine RP (1971). Genetic transcription and translation specifying chloroplasts components in Chlamydomonas reinhardi. Biochemistry 10: 692–701 Arnon DL, Whatley FR and Allen, MB. (1954). Photosynthesis by isolated chloroplasts. II. Photosynthetic phosphorylation, the conversion of light into phosphate bond energy. J Am Chem Soc 76: 6324–6329 Beck DP and Levine RP (1974). Synthesis of chloroplast membrane polypeptides during synchronous growth of Chlamydomonas reinhardi.. J. Cell Biol 63: 759–772 Bendall DS, Sanguansermsri M, Girard-Bascou J and Bennoun P (1986) Mutations of Chlamydomonas reinhardtii affecting the cytochrome complex. FEBS Lett 203: 31–35 Bennoun P and Levine RP (1967 ). Detecting mutants that have impaired photosynthesis by their increased level of fluorescence. Plant Physiol 42: 1284–1287 Bassham JA and Calvin M (1957) The path of carbon in photosynthesis. Prentice Hall, Englewood Cliffs Bishop NI (1967) Analysis of photosynthesis in green algae through mutation studies. In: Giese AC (ed) Photophysiology, Vol 8, pp 65–96. Academic Press, New York Brand JJ, Curtis VA, Togasaki RK and San Pietro A (1975) Partial reaction of photosynthesis in briefly sonicated Chlamydomonas. II Photophosphorylation activities. Plant Physiol 55: 187–191 Chua N and Levine RP (1969) The photosynthetic electron transport chain of Chlamydomonas reinhardi. VIII. The 520 nm light induced absorbance change in the wild type and mutant strains. Plant Physiol 44: 1–6 Duysens LNM, Amesz J and Kamp BM (1961) Two photo­ chemical systems in photosynthesis. Nature 190: 157–161 Edwards G and Walker DA (1983) C3, C4: Mechanisms and cellular and environmental regulation of photosynthesis. University of California Press, Berkeley Emerson R and Lewis CM (1943) The dependence of the quantum yield of Chlorella photosynthesis on wave length of light. Am J Bot 30: 126–139 Emerson R, Chalmers RV and Cederstand C (1957) Some factors influencing the long-wave limit of photosynthesis. Proc Natl Acad Sci USA 43: 133–143 Fork DC and Urbach W (1965) Evidence for the localization of plastocyanin in the electron transport chain of photosynthesis. Proc Natl Acad Sci USA 53: 1307–1315 Garnier J (1967) Une methode de detection, par photographie, de souches d’Algues vertes emettant in vivo une fluorescence anormale. CR Acad Sci, Ser D 265: 874–877 Gillham NW (1965) Linkage and recombination between non­ chromosomal mutations in Chlamydomonas reinhardi. Proc Natl Acad Sci USA 54: 1560—1567 Goodenough U W and Levine RP (1969) chloroplast ultrastructure in mutant strains of Chlamydomonas reinhardtii lacking components of the photosynthetic apparatus. Plant Physiol 44: 990–1000 Goodenough UW and Levine RP (1970) Chloroplast structure a mutant strain of Chlamydomonas and function in reinhardtii. III. Chloroplast ribosomes and membrane organization. J Cell Biol 44: 547–562

Robert K. Togasaki and Stefan J. Surzycki Goodenough UW, Armstrong JJ and Levine RP (1969) Photosynthetic properties of a mutant strain of Chlamydomonas reinhardtii devoid of chloroplast membrane stacking. Plant Physiol. 44: 1001–1012 Gorman DS and Levine RP (1965) Cytochrome f and plastocyanin: Their sequence in the photosynthetic electron transport chain of Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 54: 1665–1669 Gorman DS and Levine RP (1966a) Photosynthetic electron transport chain of Chlamydomonas reinhardtti. III. Lightinduced absorbance changes in chloroplast fragments of the wild type and mutant strains. Plant Physiol 41. 1293–1300 Gorman DS and Levine RP (1966b) Photosynthetic electron transport chain of Chlamydomonas reinhardtti. VI. Electron transport in mutant strains lacking either cytochrome 553 or plastocyanin. Plant Physiol 41: 1648–1656 Harris EH (1988) The Chlamydomonas Sourcebook. A Comprehensive Guide to Biology and Laboratory Use. Academic Press, New York Hastings PJ, Levine EE, Cosbey E, Hudock MO, Gillham NW, Surzycki SJ, Loppes R and Levine RP (1965) The linkage groups of Chlamydomonas reinhardi. Microb Genet Bull 23: 17–19 Hill R and Bendall F (1960) Function of the two cytochrome components in chloroplasts: A working hypothesis. Nature 186: 155–156 Hind G and Jagendorf A (1963) Separation of light and dark stages in photophosphorylation. Proc Natl Acad Sci USA 49: 715–722 Katoh S, Suga I, Shiratori I and Takamiya A. (1961) Distribution of plastocyanin in plants, with special reference to its local­ ization in Chloroplasts. Arch Biochem Biophys 94: 136–141 Kimimura M and Katoh S (1972) Studies on electron transport associated with Photosystem I.I. Functional site of plastocyanin: Inhibitory effects of HgCl 2 on electron transport and plastocyanin in chloroplasts. Biochim. Biophys. Acta, 283: 279–292 Kortschak HP, Hartt CE and Burr GO (1965) Carbon dioxide fixation in sugarcane leaves. Plant Physiol 40: 209–213 Levine RP (1960a) A screening technique for photosynthetic mutants in unicellular algae. Nature 188: 339–340 Levine RP (1960b) Genetic control of photosynthesis in Chlamydomonas reinhardtii Proc Natl Acad Sci USA 46: 972–977 Levine RP (1968) Genetic Dissection of photosynthesis in Chlamydomonas reinhardtii. Science 162: 768–771 Levine RP (1969a) A light-induced absorbance change at 564 nm in wild-type and mutant strains of Chlamydomonas reinhardi. In: MetzneR H (ed) Proc. International Congress of Photosynthesis Research. Progress in Photosynthesis II, pp 971–977. International Union of Biological Sciences, Tuebingen Levine RP (1969b) The analysis of photosynthesis using mutant strains of algae and higher plants. Annual Rev Plant Physiol 20: 523–543 Levine RP and Ebersold WYE (1958) Gene recombination in Chlamydomonas reinhardi. Cold Spring Harbor Symp Quant Biol. 23: 101–109 Levine RP and Goodenough UW (1970) The genetics of photosynthesis and of the chloroplast in Chlamydomonas reinhardi. Annual Rev Genetics 4: 397–408

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Levine RP and Smillie RM (1962) The pathway of triphospho­ pyridine nucleotide photoreduction in Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 48: 417–421 Levine RP and Togasaki RK (1965) A mutant strain of Chlamydomonas reinhardi lacking ribulose diphosphate carboxylase activity. Proc Natl Acad Sci USA 53: 987–990 Merchant S and Bogorad L (1986) Regulation by copper of the expression of plastocyanin and cytochrome c552 in Chlamydomonas reinhardti. Mol Cell Biol 6: 462–469 Miles CD and Daniel DJ (1973) A rapid screening technique for photosynthetic mutants of higher plants. Plant Science Letters 11: 237–240 Mets LG and Bogorad L (1971) Mendelian and uniparental alterations in erythromycin binding by plastids ribosomes. Science 174: 707–709 Moll B and Levine RP (1970) Characterization ofa photosynthetic mutant strain of Chlamydomonas reinhardti deficient in phosphoribulokinase activity. Plant Physiol 46: 576–580 Rochaix JD (1972) Cyclization of chloroplast DNA fragments of Chlamydomonas reinhardi. Nature New Biol 238: 76–78 San Pietro A. and Lang HM (1957) Photosynthetic pyridine nucleotide reductase. 1 Partial purification and properties of the enzyme from spinach. J Biol Chem 231: 211–227 Sato VL, Levine RP and J. Neumann J (1971) Photosynthetic phosphorylation in Chlamydomonas reinhardti. Effects of a mutation altering an ATP synthesizing enzyme. Biochim BiophysActa 253: 437–448 Sager R (1960) Genetic Systems in Chlamydomonas. Science 132: 1459–1465 Spreitzer RJ and Mets LJ (1980) Non-mendelian mutation affecting ribulose-1,5-bisphosphate carboxylase structure and activity. Nature 285: 114–115 Surzycki SJ (1969) Genetic functions of the chloroplast of Chlamydomonas reinhardi: Effects of rifampin on chloroplast DNA-dependent RNA polymerase. Proc Natl Acad Sci USA 63: 1327–1334

23 Surzycki SJ (1971) Synchronously grown culture of Chlamy­ domonas reinhardi. Methods Enzymol 23: 67—73 Surzycki SJ and Hastings PJ (1968) Control of chloroplast RNA synthesis in Chlamydomonas reinhardi. Nature 220: 786–787 Surzycki SJ and Rochaix JD (1971) Transcriptional mapping of ribosomal RNA genes of the chloroplast and nucleus of Chlamydomonas reinhardi. J Mol Biol 62: 89–109 Surzycki SJ, Goodenough UW, Levine RP and Armstrong JJ (1970) Nuclear and chloroplast control of chloroplast structure and function in Chlamydomonas reinhardti. Symposia Soc Exp Biol 24: 13–37 Teichler-Zallen, D. and R. P. Levine. (1969). The effect of manganese on chloroplasts structure and photosynthetic ability of Chlamydomonas reinhardi. Plant Physiol 44:701–710 Togasaki RK and Botos CR (1971) Enhanced dark fixation by preilluminated algae: A tool for analysis of photosynthetic mechanisms in vivo. In: Forti G, Avron M and Melandri A (eds) Proceedings IInd International Congress on Photo­ synthesis Research, Vol 3, pp 1759–1772. Dr. W. Junk N.V. Publishers, The Hague Togasaki RK and Levine RP (1970) Chloroplast structure and function in ac-20, a mutant strain of Chlamydomonas reinhardi. I. CO2. fixation and riublose-l,5-diphosphate carboxylase synthesis. J Cell Biol 44: 531–539 Togasaki RK and Whitmarsh J. (1986) Multidisciplinary research in photosynthesis: a case history based on the green alga Chlamydomonas. Photosynthesis Research 10: 415–422 Whitmarsh J and Levine RP (1974) Excitation energy transfer and chlorophyll orientation in the green alga Chlamydomonas reinhardi. Biochim Biophys Acta 368: 199–213 Wood PM (1978) Interchangeable copper and iron proteins in algal photosynthesis. Studies on plastocyanin and cytochrome c552 in Chlamydomonas. Eur J Biochem 87: 9–19

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Chapter 3 Organization of the Nuclear Genome Carolyn D. Silflow Department of Genetics and Cell Biology, Department of Plant Biology, Plant Molecular Genetics Institute, University of Minnesota, 1445 Gortner Ave., St. Paul, MN 55108, U.S.A.

Summary I. Introduction and Scope II. General Characteristics of the Nuclear Genome Organization of the Genome A. Interspersed Repeated Sequences 1. Transposable Elements 2. Transcribed Repetitive DNA Sequences B. Tandemly Repeated Sequences 1. Large Tandem Repeats 2. Simple Sequence Repeats 3. Telomere Repeat Sequences 4. Ribosomal DNA C. Low Copy Number Sequences (Gene Families) D. The Mating-Type Locus IV. Characteristics of Chlamydomonas Genes Transcribed by Polymerase II A. Promoters B. Codon Bias C. Introns D. Translation Start Codon and Stop Codon Sequence Context E. 3´ Noncoding Sequences IV. Physical Mapping of the Chlamydomonas Genome VI. Future Prospects Acknowledgments References

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Summary The unicellular green alga Chlamydomonas reinhardtii is an outstanding system for investigation of numerous cellular processes including photosynthesis and other metabolic pathways, biogenesis of organelles, assembly and motility of flagella, gametogenesis and mating, phototaxis and circadian rhythms. Genetic studies have generated mutations at more than 200 nuclear loci whose products function in these processes. Recent advances in molecular genetic techniques including transformation, expression of selectable marker genes, insertional mutagenesis, and genetic rescue methods have facilitated the isolation of genes identified by mutation. The nuclear genome has a sequence complexity approximately 20–30 times that of Escherichia coli, a GC content of 62%, and a large proportion of unique sequences. Among the classes of repeated sequences in the genome are several families of transposable elements which have proven useful for gene tagging, and the interspersed simple sequence repeat DNA sequences in the mating-type locus are highly rearranged between the genomes. DNA sequences obtained from approximately 200 genes and cDNAs have led to a general and picture of gene structure that includes features such as biased codon usage, a high frequency of small introns, J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 25–40. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

Carolyn D. Silflow

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and a putative polyadenylation signal that differs from the consensus in other eukaryotes. Genes located in the mating-type locus lack some of these features. The construction of a molecular map of the genome has utilized polymorphic interfertile strains to align molecular markers with the genetic map. The molecular map, together with genomic libraries in YAC (yeast artificial chromosome) and cosmid vectors, presents opportunities for further physical mapping of the genome.

I. Introduction and Scope This review will focus on the nuclear genome of Chlamydomonas reinhardtii, a species that provides distinct experimental advantages for the investigation ofnumerous biological processes. Outside the scope of this article are several other Chlamydomonas species whose phylogenetic relationships to C. reinhardtii are relatively distant. Also outside the present scope are the genomes of Volvox and other related multicellular species. The fascinating questions that remain regarding the evolutionary relationships among the genomes of this group of algal species have been discussed (Schmitt et al., 1992; Liss et al., 1997). Many aspects of the C. reinhardtii genome were reviewed previously by Harris (1989) and by Rochaix (1995). This review will summarize work that has provided insight into the organization of the nuclear genome and the structure of Chlamydomonas genes. Information gained from these studies may be particularly relevant for further development of molecular genetic tools for this system.

II. General Characteristics of the Nuclear Genome Estimates of the DNA content ofthe haploid genome (Harris, 1989). The range from genetic map consists of 17 linkage groups on which nearly 200 molecular markers have been mapped (Dutcher et al., 1991; Harris, 1993). Chromatin made from isolated nuclei contains polynucleosomes, the complement ofhistone proteins typical of eukaryotes, and a nucleosomal DNA repeat length of 189 bp (Morris et al., 1990). One unusual feature of the genome is its high GC content of 62% (Harris, 1989). In contrast to DNA from higher plants, relatively little methylation of cytosine residues in Abbreviations: PCR – polymerase chain reaction; RFLP – restriction fragment length polymorphism; YAC – yeast artificial chromosome

the CpG dinucleotide was detected in Chlamy­ domonas nuclear DNA (Day et al., 1988). Con­ clusions from early studies of genome organization differed as to the content of repetitive DNA in the nuclear genome, ranging from estimates of almost none to 30% (reviewed by Harris, 1989). Studies in the past dozen years have characterized numerous classes of repetitive sequences.

III. Organization of the Genome

A. Interspersed Repeated Sequences 1. Transposable Elements The Chlamydomonas genome contains examples of each of two groups of transposons as classified by the mechanism oftransposition (Finnegan, 1989). Class I elements or retrotransposons are flanked by long terminal repeats (LTRs) at each end and transpose via an RNA intermediate using the activity ofreverse transcriptase. Class II elements have inverted terminal repeats at each end and transpose via a DNA intermediate. The first Chlamydomonas sequence with charac­ teristics of a transposable element was discovered as an insertion in the gene encoding the oxygen-evolving enhancer 1 (OEE1) protein (Day et al., 1988). The TOC1 element is a 5.7 kb sequence found in variable copy number (2 to more than 30) in different strains. The element is dispersed in the genome, as shown by its hybridization to several chromosomes when used as a probe for in situ hybridization experiments (Hall and Luck, 1995). The element contains LTRs as found in Class I elements; however, the repeats are arranged in an unusual manner with part of the leftend LTR found at the right end of the element (Day et al., 1988). The 4.6 kbp internal region of TOC1 was found to share certain characteristics with nonviral retrotransposons including several copies of a 76 bp repeat sequence and the absence of large open reading frames encoding retroviral proteins (Day and Rochaix, 1991 a). Most TOC1 elements appear to

Chapter 3

The Nuclear Genome

be relatively recent amplification products of a single progenitor element (Day and Rochaix, 1991b). A possible retrotransposition intermediate ofthe TOC1 element was represented by a nearly full-length sense transcript (Day and Rochaix, 1991c). Comparison of two members of the TOC1 family led to the discovery of a 1.2 kb sequence inserted into a TOC1 element (Day, 1995). This novel element, termed TOC2, was shown to contain short 14-bp imperfect inverted terminal repeats and was found in ten or more copies in the DNA of several Chlamy­ domonas strains, some of which contain only one or two copies of TOC1. These results suggest that TOC2 sequences represent an independent family of Class II transposable elements. The Gulliver element, which shares several features with Class II elements in other systems, was discovered by Ferris (1989) who conducted a genomic genomic region walk in the mating-type plus on linkage group VI. This 12 kb element was shown to contain a 15-bp imperfect inverted repeat at its termini and to create an 8-bp target site duplication upon insertion. Twelve copies of the element were found dispersed in the genome in a laboratory strain, but the element was absent in the field isolate S1-D2 (Gross et al., 1988). Analysis of several independent transposition events indicated that the site of insertion was often genetically linked to the donor site (Ferris, 1989). The Gulliver element was used by Schnell and Lefebvre (1993) in a transposon-tagging strategy to isolate the Nit2 gene. They found that two of 14 spontaneous nit2 mutations were caused by insertion of a Gulliver element. Additional transposable element families have been discovered through analysis ofspontaneous mutations in several Chlamydomonas genes. The Tcr1 and Tcr2 elements were found to be associated with mutations in the Nit2- gene (Schnell and Lefebvre, 1993) and the Tcr1 element also caused a fus1 mutation (Ferris et al., 1996). Both the Tcr1and Tcr3 elements appear to be Class II transposons. Analysis of the fus1-1 mutation caused by insertion of a 9.4 kb Tcr1 element showed that it created an 8-bp target site duplication and contained terminal 140-bp perfect inverted repeats (Ferris et al., 1996). The Tcr3 element was associated with two additional fus1 mutations (Ferris et al., 1996) as well as a nit8 mutation (S.-C. Wang and P. Lefebvre, personal communication). The Tcr3 insertions contained 56 or 58-bp imperfect inverted terminal repeat sequences and created a 2-bp target site duplication upon insertion. Estimates of copy

27 number for the Tcr3 element in several Chlamy­ domonas strains ranged from two to approximately twenty copies per genome (S.-C. Wang and P. Lefebvre, personal communication). An element termed Pioneer 1 was isolated after selection for spontaneous mutations in the NIT1 gene in the field isolate strain JG224 (Graham et al., 1995). This 2.8 kb element appeared to be associated with a 2-bp duplication of the target sequence, but it did not contain other features oftransposable elements such as terminal repeats. The Pioneer element was present in low copy number in several field isolate strains of Chlamydomonas but was not present in laboratory strains 137c and 21gr.

2. Transcribed Repetitive DNA Sequences A starting point for studying repetitive DNA sequences transcribed by RNA polymerase II is the isolation of cDNA clones that hybridize to repeated sequences in nuclear DNA. Three clones isolated in this way were found to hybridize to multiple transcripts in polyadenylated RNA and to different classes of short (.5 kb) interspersed repetitive DNA sequences (Day and Rochaix, 1989). Because the cDNA clones in this study had small inserts of less than 400 bp, the repetitive elements were likely to be located in 3´ noncoding sequences of the corres­ ponding transcripts. The hybridizing DNA fragments were polymorphic between a C. reinhardtii laboratory strain and two interfertile strains, C. smithii and the field isolate S1-D2 (Gross et al., 1988), providing multiple RFLP markers for a single hybridization probe. Other examples of highly repeated sequences within cloned genes have been noted (e.g. Debuchy et al., 1989)

B. Tandemly Repeated Sequences 1. Large Tandem Repeats Evidence for large arrays of tandemly repeated sequences in the Chlamydomonas nuclear genome was found by analyzing discrete fragments of high molecular weight DNA (50–500 kb) resulting from digestion of DNA with enzymes that cut nuclear DNA with relatively high frequency (Hails et al., 1993). One class of repeat (represented by the clone pTANC1.5) was found associated with three large DNA fragments ranging in size from 200 to 700 kb and representing approximately 1% of the nuclear

Carolyn D. Silflow

28 genome by mass. Within these large arrays, four different repeat units of 1.5 to 2.5 kb were observed by digestion with BamH I, an enzyme that cleaves once within each repeat. The presence of the four repeat units varied among different C. reinhardtii strains; some interfertile strains including C. smithii and various field isolates were deficient in all four units, indicating that the sequence is an unstable or rapidly evolving component of the nuclear genome. Analysis of tetrad progeny from crosses of these polymorphic strains indicated that the large arrays represented at least five different loci and that the repeat units were not extensively interspersed with each other. Ferris and Goodenough (1994) in a genomic walk through the MT locus found a region of 1.1 kb repeat units that also were found in other genomic locations.

2. Simple Sequence Repeats The simple sequence repeat or microsatellite has been found in the genomes of numerous eukaryotic organisms including Chlamydomonas (Morris et al., 1986). Because the number of repeated motifs in each element may be highly variable, these elements can serve as a source of polymorphic DNA markers. An estimated reiteration frequency of 25 elements per 100 kb ofChlamydomonas DNA (based on genomic DNA hybridization) was found to be approximately 200-fold and 3.4-fold higher than the frequency estimates obtained for maize and mouse DNA, respectively (Morris et al., 1986). Based on repeats in a library of the frequency of cloned genomic DNA fragments, Kang and Fawley (1997) estimated a frequency of 5–6 elements per 100 kb of Chlamydomonas DNA. They found that elements is the presence and frequency of highly variable among different Chlamydomonas species. This group also demonstrated that the polymerase chain reaction (PCR) could be used to amplify small genomic fragments surrounding elements which proved to be specific polymorphic in size among different isolates of C. reinhardtii.

3. Telomere Repeat Sequences As is true for most eukaryotic species studied to date, telomeres in C. reinhardtii are composed of short tandem repeat sequences. Petracek et al. (1990) identified the telomere sequences from Chlamy-

domonas by hybridization with telomere repeats from Arabidopsis thaliana. DNA fragments containing these repeats were sensitive to Bal-31 exonuclease digestion, indicating that they were located at the ends of chromosomes. Analysis of cloned DNA containing the putative telomere sequences revealed which was a repeat sequence of estimated to be present in tracts at least 300 bp in length in genomic DNA. The telomere-associated sequences adjacent to the telomere repeats from three different chromosome ends were conserved, with 78% sequence identity over a distance of approximately 100 bp. Although the Chlamydomonas telomere repeat sequence is quite divergent from that found in Sacharomyces cerevisiae it is apparently recognized as a telomere in S. cerevisiae. After ligation of Chlamydomonas DNA fragments to a linearized S. cerevisiae plasmid and transformation of the ligated DNA into S. cerevisiae cells, Hails et al. (1995) recovered a transformant containing a linear plasmid in which Chlamydomonas telomere repeats were capped by telomere repeats of S. cerevisiae.

4. Ribosomal DNA Hybridization of cytosolic ribosomal RNA to DNA fractionated by equilibrium density centrifugation showed that the nuclear ribosomal DNA bands with a slightly lower density than the major band of nuclear DNA, indicating that it has a lower GC content than the majority of nuclear DNA (Harris, 1989), Approximately 400 copies of the ribosomal RNA genes were estimated to be present in the genome, based on saturation hybridization experi­ ments (Howell, 1972). The genomic repeat unit encoding the ribosomal RNAs was cloned and characterized by Marco and Rochaix (1980). They found that the repeat unit is approximately 8.0 kb in length and that it contains genes encoding 18 S and 25 S rRNAs, two internal transcribed spacer regions between the genes as well as a gene encoding 5.8 S ribosomal RNA in the smaller spacer. The DNA sequences and conserved secondary structure of the spacer regions of ribosomal RNA genes have been useful for phylogenetic studies ofgreen algal species (Mai and Coleman, 1997). Cytoplasmic 5S RNA hybridized with the bulk of nuclear DNA in density gradient fractions, indicating that the 5S genes are not a part of the ribosomal DNA repeat unit (Marco and Rochaix, 1980).

Chapter 3

The Nuclear Genome

C. Low Copy Number Sequences (Gene Families)

The minimal number of each of the four genes encoding core histones in the C. reinhardtii genome was estimated at 15, whereas histone H1 genes were estimated to be present in only one or two copies (Fabry et al., 1995). Seven different genomic regions containing core histone genes were examined, revealing that the four core histone genes usually are clustered, with a histone H3, H4 gene pair separated by about 1 kb from a histone H2A, H2B gene pair. DNA sequences from five different histone gene clusters (16 histone genes) have been reported (Fabry et al., 1995; Walther et al., 1995). These data show that each gene pair is separated by an intergenic region of about 300 bp and the genes are transcribed divergently. A single H1 histone gene was found separated by 1.6 kb from a H2A, H2B gene pair (Fabry et al., 1995). Several conserved promoter elements were found in the Chlamydomonas histone genes including a TATA-box and three other motifs. Walther and Hall (1995) showed that at least two histone H4 genes are likely to be located on linkage group XIX and that the other histone genes are probably dispersed on other chromosomes. Several small gene families have been described in Chlamydomonas. The genes encoding lightharvesting complex (LHC) II proteins (CabII genes) have been reported to exist as a small gene family composed of 3–7 members (Imbault et al., 1988) which are dispersed throughout the genome (P. Kathir, C. Silflow, unpublished). Similarly, several genomic fragments hybridized to the L1818 gene which encodes a protein distantly related to other LHC proteins (Savard et al., 1996). The genes encoding dynein heavy chains (Dhc) were analyzed by Porter et al. (1996) who used a PCR-based strategy to clone 11 Dhc genes, eight of which mapped to positions on five different linkage groups, with only two of the genes being closely linked. Pairs of closely linked but differentially regulated genes encode carbonic anhydrase (Fujiwara et al., 1990), Rubisco small subunit (Goldschmidt-Clermont and Rahire, 1986), and a cysteine-rich protein which may be a component of the zygote cell wall (Matters and Goodenough, 1992). In the latter case, evidence for expression of only one gene was obtained, suggesting that one of the genes may be a pseudogene (Matters and Goodenough, 1992). Two genes encoding identical proteins are separated by 12 cM on linkage

29 group XII; two genes encoding identical proteins are located on linkage groups III and IV (Ranum et al., 1988; James et al., 1993). Although the genomes of Chlamydomonas and the higher plant Arabidopsis thaliana are of similar size, Arabidopsis contains six genes encoding (Kopczak et al., and nine genes encoding 1992;Snustad et al., 1992). In some instances, genes with related function have been shown to be located in close physical proximity. For example, five genes involved in nitrate assimilation are linked within a region of 32 kb (Quesada et al., 1993; Chapter 33, Fernández et al.). The Rsp4 and Rsp6 genes encoding related protein components of the flagellar radial spoke heads are closely linked, with the 3´ end of the Rsp4 gene being only 435 bp from the 5´ end of the Rsp6 gene (Curry et al., 1992). Several genes involved in mating are located in the MT locus, (Goodenough et al., 1995).

D. The Mating-Type Locus The most detailed characterization ofthe organization of a genomic region has come from the results of an extraordinary genomic walk through the matingtype locus on linkage group VI that covered a distance of 1.1 Mb (Ferris and Goodenough, 1994; reviewed by Goodenough et al., 1995). This region, which is notable for suppression of recombination for numerous genes that map to the MT locus, was found to contain three domains. The centromereproximal C domain (525 kbp) and the telomereproximal T domain (109 kbp) are colinear in the genomes. In contrast, comparison of the and strain with the central R domain of 190 kbp in the strain showed that this region R domain in the contains major translocations, inversions, deletions and duplications, most likely leading to suppressed recombination. Several genes have now been cloned from this region including Ezy1 which is found in multiple copies in the C domain and which encodes a protein that participates in uniparental inheritance of the chloroplast genome (Armbrust et al., 1993; Chapter 6, Armbrust). The Fus1 gene, which is locus, encodes found only in the R domain of the, a glycoprotein expressed on mating structures (Ferris et al., 1996). The Mid gene, which is located locus, encodes a putative in the R domain ofthe transcription factor that controls the program of gametic differentiation (Ferris and Goodenough, 1997). Three additional genes which reside in the

Carolyn D. Silflow

30 C and T domains but do not play a role in mating functions have been localized on cloned fragments from the walk (Ferris, 1995).

IV. Characteristics of Chlamydomonas Genes Transcribed by Polymerase II

A. Promoters The expression of the four tubulin genes and other genes encoding flagellar proteins is induced coordinately following deflagellation of Chlamy­ domonas cells (Brunke et al., 1982; Schloss et al., 1984), and this induction is partly due to increased rates of transcription of these genes (Keller et al., 1984). The tubulin gene promoters have been studied extensively to identify cis-acting elements necessary for this induction. A TATA box followed by a GCrich region has been found approximately 30 bp upstream of the transcription start site in numerous Chlamydomonas genes including the tubulin genes (reviewed in Davies and Grossman, 1994). Although the GC-rich sequence was essential for transcription gene constructs injected into Xenopus of oocyte nuclei (Bandziulis and Rosenbaum, 1988), its role in transcription in Chlamydomonas cells is unclear. When this sequence was changed to an ATpromoter driving rich sequence in a expression of the arylsulfatase reporter gene, no effect was observed on the constitutive activity ofthe promoter nor on the induction of transcription following deflagellation (Davies and Grossman, 1994). Similarly, a study in which the TATA box and upstream sequences were deleted from the gene promoter construct showed that the TATA box was not required for basal levels of expression (Periz and Keller, 1997). Results from these studies indicate that basal transcription from the tubulin promoters is controlled by sequences within 35 bp upstream of the transcription start site and that the TATA box and the GC-rich regions may not be essential for basal level activity of the promoters, although these elements may be involved in selecting the site of transcription initiation. Comparison of promoter regions for the four tubulin genes identified several copies of a 10-bp motif termed a ‘tub’box (Brunke et al., 1984). Davies and Grossman (1994) examined the role of the tub boxes in induction of promoter gene after deflagellation activity for the and concluded that inclusion of at least two groups of

tub box elements was essential for the induction (i.e. four tub boxes). Studies with the gene promoter showed that two different regions of the promoter, one of which contains two tub boxes, are essential for maximal induction and suggested that promoter elements in addition to tub boxes are important for the response (Periz and Keller, 1997). The analysis of several other promoters has provided insight into gene regulation as well as useful tools for driving the expression of reporter genes. For example, the Cyc6 gene encoding cytochrome is transcribed only under conditions of copper deficiency. Mutational analysis ofthe Cyc6 promoter showed that two different sequences within 127 bp upstream of the transcription start site were sufficient for copper-responsive expression (Quinn and Merchant, 1995). Analysis of the Hsp70 gene promoter showed that the elements responsible for light induction and for heat shock response reside in different regions of a sequence 140 upstream of the transcription start site (Kropat et al., 1995). A 2.5 kb region upstream of the CabII-1 gene was able to confer light-regulated expression when fused to a reporter gene (Blankenship and Kindle, 1992). In addition, a 4 kb region upstream from the CabII-1 gene was able to drive the expression of reporter genes with a circadian rhythm similar to that seen for the expression of the CabII gene family, indicating that the circadian clock regulates transcription ofthis gene (Jacobshagen et al., 1996). A promoter region from the Nit1 gene encoding nitrate reductase conferred ammonium-repressible expression on different reporter genes (Ohresser et al., 1997; Zhang and Lefebvre, 1997). High-level expression of reporter genes has been facilitated by fusion with the promoter of the RbcS2 gene (Kozminski et al., 1993; Nelson et al., 1994; Stevens et al., 1996; Cerutti et al., 1997).

B. Codon Bias A feature of most Chlamydomonas genes is the pronounced codon bias which favors codons with C or G in the third position and reflects the high GC content of the genome. The bias is, however, more than a reflection of the high GC content. For 89 Chlamydomonas sequences, LeDizet and Piperno (1995a) calculated the ‘standardized synonymous codon bias’ or B value which may vary from 0 (no codon bias) to 1 (only one codon used per amino acid)(Long and Gillespie, 1991). They found that the

Chapter 3 The Nuclear Genome B values ranged from 0.316 to 0.78, with a median of 0.617. It is possible that the codon bias is at least partly responsible for the low levels of expression observed when heterologous genes are introduced by transformation into Chlamydomonas. Indeed, a bacterial phleomycin-resistance gene (ble), which has a codon bias similar to that seen in most Chlamydomonas genes, was shown to be expressed when transformed into Chlamydomonas cells (Stevens et al., 1996). Striking exceptions to the common codon bias were found in two genes located in the mating-type (MT) locus. The Fus 1 gene shows no codon preference (B value = .05) and all regions of the gene contain a reduced GC content (coding region = 47.7% GC) (Ferris et al., 1996). Another gene, Mid, also was found to have a very low codon bias (B value = 0.161) and a coding region GC content of 50% (Ferris and Goodenough, 1997). These authors suggest that the dramatic rearrange­ ments that have occurred in the mating-type locus R domain (Section III.D) have led to the changes in nucleotide composition and changes in codon bias that are apparent in genes located in this region. In addition to the lack of codon bias observed in these endogenous genes, further evidence that codon bias is not essential for gene expression in Chlamy­ domonas comes from studies in which a bacterial dominant selectable marker gene which lacks the Chlamydomonas codon bias was able to confer spectinomycin resistance to transformed Chlamy­ domonas cells (Cerutti et al., 1997). Two databases that maintain codon usage tables for C. reinhardtii and other organisms are TransTerm (Dalphin et al., 1996) and CUTG (Nakamura et al., 1996).

C. Introns Based on DNA sequence data for genes and/or cDNAs, one general feature ofChlamydomonas genes that has emerged is that many small introns are often present at high frequency (Table 1). On average, four introns are located within each kilobase of coding sequence and the average intron size is 219 bp. At least two introns are located in untranslatedregions of genes (Mitchell and Kang, 1993; Sugase et al., 1996). Within an average gene, the total sequence contained in introns is nearly equal to the total coding sequence (Table 1). The frequency of introns in Chlamydomonas genes may provide a large target for mutations that affect intron splicing, as suggested by the finding that three different ida4 alleles contain

31

alterations in splice site sequences (LeDizet and Piperno, 1995b). Although most Chlamydomonas genes apparently do contain introns, some intronless genes have been noted including several core histone genes (Fabry et al., 1995). In addition, introns were shown to be unnecessary for the normal expression of the Rsp3 gene, as minigenes consisting of 5´ and 3´ genomic regions fused to a Rsp3 cDNA were able to rescue the mutant phenotype when transformed into pf14 mutant cells (Diener et al., 1993). The possible role of intron splicing in regulating expression of other genes in Chlamydomonas has not been examined rigorously. However, in studies with Volvox carteri, a related alga that also has a high frequency of introns, the presence of at least one intron in cDNA constructs from the nitA gene was shown to enhance the transformation frequency of nitA mutants by ten-fold over intron-less constructs. These results suggest a role for splicing in regulating the expression of the nitA gene which normally contains 10 introns (Gruber et al., 1996). Nucleotide sequences at the 5´ and 3´ splice junctions of Chlamydomonas introns conform in general to the eukaryotic consensus for these sequences (Table 2; Lee et al., 1991; Ledizet and Piperno, 1995b). A consensus sequence (NCTAG) for the putative branch site located 15–51 bp upstream of the 3´ splice site in 56 Chlamydomonas nuclear introns was also observed (Lee et al., 1991). The presence of these consensus sequences suggests that the splicing machinery in Chlamydomonas cells is similar to that in other

32

eukaryotes. Some evidence supporting this con­ clusion has come from experiments reported by Jarvik et al. (1996) who developed a ‘CD-Tagging’ method for inserting small sequences encoding epitopes into various locations in genes of interest. The method involves insertion of engineered miniexons into existing introns to produces genes encoding epitope-tagged proteins that can be localized in cells using specific antibodies. In these experiments, engineered constructs based on intron splice site sequences of Chlamydomonas were able to function in Drosophila cells. Information on the positioning of introns within Chlamydomonas genes has been used in the debate over the origins of spliceosome-dependent introns. Some evidence showing conserved intron positions within ancient genes such as glyceraldehyde-3phosphate dehydrogenase (Kersanach et al., 1994) have supported the hypothesis that introns arose early in the evolution of genes (Dorit et al., 1990). However, most introns in Chlamydomonas genes occur at unique positions when compared with homologous genes in animals, higher plants, fungi and other protists (Dibb and Newman, 1989; Lee et al., 1991; Kopczak et al, 1992; Snustad et al., 1992;

Carolyn D. Silflow

Sugase et al., 1996). These data lend support to the hypothesis that multiple intron insertion events occurred late in the evolution of eukaryotic genes (reviewed by Palmer and Logsdon, 1991). The presence of conserved exon sequences adjacent to introns would not be expected unless these sequences were involved in some way in the acquisition or splicing of introns. Dibb and Newman (1989) proposed that the sequence (A/C) AG(A/G) is a proto­ splice site, where introns are inserted between the conserved G and (A/G) nucleotides. The consensus sequence is clearly present in exon positions adjacent to Chlamydomonas introns (Table 2). Intron sequences have been exploited as tools to evaluate phylogenetic relationships among groups of related algae in the order Volvocales (Liss et al., 1997). Several studies have noted that intron positions are often conserved between genes in C. reinhardtii and Volvox carteri, implying intron insertion into common ancestral genes (Harper and Mages, 1988; Mages et al., 1988; Dietmaier et al., 1995; Fabry et al., 1995). However, DNA sequences in introns have diverged much more rapidly than those in protein coding regions. The rapidly evolving intron sequences provided the resolution necessary to describe the

Chapter 3

The Nuclear Genome

relationships among several strains of Volvox and among several interfertile strains ofChlamydomonas which had been difficult to resolve using rDNA or protein coding sequences (Liss et al., 1997). These authors estimated that the substitution rate in intron sequences is approximately ten-fold higher than the synonymous substitution rate in protein-coding sequences.

D. Translation Start Codon and Stop Codon Sequence Context Because the sequence context surrounding start codons and stop codons affects the efficiency of translation, the TransTerm database was created to compile sequence context data for numerous species including C. reinhardtii (Dalphin et al., 1996). Comparisons of 133 sequences showed, for example, a consensus of (A/C) A (A/C) (A/C) A T G (G/C) for the context ofthe start codon (Table 3). Similarly, the consensus of (G/C) TAA (G/A) for the context ofthe stop codon was obtained from analysis of 149 cloned sequences (Table 3). The stop codon TAA was found in 70% of genes; TGA and TAG stop codons were each found in 15% of the genes surveyed.

33

E. 3´ Noncoding Sequences The 3´ noncoding regions of Chlamydomonas genes are often several hundred bp in length. The most common putative polyadenylation signal is TGTAA, located 10–20 bp upstream of the polyadenylation site in approximately 90% ofChlamydomonas cDNA sequences reported to date. Variations ofthis sequence including TGTAG and TGTTA in the appropriate location have been reported for a few cDNA clones. Two genes found in the mating-type locus, Fus1 and Mid, have among other unusual characteristics (see Sections III.D, IV.B), an absence of the canonical polyadenylation signal, leading to transcripts with a variety of polyadenylation sites (Ferris et al., 1996; Ferris and Goodenough, 1997). The histone genes (including the histone H1 gene) do not contain the canonical Chlamydomonas polyadenylation signal at the 3´ end but do contain a 27-bp palindromic sequence (Fabry et al., 1995; Walther and Hall, 1995). Similar palindromes are involved in regulating the stability of histone mRNA molecules in metazoans and were observed in the histone genes of the related alga Volvox, but not in those of land plants.

34

Carolyn D. Silflow

Chapter 3 The Nuclear Genome

35

36 IV. Physical Mapping of the Chlamydomonas Genome The genetic map of Chlamydomonas consists of 17 linkage groups (Dutcher et al., 1991; Harris, 1993). Although progress has been made in developing techniques for visualizing Chlamydomonas chromo­ somes and for in situ hybridization to specific chromosomes, it has not been possible to verify chromosome number using this approach (Hall and Luck, 1995). In addition, only partial resolution of the chromosomes has been achieved using pulsed field gel electrophoresis (Hall et al., 1989; A. Day, personal communication). The availability of Chlamydomonas strains interfertile with C. rein­ hardtii but divergent in genomic DNA sequence has facilitated the mapping ofrestriction fragment length polymorphisms (RFLP) (Ranum et al., 1988). The progeny of a cross between C. reinhardtii strains containing various genetic markers and the interfertile C. smithii strain were analyzed to align molecular markers with the genetic map. Further mapping has utilized a wild isolate S1-D2 which exhibits RFLPs with C. reinhardtii more frequently than does the C. smithii strain (Gross et al. 1988; Porter et al., 1996). A molecular map currently under construction has analyzed the segregation of markers among 136 random progeny of a cross between C. reinhardtii and the S1-D2 strain (Fig. 1; P. Kathir, P. Lefebvre, and C. Silflow, unpublished data). Currently, this map contains approximately 170 molecular markers including RFLP markers and sequence tagged site (STS) markers that utilize specific primers for the polymerase chain reaction (PCR) to amplify genomic fragments that vary in length between the two polymorphic strains (Silflow et al. 1995). The molecular markers, which include random small Pst I genomic fragments, random cDNA clones, and specific cloned genes or cDNAs contributed by Chlamydomonas researchers, are distributed among all 17 known linkage groups. For most ofthe linkage groups, it has been possible to orient the groups of molecular markers with the genetic map by utilizing cloned genes known to correspond to mutations for which genetic map location was determined previously or by using a C. reinhardtii parent strain containing multiple mutations whose genetic map locations were known (Ranum et al., 1988). In addition to the groups of linked markers shown in Fig. 1, we have identified two groups of markers which do not appear to be linked to known linkage

Carolyn D. Silflow groups. Further analysis will be required to determine whether these groups ofmarkers represent previously unidentified chromosomes or whether they map to a known linkage group but sufficiently distant from the current set of markers that linkage has not been detected. The Chlamydomonas molecular map has been used to determine whether cloned genes/cDNAs correspond to known genetic loci (e.g. Porter et al., 1996). It has proven useful for mapping mutations generated by insertional mutagenesis by using DNA fragments flanking the inserted plasmid. Mapped probes also have been used as the starting points to ‘walk’to nearby genes of interest (e.g. Walther et al., 1994). A large set of molecular markers will facilitate the ordering of cloned genomic fragments into ‘contigs’ of overlapping fragments during the construction ofa physical map ofthe Chlamydomonas genome (see below). By comparing the genetic map containing approximately (Harris, 1993) with the genome complexity (see above), a rough estimate for the physical distance of a recombination unit of 100– 160 kb per cM is obtained. The results ofVashishtha et al. (1996) suggest that this may be an overestimate. They found that severalYAC clones containing inserts of approximately 130 kb encompassed intervals defined by one or two recombination events among 15–20 tetrads tested. The physical distance corres­ ponding to a recombination unit can be much smaller than this estimate, as shown by Debuchy et al. (1989) who found that a genomic fragment of 7.8 kb containing the argininosuccinate lyase gene was able to rescue the phenotype of three mutations, arg2, arg 7-3, and arg7, which had been ordered previously in genetic experiments (Matagne, 1978). The genetic distance over this interval was calculated to be 1.0 to 1.6 map units. Physical distances could be larger in regions of suppressed recombination such as telomeric and centromeric regions. In addition to the molecular maps described above, other necessary tools have been prepared to facilitate gene cloning in Chlamydomonas. Hall and co­ workers have generated a genomic library in a yeast artificial chromosome (YAC) vector which contains approximately four genome equivalents in clones with an average size of 125 kb (Infante et al., 1995; Vashishtha et al., 1996). In another project, an ‘indexed’ genomic library ofcosmid clones has been prepared, with the clones plated individually in wells so that pools of clones can be analyzed in gene cloning experiments (Zhang et al., 1994, Chapter 4,

Chapter 3

The Nuclear Genome

Kindle). An indexed library of genomic fragments also has been constructed using a bacterial artificial chromosome (BAC) vector (P. Lefebvre, personal communication).

VI. Future Prospects To take full advantage of Chlamydomonas as a genetic system, a major goal of future research efforts in many areas will be the cloning of genes identified by mutation. The insertional mutagenesis technique is a powerful new tool that relies on random integration of plasmids containing selectable marker genes into the nuclear genome (Tam and Lefebvre, 1993; Gumpel et al., 1995; Pazour et al., 1995; Davies et al., 1996; Smith and Lefebvre, 1996). This method generally causes gene disruption or deletion, producing null mutations. Mutations in genes essential for viability are not expected to be recovered, nor are mutations with conditional phenotypes. Cloning genes by transposon tagging is another potentially fruitful approach (Day et al., 1988; Schnell and Lefebvre, 1993; Ferris et al., 1996), but more research is needed to characterize the classes of transposons present in the genome and to define the conditions that may influence transposition. For certain nuclear genes that have homologs in E. coli, cloning has been accomplished by functional complementation of the appropriate E. coli mutant with sequences from a Chlamydomonas cDNA library (Matters and Beale, 1994, 1995; Yildiz et al., 1996). For mutations that result in selectable phenotypes, gene cloning may be accomplished by transformation of mutant cells with genomic libraries (Purton and Rochaix, 1994; Zhang et al., 1994). However, the size of the genome and the current transformation efficiency preclude this method in cases where a strong selection method is not available. The phenotypes of many interesting mutations are subtle and require labor-intensive screening methods. The cloning of the affected genes in such mutants will be facilitated by the development of better methods for genome walking from nearby molecular markers. To facilitate these approaches, an important goal is the further development of the physical map in which cloned genomic fragments have been ordered in ‘contigs’ that cover large regions of the genome. A puzzling aspect of molecular genetic studies with Chlamydomonas has been the inability of these cells to express foreign genes including many reporter

37 genes that are expressed routinely in other eukaryotic systems. Speculation about the molecular basis of the block to expression has included unusual aspects of Chlamydomonas genes including the codon bias, intron sequences, and the polyadenylation signal. Some success in solving the problem of lack of expression has been reported (e.g. Stevens et al., 1996; Cerutti et al., 1997). The latter group found evidence that the lack of stable expression of the bacterial spectinomycin resistance gene may have been due to gene silencing or transcript instability. Further investigation into the basic mechanisms involved in control of gene expression in Chlamy­ domonas are important for gaining insight into this problem and for developing the strengths of Chlamydomonas as an experimental system.

Acknowledgments Work in the author’s laboratory was funded by the National Institutes of Health GM51995. I thank O. Sodeinde, S. Merchant and P. Lefebvre for commun­ icating unpublished results and P. Lefebvre for comments on the manuscript.

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39 Mai JC and Coleman A W (1997) The internal transcribed spacer 2 exhibits a common secondary structure in green algae and flowering plants. J Mol Evol 44:258–271 Marco Y and Rochaix J-D (1980) Organization of the nuclear ribosomal DNA of Chlamydomonas reinhardtii, Mol Gen Genet 177:715–723 Matagne RF (1978) Fine structure of the arg-7 cistron in Chlamydomonas reinhardtii. Molec Gen Genet 160:95–99 Matters GL and Beale SI (1994) Structure and light-regulated expression of the gsa gene encoding the chlorophyll biosynthetic enzyme, glutamate 1-semialdehyde aminotransferase, in Chlamydomonas reinhardtii. Plant Mol Biol 24:617–629 Matters GL and Beale SI (1995) Structure and expression of the Chlamydomonas reinhardtii alad gene encoding the chlorophyll biosynthetic enzyme, acid dehydrase (porphobilinogen synthase). Plant Mol Biol 27:607–611 Matters GL and Goodenough UW (1992) A gene/pseudogene tandem duplication encodes a cystein-rich protein expressed during zygote development in Chlamydomonas reinhardtii. Molec Gen Genet 232:81–88 Mitchell DR and Kang Y (1993) Reversion analysis of dynein intermediate chain function. J Cell Sci 105:1069–1078 Morris J, Kushner SR and Ivarie R (1986) The simple repeat poly(dT-dG)-poly(dC-dA) common to eukaryotes is absent from eubacteria and archaebacteria and rare in protozoans. Mol Biol Evol 3:343–355. Morris RL, Keller LR, Zweidler A and Rizzo PJ (1990) Analysis of Chlamydomonas reinhardtii histones and chromatin. J Protozool 37:117–123. Nakamura Y, Wada K, Wada Y, Doi H, Kanaya S, Gojobori T and Ikemura T (1996) Codon usage tabulated from the international DNA sequence databases. Nucl Acids Res 24:214– 215. Nelson JAE, Savereide PB and Lefebvre PA (1994) The CRY1 gene in Chlamydomonas reinhardtii: Structure and use as a dominant selectable marker for nuclear transformation. Mol Cell Biol 14:4011–4019 Ohresser M, Matagne RF and Loppes R (1997) Expression of the arylsulphatase reporter gene under the control of the nit1 promoter in Chlamydomonas reinhardtii. Curr Genet 31:264– 271 Palmer JD and Logsdon JM (1991) The recent origins of introns. Curr Opin Genet Develop 1:470–77 PazourGJ, Sineschekov OA and Witman GB (1995) Mutational analysis of the phototransduction pathway of Chlamydomonas reinhardtii. J Cell Biol 131:427–440 Periz G and Keller LR (1997) DNA elements regulating gene induction during regeneration of eukaryotic flagella. Mol Cell Biol 17:3858–3866 Petracek ME, Lefebvre PA, Silflow CD and Berman J (1990) Chlamydomonas telomere sequences are A+T-rich but contain three consecutive G-C base pairs. Proc Natl Acad Sci USA 87:8222–8226 Porter ME, Knott JA, Myster SH and Farlow SJ (1996) The dynein gene family in Chlamydomonas reinhardtii. Genetics 144:569–585 Purton S and Rochaix J-D (1994) Complementation of a Chlamydomonas reinhardtii mutant using a genomic cosmid library. Plant Mol Biol 24:533–537 Quesada A, Galvan A, Schnell RA, Lefebvre PA and Fernandez E (1993) Five nitrate assimilation-related loci are clustered in

40 Chlamydomonas reinhardtii. Mol Gen Genet 240:387–394 Quinn JM and Merchant S (1995) Two copper-responsive elements associated with the Chlamydomonas Cyc6 gene function as targets for transcriptional activators. Plant Cell 7:623–638 Ranum LPW, Thompson MD, Schloss JA, Lefebvre PA, and Silflow CD (1988) Mapping flagellar genes in Chlamydomonas using restriction fragment length polymorphisms. Genetics 120:19–122 Rochaix J-D (1995) Chlamydomonas reinhardtii as the photosynthetic yeast. Annu Rev Genet 29:209–230 Savard F, Richard C and Guertin M (1996) The Chlamydomonas reinhardtii L1818 gene represents a distant relative of the Cab1/II genes that is regulated during the cell cycle and in response to illumination. Plant Mol Biol 32:461–473 Schloss JA, Silflow CD and Rosenbaum JL (1984) mRNA abundance changes during flagellar regeneration in Chlamy­ domonas reinhardtii. Mol Cell Biol 4:424–434 Schmitt R, Fabry S and Kirk DL (1992) In search ofthe molecular origins of cellular differentiation in Volvox and its relatives. Int Rev Cytol 139:189–265 S c h n e l l RA and Lefebvre PA (1993) Isolation of the Chlamydomonas regulatory gene NIT2 by transposon tagging. Genetics 134:737–747 Senapathy P, Shapiro MB and Harris NL (1990) Splice junctions, branch point sites, and exons: Sequence statistics, identification, and applications to genome project. Methods Enzymol 183:252–278 Silflow CD, Kathir P and Lefebvre PA (1995) Molecular mapping of genes for flagellar proteins in Chlamydomonas. Methods in Cell Biol 47:525–530 Smith EF and Lefebvre PA (1996) PF16 encodes a protein with armadillo repeats and localizes to a single microtubule of the central apparatus in Chlamydomonas flagella. J Cell Biol 132:359–370

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Chapter 4

Nuclear Transformation: Technology and Applications Karen L. Kindle

Plant Science Center, Biotechnology Program, Biotechnology Building,

Cornell University, Ithaca, NY 14853

Summary I. Introduction II. A Brief History of C. reinhardtii Nuclear Transformation III. Selectable Markers A. Complementation of C. reinhardtii Mutations B. Drug Resistance Markers IV. Methods for Introducing DNA into the Nuclear Genome of C. reinhardtii A. Particle Bombardment B. Glass Beads, Silicon Whiskers C. Electroporation V. Reporters and Promoters A. Arylsulfatase as a Reporter B. Other Reporters C. Constitutive Promoters D. Regulated Promoters VI. Characteristics of Transformation Events A. Nature and Stability of Introduced DNA B. Insertion Events C. Cotransformation and Expression of C. reinhardtii Genes D. Expression of Foreign Genes/Silencing VII. Insertional Mutagenesis and Gene Tagging A. Development and Application of the Approach B. Technical Considerations C. Isolation of the Gene Responsible for the Mutant Phenotype D. Perspectives VIII. Gene Isolation by Complementation of a Mutant Phenotype IX. Homologous Recombination and Gene Targeting X. The Use of Nuclear Transformation to Study Promoter Function XI. Conclusion Acknowledgments References

J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 41–61. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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Karen L. Kindle

Summary Nuclear transformation is now simple and efficient, and it has revolutionized the kinds of questions that can be addressed using Chlamydomonas reinhardtii as a model system. The highest rates of transformation are obtained using C. reinhardtii genes that complement auxotrophic mutations as selectable markers. Cotransformation is efficient, so effects of mutations engineered in vitro can readily be tested by reintroducing the altered genes into an appropriate mutant strain together with a selectable marker. Because nearly all nuclear transformation events result from apparently random integration of the introduced plasmid into chromosomal DNA, it is possible to generate large numbers of insertional mutants. The subsequent isolation of the disrupted genes by virtue of the molecular tag provides a very powerful means for cloning genes with known mutant phenotypes. Alternatively, since nuclear transformation is so efficient, genes with good selectable phenotypes can be isolated by complementing appropriate recessive mutations with pools of DNA from an indexed genomic library. Recently, expression of eubacterial genes that confer antibiotic resistance to C. reinhardtii has been reported; these genes should be useful as dominant selectable markers in any genetic background. However, transformants are recovered with these bacterial genes at only 1 % the rate obtained by transformation with C. reinhardtii genes, and in about half the cases the introduced genes are silenced under nonselective conditions, sometimes a problem with C. reinhardtii genes as well. Although an active homologous recombination system allows efficient recombination between simultaneously introduced plasmids, the rate of gene-targeted homologous integration events is very low. Further understanding of the factors that limit the expression of reintroduced genes and the rate of gene-targeting could lead to substantial improvements in the capability of manipulating the nuclear genome and of generating or phenocopying mutations corresponding to cloned genes.

I. Introduction Although a subset of chloroplast and mitochondrial proteins is encoded by organellar genes, the biogenesis and functions of these organelles are to a large extent dependent on structural and regulatory proteins encoded in the nucleus. The functions of these nucleus-encoded polypeptides and how they are correctly targeted to the appropriate subcellular compartment are some of the most intriguing questions in organelle ontogeny. The capability of introducing DNA into the nuclear genome is essential for a variety of ‘reverse’ genetic approaches to investigate organellar biogenesis, structure, and function. For example, site-directed mutations can be engineered into a cloned gene in vitro and tested in vivo by transforming the gene into an appropriate mutant strain. With a sufficiently high transformation efficiency, it is possible to isolate a gene by its ability to complement a particular mutant phenotype. Nuclear transformation can also be used as a mutagen, either in the course of random insertion events, or Abbreviations: Ars – arylsulfatase; CabII-1 – chlorophyll a/b binding protein gene; nos – nopaline synthetase gene; nptII – neomycin phosphotransferase gene; ocs – octopine synthetase gene; PEG – polyethylene glycol; Rubisco – ribulose bisphosphate carboxylase/oxygenase; RFLP – restriction fragment length polymorphism; SV40 – simian virus 40; Tn5, 9 – transposons 5 and 9

potentially, by generating gene-targeted disruptions. In the last decade, simple, efficient, and reliable methods for transforming the nucleus of Chlamy­ domonas reinhardtii have been developed. For those who have watched the field since the mid 1980s, it is gratifying to see this technique evolve from frustratingly elusive to so simple that it has been incorporated into many introductory laboratory courses. This review will briefly describe the history of nuclear transformation, then discuss the selectable markers and other molecular tools that are available. Alternative methods for introducing DNA into the nucleus will be presented, and the molecular events that appear to occur during transformation will be reviewed. Finally, some of the limitations and perspectives for the future will be discussed, along with a brief summary of results on promoter structure and function that have been deduced from transformation experiments.

II. A Brief History of C. reinhardtii Nuclear Transformation Many individuals and laboratories have contributed to the development of nuclear transformation technology, a process that is ongoing, since certain technical challenges remain (see below). The first

Chapter 4 Nuclear Transformation reports of nuclear transformation utilized ‘foreign’ (non-C. reinhardtii) genes. Thus, the yeast ARG4 gene was shown to complement the arg7 mutation in C. reinhardtii (Rochaix and vanDillewijn, 1982). Unfortunately, the transformation rate ( cell) was not much higher than the frequency of reversion. Although transformants apparently contained integrated copies ofthe introduced plasmid DNA, genetic crosses to demonstrate that the phenotype was due to the introduced gene were not performed, and the hybridization pattern changed when transformed strains were maintained under nonselective conditions. Putative replication origins were identified by cloning C. reinhardtii restriction fragments into the ARG4 transformation vector (Rochaix et al., 1984). A number of C. reinhardtii transformants were isolated that maintained the introduced DNA in an unintegrated state. The unintegrated DNA was gradually lost when the cells were cultured under nonselective conditions, however, and the recovered plasmids, which presumably contained functional replication origins, did not result in higher transformation rates than the original transformation vector, suggesting that integration into the genome is not the step that limits nuclear transformation rates (Rochaix et al., 1984). Shortly thereafter, the E. coli nptII gene from Tn5 (encodes neomycin phosphotransferase) was reported to confer resistance to G418 and kanamycin and appeared to have promise as a dominant selectable marker (Hasnain et al., 1985). Expression of this gene was driven by the SV40 promoter in plasmids that contained the yeast 2 micron origin of replication. It and conferred G418 resistance at the rate of the plasmid DNA appeared to remain unintegrated. Unfortunately, this marker turned out not to be very useful, possibly because of the high rate of spontaneous resistance to kanamycin and G418 or silencing of the introduced DNA (see below). The breakthrough in C. reinhardtii nuclear transformation resulted from two factors: the use of C. reinhardtii genes as selectable markers and the development of efficient procedures for introducing the DNA into the nucleus. The isolation of genes encoding argininosuccinate lyase (Arg7; Debuchy et al., 1989) and nitrate reductase (Nit1; Fernández et al., 1989) provided markers with excellent selectable phenotypes. Particle bombardment allowed the recovery of small numbers of transformants ( cell) with fairly high numbers of integrated copies (Debuchy et al., 1989; Kindle et al., 1989). The rate

43 of cotransformation was very high (Kindle et al., 1989; Day et al., 1990), which facilitated the isolation of transformants expressing genes that had no selectable phenotype (Diener et al., 1990). The development of the glass bead method, in which cells are transformed simply by agitating them vigorously with DNA in the presence ofpolyethylene glycol and glass beads, made the technology accessible to anyone with a vortex mixer (Kindle, 1990). Since then, methods for electroporation (Brown et al., 1991; Tang et al., 1995) and agitation in the presence of silicon carbide whiskers (Dunahay, 1992) have been developed, as well as a number of molecular tools and specific applications, which are discussed in detail below.

III. Selectable Markers The most effective selectable markers are those for which there is a strong phenotypic selection, so that there is little growth of nontransformed cells and vigorous growth of transformant colonies. Further­ more, the rate of reversion (for complementation of auxotrophic or nonphotosynthetic mutants) or spontaneous resistance (for drug resistance markers) should be as low as possible. A relatively short coding region facilitates molecular manipulations to create new vectors, and analysis of insertion events following nuclear transformation is more straight­ forward if the gene does not contain repeated sequences. Depending on the application, it may be advantageous to have a marker where even a low level of gene expression affords a robust growth phenotype.

A. Complementation of C. reinhardtii Mutations The Arg7 gene encodes argininosuccinate lyase. It spans 7 kb and is interrupted by 12 introns, which contain highly repeated sequences (Debuchy et al., 1989; Purton and Rochaix, 1994a). It complements arg7 and arg2 mutations, which map to the two ends of the ARG7 locus, and transformants are selected on acetate plates that lack arginine. The insertion of a 400 bp fragment from bacteriophage into an intron of the Arg7 gene provides a molecular marker that facilitates DNA blot analysis of putative Arg7 transformants (Gumpel and Purton, 1994). An Arg7­ based cosmid has been constructed, and was used to independent produce a genomic library with

Karen L. Kindle

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clones, representing approximately 250 genome equivalents (Purton and Rochaix, 1994a). The Nit1 gene encodes nitrate reductase, which alleviates the ammonium requirement of nit1 mutants, to allow transformants to grow on media with nitrate as the sole nitrogen source (Fernández et al., 1989; Kindle et al., 1989; see Chapter 33, Fernández et al.; Nia1 is the same as Nit1). It contains 15 introns and spans a region of about 8.5 kb (D. Zhang, M. Lavoie, S. Christenson and P.A. Lefebvre, unpublished). For transformation with Nit1, a nit1 NIT2 strain must be used; Nit2 is a regulatory gene that is required for expression of Nit1. Many of the strains in the Chlamydomonas Genetics Center collection are derived from a ‘wild-type’ background that actually carries nit1 and nit2 mutations and hence are inappropriate for transformation with Nit1 (Harris, 1989). Mutants defective at the nit8 (nar2) locus also require ammonium for growth (Nelson and Lefebvre, 1995a), and transformants can be selected on nitrate plates following introduction of a 3.2-kb DNA fragment carrying the Nit8 (Nar2) gene (S.-C. Wang and P. A. Lefebvre, personal communication). The Nic7 gene complements a nicotinamiderequiring mutation (Ferris, 1995). Although it has not been used extensively, it should be a good transformation marker, since reversion of the nic7 mutation is rare and background growth of mutant cells can be reduced by growing them in the presence of 3-acetylpyridine. The Thi10 gene has also been used to complement a thiamine auxotroph, though untransformed cells apparently die slowly (Ferris, 1995). The CRlpcr-1 (Por1) gene encodes light-dependent NADPH:protochlorophyllide oxidoreductase, which carries out the final step in chlorophyll synthesis. It complements the yellow, light-sensitive phenotype of mutants such as pc1y7, which are defective in both light-independent and light-dependent protochloro­ phyllide reductase (Li and Timko, 1996). The pc1 and the gene is mutation reverts at a low rate relatively small, since a 3.6 kb fragment retains the ability to complement the pc1 mutation (our unpublished observations). A number of cloned genes complement the acetaterequiring phenotype of nonphotosynthetic mutants. For example, Oee1 encodes the 33 kD protein of the oxygen evolving complex and complements a transposon-induced mutation (Mayfield and Kindle, 1990). AtpC encodes the gamma subunit of chloroplast ATPase, and complements an insertion

mutation (Smart and Selman, 1991, 1992). RbcS1 and RbcS2 are linked genes that encode the small subunit of Rubisco and each complement a deletion mutation that was generated in the course of insertional mutagenesis (Khrebtukova and Spreitzer, is defective in chloroplast petA mRNA 1996). accumulation, and the gene that complements the defect has been cloned. (Gumpel et al., 1995). Finally, cytochrome the Ccs1 gene is required for synthesis and complements an insertion mutation (Inoue et al., 1997). Although all ofthese genes have good selectable phenotypes, they are not particularly useful as coselectable markers or for insertional mutagenesis in investigations of chloroplast biogenesis, since mutations that affect chloroplast biogenesis often impair photosynthesis themselves.

B. Drug Resistance Markers Drug resistance markers have a substantial advantage over markers that complement C. reinhardtii mutations since they can be used in any genetic background, thereby circumventing the need to construct appropriately marked strains as trans­ formation recipients. One such marker is the C. reinhardtii Cry1 gene, which confers resistance to the cytosolic translation inhibitors cryptopleurine and emetine, due to a mutation in the coding region of cytosolic ribosomal protein S14. Although the Cry1 gene confers cryptopleurine resistance at a high frequency, it is semi-dominant, so existing ribosomes must be depleted by a period of nitrogen starvation and regenerated with the mutant gene product before the resistance can be expressed (Nelson et al., 1994). A number of studies have utilized bacterial genes that confer resistance to aminoglycoside antibiotics, such as the nptII gene. When nptII was fused to the nopaline synthetase promoter and 3´ flanking regions from the Agrobacterium tumefaciens Ti plasmid, about half the transformants selected for growth on nitrate plates following cotransformation with Nit1 were resistant to low levels of kanamycin. At least one of these transformants appeared to synthesize a polypeptide with enzymatic activity (Hall et al., 1993). A very low frequency transformation event was observed with an octopine synthase/ nptII chimeric gene. In this transformant, the polypeptide was larger than expected, suggesting that it may have been a fusion protein resulting from a rare in-frame insertion into an expressed coding

Chapter 4 Nuclear Transformation region. Recently, it was reported that stable paromomycin-resistant C. reinhardtii transformants using were recovered at a low frequency the aminoglycoside 3´-phosphotransferase typeVIII gene from Streptomyces rimosus, which has a high GC content and weakly biased codon usage (Sizova et al., 1996). Higher frequency transformation (~ 1 % of the rate with endogenous C. reinhardtii genes) was recently reported when the Streptoalloteichus hindustanus ble gene, which encodes a bleomycin binding protein and confers resistance to phleomycin, was linked with 5´ and 3´ regulatory sequences from the RbcS2 gene (Stevens et al., 1996). Since antibiotic resistance seems to vary with different strains, media, and lots of antibiotics, it should be empirically determined for each application (J. Moseley and S. Merchant, unpublished; our unpublished observations). A similar construct containing the E. coli aadA coding region, which encodes aminoglycoside adenine transferase, confers resistance to spectinomycin and streptomycin (Cerutti et al., 1997b). High frequency transformation was reported with a construct in which the chloramphenicol acetyl transferase gene from Tn9 was cloned between the 35S promoter of cauliflower mosaic virus and the 3´ region of nopaline synthetase; it was introduced into C. reinhardtii cells by electroporation (Tang et al., 1995). Finally, when the hexose transporter gene from Chlorella was fused to the constitutive Volvox tubulin promoter and transformed into Volvox, it conferred increased sensitivity to the toxic glucose/ mannose analog 2-deoxyglucose. In Volvox the gene can be selected either negatively, as just mentioned, or positively, since the cells survived for prolonged periods in the dark in the presence of glucose when the gene was expressed (Hallman and Sumper, 1996). This could be a very useful negative selectable marker, if it can be expressed in C. reinhardtii. Thus, a wide variety of C. reinhardtii genes and bacterial drug resistance markers are available for nuclear transformation, and the number will certainly increase as wild-type versions ofthe genes disrupted by insertional mutagenesis are isolated (see below).

IV. Methods for Introducing DNA into the Nuclear Genome of C. reinhardtii Each of the methods that have been developed for nuclear transformation of C. reinhardtii has certain

45 advantages, depending on the application.

A. Particle Bombardment Reliable nuclear transformation was first reported using particle bombardment (Debuchy et al., 1989; Kindle et al., 1989; Diener et al., 1990; Mayfield and Kindle, 1990). In this method, DNA is precipitated onto the surface ofsmall particles (usually tungsten) and accelerated towards the target cells by a gun powder charge or high-pressure helium. The number of colonies per bombarded plate was fairly small with the gun powder version of the device (5–50); somewhat higher levels of transformation can be achieved with high pressure helium (~200 colonies per plate). A fairly large amount of DNA is probably delivered to cells that survive the trauma, and the number of gene copies that integrate into nuclear DNA is quite high. The particle gun works better with walled than wall-less strains, and seems to result in a slightly higher rate of homologous recombination than the glass bead method described below (Kindle, 1990; Sodeinde and Kindle, 1993).

B. Glass Beads, Silicon Whiskers High frequency transformation can be obtained by agitating cells on a vortex mixer in the presence of DNA, glass beads, and 5% polyethylene glycol (PEG; Kindle, 1990). The simplicity and efficiency of this procedure, as well as the fact that it requires no specialized equipment, has made it the method of choice for most applications. This method requires cell wall-deficient cells, either a cw mutant, or cells that have been rendered cell wall-deficient by treatment with gamete lytic enzyme (autolysin). Glass bead transformation with Nit1 can be carried out with walled cells, as long as the cells are incubated in ammonium-deficient medium for several hours prior to transformation (Nelson and Lefebvre, 1995b; Kindle, 1998). The conditions under which PEG stimulates transformation are still controversial. Stevens et al. (1996) have suggested that the nature of the cell wall mutation determines whether PEG improves transformation rates, though we have not found a strain or marker for which PEG is not effective (unpublished observations). Glass bead transformation results in a substantially lower number of integrated gene copies than particle bombardment, and the number of copies appears to be proportional to the amount of added DNA (Kindle, 1990). Single

Karen L. Kindle

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or low copy number integrations are highly advantageous for certain applications such as insertional mutagenesis, where a single insertion event greatly facilitates subsequent analysis and gene isolation. Silicon carbide whiskers can be employed in place of glass beads, and their use results in transformation with substantially lower cell rates of lethality during the vortexing period (Dunahay, 1992; Dunahay, 1993). Furthermore, it is not necessary to remove cell walls prior to whisker transformation. The main disadvantage ofthis method is the potential health hazard and waste disposal problem caused by the biohazardous nature of the silicon carbide microfibers.

C. Electroporation Low frequency stable nuclear trans­ formation has been reported by using single or double electrical discharges during electroporation (Brown et al., 1991; Keller, 1995) for wall-less or walled cells, respectively. Very high frequency nuclear was reported with a transformation bacterial gene encoding chloramphenicol acetyl transferase, with asomewhatdifferentelectroporation set-up involving a DC-shifted radio frequency wave (Tang et al., 1995). A fraction of transformants generated by this technique contained unintegrated copies of introduced plasmid DNA, though the unintegrated DNA declined after prolonged periods in culture. Among stable transformants, the number of integrated copies was low in most cases. Recently, an abstract also reported very high frequency transformation by electroporation (up to cell) ofcell wall-deficient strains with a 12 kb plasmid carrying Arg7 (Shimogawara et al., 1998). V. Reporters and Promoters The analysis of gene expression and the isolation of regulatory mutants is greatly facilitated by reporter genes encoding proteins that can easily be assayed, screened, or selected. For analysis ofprotein function or to make dominant negative mutants, it is important to express an altered coding region at a high level. If the expression of a given gene is anticipated to have a negative effect on the cell, it may be necessary to use a regulated promoter. The expression of antisense constructs or genes expected to have a dominant

negative effect has not yet been reported for Chlamydomonas, though a number of useful molecular tools that may have utility for these applications have emerged.

A. Arylsulfatase as a Reporter Probably the most useful reporter described to date is the periplasmic enzyme arylsulfatase (Ars), which functions in sulfur acquisition from aromatic compounds under conditions where other sulfur sources are unavailable (de Hostos et al., 1989; Chapter 32, Davies and Grossman). Expression of the Ars gene is completely repressed under normal sulfur-replete growth conditions, and sensitive, chromogenic substrates are available for plate and enzymatic assays (Davies et al., 1992; Ohresser et al., 1997). The Ars gene has been used as a reporter for gene expression from the promoters of TubB2 Davies et al., 1992), Cyc6 (cytochrome Quinn and Merchant, 1995), CabII-1 (Jacobshagen (mitochondrial carbonic et al., 1996), anhydrase; Villand et al., 1997), and Nit1 (nitrate reductase; Ohresser et al., 1997). Transformants containing the TubB2/Ars fusion gene expressed it at a much lower level than the endogenous TubB2 gene. Furthermore, no increase in Ars enzymatic activity was detected following deflagellation, although the abundance of the chimeric mRNA increased transiently (Davies et al., 1992), with a time course identical to endogenous TubB2 transcripts. The arylsulfatase protein is very stable, which may explain why Ars activity did not vary as much during a 24 h light-dark cycle as chimeric CabII-1/Ars mRNA (Jacobshagen et al., 1996). However, Ars enzyme activity increased in response to a shift from Cureplete to Cu-free medium in transformants containing a construct in which the Ars coding region had been fused to the Cyc6 promoter (Quinn and Merchant, 1995). Furthermore, a dramatic (> 100­ fold) increase in Ars activity was measured within four hours of a shift from ammonium-containing to nitrogen-free medium in transformants carryingNit1/ Ars chimeric constructs (Ohresser et al., 1997). A very sensitive method for detecting Ars activity has been reported recently in which is used as the substrate and a diazonium salt as the post-coupling reagent. This resulted in a 250-fold increase in sensitivity over 5-bromo-4-chloroindolylsulfate ( Ohresser et al., 1997). Because of the increase in sensitivity, it was possible to study

Chapter 4 Nuclear Transformation promoter function in pools of co-transformed colonies, rather than isolating and analyzing individual transformants, which should correct for variations in transcription activity due to different genomic insertion sites. and can be used to Both stain cells on agar plates to distinguish colonies that express Ars from those that do not. This is useful in identifying both transformants that express the reporter gene as well as mutants that fail to express it. Davies et al. (1994) have used this screen to isolate mutants defective in the expression ofthe endogenous Ars gene.

B. Other Reporters Another extracellular enzyme with the potential to be a good reporter is alkaline phosphatase, since sensitive enzymatic and plate assays are available. C. reinhardtii alkaline phosphatases have recently been characterized (Quisel et al., 1996). Expression of the gene encoding protochlorophyllide oxidoreductase is an easily screened visual marker that can be used both for selection (Li and Timko, 1996), as described above, or potentially as a reporter. The Rsp3 gene, which complements a motility defect in the paralyzed flagellar mutant pf14 (Diener et al., 1990), has recently been used as a reporter to identify mutants that express the Nit1 gene constitutively (Zhang and Lefebvre, 1997). A Nit1/ Rsp3 gene fusion was shown to confer motility when transformed into the pf14 strain, but only in nitrate medium, conditions under which the Nit1 gene is expressed. Twenty-one mutants that swam in the presence of ammonium were found to express both the reporter gene and the endogenous Nit1 gene constitutively, identifying at least two loci that regulate genes in the nitrate assimilation pathway.

C. Constitutive Promoters The most widely used constitutive promoter to date has been that from RbcS2 (Goldschmidt-Clermont and Rahire, 1986), which was used to express an epitope-tagged, non-acetylatable to 40– 70% of the total a five-fold increase over expression from the TubA1 promoter (Kozminski et al., 1993). The RbcS2 promoter has since been used in constructs to express the Cry1, ble, and aadA genes (Nelson et al., 1994; Stevens et al., 1996; Cerutti et al., 1997b). Other constitutively expressed

47 promoters that have been shown to be active in chimeric constructs include those from PetE (or Pcy1, encodes plastocyanin) and AtpC (encodes the gamma subunit of chloroplast ATPase) which have been fused to the coding region of Cyc6 (Quinn and Merchant, 1995), CabII-1, which has been fused to the nitrate reductase coding region (Blankenship and Kindle, 1992), and the and CabII-1 promoters, which were fused to arylsulfatase, as described above. RbcS2 is generally regarded as the strongest nuclear promoter tested to date. Although CabII-1 mRNA accumulates to a high level in vivo, transformants carrying a tagged version of this gene accumulated only a very low level of the marked transcript. Furthermore, chimeric CabII-1/Nit1 constructs that included up to 8.5 kb of upstream sequence were expressed only at a low level. However, wild-type levels of cytochrome accumulated when this gene was derived by the AtpC and PetE promoters (Quinn and Merchant, 1995). A comparison of the expression of the same reporter gene driven by each of these promoters would clearly be useful.

D. Regulated Promoters Many genes are regulated in response to environ­ mental conditions or during development and are therefore sources of potentially useful promoters for nuclear transformation. Some of these are expressed constitutively, but further induced in response to which is induced certain conditions, such as when cells are deflagellated. Others are expressed at a very low level until they are induced in response to specific stimuli. Table 1 shows a list of regulated promoters whose function has been tested by fusion to a reporter gene. The CabII-1 gene is expressed constitutively in mixotrophically grown cells, but is induced in response to light in synchronized cells grown under phototrophic conditions (Shepherd et al., 1983; Kindle, 1987), apparently by a circadian rhythm (Hwang and Herrin, 1994; Jacobshagen and Johnson, 1994). The Nit1 gene is induced by growth in nitrate medium and derepressed in nitrogen-deficient medium (Fernández et al., 1989; Quesada and Fernández, 1994; Chapter 33, Fernández et al.; Nia1 is the same as Nit1). Cyc6 expression is regulated at the level of transcription, which is induced when copper is limiting (Merchant et al., 1991; Quinn and Merchant, 1995). In addition to its response to gene expression varies deflagellation,

48

during the cell cycle (Silflow and Rosenbaum, 1981; Nicholl et al., 1988), while the mitochondrial carbonic anhydrase genes Ca1 and Ca2 are among a number of nucleus-encoded genes whose expression is (Villand et al., induced by low (air) levels of 1997). Conditional gene expression would be particularly important for certain experimental approaches. For example, with dominant negative mutants or antisense constructs, where the gene product might have a deleterious phenotype, it may be essential to keep the gene silent while the cells are growing. The gene could then be induced to determine its phenotypic effect. Alternatively, ifa gene is required for viability, one could introduce a conditionally expressed copy and then generate an insertion mutation in the endogenous (constitutively expressed) gene. It might then be possible to assess the phenotype of a null mutation by transferring the cells to conditions that prevent the expression ofthe introduced gene. Neither of these approaches has yet been taken to examine gene function in C. reinhardtii, partially due to problems with gene silencing and the difficulties in generating gene-targeted disruption mutants. However, the characterization of these regulated promoters is a first step toward developing these sophisticated genetic approaches. Several issues are important for the use of conditional promoters. The first is how much residual transcription takes place under repressed or noninduced conditions. This is critical if expression of the gene product is expected to be deleterious, since it might determine whether transformants capable of expressing the gene under inducing conditions can be recovered. The second issue is how highly expressed the genes are when they are induced/ derepressed, which may determine whether a phenotypic effect can be observed. Finally, since gene silencing seems to be a significant problem in C. reinhardtii (see below), it would be useful to know

Karen L. Kindle

how long a gene that is maintained under noninducing or repressing conditions maintains the ability to be activated upon transfer to appropriate conditions. When the Nit1 (Ohresser et al., 1997) and Cyc6 (Quinn and Merchant, 1995) promoters were fused to the arylsulfatase coding region and transformed into C. reinhardtii, the expression of arylsulfatase activity under non-inducing conditions varied from undetectable to very low. Ars activity increased by a factor of 100–1000 upon induction using the Nit1 promoter and up to 80-fold when the Cyc6 promoter was induced. Similarly, when the Ca1/Ca2 promoter was fused to Ars, both mRNA and enzymatic activity and increased were undetectable in 5% dramatically upon a shift to low (Villand et al., 1997). It would be interesting to compare expression at the mRNA level ofthese constructs under inducing and repressing conditions, to determine which promoters exhibit the tightest regulation and highest levels of induced gene expression. A priori Nit1, Cyc6 (and perhaps Ars) would seem to be good promoters for genes to be kept silent, since they are turned off in normal culture medium, which contains ammonium, copper, and sulfur. Alternatively, the Ca1 or Ca2 promoter would be useful for genes that should be maintained in an expressed state, since they are induced in air, but repressed in 5% (Villand et al., 1997).

VI. Characteristics of Transformation Events In order to survive selection, transformed cells must replicate the introduced DNA and express the selectable marker at a level high enough to allow cell growth. Whether expression of the introduced DNA remains stable appears to depend to some extent on whether it remains under selection. The difficulty in achieving stable expression of foreign and C.

Chapter 4 Nuclear Transformation reinhardtii genes at a high level seems to have several facets, which are discussed below.

A. Nature and Stability of Introduced DNA The best characterized markers (Nit1 and Arg7) and methods for introducing DNA into C. reinhardtii cells (glass bead transformation and particle bombardment) result primarily in transformants that contain integrated copies of the introduced DNA (Debuchy et al., 1989; Kindle et al., 1989; Kindle, 1990). The restriction pattern of introduced DNA is for the most part stable following extended periods in culture, whether or not the cells are maintained under selection (Kindle et al., 1989; Day et al., 1990; Cerutti et al., 1997a), though minor differences have occasionally been observed (Cerutti et al., 1997a). The restriction pattern of the introduced DNA is also stable through meiosis (Kindle et al., 1989; Diener et al., 1990). A recent report suggested that some transformants generated by electroporation contained unintegrated plasmid DNA (Tang et al., 1995). When undigested DNA from these transformants was separated by electrophoresis, a band that hybridized to the introduced plasmid migrated considerably ahead of chromosomal DNA. Furthermore, the introduced plasmid could be separated from the main nuclear DNA on CsCl gradients and recovered by electro­ poration into E. coli. It is unknown whether the observation of unintegrated DNA is a consequence of the electroporation method or the selectable marker. It is tempting to speculate that the high rate of may transformation with a foreign gene have resulted because a large fraction oftransformants harbor unintegrated gene copies, which might be less subject to silencing by chromosomal mechanisms (see below). If this electroporation method also results in the transient maintenance of introduced C. reinhardtii DNA in an unintegrated state, the isolation of genes by functional complementation could be facilitated, since it might then be possible to rescue into E. coli the gene responsible for complementing the defect.

B. Insertion Events Although the number of insertion events appears to depend on the amount of DNA delivered into the nucleus, the nature of the insertions appears to be similar, regardless of whether particle bombardment

49 or glass bead-mediated transformation is used to introduce DNA into the cell. Insertions nearly always take place in ectopic locations, since DNA blot analysis indicates that the endogenous gene is unaltered, and the introduced gene(s) segregate independently of the endogenous one in genetic crosses (Kindle et al., 1989; Diener et al., 1990; Mayfield and Kindle, 1990; Nelson and Lefebvre, 1995b). From these observations, it has been suggested that insertion occurs at random ectopic locations, though the high rate at which some insertional mutations are recovered suggests that integration may occur preferentially in some areas of the genome (Wilkerson et al., 1995). The insertion of linearized plasmid DNA appears to occur through its ends, usually with relatively small terminal deletions (Blankenship and Kindle, 1992; Cerutti et al., 1997b). Insertions can result in arrays with multiple copies of the introduced DNA or alternatively in a single integrated gene copy. Several unlinked single copy insertions have been documented in some transformants (Pazour et al., 1995). Multiple gene copies are usually inherited together in the progeny of genetic crosses, although occasionally one or more of multiple integrated DNA copies will segregate independently (Kindle et al., 1989; Pazour et al., 1995). A simple tandemly repeated array is ruled out by the complexity of hybridization patterns in DNA blots. Markers that are cotransformed on independent replicons are often genetically linked (Diener et al., 1990; Zhang and Lefebvre, 1997). It is likely that when the introduced plasmid DNA is present in the cell at a sufficiently high concentration, the molecules join together prior to integration. To determine whether homologous recombination between introduced molecules is efficient, two different truncated Nit1 or Arg7 genes were cotransformed into an appropriate mutant strain. Neither truncated gene could by itself complement the mutation in the recipient strain, but an extrachromosomal homologous recombination event would generate a fully functional gene. Using the two truncated genes, cotransformants were recovered at nearly as high frequency as with a single intact gene, suggesting that extrachromosomal homologous recombination is efficient (Sodeinde and Kindle, 1993; Gumpel et al., 1994). In high copy number transformants generated with the RbcS2/aadA/RbcS2 construct, genomic DNA fragments consistent with tail to tail, head to head, and head-to-tail repeats were observed (Cerutti et al., 1997b). Evidence for a small

50

number of tandemly repeated copies in low copy number transformants has been obtained both for targeted (homologous) integration events (Sodeinde and Kindle, 1993) and for integrations at ectopic sites (Cerutti et al., 1997b). Together, these results suggest considerable end-to-end intermolecular joining of the introduced plasmid DNA prior to integration.

C. Cotransformation and Expression of C. reinhardtii Genes Because both particle bombardment and glass bead transformation can result in the introduction of multiple copies of the transforming DNA, the rate of cotransformation of an unselected DNA is very high (Kindle et al., 1989; Day et al., 1990; Diener et al., 1990; Kindle, 1990). This is important for identifying transformants that contain genes that have an unknown or unselectable phenotype. In the case of C. reinhardtii genes, most cotransformants that contain the unselected DNA appear to express it, at least initially. However, the rate of expression of an gene in introduced non-functional transformants was increased significantly by putting both the selected and unselected genes on the same plasmid (Periz and Keller, 1997). We have noted that cotransformants that initially express plastocyanin genes with mutations that prevent chloroplast protein import and accumulation ofthe mature protein fail to express the gene at the mRNA level after various periods in culture (unpublished observations). Introduced genes may tend to be silenced during culture, so that only when gene expression confers a selective advantage, would gene expression persist. If gene expression is deleterious, cells in which gene expression has been silenced would be selected rapidly. Genes on the same piece ofDNA presumably integrate into the same locus, so that maintaining selection for one gene may prevent silencing of the unselected gene. There is a substantial difference in the levels of gene expression among different transformants with the same construct, e.g. (Blankenship and Kindle, 1992; Quinn et al., 1993; Davies et al., 1994), presumably due to differences in transcription caused by the particular genomic insertion site, so called position effects. In some cases, actively expressed reintroduced C. reinhardtii genes accumulate mRNA at a level comparable to the endogenous gene, e.g. Nit1 (Kindle et al., 1989) and PetE (or Pcy1,

Karen L. Kindle plastocyanin; Quinn et al., 1993) while for other genes, the level of mRNA or protein is much lower, e.g. CabII-1 (Blankenship and Kindle, 1992), Cyc6 (Quinn and Merchant, 1995), TubA Kozminski et al., 1993; Periz and Keller, 1997), and Davies et al., 1994). In the case of TubB2 chimeric genes containing the C. reinhardtii CabII­ 1 promoter linked to the Ars and Nit1 coding regions, the level of mRNA accumulation was somewhat higher than that of Ars and Nit1 under induced conditions, but significantly lower than CabII-1 (Blankenship and Kindle, 1992; Jacobshagen et al., 1996), even though 8.5 kb of upstream CabII-1 promoter sequences were integrated in some transformants. In most cases, the level ofexpression of introduced C. reinhardtii genes is not correlated with the number of introduced gene copies (Blankenship and Kindle, 1992; Quinn et al., 1993). An interesting exception was the Oee1 gene, which encodes the 33 kD polypeptide of the oxygen evolving complex. In this case, gene expression was proportional to the number of introduced gene copies, so that transformants harboring three to five introduced copies accumulated the polypeptide in excess of wild-type (Mayfield, 1991). Perhaps a locus control region, defined as an element that results in gene expression that is independent of integration site but proportional to the number of introduced gene copies (Dillon and Grosveld, 1993), lies close to Oee1 and was included in the cloned DNA that was used for transformation.

D. Expression of Foreign Genes/Silencing The difficulty inestablishing anucleartransformation system for C. reinhardtii was largely due to the difficulty, not yet fully understood, in expressing foreign genes. The failure to express non-C. reinhardtii genes may be attributed to a number of factors, including the inability to recognize regulatory sequences from heterologous sources, gene silencing of integrated foreign gene copies, or inefficient translation and/or instability of foreign transcripts, which could be a consequence of differences in codon usage compared to the very biased codon usage of C. reinhardtii nuclear genes. It seems likely that some of the problem with foreign gene expression is due to the coding region, since chimeric foreign genes with C. reinhardtii promoters and 5´ and 3´ untranslated regions are expressed at a low frequency (Stevens et al., 1996; Cerutti et al., 1997b; see below)

Chapter 4 Nuclear Transformation or at a frequency too low to be detected (Blankenship and Kindle, 1992). However, differences in codon usage seem unlikely to be the major reason for the difficulty in expressing foreign coding regions. The ble gene from Streptoalloteichus hindustanus has codon usage similar to that in C. reinhardtii, and although phleomycin-resistant transformants could be selected directly, the number of transformants was still about two orders of magnitude lower than those obtained using a homologous C. reinhardtii transformation marker (Stevens et al., 1996). A similarly low recovery of spectinomycin-resistant transformants was reported when the RbcS2 promoter was used to drive expression of aadA (Cerutti et al., 1997b). Furthermore, a careful study of aadA transformants indicated that the spectinomycin­ resistant phenotype of many of them was unstable when cells were grown under nonselective conditions, indicating that gene silencing can occur even after the initial selection (Cerutti et al., 1997b). Thus, there appear to be at least two levels at which foreign genes may fail to be expressed. Assuming that the same kinds of integration events occur regardless of the nature ofthe introduced DNA, it appears that 97– 99% of insertion events fail to lead to the expression of foreign genes. Alternatively, it is possible that expression of foreign genes is affected in all locations, and it is only those that integrate into extremely active sites in the genome that are expressed well enough to survive selection. The instability of expression following the initial integration and selection appears to be an independent problem, which affects about half of the selected transformants (Cerutti et al., 1997a). As mentioned above, even reintroduced C. reinhardtii genes may be subject to this kind of silencing. The extent of methylation of the CabII-1 promoter, assessed by digestion with methylation-sensitive restriction enzymes, was related to the expression status ofintroduced chimeric constructs: unexpressed ) CabII-1/uidA (uidA encodes constructs were hypermethylated relative to CabII1/Nit1 constructs that were expressed at a level high enough to allow growth on nitrate medium (Blankenship and Kindle, 1992). Cerutti and coworkers have analyzed gene silencing using the RbcS2/aadA gene. They have shown that silencing and reactivation are reversible and that they are not a consequence of loss or rearrangement of the integrated gene copies, as determined by digestion with restriction enzymes. Neither were the instances

51 of gene silencing examined correlated with changes in methylation status or chromatin structure, as assessed by sensitivity to DNase (Cerutti et al., 1997a). An intriguing possibility is to use the unstable expression of aadA to search for mutants that lose the ability to silence introduced gene copies. It might be possible to do this by selecting for strains that reactivate an unstably silenced gene copy to a stably expressed state. Such a mutant might be defective in gene silencing and therefore transformable by foreign genes at a high rate.

VII. Insertional Mutagenesis and Gene Tagging

A. Development and Application of the Approach The observation that transforming DNA integrates into the nuclear genome at apparently random locations by nonhomologous recombination sug­ gested that these insertions might be mutagenic if they disrupted a gene. Furthermore, the inserted DNA would act as a molecular tag that would allow the mutant gene to be isolated. (See Fig. 1 for an outline of this approach.) Tam and Lefebvre (1993) pioneered this powerful approach by generating a series of motility-defective mutants following transformation with theNit1 gene. They demonstrated phenotype that for more than 85% of them, the cosegregated with the motility defect in genetic crosses, suggesting that the insertion of the selectable marker was the mutagenic event. The C. reinhardtii DNA that flanked the inserted DNA was cloned for two mutants, using two different approaches. In the first case, the plasmid DNA was intact, so that it could be recovered in E. coli by transformation using genomic DNA that had been digested with an appropriate restriction enzyme and self-ligated. In the second instance, the vector sequences were not intact, but a partial genomic library was prepared and hybridized with a Nit1 probe to take advantage of the tightly linked Nit1 gene. In both cases, the wild-type alleles were screened from a genomic library, and the functional gene was demonstrated to complement the mutant phenotype. An analysis of three of the insertion mutations revealed that in all three cases, there were rearrangements at the tagged locus. In two cases, the rearrangements were deletions, which ranged in size from 3–4 kb to over 23kb.

52

Insertional mutagenesis has been utilized to generate mutants defective in many cellular processes, including photosynthesis (Adam et al., 1993; Gumpel et al., 1995; Khrebtukova and Spreitzer, 1996; Inoue et al., 1997; M. Goldschmidt-Clermont, unpublished), motility (Wilkerson et al., 1995; Smith and Lefebvre, 1996), phototaxis (Pazour et al., 1995), sulfur and nitrate assimilation (Davies et al., 1994; Prieto et al., 1996a), and sensitivity to cadmium or high salt (McHugh and Spanier, 1994; Tang et al., 1995; Prieto et al., 1996b). The efficiency with which mutants were recovered varied dramatically in different studies, depending in part on the number of genes involved in a given process, i.e. the target size for mutagenesis, and on the conditions used for selection. For example, the recovery of nonphotosynthetic mutants has been reported to vary between 0.22% of transformants, when transformants were selected in dim light and then screened for failure to grow without acetate (Adam et al., 1993) to 1.5%, when transformants were selected in the dark and then screened for an acetate-requiring phenotype (Khrebtukova and Spreitzer, 1996). Many of the

Karen L. Kindle

mutants recovered in the latter study could be maintained only in the dark, suggesting that selection in the dark is important for recovering certain kinds of nonphotosynthetic mutants. The recovery of motility-impaired mutants was exceptionally high in one study (2.5%; Tam and Lefebvre, 1993) and the number of mutations in the single copy gene that encodes IC78, the intermediate chain of outer arm transformants) dynein, was so high (0.25% of the that it must represent a hotspot for integration and/or deletion (Wilkerson et al., 1995). Thus, it is very likely that insertional mutagenesis is not as random as once thought. A priori, there is no way to tell whether a gene is more or less likely than average to be disrupted. Therefore, it is impressive that mutations sought in two single copy genes required for photosynthesis, PsaF and AtpC, were recovered using this method. In both studies, the authors were originally trying to target mutations to these genes by homologous recombination. The mutations that were recovered, a deletion in PsaF (Farah et al., 1995) and what appears to be an insertion in AtpC (Smart and Selman, 1991), were probably initiated

Chapter 4 Nuclear Transformation by a random insertion event, though the deletion in PsaF could have resulted from abortive homologous recombination.

B. Technical Considerations It is advantageous to carry out insertional mutagenesis in such a way as to maximize the number of mutants in which a single insertion event has occurred, and in which both the marker and the vector sequences of the introduced DNA are intact. This facilitates genetic analysis, since the mutant phenotype and selectable marker should cosegregate, and increases the chance that the genomic DNA that flanks the insertion can be cloned by a plasmid rescue strategy. To accomplish this, the plasmid DNA should be linearized before transformation. This increases the likelihood that recombination will occur through the ends, rather than at a random break in the vector sequences (J. Woessner and M. Campbell, personal commun­ ication); the selectable marker must be intact in order to recover the transformant. Furthermore, the minimum amount of DNA necessary to generate a reasonable number of transformants should be used, to decrease the likelihood of concatemer formation prior to insertion or multiple integration events (Kindle, 1990). The Nit1 and Arg7 genes have both been used for insertional mutagenesis, and so far the rate of tagging for Nit1 appears to be quite high, generally about 80% (Tam and Lefebvre, 1993; Pazour et al., 1995). In some of the mutants characterized in detail by Pazour et al. (1995), the mutant phenotype cosegregated with an integrated piece of DNA, even though the functional Nit1 gene integrated elsewhere. In fact, it seems likely that even untagged mutations are due to deletions generated as a byproduct of insertion events, since the likelihood of a random mutation among the relatively small number of transformants examined in most studies (2000– 20,000) is very low. As already noted, deletions of the target locus are often associated with insertions of plasmid DNA (Tam and Lefebvre, 1993; Wilkerson et al., 1995).

C. Isolation of the Gene Responsible for the Mutant Phenotype Before embarking on a strategy to isolate the C. reinhardtii DNA that flanks the plasmid insertion, it

53 is important to establish that the selectable marker, or at least a part of the introduced plasmid DNA, cosegregates with the mutant phenotype in genetic crosses. A restriction analysis of the inserted DNA can then establish whether the vector is intact so that a plasmid rescue approach is likely to succeed or whether a hybridization strategy must be employed. Once the DNA flanking the insertion has been isolated, it can be used as a hybridization probe to demonstrate a restriction fragment length poly­ morphism (RFLP) between the mutant strain and the transformation recipient and also to isolate a wildtype genomic clone from an appropriate library. If there has been no deletion of genomic DNA at the insertion site, the cloned wild-type DNA should be able to complement the mutant phenotype, providing that the entire gene is present on the cloned fragment. If there has been a deletion, then the gene responsible for the mutant phenotype could be some distance away from the fragment that flanks the insertion site. It might therefore be necessary to perform a short walk to find the DNA fragment capable of complementing the mutant phenotype. If the mutant locus is tagged by an intact plasmid copy, the entire mutagenesis and gene isolation can be carried out within a few months, though in practice it often takes a while to hit upon the ideal screening strategy.

D. Perspectives The real power of insertional mutagenesis by nuclear transformation lies in the ease with which huge numbers of potentially tagged mutants can be created. It is easy to generate 10,000 transformants and not unreasonable to contemplate making 100,000. Much more labor may be involved in screening this large pool of potential mutants, so visual screens, such as those for high chlorophyll fluorescence are highly advantageous (Farah et al., 1995; Bennoun and Béal, 1997; Chapter 23, Bennoun and Béal). Because of the large number of mutants that can be generated, it should be possible to tag every gene in a given pathway that is not required for viability, assuming that insertional mutagenesis is reasonably random. Even if some genes cannot be tagged because they lie in a part of the genome where insertions do not readily occur or near genes that are required for viability, the number of mutations in interesting genes is likely to be large, as has already been demonstrated by information generated in the few

54 years since the technique has been available. Despite the potential of insertional mutagenesis, not all interesting genes can be isolated using this approach, which for the most part generates null mutations. Fortunately, alternative strategies for isolating genes by their ability to complement recessive mutations have been developed, and are described further below.

VIII. Gene Isolation by Complementation of a Mutant Phenotype Shotgun cloning requires a high efficiency trans­ formation system as well as a representative genomic library with large inserts. Complementation of the arg7 mutation with a total genomic cosmid library demonstrated the feasibility of selecting C. reinhardtii genes from a complex mixture by function (Purton and Rochaix, 1994b). Isolation of the transforming DNA from such a rescued mutant would be easier if it replicated autonomously in the host and could be recovered by transformation in E. coli. Since DNA introduced into the nuclear genome of C. reinhardtii usually integrates, an alternative strategy has been employed for isolating genes by function: the use of an indexed cosmid library. Zhang et al. (1994) produced a genomic cosmid library containing 11,280 clones with an average insert size of 38 kb. The individual cosmids were picked into the wells of 120 microtiter dishes and cosmid DNA was prepared from pools of clones and used in transformation experiments to complement the arg7 mutation. By identifying the individual microtiter dish that contained the DNA that gave rise transformants and then pinpointing the row to and column, a cosmid containing a functional Arg7 gene was identified. This library has recently been used to isolate the gene that complements the high -requiring mutant ca-1 (Funke et al., 1997). The gene that was recovered encodes an intracellular carbonic anhydrase and is identical to the previously identified Cah3 gene (Karlsson et al., 1995). The same library has also been used to isolate genes that rescue a nonphotosynthetic mutant that lacks psbB mRNA (F. Vaistij, M. Goldschmidt-Clermont, J.-D. Rochaix, unpublished) and a psaA trans-splicing mutant (C. Rivier, M. Goldschmidt-Clermont, and J.-D. Rochaix, unpublished). A genomic clone capable of complementing an arg7 nac2-26 cw15 mutant (Kuchka et al., 1989) and another one complementing

Karen L. Kindle a psaA trans-splicing mutant were isolated from a genomic cosmid library prepared in an Arg7 vector (Purton and Rochaix, 1994a; J. Nickelsen, K. Perron, M. Goldschmidt-Clermont and J.-D. Rochaix, unpublished) Although constructing the cosmid library and preparing the DNA pools necessary for this approach are rather labor-intensive, the transformations are straightforward, providing that the mutation is recessive and has a low reversion rate, and that transformants have a good selectable phenotype. Identifying the cosmid that complements the mutation is a matter of only two or three rounds of transformations. It is in some respects simpler than insertional mutagenesis strategies, since relatively little characterization of the mutant is required. Furthermore, this approach should allow the isolation of genes that complement mutations generated by chemical or UV mutagenesis, as well as insertional mutants that are not tagged or that have a complex insertion pattern. If the mutation of interest is dominant, e.g. a dominant suppressor mutation, the same strategy can be used, though it would be necessary in this case to prepare the genomic library from the suppressor strain and transform the DNA into the appropriate mutant strain.

IX. Homologous Recombination and Gene Targeting The ability to target endogenous genes for homolo­ gous recombination raises the possibility of disrupting their function or replacing them with altered versions. Therefore, considerable effort has been expended to develop technology for gene targeting in C. rein­ hardtii. As mentioned earlier, it is clear that C. reinhardtii contains the enzymatic machinery necessary for efficient homologous recombination, since transformation is nearly as frequent using two plasmids with non-overlapping mutations in Nit1 or Arg7 as with intact versions of the gene (Sodeinde and Kindle, 1993; Gumpel et al., 1994). Moreover, homologous recombination has been shown to occur between extrachromosomal DNA and the cognate endogenous gene, by using a Nit1 gene truncated at the 5´ end to repair the genomic copy. Molecular evidence for homologous recombination was presented and was consistent with single or multiple insertion events as well as gene replacement events. The estimated ratio of targeted integration and

Chapter 4 Nuclear Transformation replacement events to random insertion events was significantly higher for the particle gun (1:40) than for glass bead transformation (1:1000), though the reason for this was not established (Sodeinde and Kindle, 1993). For glass bead transformation, 70% of the events were consistent with single or multiple insertions, while about 30% were replacement events (Kindle and Sodeinde, 1994). Rare gene-targeted insertions have also been documented at the Arg7 locus (Gumpel et al., 1994). Several groups have tried to generate mutations by gene-targeting strategies, but have met with very limited success. Attempts to obtain a mutant with a disrupted AtpC gene by performing glass bead transformation in the presence of excess herring sperm DNA resulted in the isolation of a mutant that was apparently the consequence of insertion of the herring sperm DNA within the AtpC gene (Smart and Selman, 1991; our unpublished observations). Similarly, Farah et al. (1995) tried to target a mutation to PsaF by cotransforming an arg7 strain with the Arg7 gene and a disruption plasmid that contained two short deletions at either end of the PsaF gene. The disruption plasmid was linearized between these deletions so as to produce a gap. Among 22,000 transformants, 110 colonies with anomalous fluorescence induction kinetics suggestive of a possible PsaF defect were identified. They were then screened by hybridization to identify those that carried a PsaF deletion corresponding to the gap in the original transforming plasmid. The PsaF deletion mutant that was isolated did not have a structure consistent with a targeting event and probably arose as a consequence of a deletion following random insertion of one of the transforming plasmids or possibly from an abortive homologous recombination event. The gap would probably have been repaired during accurate homologous recombination in C. reinhardtii, as it is in yeast and mammals (Szostak et al., 1983; Valancius and Smithies, 1991). The only convincing evidence for a mutagenic gene targeting event was reported by Nelson and Lefebvre (1995a), who used a targeting plasmid in which the Nit8 gene was interrupted by the selectable Cry1 gene. Two substrates were used: 1) a fragment containing only the disrupted C. reinhardtii Nit8 gene; and 2) a plasmid linearized within C. rein­ hardtii DNA downstream of the disrupted Nit8 gene. The linearized plasmid would be expected to integrate by a single crossover, giving rise to a gene duplication that would not be mutagenic unless recombination

55

subsequently took place between the duplicated copies to eliminate the wild-type copy. The isolated fragment would be expected to replace the endo­ genous gene by a double recombination or gene conversion event, which would be mutagenic. (See Fig. 2 for an illustration of mutagenesis by insertion and replacement events.) Potential nit8 strains were enriched for by selecting for chlorate resistance and screened for a phenotype. No nit8 strains were recovered using linearized plasmid DNA, while eight nit8 mutants were recovered using the isolated fragment. Three of them showed evidence of a homologous recombination event, but only on one

56 side of the Nit8 gene. In each case the other end was generated by a more complex process, probably secondary to a nonhomologous recombination event. There were no simple homologous replacements. Double recombination events may be extremely rare in C. reinhardtii. Earlier work (Sodeinde and Kindle, 1993; Kindle and Sodeinde, 1994), suggested that single; recombination events are more frequent than homologous replacements. Furthermore, even the homologous replacement events that were observed may have arisen in two steps: an insertion followed by homologous recombination between the integrated copies. In the work with Nit1, either insertion or phenotype, replacement would have resulted in a while for disruption of Nit8, only a complete Nit8 replacement would have led to a chlorate-resistant phenotype. The frequency of such double recom­ bination events may be so low that even random insertions into the single copy gene or insertioninduced deletions are more frequent. It is interesting that in mammalian systems, insertion and replacement vectors target with similar efficiency (Thomas and Capecchi, 1987). In conclusion, it is clear that gene-targeting events in C. reinhardtii are very rare, occurring with a frequency comparable to that for random insertion in a specific gene. Perhaps the efficiency could be improved by performing transformation in syn­ chronized cells at a specific time in the cell cycle, when the chromatin might be less condensed and more likely to interact with exogenous DNA. Gene targeting might also be enhanced by transforming gametes and then immediately mating them, to increase the chance that they would contain a relatively high concentration of introduced DNA during meiosis, a time when chromosomes are known to undergo homologous recombination. It might also be helpful to induce DNA-repairpathways by treating cells with DNA damaging agents. The best template for gene targeting appears to be a linearized or gapped plasmid, with mutations engineered upstream and downstream of the linearization point, as shown in Figure 2. However, the introduced mutations should not lie too close to the linearization site or gap, since the targeting DNA may be digested from the ends prior to double-strand gap repair, which could correct the introduced mutations. Nelson and Lefebvre’s results (1995a) suggest that it may be advantageous to use plasmid DNA prepared from E. coli strains that are deficient in methylation. A strong selection or screen for targeting events is essential, given their

Karen L. Kindle low incidence. A scheme that would select for very rare homologous recombination events in genes for which there is no easy screen would clearly be very useful (Mansour et al., 1988).

X.The Use of Nuclear Transformation to Study Promoter Function Nuclear transformation has the potential to facilitate the analysis of a wide variety of cellular processes, some ofwhich are discussed elsewhere in this volume. Since promoter structure and function are important issues for technical aspects ofnuclear transformation, I will conclude this review with a more detailed examination of promoter structure and function, as assessed by transforming various constructs into the C. reinhardtii nucleus. As mentioned above and summarized in Table 1, a number of promoters have been linked to reporter genes and shown to confer transcriptional regulation. Regulatory sequences appear to be located fairly close to the transcription start sites, within 200–300 bp, in most cases (Davies et al., 1994; Quinn and Merchant, 1995; Ohresser et al., 1997; Periz and Keller, 1997; Villand et al., 1997). The promoters of four genes have been examined ), TubA1 in some detail: TubB2 (encodes ), Cyc6, and Nit1. The TubB2 (encodes promoter contains seven copies of a repeat, termed the tub box, which is also found upstream of other flagellar genes. Davies and Grossman (1994), made fusions of the TubB2 promoter and part of the 5´ untranslated region to the arylsulfatase coding region and examined accumulation of Ars mRNA in response to deflagellation and during the cell cycle in individual transformants that expressed Ars protein. They found that sequences between –95 and +65 relative to the mRNA 5´ end were sufficient for accumulation of Ars mRNA in response to deflagellation, which occurred with a time course indistinguishable from transcripts derived from the endogenous TubB2 gene (see Table 2). Transformants with sequences between –64 and +65 expressed the gene at a very low level, and there was no response to deflagellation. The –95 to +65 region contains 4 copies of the tub box, while the –64 to +65 region has two copies. By testing constructs in which various tub boxes were removed or mutated, it appeared that multiple copies ofthe tub box were necessary for regulated expression, and that the pair in the central region was most critical.

Chapter 4 Nuclear Transformation

Transformants expressing chimeric TubB2/Ars genes accumulated Ars mRNA during a specific part of the cell cycle, but considerably earlier than the peak of TubB2 mRNA. Constructs that contained sequences from –64 to +65 accumulated Ars mRNA, but showed a dampened rhythmicity during the cell cycle, while the region from –36 to +65 allowed low level constitutive expression. Thus, the region required for basal expression is very small, while the sequences required for cell cycle regulation and response to deflagellation are partially overlapping but distinct, lying within 100 bp of the transcription start site. Since only steady-state mRNA levels were examined, the possibility of differential mRNA stability, due to sequences from +1 to +65 cannot be eliminated. Since the differences in expression among individual transformants from a single construct were greater than the variation between constructs, it was not possible to define regions that affect TubB2 expression quantitatively. Periz and Keller made a series of 5´ deletions in a TubA1 gene that had been marked by a 233 bp deletion in the coding region (Periz and Keller, 1997).

57

The plasmids also contained the Arg7 gene, and 60– 70% of transformants expressed the test gene. By testing both pools of transformants and individual transformants for induction ofthe test and endogenous TubA1 genes in response to acid-induced deflagel­ lation, two regions were found to be important for the deflagellation response. An element between –176 and –122 was important for determining the magnitude of the response, but not the time course, while deleting a region between –85 and –56 abolished the deflagellation response. The region between –85 and –16 was sufficient to confer a deflagellation response on a Cyc6/Ars chimeric construct. Addition of tub box elements to this construct had no apparent effect, but it was noted that the –85 to –16 region contains elements common to other rubulin promoters that could play an important role in the deflagellation response, including an ATB (tub-associated) box (TTCGGGG), and a GC-like box (CGGGCG.) Quinn and Merchant (1995) identified two elements between –127 and –56 relative to the transcription start site that regulate Cyc6 transcription in response to copper limitation (see Table 2). These elements

58 appear to be transcriptional activators, since when they were fused to the basal TubB2/Ars construct, they stimulated expression in response to copper limitation, but did not reduce expression in the presence of copper. Sequences upstream of–127 and within the Cyc6 coding region played no role in the copper response, while constructs that contained the region between –54 and –7 did not express the reporter. Because ofthe wide variability in expression of the gene in individual transformants, it was not possible to define additional sequence elements that played a role in quantitative, as opposed to copperregulated, levels of Cyc6 expression. Because of the sensitivity of the chromogenic substrate for assaying Ars activity, Ohresser et al. (1997) were able to measure enzymatic activity in pools of transformed colonies, which circumvents the need to analyze multiple individual transformants to correct for variations in gene expression due to position effects. For this approach to be meaningful, the co-transformation frequency must be similar for different constructs. By comparing pools of transformants generated with different promoter deletion mutations, they were able to identify elements that affected expression of the reporter gene quantitatively. Analysis of transformants expressing fusions of the Nit1 promoter to the Ars coding region suggested that positive enhancer-type elements are localized between –751 and –353, and between –282 and –198, while a negative element resides between –342 and –282 (see Table 2). The role of introns and 5´ and 3´ untranslated regions of nuclear transcripts in regulating gene expression has not been investigated systematically in C. reinhardtii, to my knowledge. However, a nitA (nitrate reductase) mutation in Volvox carteri was shown to affect the 5´ splice site of intron 2, leading to non-functional splice variants. Furthermore, nuclear transformation of a nitA cDNA into Volvox was increased 10-fold by including intron 1 or introns 9 and 10, suggesting that introns may play an important regulatory role in gene expression in this closely related alga (Gruber et al., 1996). A radial spoke protein (Rsp3) minigene, which included the cDNA sequence and ~665 and 380 bp from the 5´ and 3´ flanking regions of genomic Rsp3 DNA, successfully complemented a motility defect due to a mutation at PF14 (Diener et al., 1993). Although the level of gene expression was not directly examined, the gene functions well enough that it does not limit the synthesis or assembly of flagellar proteins.

Karen L. Kindle XI. Conclusion The development of efficient methods and molecular tools for nuclear transformation has revolutionized the analysis of cellular processes in C. reinhardtii. Nuclear transformation provides powerful approaches for assessing gene function, generating large numbers of mutants, and isolating the genes that are affected in these mutants. However, technical challenges remain. Despite recent advances, the expression of foreign genes is still problematic. Gene silencing may be part of the problem, which may also affect the stability of expression of unselected genes, both from C. reinhardtii and other organisms. Furthermore, there is, as of yet, no good way to mutate or downregulate the expression of endogenous copies of cloned genes. An effective gene targeting strategy or development of antisense technology would be very helpful. The progress in the last decade has been dramatic, and the enthusiasm for this organism as a model for a variety of developmental and metabolic pathways suggests that this progress will continue.

Acknowledgments I am grateful to the National Science Foundation for supporting my research program, most recently through grants from the Cell Biology Program (MCB­ 9406540) and the Biochemical Genetics Program (with David Stern; MCB-9406550). I am also grateful to former and present members of the Stern and Kindle labs for helpful discussions and to Donna Esposito for critiquing the manuscript.

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60 Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 87: 1228–1232 Kindle KL (1998) High-frequency nuclear transformation of Chlamydomonas reinhardtii. Meth Enzymol (in press) Kindle KL and Sodeinde OA (1994) Nuclear and chloroplast transformation in Chlamydomonas reinhardtii: strategies for genetic manipulation and gene expression. J Appl Phycol 6: 231–238 Kindle KL, Schnell RA, Fernández E and Lefebvre PA (1989) Stable nuclear transformation of Chlamydomonas using the Chlamydomonas gene for nitrate reductase. J Cell Biol 109: 2589–2601 Kozminski KG, Diener DR and Rosenbaum JL (1993) High level expression of non-acetylatable alpha tubulin in Chlamydomonas reinhardtii. Cell Motil Cytoskeleton 25: 158–170 Kuchka MR, Goldschmidt-Clermont M, van Dillewijn J and Rochaix J-D (1989) Mutation at the Chlamydomonas nuclear NAC2 locus specifically affects stability of the chloroplast psbD transcript encoding polypeptide D2 of PS II. Cell 58: 869–876 Li J and Timko MP (1996) The pc-1 phenotype of Chlamydomonas reinhardtii results from a deletion mutation in the nuclear gene for NADPH:protochlorophyllide oxidoreductase. Plant Mol Biol 30: 15–37 Mansour SL, Thomas KR and Capecchi MR (1988) Disruption of the proto-oncogene int-2 in mouse embryo-derived stem cells: a general strategy for targeting mutations to non-selectable genes. Nature 336: 348–352 Mayfield SP (1991) Over-expression of the oxygen-evolving enhancer 1 protein and its consequences on Photosystem II accumulation. Planta 185: 105–110 Mayfield SP and Kindle KL (1990) Stable nuclear transformation of Chlamydomonas reinhardtii by using a C. reinhardtii gene as the selectable marker. Proc Natl Acad Sci USA 87: 2087– 2091 McHugh JP and Spanier JG (1994) Isolation of cadmium-sensitive mutants in Chlamydomonas reinhardtii by transformationinsertional mutagenesis. FEMS Microbiol Lett 124: 239–244 Merchant S, Hill K and Howe G (1991) Dynamic interplay between two copper-titrating components in the transcriptional EMBO J 10: 1383–1389 regulation of cytochrome Nelson JA and Lefebvre PA (1995a) Targeted disruption of the N I T 8 gene in Chlamydomonas reinhardtii. Mol Cell Biol 15: 5762–5769 Nelson JAE and Lefebvre PA (1995b). Transformation of Chlamydomonas reinhardtii. In: Dentler W and Witman G (eds) Methods in Cell Biology, Vol 47: Cilia and Flagella, pp 513–517. Academic Press, New York Nelson JAE, Savereide PB and Lefebvre PA (1994) The CRY1 gene in Chlamydomonas reinhardtii: Structure and use as a dominant selectable marker for nuclear transformation. Mol Cell Biol 14: 4011–4019 Nicholl DST, Schloss JA and John PCL (1988) Tubulin gene expression in the Chlamydomonas reinhardtii cell cycle: Elimination of environmentally induced artifacts and the measurement of tubulin mRNA levels. J Cell Sci 89: 397–403 Ohresser M, Matagne RF and Loppes RL (1997) Expression of the arylsulfatase reporter gene under the control of the Nit1 promoter in Chlamydomonas reinhardtii. Curr Genet 31: 264– 271 Pazour GJ, Sineshchekov O and Witman GB (1995) Mutational

Karen L. Kindle analysis ofthe phototransduction pathway of Chlamydomonas reinhardtii. J Cell Biol 131: 427–440 Periz G and Keller LR (1997) DNA elements regulating tubulin gene induction during regeneration of eukaryotic flagella. Mol Cell Biol 17: 3858–3866 Prieto R, Dubus A, Galván A and Fernández E (1996a) Isolation and characterization of two new negative regulatory mutants for nitrate assimilation in Chlamydomonas reinhardtii obtained by insertional mutagenesis. Mol Gen Genet 251: 461–471 Prieto R, Pardo JM, Niu X, Bressan RA and Hasegawa PM (1996b) Salt-sensitive mutants of Chlamydomonas reinhardtii isolated after insertional tagging. Plant Physiol 112: 99–104 Purton S and Rochaix J-D (1994a) Characterization of the ARG7 gene of Chlamydomonas reinhardtii and its application to nuclear transformation. Eur J Phycol 30: 141–148 Purton S and Rochaix J-D (1994b) Complementation of a Chlamydomonas reinhardtii mutant using a genomic cosmid library. Plant Mol Biol 24: 533–537 Quesada A and Fernández E (1994) Expression of nitrate assimilation related genes in Chlamydomonas reinhardtii. Plant Mol Biol 24: 185–194 Quinn JM and Merchant S (1995) Two copper-responsive elements associated with the Chlamydomonas Cyc6 gene function as targets for transcriptional activators. Plant Cell 7: 623–638 Quinn J, Li HH, Singer J, Morimoto B, Mets L, Kindle K and Merchant S (1993) The plastocyanin-deficient phenotype of Chlamydomonas ac-208 results from a frame-shift mutation in the nuclear gene encoding preapoplastocyanin. J Biol Chem 268: 7832–7841 Quisel JD, Wykoff DD and Grossman AR (1996) Biochemical characterization ofthe extracellular phosphatases produced by phosphorus-deprived Chlamydomonas reinhardtii. Plant Physiol 1 1 1 : 839–848 Rochaix J-D and vanDillewijn J (1982) Transformation of the green alga Chlamydomonas reinhardtii with yeast DNA. Nature 296: 70–73 Rochaix J-D, van Dillewijn J and Rahire M (1984) Construction and characterization of autonomously replicating plasmids in the green unicellular alga Chlamydomonas reinhardtii. Cell 36: 925–931 Shepherd HS, Ledoigt G and Howell SH (1983) Regulation of light-harvesting chlorophyll-binding protein (LHCP) mRNA accumulation during the cell cycle in Chlamydomonas reinhardtii. Cell 32: 99–107 Shimogawara K, Fujiwara S, Grossman AR and Usuda H (1998) High efficiency transformation of Chlamydomonas reinhardtii by electroporation. Genetics, in press Silflow CD and Rosenbaum JL (1981) Multiple and genes in Chlamydomonas and regulation of tubulin mRNA levels after deflagellation. Cell 24: 81–88 Sizova IA, Lapina TV, Frolova O, Alexandrova NN, Akopiants KE and Danilenko VN (1996) Stable nuclear transformation of Chlamydomonas reinhardtii with a Streptomyces rimosus gene as the selective marker. Gene 181: 13–18 Smart EJ and Selman BR (1991) Isolation and characterization of a Chlamydomonas reinhardtii mutant lacking the of Mol Cell Biol 1 1 : 5053– chloroplast coupling factor 1 5058 Smart EJ and Selman BR (1992) Complementation of a Chlamydomonas reinhardtii mutant defective in the nuclear

Chapter 4 Nuclear Transformation gene encoding the chloroplast coupling factor 1 subunit (atpC). J Bioenerg Biomembr 25: 275–284 Smith EF and Lefebvre PA (1996) PF16 encodes a protein with armadillo repeats and localizes to a single microtubule of the central apparatus in Chlamydomonas flagella. J Cell Biol 132: 359–370 Sodeinde OA and Kindle KL (1993) Homologous recombination in the nuclear genome of Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 90: 9199–9203 Stevens DR, Rochaix J-D and Purton S (1996) The bacterial phleomycin resistance gene ble as a dominant selectable marker in Chlamydomonas. Mol Gen Genet 251: 23–30 Szostak JW, Orr-Weaver TL and Rothstein RJ (1983) The doublestrand-break repair model for recombination. Cell 33: 25–35 Tam L-W and Lefebvre PA (1993) The use of DNA insertional mutagenesis to clone genes in Chlamydomonas. Genetics 135: 375–384 Tang DKH, Qiao S-Y and Wu M (1995) Insertion mutagenesis of Chlamydomonas reinhardtii by electroporation and hetero­ logous DNA. Biochem Molec Biol Int 36: 1025–1035 Thomas KR and Capecchi MR (1987) Site-directed mutagenesis

61 by gene targeting in mouse embryo-derived stem cells. Cell 51: 503–512 Valancius V and Smithies O (1991) Double-strand gap repair in a mammalian gene targeting reaction. Mol Cell Biol 11: 4389– 4397 Villand P, Eriksson M and Samuelsson G (1997) Carbon dioxide and light regulation of promoters controlling the expression of mitochondrial carbonic anhydrase in Chlamydomonas reinhardtii. Biochem J 327: 51–57 Wilkerson CG, King SM, Koutoulis A, Pazour GJ and Witman GB (1995) The 78,000 Mr intermediate chain of Chlamy­ domonas outer arm dynein is a WD-repeat protein required for arm assembly. J Cell Biol 129: 169–178 Zhang D and Lefebvre PA (1997) FAR1, a negative regulatory locus required for the repression of the nitrate reductase gene in Chlamydomonas reinhardtii. Genetics 146: 121–133 Zhang H, Herman P and Weeks DP (1994) Gene isolation through genomic complementation using an indexed library of Chlamydomonas reinhardtii DNA. Plant Mol Biol 24: 663– 672

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Chapter 5 Modes and Tempos of Mitochondrial and Chloroplast Genome Evolution in Chlamydomonas: A Comparative Analysis Aurora M. Nedelcu and Robert W. Lee Department of Biology, Dalhousie University, Halifax, Nova Scotia B3H 4J1, Canada

Summary I. Introduction II. Phylogenetic Position of Chlamydomonas Ill. Monophyletic versus Polyphyletic Origin of Mitochondria and Plastids: The Chlamydomonas Case IV. Evolution of Mitochondrial and Chloroplast Genome Size in Chlamydomonas A. Factors Contributing to Variation in Genome Size 1. Changes in Intergenic Spacer Size 2. Changes in Intron Number 3. Changes in Gene Content 4. Changes in the Amount of Repeated DNA B. Mechanisms Possibly Involved in the Evolution of Genome Size 1. Length Mutations 2. Intron Mobility 3. Gene Transfer V. Evolution of Mitochondrial and Chloroplast Genome Organization in Chlamydomonas A. Mitochondrial and Chloroplast Genome Structure B. Mitochondrial and Chloroplast Gene Order 1. Factors Contributing to Gene Rearrangement 2. Mechanisms Possibly Involved in Gene Rearrangement VI. Evolution of Mitochondrial and Chloroplast Gene Structure and Organization in Chlamydomonas A. Intron-containing Coding Regions B. Fragmented Coding Regions VII. Evolution of Mitochondrial and Chloroplast DNA Sequences in Chlamydomonas VIII. Conclusions Acknowledgments References

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Chlamydomonas mitochondrial and chloroplast genomes, in contrast to the land plant counterparts, exhibit concerted modes and tempos of evolution. The 1.5-fold variation currently observed in the size of both organelle genomes is mostly accounted for by changes in the spacer DNA and intron number, with less contribution from changes in gene content and amount of repeated DNA. Gene order is highly variable in both mitochondrial and chloroplast genomes of Chlamydomonas, the level of gene rearrangement being correlated with the abundance of short dispersed repeated sequences throughout the genome. Intron-containing-, J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 63–91. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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fragmented-, and fragmented and scrambled coding regions are common features of mitochondrial and chloroplast gene structure and organization within the group. The level of ribosomal RNA gene sequence divergence in both mitochondrial and chloroplast genomes is higher in the Chlamydomonas lineage than in land plants and is most likely due to higher rates of nucleotide substitution in Chlamydomonas organellar DNAs. The mechanisms as well as the selective pressures that shaped the organellar genomes in the Chlamydomonas lineage remain to be explained. I. Introduction The increasing accumulation of information on organellar genome sequence, structure and organi­ zation in various lineages makes it possible to address questions such as: (i) how conserved are the organellar genomes in terms of DNA sequence, structure and organization among different evolutionary lineages; (ii) what are the mechanisms underlying the evolutionary processes in organellar genomes; (iii) are the mechanisms acting on mitochondrial and chloroplast genomes similar or different; (iv) do the mitochondrial and chloroplast genomes have the same tempo and mode of evolution in a given lineage; (v) what are the evolutionary forces shaping the organellar genomes. Land plants and green algae are the only two groups that possess both mitochondria and chloro­ phyll a/b-containing chloroplasts. As the complete DNA sequence of several land plant chloroplast (see Shimada and Sugiura, 1991 for references) and mitochondrial (Oda et al., 1992) genomes has become available and more and more genes have been mapped and sequenced it has become obvious that the two organellar genomes exhibit different modes and tempos of evolution in this group: chloroplast genomes are more conserved in size and gene order but more variable in DNA sequence than the mitochondrial counterparts (Palmer, 1990). No extensive analyses have been done, however, to assess the rates and patterns of evolutionary change of green algal organellar genomes, mainly due to incompleteness or disparity of the available data. Given that Chlamydomonas is the only green algal lineage for which information on the structure, organization and DNA sequence of both mitoAbbreviations: cpDNA – chloroplast DNA; CW – clockwise flagellar configuration; DIR – direct inverted repeat; DO – directly opposed flagellar configuration; IR – inverted repeat; ITS – internal transcribed spacer; LSU – large subunit; mtDNA – mitochondrial DNA; ORF – open reading frame; rDNA – ribosomal RNA coding sequence; SSU – small subunit; TIR – terminal inverted repeat

chondrial and chloroplast genomes is available, it provides us with the opportunity to address such issues. This chapter is not intended to review the vast amount of information on Chlamydomonas organellar genomes; rather, it will present only the information considered to have evolutionary significance from a comparative point of view. Extensive general reviews dealing with the structure, organization and evolution of both mitochondrial and chloroplast genomes have been published in the last decade (Palmer et al., 1985; Palmer, 1987, 1990, 1991; Gray, 1992, 1993, 1995;Gillham, 1994;Rochaix, 1995; Wolstenholme and Fauron, 1995; Gray and Spencer, 1996). Any attempt to assess features of organelle genome evolution in a given group requires a good understanding of the phylogeny of that group. On the other hand, a better understanding of phylogeny grows from knowledge about organellar genome evolution. The second section of this chapter, therefore, will present a phylogenetic framework of green algae focusing, however, only on the information necessary for understanding the phylogenetic position of Chlamydomonas within the group. The third section will address the issue of the mono- versus polyphyletic origin of mitochondria and plastids with reference to the Chlamydomonas case. In the remaining sections of this chapter we will attempt to (i) define evolutionary trends in the two organellar genomes of Chlamydomonas; (ii) compare the mitochondrial genome mode and tempo of evolution to that of the chloroplast counterpart within the Chlamydomonas lineage and among green algae; and (iii) contrast the rates and patterns of evolutionary change in the organellar genomes of Chlamydomonas and land plants. II. Phylogenetic Position of Chlamydomonas The green algal genus Chlamydomonas Ehrenberg consists of over 450 species (Ettl, 1976) with considerable morphological, physiological and reproductive interspecific variability (Schlösser,

Chapter 5 Mitochondrial and Chloroplast Genome Evolution 1984). Systematic studies of the genus defined nine distinct morphological groups that differ primarily in chloroplast morphology (Ettl, 1976), as well as 15 distinct sporangial autolysin groups (Schlösser, 1984). The flagellate genus Chlamydomonas belongs to the order Chlamydomonadales of the class Chloro­ phyceae sensu Mattox and Stewart (1984). Mattox and Stewart’s classification system divides the green algae (Chlorophyta) into five classes: Chlorophyceae, Pleurastrophyceae, Ulvophyceae, Charophyceae and Micromonadophyceae. Cladistic analyses using organismal data are consistent with the evolutionary hypotheses underlying Mattox and Stewart’s classification system, and indicate that (i) the Chloro­ phyceae is a sister group to the Pleurastrophyceae; (ii) the Ulvophyceae is a sister group to the Chloro/Pleurastrophyceae clade; and (iii) the charophycean group, the Ulvo-/Chloro-/Pleurastrophyceae clade, and the micromonadophycean taxa all emerge from an unresolved node (Kantz et al., 1990). The phylogeny of green algae is being progressively deciphered and the new information gathered through molecular approaches will probably trigger the reconsideration of traditional green algal systematics (Chapman and Buchheim, 1991; Friedl, 1995). Phylogenetic analyses based on nuclear ribosomal RNA gene sequence (rDNA) data confirm the presence of five main evolutionary lineages among green algae, but they increasingly reveal more inconsistencies between the phylogenetic position and the polyphyly of many generic- and ordinal-level lineages on the one hand, and the traditional taxonomy on the other (Buchheim and Chapman, 1992; Friedl, 1995). Similar analyses indicate that the class Chlorophyceae itself consists of two distinct evolutionary lineages (Steinkötter et al., 1994; Friedl, 1995) that are consistent with the two flagellar apparatus configurations described among flagellate chlorophycean taxa, the directly opposed (DO) and clockwise (CW) types. The same phylogenetic analyses, however, suggest that while some members of the non-flagellate (autosporic) taxa (such as Scenedesmus obliquus) traditionally included in the chlorophycean order Chlorococcales, affiliate with DO chlorophycean taxa, other members (e.g., Prototheca wickerhamii) form amonophyletic group with advanced lineages of the class Pleurastrophyceae (sensu Mattox and Stewart, 1984); this latter group has been recently defined by Friedl (1995) as a new class, the Trebouxiophyceae (Fig. 1a). In addition, the primitive green flagellates grouped by Mattox

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and Stewart (1984) and Moestrup and Throndsen (1988) into the class Micromonadophyceae and Prasinophyceae, respectively, do not form a monophyletic assemblage of taxa (Steinkötter et al., 1994). Furthermore, phylogenetic analyses using nuclear and chloroplast rDNA sequences clearly show that the flagellate genus Chlamydomonas is not a natural assemblage of taxa: multiple lineages exist within the group, some of them containing both Chlamydomonas and non-Chlamydomonas taxa from distinct families or orders (Buchheim et al., 1990, 1996). Despite the continuous reconsideration of the phylogenetic relationships among green algae, there is no question that the Chlorophyta (green algae) and Embryophyta (land plants) form a monophyletic group (see McCourt, 1995 for a review). Nevertheless, phylogenetic analyses based on mitochondrial data, both rDNA nucleotide and cytochrome oxidase subunit 1 amino acid (COXI) sequences, reveal an unexpected dichotomy among green algae with respect to their relationships with land plants: while at least in COXI phylogenetic analyses, some green algae most closely affiliate, as expected, with the land plant group, the chlamydomonadalean lineage branches inconsistently with ciliates, fungi or animal counterparts in both rDNA and COXI trees (Wolff et al., 1993; Antamarian et al., 1996; Denovan-Wright et al., 1996) (Fig. 1b).

III. Monophyletic versus Polyphyletic Origin of Mitochondria and Plastids: The Chlamydomonas Case It is well accepted now that at least two of the eukaryotic cell’s organelles, namely the mitochondria and plastids, have a eubacterial and cyanobacterial, respectively) endosymbiotic origin, although their mono-or polyphyletic ancestry is still debated (see Gray, 1992 for a review). Single endosymbiotic events accounting for the origin of mitochondria and plastids, respectively, would imply that some common ancestral characters should be present in all the extant lineages, and that distinct derived traits should be developed within and shared among related lineages. In addition, monophyletic origins for the mitochondria and plastids, respectively, would also require that phytogenies based on organellar traits be consistent with the ones based on nuclear or nucleus-encoded features; in other words

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68 all the compartments within a eukaryotic cell should resemble their corresponding counterparts in the same compared lineage. Although the phylogenetic relationships within the chlamydomonadalean group as well as between the group and other green algal lineages are not fully deciphered, Chlamydomonas taxa appear to be legitimate members of green algae. However, in an early archaebacterial-eubacterial-chloroplastmitochondrial-nuclear phylogenetic tree inferred from small subunit ribosomal RNA (SSU rRNA) gene sequence data, the Chlamydomonas reinhardtii nuclear and mitochondrial sequences suggested different phylogenetic affiliations when compared to the closest relatives of green algae, namely, the land plants (Gray et al., 1989). In the nuclear subtree, C. reinhardtii formed a clade with the plant sequences (as it did also in the chloroplast subtree) and branched off at about the same point as animals and fungi. In contrast, in the mitochondrial subtree, C. reinhardtii branched with the ciliate/fungal/animal sequences, far away from higher plants, which clustered very clade. near the root, close to the The affiliation of the nuclear SSU rRNA gene sequences of higher plants and C. reinhardtii was seen as consistent with traditional phylogenies that consider green algae as being the closest relatives of land plants (Chapman and Ragan, 1980; Chapman and Buchheim, 1991), whereas the green algal/land plant dichotomy in the mitochondrial tree was interpreted as an anomaly. This anomaly in branching topology was, however, attributed to the plant rather than C. reinhardtii mitochondrial sequences and was considered not to be a treeing artifact due to the relatively rapid rate of sequence divergence of nonplant mitochondrial rRNA sequences (Gray et al., 1989). To explain the different branching position of plants within the nuclear and mitochondrial lineages, respectively, and to account for the strong eubacterial features of their mitochondrial rRNAs, Gray et al. (1989) suggested two possibilities: either (i) the mitochondrial rRNA genes of plants have diverged relatively little from the rRNA genes of the ancient eubacterial ancestor of all mitochondria (mono­ phyletic origin) or (ii) the higher plant mitochondrial rRNA genes or the mitochondria itself have been acquired more recently than those ofother eukaryotic lineages (biphyletic origin). In addition, due to the very different way in which genes are organized and expressed in the mitochondrial genomes of C. rein­ hardtii and land plants, the authors concluded that

Aurora M. Nedelcu and Robert W. Lee there is no indication that the two shared a common mitochondrial ancestor as recently as they shared a common nuclear (or chloroplast) ancestor. Because all the mitochondria investigated seemed to affiliate with only one subgroup of the biphyletic or polyphyletic concept as used by Gray et al. (1989) did not, however, imply more than one primary original endosymbiosis but rather a more recent secondary endosymbiotic event for the land plant mitochondria or its rRNA genes. In contrast to these results, Van de Peer et al. (1990) presented a phylogenetic tree based on SSU rRNA gene (rrnS) sequences of eukaryotic, archaebacterial, eubacterial, chloroplast, and mitochondrial origin and argued that mitochondria appeared polyphyletic: one cluster contained all the animal mitochondria; a second cluster, was formed by the C. reinhardtii, fungal and ciliate mitochondria; and the third cluster was comprised of the land plant mitochondria and was embedded in the eubacterial as the cluster with the Proteobacteria closest relative. The input of other green algal mitochondrial rDNA sequences, namely, of Prototheca wickerhamii (Wolff and Kück, 1990; Wolff et al., 1993) and Chlamy­ domonas eugametos (Denovan-Wright et al., 1996) did not resolve Chlamydomonas and P. wickerhamii sequences as a green algal clade sharing a most recent common ancestor with the land plants to the exclusion of other groups (Fig. 1b). Phylogenetic trees based on COXI amino acid sequence suggested, however, that the plant and green algal mitochondrial lineage including P. wickerhamii (Wolff et al., 1993) and the prasinophycean (sensu Moestrup and Throndsen 1988) Platymonas (Tetraselmis) subcordi­ formis (Kessler and Zetsche, 1995) do form a monophyletic group. The expected congruency of nuclear, plastid and mitochondrial phylogenetic trees appears thus verified in the case of trebouxiophycean (sensu Friedl, 1995) and prasinophycean (sensu Moestrup and Throndsen, 1988) green algae and land plants, whereas the chlamydomonadalean taxa branch with land plants in nuclear and chloroplast trees and with unrelated taxa (e.g., fungi or ciliates) in mitochondrial trees. To explain such findings, a polyphyletic origin for the green algal mitochondria was suggested (Wolff and Kück, 1993). Gray and Spencer (1996) also considered that there is little or no evidence that land plants and Chlamydomonas shared a common mitochondrial ancestor as recently as they shared a common chloroplast or nuclear ancestor; however, they proposed that the differences

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between the Chlamydomonas and Prototheca/land plant mitochondrial genome types are ‘best explained by a relatively rapid and extreme evolution’ of the former genome from the ancestral pattern represented by the more conservative genomes in the latter group. Similarly, current evidence seems to favor the view of a primary monophyletic cyanobacterial origin of plastids followed by an early subsequent diversification of the accessory pigments (Gray, 1993). Nevertheless, it appears that the rhodophyte, cryptophyte and chromophyte plastids are more closely related to each other than to their chlorophyte and land plant counterparts (Douglas, 1994). Phylogenetic trees constructed from the small and large subunits of ribulose-1,5-bisphosphate carboxylase/oxygenase (rbcS and rbcL) amino acid sequences indicated that the plastids of non-green plants are most closely related to proteobacteria whereas the green algal and land plant counterparts are most closely related to cyanobacteria (Morden et al., 1992; Delwiche et al, 1995); potential explan­ ations for this apparent dichotomy are discussed by Gray and Spencer (1996). Phylogenetic analyses using either SSU rDNA sequences or translation elongation factors and other protein amino acid sequences are contradictory and do not provide confident support for either a monophyletic or polyphyletic origin of plastids (reviewed by Douglas, 1994 and Gray and Spencer, 1996). Although the chlorophyte/embryophyte connection is relatively well supported in all chloroplast phylogenetic analyses, it is noteworthy that in a recent phylogenetic analysis, Chlamydomonas SSU rDNA nucleotide sequences did not branch as expected with other green algae, but rather suggested a very early divergence of this lineage relative to all other chloroplast counterparts examined (Gray and Spencer, 1996) (Fig. 1c).

IV. Evolution of Mitochondrial and Chloroplast Genome Size in Chlamydomonas The size of mitochondrial and chloroplast genomes appears quite different, both in the same lineage, as well as among lineages. Land plant mitochondrial genomes are large and extremely variable in size, ranging from 186 kb to 2500 kb (reviewed by Palmer, 1990). In contrast, land plant chloroplast genomes are smaller and seem to be rather conservative in size, varying from 120 kb to 160 kb, in only few

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cases reaching 220 kb (reviewed by Palmer, 1991). On the other hand, animal mitochondrial genomes are very small and extremely conserved in size, varying generally from 15.7 kb to 21 kb (see Wolstenholme and Fauron, 1995 for a review), with the exception of scallop mitochondrial genomes that vary from 16.2 kb to 41 kb (see Gjetvaj et al., 1992 for references). Surprisingly, the chlamydomonadalean mito­ chondrial genomes are 10–100-fold smaller than their land plant homologs, thus approximating the size of their metazoan counterparts, whereas the chlamydomonadalean chloroplast genomes are slightly larger than most of their angiosperm homologs. Our analysis of the known mitochondrial and chloroplast genome sizes within the chlamy­ domonadalean group suggests a 1.5-fold variation in both organelles, from a 15.8-kb mitochondrial DNA (mtDNA) in C. reinhardtii (Michaelis et al., 1990) to a 22.9-kb counterpart in C. eugametos (DenovanWright et al., in press), and from a 187-kb chloroplast DNA (cpDNA) in Chlamydomonas pitschmannii (Boudreau and Turmel., 1995) to a 292-kb homolog in Chlamydomonas moewusii (Boudreau et al., 1994). It is noteworthy that the same level ofchloroplast and mitochondrial genome size variation is also observed in each of the two very divergent evolutionary lineages within the Chlamydomonas group, i.e., the ‘C. rein­ hardtii’ and the ‘C. eugametos’ lineages. Exceptional variation in genome size has, however, been described among the sexually incompatible members of the colonial chlamydomonadalean taxon, Pandorina morum, whose mitochondrial and chloroplast genome sizes vary from 20 kb to 38 kb, and from 150 kb to 450 kb, respectively (Moore and Coleman, 1989; Moore, 1990). It appears, therefore, that mito­ chondrial genome size is more conservative among chlamydomonadalean taxa than among land plants (1.5-fold in the former, relative to 12.5-fold in the latter), whereas chloroplast genomes vary slightly more (1.5-fold) than do their land plant counterparts (1.3-fold). It is noteworthy that in contrast to land plants, in Chlamydomonas both organellar genomes exhibit the same level of size variation. Furthermore, it appears that the two organelle genomes followed parallel evolutionary pathways not only within the group but also within a given species: the mito­ chondrial and chloroplast genomes are both either small (e.g., in C. reinhardtii and C. pitschmannii) or large (e.g., in C. eugametos and C. moewusii) relative to the currently known size range in the Chlamy­ domonas group.

70 Among green algae, however, there is a five-fold variation in mitochondrial genome size (excluding the 220-kb mitochondrial genome of Bryopsis), from 15.8 kb in C. reinhardtii (Michaelis et al., 1990) to 80 kb in Chlorella pyrenoidosa (Bayen and Rode, 1973). Similarly, there is at least a five-fold variation in chloroplast genome size among the green algal lineages investigated so far, from 89 kb in Codium fragile to 400 kb in a few members of three out of the five green algal classes, namely, the Charophyceae, Ulvophyceae, and Chlorophyceae (see Palmer, 1991 for references). Although the degree of variation in organelle genome size is overall higher among green algae compared to Chlamydomonas, it is interesting that both genome types seem to exhibit the same level of variation in size, a situation similar to that observed in Chlamydomonas but in contrast to that noted among land plants. Because the mitochondrial genome in Platymonas subcordiformis (Kessler and Zetsche, 1995), a green flagellate that retains ancestral-like features, is larger (i.e., 42.8 kb) than in chlamydomonadalean taxa, it is most likely that the reduced genome size in the latter represents a derived condition among green algae. Moreover, one can hypothesize an evolutionary trend towards a smaller mitochondrial genome within the chlorqphycean group, from a 45-kb mitochondrial genome as in, for example, Scenedesmus obliquus (Kück, 1989), to a 15.8 kb homolog in C. reinhardtii. Although limited, the current data do not support a similar trend among other green algal lineages: at 42.8 kb, the mitochondrial genome of the primitivelike green flagellate Platymonas subcordiformis is smaller than the 55.3-kb homolog in Prototheca wickerhamii (Wolff et al., 1994) and the 80-kb counterpart in Chlorella pyrenoidosa. There is no indication of a tendency towards a smaller chloroplast genome among chlorophycean taxa as suggested for the mitochondrial counterparts; however, chloroplast genome sizes much smaller than the average have been reported among ulvophycean (89 kb in Codium fragile, Manhart et al., 1989) and charophycean (130 kb in Spirogyra maxima, Manhart et al., 1990) taxa.

A. Factors Contributing to Variation in Genome Size Generally, changes in genome size are the result of changes in sequence complexity and/or changes in the amount of repeated DNA (Palmer, 1990). Changes in genome complexity occur through the deletion

Aurora M. Nedelcu and Robert W. Lee and insertion of unique sequences (intergenic regions, introns and open reading frames). The great variation in mtDNA size among land plants is mostly accounted for by changes in the complexity of spacer DNA. In contrast, cpDNAs in the same group vary relatively little in size, with less contribution from changes in intergenic region size, intron number or gene content, but more (i.e., nearly two-thirds of cpDNA size variation) from the expansion/contraction of the inverted repeat (see Palmer 1991, for a review).

1. Changes in Intergenic Spacer Size The intergenic spacers are very reduced in Chlamy­ domonas mitochondrial genomes (i.e., 16–17% of the genome). In C. eugametos, with the exception of two large intergenic regions of 902 bp and 1057 bp, respectively, the intergenic spacers range from 0 to 466 bp, with most of them being smaller than 72 bp. In the more compact mitochondrial genome of C. reinhardtii, with the exception of the terminal noncoding regions of about 530 bp each, most of the intergenic regions are smaller than 200 bp and missing whenever two rRNA gene pieces are adjacent. On the other hand, the intergenic regions in the mitochondrial genome of P. wickerhamii represent 29% of the genome, and in the majority of cases are 100-150 bp long with only two considerably longer regions (1118 bp and 1993 bp) (Wolff et al., 1994). Only about 12% of the 7-kb difference in size between the two completely sequenced Chlamydomonas mitochondrial genomes can be accounted for by differences in intergenic region size, whereas the 30­ kb difference in mitochondrial genome size between the mitochondrial genomes of Chlamydomonas and Prototheca is a consequence of variation in both the intergenic region size as well as gene content (discussed later). In contrast, the substantial difference in chloroplast genome size between closely related Chlamy­ domonas taxa (i.e., C. moewusii and C.pitschmannii as well as C. reinhardtii and C. gelatinosa) is mainly a consequence of multiple deletions/additions in the intergenic spacers (Boudreau and Turmel, 1995, 1996). Similarly, most of the observed differences in cpDNA restriction patterns between the interfertile C. reinhardtii and C. smithii are correlated with insertions/deletions of short dispersed repeated sequences of 50 bp to 200 bp, which are ubiquitous in the intergenic regions of C. reinhardtii cpDNA (Rochaix, 1978; Gelvin and Howell, 1979; Palmer et

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al., 1985). On the other hand, 12% of the 50-kb difference in size between the interfertile taxa C. moewusii and C. eugametos is accounted for by a 6-kb insertion in one of the single copy-regions of the C. moewusii chloroplast genome (Lemieux et al., 1985); the rest is the consequence of a 21 -kb insertion in its large inverted repeat (Turmel et al., 1987). Another type of insertion is represented by the two copies of a 2.4 kb DNA sequence, the Wendy element, that has many of the features of transposable elements (discussed later) and has only been described in the chloroplast genome of C. reinhardtii (Fan et al., 1995). The absence of a counterpart in any of the other chlamydomonadalean or land plant chloroplast genomes examined to date suggests that Wendy is a relatively recent acquisition in the C. reinhardtii lineage (Fan et al., 1995). The difference in chloroplast genome size between Chlamydomonas and land plants is correlated with the presence of enlarged intergenic spacers in the former relative to the latter (Boudreau et al., 1994).

2. Changes in Intron Number There is quite a variation among chlamydo­ monadalean taxa in terms of the number of introns present in their organellar genomes. The difference in size between the mitochondrial genomes, otherwise co-linear (Boynton et al., 1987), of the two interfertile taxa, C. reinhardtii and C. smithii, is solely the result of a unique 1-kb intron inserted in the cob sequence of C. smithii (Matagne et al., 1988; Colleaux et al., 1990). In addition, it appears that the difference in mitochondrial genome size between the members of the other pair of interfertile Chlamydomonas taxa, that is C. eugametos and C. moewusii, might also be correlated with the presence of optional introns (Denovan-Wright and Lee, 1993). Moreover, 88% of the difference in size between the C. reinhardtii and C. eugametos mitochondrial genomes can be accounted for by the presence of nine intervening sequences in the coding regions of the latter (Fig. 2). There seems to be a variation also in the number of introns harbored by homologous genes among chlamydomonadalean taxa: two introns are present in the cob sequence of a chlamydomonadalean taxa more closely related to C. eugametos than to C. reinhardtii (Buchheim et al., 1996), namely Chlorogonium elongatum (Kroymann and Zetsche, 1997), whereas only one intron is found in C. eugametos (Denovan-Wright et al., 1998) and C.

71

smithii (Colleaux et al., 1990) cob, but none in the homologous C. reinhardtii gene (Michaelis et al., 1990). More data have to become available before one can decide whether the Chlamydomonas ancestor had or did not have introns in its mitochondrial genome, but variation in intron content is clearly a significant factor responsible for the observed variation in mitochondrial genome size among Chlamydomonas taxa. Similarly, there is quite a variation in intron content among Chlamydomonas chloroplast genomes (Turmel et al., 1991). It is noteworthy that the chloroplast large subunit (LSU) rRNA genes from 17 Chlamydomonas taxa investi­ gated contain a total of 39 group I introns representing 12 insertion sites (Turmel et al., 1993). While C. pitschmannii and C. reinhardtii have no and only one intron, respectively, in their chloroplast LSU rRNA gene, C. moewusii and C. eugametos have five and six, respectively. On the other hand, four introns are present in the C. reinhardtii psbA gene, which is intronless in C. eugametos (Erickson et al., 1984; Lemieux et al., 1985), and one of the four introns present in C. reinhardtii is missing in the interfertile C. smithii (Palmer et al., 1985); additional optional introns are also present in rrnS, psaB and psbC (see Turmel et al., 1993 for references). Another case of optional insertions contributing to variation in size of the open reading frames in Chlamydomonas chloroplast genes is represented by the presence of (i) one or two large insertion sequences that are not spliced out at the mRNA level in the genes coding for the catalytic subunit of the ATPdependent Clp protease (clpP) of C. reinhardtii and C. eugametos, respectively (Huang et al., 1994), and (ii) in-frame long sequences of unknown identity juxtaposed within the gene coding for the RNA polymerase subunit C (rpoC2) of C. reinhardtii cpDNA (Fong and Surzycki, 1992).

3. Changes in Gene Content One of the most distinctive features of the mitochondrial genome in Chlamydomonas is its very reduced gene content relative to other green algal and land plant counterparts. The two Chlamydomonas mitochondrial genomes completely sequenced to date, i.e., those of C. reinhardtii (Michaelis et al., 1990, and references therein) and C. eugametos (Denovan-Wright et al., 1998) have the same set of standard genes: seven respiratory protein-, three transfer RNA (tRNA)- as well as fragmented and

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Aurora M. Nedelcu and Robert W. Lee scrambled SSU and LSU rRNA-coding regions (Fig. 2). However, a reverse transcriptase-like gene (rtl) (Boer and Gray, 1988b) and an additional tRNA coding region (Denovan-Wright et al., 1998) have been identified in C. reinhardtii and C. eugametos, respectively. Nevertheless, different codon usage and deduced amino acid composition of the reverse transcriptase-like coding region led Boer and Gray (1988b) to suggest that rtl in C. reinhardtii had an independent more recent origin relative to the standard mitochondrial genes. Although in Oenothera berteriana mtDNA an independent open reading frame showing reverse transcriptase-like similarity has also been described (Schuster and Brennicke, 1987), it is noteworthy that the rtl in C. reinhardtii mtDNA is flanked by intergenic regions that contain sequence motifs present at the splice sites of group II introns (Nedelcu and Lee, unpublished). The additional coding region present in C. eugametos mtDNA flanks one of the two copies of a large direct repeat (Fig. 2) and may be the result of a duplication/inversion-related event given that the other end of the large direct repeat is also flanked gene. Duplicated tRNA genes or tRNA by a pseudogenes have been previously reported at inversion ends in wheat and rice chloroplast genomes (Howe et al., 1988; Shimada and Sugiura, 1989). The presence of virtually the same set of coding regions in the mitochondrial genomes of two Chlamydomonas taxa that belong to lineages considered to have diverged very early in the evolution of the Chlamydomonas group allows us to speculate that (i) the feature ofa very reduced gene content was already present in the most recent chlamydo­ monadalean common ancestor and (ii) gene content is not a significant contributor to the mitochondrial genome size variation within the Chlamydomonas group. No Chlamydomonas chloroplast genome is fully described, but half of the C. moewusii and C. eugametos chloroplast genomes have been sequenced and the analysis of these genomes revealed half of the land plant cpDNA gene content (Boudreau

Chapter 5

Mitochondrial and Chloroplast Genome Evolution

et al., 1994). Moreover, the two very divergent Chlamydomonas taxa, C. reinhardtii and C. moewusii, appear to share a similar gene complement (Fig. 3) (Boudreau et al., 1994). Among the 75 genes mapped on the two Chlamydomonas chloroplast genomes, five coding regions have not been reported in any land plant cpDNA, and four of these have also not been identified in other green algal counterparts (Boudreau et al., 1994). On the other hand, the genes encoding the chlororespiratory NADH dehy­ drogenase subunits present in land plant cpDNA may be lacking in the chloroplast genome of Chlamydomonas. The information accumulated so far led Boudreau et al. (1994) to suggest that the presence of additional genes accounts only for a very small fraction of the increased size of the chlamy­ domonadalean chloroplast genome relative to the land plant homologs. It is noteworthy that some chloroplast genes in C. reinhardtii display unusual structures and the corresponding transcripts are undetectable, suggesting that they might not be functional (Fong and Surzycki, 1992). Although there seems to be some variation in gene content among green algal chloroplast genomes (e.g., several additional genes involved in nucleotide metabolism reported in the cpDNA of the ulvo­ phycean taxon Acetabularia mediterranea and a more reduced gene content in that of Codium fragile, see Palmer, 1991 for discussion), the most striking differences in gene content have been observed among mitochondrial genomes. Unexpectedly, the mito­ chondrial genomes of green algal taxa outside the chlamydomonadalean group do not seem to share the very reduced gene content of their Chlamydo­ monas counterparts. The Platymonas subcordiformis mtDNA encodes at least 12 respiratory proteins, seven tRNAs, two ribosomal proteins as well as continuous LSU and SSU rRNAs (Kessler and Zetsche, 1995). Moreover, the mitochondrial genome of Prototheca wickerhamii codes for 16 respiratory proteins (including three subunits of the ATPase complex which is entirely non-mitochondriallyencoded in Chlamydomonas), 26 tRNAs, 13 ribosomal proteins as well as 5S, LSU and SSU rRNAs (Wolff et al., 1994). Given that the mitochondrial genome of the ancestral-like green algal lineage Platymonas as well as of the advanced trebouxiophycean (sensu Friedl 1995) lineage, Prototheca, has a gene content more similar to the land plant counterparts, one can hypothesize that the most recent common ancestor

73

of green algae and land plants contained quite a large number of genes in its mtDNA and that a reduction in gene content occurred in the chlorophycean lineage leading to Chlamydomonas. However, given that Chlamydomonas and Prototheca may not be as closely related as previously thought (Friedl, 1995) (i.e., they are in fact members of the chlorophycean and trebouxiophycean lineages [sensu Friedl, 1995], respectively, whose divergence is probably very old) the differences in gene content between Chlamy­ domonas and Prototheca are less surprising. The dichotomy in mitochondrial gene content within the green algal group is also reflected in different degrees of resemblance to their counterparts in land plants, the group considered their closest relatives at the nucleo-cytosolic level (Chapman and Ragan, 1980; Chapman and Buchheim, 1991). While the mitochondrial genomes of C. reinhardtii and C. eugametos (and most likely of all the chlorophycean taxa) resemble more closely their ciliate/fungal/ animal counterparts in terms of gene content, those of Platymonas subcordiformis and Prototheca wickerhamii (and most likely of all trebouxiophycean lineages) have a gene content similar to their land plant counterparts.

4. Changes in the Amount of Repeated DNA Large repeated sequences have been found in mitochondrial genomes of Chlamydomonas. The terminal non-coding regions of the C. reinhardtii linear mtDNA each contain a copy of an inverted repeat (TIR) of about 530 bp (Fig. 2) (Vahrenholz et al., 1993). On the other hand, the two largest intergenic regions in the C. eugametos circular-mapping mitochondrial genome each contain a copy of a 260­ bp direct repeat (DR) (Fig. 2) (Denovan-Wright et al., 1998). TIRs were described also in the linear mitochondrial genome of the colonial chlamy­ domonadalean taxon Pandorina morum (Moore and Coleman, 1989). Given that among sexually incompatible members of P. morum the TIRs range in size from 1.8 to 3.3 kb, it seems, at least in this taxon, that the variation in the amount of repeated DNA could be responsible in part for the observed intraspecific variation in mitochondrial genome size (from 20 kb to 38 kb). However, complete mitochondrial genome sequence data for more chlamydomonadalean taxa are needed in order to make any suggestion as to how significant, if at all, is the amount of repeated DNA in genome size evolution

74

Aurora M. Nedelcu and Robert W. Lee

Chapter 5

Mitochondrial and Chloroplast Genome Evolution

within the chlamydomonadalean group. Among the large repeated chloroplast DNA sequences in Chlamydomonas, the most common is a two-copy inverted repeat (IR) found always as two identical but oppositely oriented copies that divide the chloroplast genome into two rather equal singlecopy regions (Fig. 4a) (Rochaix, 1978; Lemieux et al., 1985; Turmel et al, 1987; Boynton et al., 1992). As in land plants, the Chlamydomonas IR varies in size by spreading or shrinkage (Turmel et al., 1991). The 50-kb difference in chloroplast genome size between the very closely related C. moewusii and C. eugametos is mostly accounted for by an enlarged inverted repeat in C. moewusii, which was shown to be the consequence of a 21 -kb insertion (Turmel et al., 1987). The expansion/contraction of the inverted repeat also accounts for the gain/loss of a 7-kb sequence containing the atpB gene between the two closely related C. reinhardtii and C. gelatinosa (Boudreau and Turmel, 1996). On the other hand, although there is a 53-kb difference in size between the chloroplast genomes of the distantly related C. eugametos and C. reinhardtii, their inverted repeats are about the same size in both taxa. Nevertheless, similar size does not necessarily imply identical sequence complexity, given that the inverted repeat in C. reinhardtii lacks the rbcL gene, which is located instead in a single-copy region (Malnoë et al., 1979; Dron et al., 1982). In contrast, although the inverted repeats of the interfertile C. reinhardtii and C. smithii are identical in gene content and similar in overall gene organization, they differ at the fine structure level by an extensive series of small deletions/ additions: a minimum of 11 length mutations (20– 1600 bp) are distributed throughout the IR of C. smithii, which makes it almost 1 kb larger that the 22-kb repeat of C. reinhardtii (Palmer et al., 1985). Comparisons between the inverted repeats of the members of each of the interfertile Chlamydomonas pairs, C. reinhardtii/C. smithii and C. eugametos/ C. moewusii, have revealed more deletion/addition differences than nucleotide substitutions (Palmer et al., 1985; Lemieux et al., 1985), which is opposite to the situation observed in higher plants (Zurawski et al., 1984). Among green algae there is even more variation than seen in Chlamydomonas in terms of the amount of repeated DNA in the organellar genomes. While there are no large repeats in the circular-mapping mitochondrial genome of Prototheca wickerhamii, a two-copy inverted repeat of ca. 1.5 kb was reported in the Platymonas subcordiformis homolog (Kessler

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and Zetsche, 1995). Similarly, among green algae, the chloroplast IR ranges in size between 20 and 41 kb and seems to be lacking in a few charophycean and ulvophycean lineages whose cpDNAs are also known to be smaller than the chloroplast genome size average (see Palmer, 1991, for references).

B. Mechanisms Possibly Involved in the Evolution of Genome Size 1. Length Mutations Changes in organellar genome sequence complexity occur primarily by length mutations, i.e., the addition of new sequences or the deletion of existing ones. Many of the small-length mutations in organelle genomes were found to be flanked by or close to short direct repeats, suggesting their occurrence during DNA replication or repair according to the ‘slippage-mispairing’ model (Takaiwa and Sugiura, 1982; Zurawski et al., 1984). The great majority of length mutations are small, only 1–10 bp in size, and occur predominantly in noncoding DNA (intergenic spacers and introns). Length mutations of 10–1,200 bp in size occur less frequently than smaller ones and are more likely to occur by recombination than by replication; unequal crossing-over between mis­ aligned tandem repeats could produce both deletions and additions, and intramolecular recombination between short direct repeats could produce deletions (see Palmer, 1991 for a review). Repetitious DNA is very abundant in the mitochondrial genome of Chlamydomonas. Short dispersed repeats have been reported in five of the intergenic spacers ofthe C. reinhardtii mitochondrial genome (Boer and Gray, 1991). A more abundant and complex set of repetitive sequences (in both direct and inverted orientation) has been found in the mitochondrial genome of C. eugametos; more than 80 repetitive elements, ranging in size from 6 bp to 17 bp, are dispersed throughout the intergenic regions as well as within several introns (Denovan-Wright et al., 1998; Nedelcu and Lee, 1998). Identical copies or closely related sequences of the same repeat are present tandemly repeated in various combinations. The C. reinhardtii chloroplast genome also possesses a family of about 40 short (100–300 bp) repeated sequences scattered throughout most of the genome (Rochaix, 1978; Gelvin andHowell, 1979), at least 13 of them being localized within the IR (Palmer et al., 1985). The short dispersed repeats account for ca. 22% of the total C. reinhardtii

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chloroplast genome and each is composed of shorter repeated motifs that occur in direct or inverted orientation and in various combinations (Gillham, 1994). Given that a number of the observed smalllength mutations mapped close to regions showing differences in the number of these small repeats, Palmer et al. (1985) suggested that enhanced recombination within and between repeat elements may be related to the increased incidence of length

Aurora M. Nedelcu and Robert W. Lee

mutations in Chlamydomonas relative to angiosperm cpDNA. Studies of experimentally induced mutants showed that the endpoints of most of the structural mutations, both deletions and inversions, mapped in the general vicinity of the 100–3 00-bp repeat elements scattered throughout the inverted repeat (Palmer et al., 1985). The high frequency of symmetrical alterations suggested the existence of a copycorrection mechanism for maintaining identity

Chapter 5 Mitochondrial and Chloroplast Genome Evolution between the two copies of the inverted repeat. The mechanism for insertion of new sequences within the intergenic regions of Chlamydomonas cpDNA is not known. Because no dispersed repeats have been detected by Southern blot hybridization in the cpDNAs of C. pitschmannii and C. eugametos, Boudreau and Turmel (1995) did not favor the proliferation of existing sequences throughout the genome through unequal recombination as a mechanism responsible for the 56-kb difference in size between the intergenic regions in the cpDNAs of these two taxa. It is noteworthy that in land plant cpDNA, repeats were thought to have been created and spread by duplicative transposition (Tsai and Strauss, 1989). No classical transposable element has been, however, isolated from any land plant chloroplast. In contrast, the repeated sequences and the ORFs (whose deduced amino acid sequence show some similarity with transposases and integrases of other mobile elements) associated with the Wendy element in the C. reinhardtii chloroplast genome argue for the existence, present or past, of a transposable element in this lineage. Short repetitive sequences are also present in the organellar genomes of other green algae. The mitochondrial genome of P. wickerhamii is rich in very complex repetitive motifs consisting of AT-rich tandem repeats in both intergenic spacers and introns; although it has been suggested that they might be implicated in transcription and/or processing, their evolutionary origin is not known (Wolff et al., 1994). In the Chlorella ellipsoidea chloroplast genome, insertions of repeated sequences as well as ORFs with terminal repeated sequences account for larger intergenic spacers in its IR relative to the C. reinhardtii counterpart (Yamada, 1991). Because the smallest chloroplast IRs known to date contain at least the rRNA (rrn) operon, Yamada (1991) suggested that the IR might have been originally created from the duplication (in an inverted orientation) of the rrn operon, followed by its expansion to incorporate additional coding regions. The mechanism proposed for the expansion of the IR involves a double reciprocal recombination during the replication step and requires the presence of repetitive sequences acting as recombination hot spots within and around the IR (Yamada, 1991). Such a model is consistent with the location of the repetitive sequences in the inverted repeat of the Chlorella ellipsoidea chloroplast genome.

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2. Intron Mobility Although gain or loss of introns could be considered a special case of length mutation (Palmer, 1991), the mechanisms underlying these processes are very different. It was proposed that the loss of introns is the result of two processes: (i) the reversetranscription of an RNA whose intron sequences have been removed by splicing and (ii) homologous recombination between the intronless cDNA and the native gene (Dujon, 1989). In mitochondria, the putative reverse transcriptases (RTs) encoded either by the group II intronic ORFs or certain non-intronic ORFs with reverse transcriptase similarity, as found in C. reinhardtii (Boer and Gray 1988b) and a few angiosperms (see Moenne et al., 1996 for references), could produce an intronless copy from a spliced RNA. Although the above mentioned processes seem more likely to account for the loss of group II introns, Wolff et al. (1993) considered that the reversetranscriptases encoded in group II introns could have also been responsible for the loss of group I introns, culminating in their complete elimination from higher land plant mtDNA. It is noteworthy that although the overexpressed gene product of C. reinhardtii rtl does not appear to have a reverse transcriptase activity (Faßbender et al., 1994), such an activity has been recently detected in the mitochondria of potato (Moenne et al., 1996). On the other hand, in the Chlamydomonas chloroplast, evidence for reverse transcriptase activity is still lacking, but recom­ bination processes seem to be well developed (Dürrenberger et al., 1996). To explain the transfer of group I introns to novel locations (intron transposition), two mechanisms have been proposed: one occurs at the DNA level and is promoted by an intron-encoded endonuclease (Dujon, 1989), whereas the other starts at the RNA level and involves (i) the introduction of an intron RNA sequence into a foreign RNA by a reversal of a selfsplicing reaction, (ii) the reverse-transcription of the recombined RNA and (iii) the integration of the cDNA sequence into the genomic DNA by homolo­ gous recombination (Woodson and Cech, 1989). Turmel et al. (1993) presented evidence that reverse self-splicing might have played the major role in the creation of novel intron insertion sites in the LSU rRNA genes (rrnL) as well as elsewhere in the chloroplast genome of Chlamydomonas. However, these authors also suggested that certain group I

78 introns might have been introduced via lateral transfer facilitated by the site-specific endonuclease encoded in these introns. Such a mechanism is considered to be responsible for the mobility (intron homing) of the chloroplast rrnL intron of C. eugametos (Lemieux and Lee, 1987; Gauthier et al., 1991; Bussières et al., 1996) and C. reinhardtii (Dürrenberger and Rochaix, 1991) as well as of the mitochondrial cob intron of C. smithii (Boynton et al., 1987; Matagne et al., 1988; Colleaux et al., 1990; Ma et al., 1992) to cognate positions within the corresponding intronless genes. Another case indicative of intron mobility is the apparent evolutionary transfer of a group I intron between the mitochondrial rrnL of Acanthamoeba castellanii and the chloroplast counterpart of Chlamydomonas, either intracellularly in a remote photosynthetic common ancestor of the two lineages, or intercellularly, as a result ofa recent lateral transfer event (Lonergan and Gray, 1994; Turmel et al., 1995b).

3. Gene Transfer Among the processes of genetic flux (such as gene transfer, reverse gene transfer, gene substitution, gene sharing, gene recruitment, and gene loss—see Palmer, 1991 for a review) that account for the differences in gene content among lineages, gene transfer seems to be the main contributor to changes in genome complexity in organellar genomes. However, Palmer (1991) considers that the incor­ poration of a foreign gene into an organelle genome (reverse gene transfer) is not very rare and includes events such as the invasion of chloroplast genes into the higher plant mtDNA as well as the recent acquisition of a number of ORFs within land plant and green algal chloroplast introns (see Palmer, 1991 for references). To explain the gene transfer from one cellular compartment to another, Obar and Green (1985) proposed a stepwise model involving: (i) duplication of an organellar gene followed by the transfer of one copy to the nucleus; (ii) activation of the nuclear copy still keeping active the organellar counterpart; and (iii) inactivation and subsequent loss of the organellar gene copy. Gene transfer from the organelle to the nuclear genome seems to have been more important in the evolution of the mitochondrial than the chloroplast genome in Chlamydomonas. The presence of a higher number of genes in the mitochondrial genome of Platymonas subcordi-

Aurora M. Nedelcu and Robert W. Lee formis, considered a descendant of the primitive green flagellates from which all the advanced green algal lineages have evolved, suggests that the feature of a very reduced gene content in Chlamydomonas is a derived trait among green algae. The mechanisms and causes responsible for such a massive reduction in the gene content of mtDNA in Chlamydomonas are not known, although a few suggestions have been made. Recombination between short direct repealed sequences was proposed to have been responsible for the ‘excision’ of mitochondrial coding regions during the evolution of the Chlamydomonas-like genomes (Nedelcu, 1997; Nedelcu and Lee, 1998). Short direct repeated sequences and short particularly recombinogenic GC-rich clusters within AT-rich spacers regions have been shown to be involved in site-specific intra­ molecular recombination events resulting in the excision of small subgenomic circles in plant and fungal mitochondrial genomes, respectively (Fig. 4b) (Zinn et al., 1988; Hartmann et al., 1994 and references therein; Benslimane et al., 1996; JamietVierny et al., 1997 and references therein). It is possible that the accumulation of GC-rich short direct repeated sequences with recombinogenic properties in the lineage leading to Chlamydomonas could have promoted recombination events responsible for the deletion of protein-coding genes as well as tRNA- or rRNA-coding regions (Fig. 4d). Nevertheless, the processes accounting for the transfer of genetic information into the nucleus remain to be deciphered. Theoretically, there are two ways one can envision such a transfer: at the DNA or RNA level. The transfer of a DNA molecule from one compartment to another could be comparable to the transfer of an episome from one eubacterial cell to another, following the excision, conjugation, and integration steps described in a transformation cycle. Alternatively, the genetic information transcribed into an RNA molecule could be reverse-transcribed into a DNA molecule either before or after leaving the organelle, and subsequently integrated into the nuclear DNA. It is noteworthy that gene transfer from the mitochondria to the nucleus seems to be an on-going process among flowering plants and at least in this group, the transfer seems to happen at the RNA level, because the nuclear copies resemble more the edited rather than unedited versions of the mitochondrial genes (Brennicke et al., 1993; Schuster and Brennicke, 1993; Gray, 1995).

Chapter 5 Mitochondrial and Chloroplast Genome Evolution V. Evolution of Mitochondrial and Chloroplast Genome Organization in Chlamydomonas

A. Mitochondrial and Chloroplast Genome Structure An unexpected dichotomy in mitochondrial genome conformation has been observed among Chlamy­ domonas taxa: linear mtDNA molecules have been isolated from C. reinhardtii (Boer et al., 1985), C. smithii (Boynton et al., 1987) and the colonial chlamydomonadalean alga Pandorina morum (Moore and Coleman, 1989) but circular-mapping mtDNAs have been reported for C. eugametos (DenovanWright and Lee, 1992), C. moewusii (Lee et al., 1991) and C. pitschmannii (Boudreau and Turmel, 1995). However, as discussed by Bendich (1993), electrophoretic migration patterns of the circularmapping mitochondrial genomes of C. moewusii and C. eugametos (Boer et al., 1985; Lee et al., 1991) leave open the possibility of their existing in vivo as linear, larger-than-unit-size genomes. Circularmapping mitochondrial genomes have also been reported in other chlorophycean taxa such as Chlorella and Scenedesmus obliquus, as well as trebouxiophycean and prasinophycean taxa like Prototheca wickerhamii and Platymonas subcordi­ formis, respectively (Kück, 1989; Moore and Coleman, 1989; Waddle et al., 1990; Kessler and Zetsche, 1995). It is note worthy that although circularmapping mitochondrial genomes have also been reported in land plants, they may exist in vivo predominantly as larger-than-unit-genome-size linear structures (Bendich, 1993, 1996). If the circular-mapping mtDNAs are circular molecules in vivo, their linearization in some chlamydomonadalean lineages could have been the consequence of a recombination event between short repeated sequences on a small linear episome and their homologs on the circular chromosome, as described during the linearization of the maize mitochondrial genome (Schardl et al., 1984). In this connection, it is intriguing that one of the long terminal inverted repeats in the linear C. reinhardtii mtDNA is flanked by small inverted repeats thus arguing for a potential previous episomal existence of this TIR (Nedelcu and Lee, 1998). In contrast, chloroplast genomes appear to be circular in conformation in all Chlamydomonas

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investigated to date; however, they exist as a 50:50 mixture of two genetically identical but physically distinct molecules that differ only in the relative orientation of their single-copy regions as a result of high-frequency intramolecular recombination events between the two copies of the IR (Fig. 4a) (Aldrich et al., 1985; Palmer et al., 1985). In addition, intermolecular recombination events involving the short repeated sequences in the C. reinhardtii cpDNA have been suggested to yield dimer and multimer cpDNA molecules (Boynton et al., 1992; Boudreau and Turmel, 1996).

B. Mitochondrial and Chloroplast Gene Order Although the Chlamydomonas mitochondrial genome size and gene content is more animal-like than plant­ like, the gene order among taxa appears to be as variable as observed among the vascular plant mitochondrial genomes. None of the coding regions is flanked by homologous counterparts in both the C. reinhardtii and C. eugametos mitochondrial genomes, i.e., there is no gene cluster common to the two genomes and the protein-coding genes are highly interspersed with tRNA genes as well as rRNA gene pieces in both Chlamydomonas genomes (Boer and Gray, 1988a; Denovan-Wright et al., 1998) (Fig. 2). Given that in C. eugametos the protein-coding genes are more interspersed with rRNA-coding regions than they are in C. reinhardtii, it seems likely that the mitochondrial genome of C. eugametos has undergone additional gene rearrangements relative to its C. reinhardtii counterpart. It is interesting that, in contrast to land plants, the chloroplast gene order is quite variable among Chlamydomonas taxa. Although the chloroplast genomes of the interfertile members of each pair of taxa, C. eugametos/C. moewusii and C. reinhardtii/ C. smithii are co-linear (Turmel et al., 1987; Boynton et al., 1992), the cpDNAs are so extensively rearranged between the two pairs (i.e., C. eugametos/ C. moewusii and C. reinhardtii/C. smithii) that rearrangements cannot be described in terms of simple individual events (Fig. 3). It has been suggested that the great evolutionary distance separating these algae might be responsible for such a high level of rearrangements although the possibility that cpDNA rearranges at a fast rate in Chlamydomonas cannot be disregarded (Lemieux and Lemieux, 1985). Interestingly, a comparative analysis of chloroplast

80 gene order in the two closely related Chlamydomonas taxa, C. moewusii and C. pitschmannii, has revealed a level of rearrangement close to that observed among all land plants: one or two inversions and possibly one or three events of expansion/contraction of the inverted repeat (Boudreau and Turmel, 1995). Unexpectedly, however, the level of rearrangements appears much more extensive between C. reinhardtii and C. gelatinosa (i.e., at least nine inversions and one expansion/contraction event of the IR), although chloroplast LSU rDNA sequence-based phylogenies suggest they are as closely related as are C. moewusii and C. pitschmannii (Boudreau and Turmel, 1996). Most changes in gene order in the two pairs of closely related Chlamydomonas are located in the single-copy region bordering the rrnS gene (Boudreau and Turmel, 1995, 1996). In the course of these rearrangements, a few cpDNA sequences have moved from one single copy region to the other, a phenomenon not observed in higher plant cpDNA. Chlamydomonas cpDNAs lack the extensive operon structure of their land plant counterparts; the best example is represented by the six atp genes, which are organized into two operons in all other chloroplasts but are scattered singly around the genome in a species-specific manner in Chlamy­ domonas (Palmer, 1991). It has been suggested (Turmel et al., 1988) that the numerous rearrange­ ments during Chlamydomonas cpDNA evolution resulted in the disruption of the polycistronic transcription units inherited from the prokaryotic ancestor; in contrast, different evolutionary pressures during land plant evolution determined an increased number of polycistronic transcription units and a more compact genome organization. Most of the ancestral operons still present in land plant cpDNAs have been lost in the Chlamydomonas lineage; of 76 genes mapped on five Chlamydomonas cpDNAs, 40 represent 15 conserved clusters, four of which are similar to the primitive operons present in land plant chloroplast genomes, and one has exactly the same gene content as the land plant equivalent (Boudreau et al., 1994; Boudreau and Turmel, 1996). The level of mitochondrial gene rearrangement among green algae is difficult to assess due to the lack of complete mitochondrial genome sequences for green algal lineages other than Chlamydomonas and Prototheca, as well as the very reduced gene content of the mitochondrial genome of the former relative to the latter. Nevertheless, among the 12 genes common to both green algal lineages, there is only one gene cluster that is common to Prototheca

Aurora M. Nedelcu and Robert W. Lee wickerhamii and C. reinhardtii, i.e., the nad5-nad4 cluster, and none between P. wickerhamii and C. eugametos. It is noteworthy that among the genes that have been mapped on the mitochondrial genome of Platymonas subcordiformis, the only gene cluster that is shared with Prototheca is the same nad5-nad4 cluster. Probably the most unexpected variation in gene order among the green algal mitochondrial genomes has to do with one of the most conserved gene clusters in the land plant counterparts, i.e, the one comprising the rRNA genes: in Platymonas, as in most other mitochondrial systems, rrnL and rrnS are located on the same DNA strand, while in Prototheca and land plant mitochondrial genomes the rrnS and rrnL are encoded on opposite DNA strands. It is also interesting that there might be at least five polycistronic units in the mtDNA of Platymonas, but as few as two in Prototheca and C. reinhardtii mtDNA, and only one in C. eugametos. Similarly, although limited, studies on cpDNA in green algae outside the chlamydomonadalean group have disclosed a highly variable gene order with only few conserved gene clusters, suggesting that the evolutionary pattern of green algal cpDNA is less conservative than that of their land plant counterparts (Palmer, 1991). The few green algal chloroplast genomes mapped to date do not share similar gene orders with either one another or with Chlamy­ domonas counterparts. Some gene clusters, however, are present in more than one lineage: for instance, the petA-petD gene cluster present in Chlamydomonas is also found in Scenedesmus obliquus (Kück, 1989; Kück et al., 1990) and it was suggested that a more extended gene cluster, including might have been present in the most recent common ancestor of Chlamydomonas and Scenedesmus (Boudreau et al., 1994). Moreover, two clusters, psaA-psaB and psbC-psbD, are shared by Spirogyra and Codium (Manhart et al., 1990). DNA rearrange­ ments involving ancestral polycistronic units also occurred d u r i n g the evolution of green algal chloroplasl genomes; rrnS and rrnL that are cotranscribed in all bacteria, as well as Chlamydomonas and land plant chloroplasts, are separately transcribed in Spirogyra (Manhart et al., 1990), Codium (Manhart et al., 1989) and Chlorella ellipsoidea (Yamada and Shimaji, 1987).

1. Factors Contributing to Gene Rearrangement Palmer (1990) suggested several factors that are likely to promote more inversional recombination in

Chapter 5 Mitochondrial and Chloroplast Genome Evolution land plant mtDNA relative to cpDNA: (i) more short dispersed repeats that could serve as points for homologous recombination; (ii) larger intergenic regions that could tolerate inversions; and (iii) more monocistronic rather than multicistronic mito­ chondrial operons. It is interesting that short repeated sequences have been found in the intergenic spacers of both C. reinhardtii and C. eugametos mtDNAs. Furthermore, the presence of fewer and more reduced gene clusters in the C. eugametos mitochondrial genome relative to the C. reinhardtii homolog, as well the disruption of the conserved nad5-nad4 gene cluster in the former, are correlated with a more abundant and complex set of short GC-rich repetitive sequences in the intergenic regions of the former relative to the latter (Nedelcu and Lee, 1998). Short repeated sequences are also present in the mito­ chondrial genome of P. wickerhamii, but they are highly AT-rich (Wolff et al., 1994). It is noteworthy in this connection that small dispersed repeated sequences of ca. 50–1000 bp have been reported in the land plant mitochondrial genome and shown to be involved in intramolecular recombination events promoting gene rearrangements (Hartmann et al., 1994 and references therein; Benslimane et al., 1996). Not surprisingly, short dispersed repeats are also particularly abundant in the highly rearranged cpDNAs of Chlamydomonas. The dispersed repeats in the C. reinhardtii and C. gelatinosa cpDNA are composed of short repeated units occurring in different combinations at different loci in the genome (Boynton et al., 1992; Boudreau and Turmel, 1996). The insertion of repeated sequences in intergenic spacers has been suggested to have been a significant factor in promoting the disruption of the ancestral polycistronic operons in the lineage leading to Chlamydomonas (Boudreau et al., 1994). The very different number of repeated sequences between the cpDNAs of the members of two apparently equally distant pairs of closely related Chlamydomonas taxa, C. reinhardtii/C. gelatinosa and C. moewusii/ C. pitschmannii, correlates directly with the very different level of gene rearrangement between the cpDNAs of the two lineages (Boudreau and Turmel, 1996). It is noteworthy that the tobacco and Marchantia chloroplast genomes, which differ in gene order by a single 30 kb inversion despite about 400 million years of evolutionary separation, contain no dispersed repeats larger than 50 bp (Palmer, 1990). However, small dispersed repeats of 50–1000 bp are unusually abundant in most of the land plant genomes that are highly rearranged. Short terminal degenerate

81

inverted repeats and four-nucleotide directly repeated sequences as well as additional degenerate copies of these sequences in direct or inverted orientation have been also found within the two copies of the Wendy element suggested to be involved in rearrangement events in the chloroplast genome of C. reinhardtii (discussed later) (Fan et al., 1995).

2. Mechanisms Possibly Involved in Gene Rearrangement In higher plant mitochondria, recombination has been invoked to explain the genomic rearrangements accounting for their complex mitochondrial genome structures. It has been proposed that the frequency of homologous recombination is related to the presence of recombinogenic repeated sequences (Palmer and Shields, 1984). A model invoking large direct and inverted repeat-mediated inter- and intramolecular recombination events involving various isomeric forms of the mitochondrial genome has been proposed to illustrate the multipartite structure and the evolution of the maize mitochondrial genome (Fauron et al., 1995). Because there is no evidence for a multipartite structure of the mitochondrial chromosome in Chlamydomonas, recombination events similar to those proposed for the land plant mitochondrial genome do not seem very likely to have happened during the evolution of the Chlamydomonas counterparts. Rather, we suggest that intramolecular recombination between one and two sets of inverted repeats, which results in the inversion and interchange of the regions flanked by the repeats, respectively, (Fig. 4b and c), might have played an important role in the evolution of mitochondrial genomes in Chlamydomonas. Moreover, we think that periods of instability associated with mtDNA replication occurring at multiple sites, first competing with, and later replacing the conventional origin of mtDNA replication could have also been contributed to the extensive gene rearrangement observed in Chlamy­ domonas, in a manner similar to that proposed for the vertebrate mitochondrial genome (Macey et al., 1997). The GC-rich palindromic sequences present in the intergenic spacers of C. reinhardtii and C. eugametos could act as surrogate origins of replication of the mitochondrial DNA (Nedelcu and Lee, 1998). In land plant chloroplast genomes, on the other hand, homologous or illegitimate recombination between short repeats (as small as 11–16 bp) and/or

82 tRNA genes (trn), both functional and pseudogenes, is considered the major cause of gene rearrangements (see Boudreau and Turmel, 1995, 1996 for references). The relative abundance of appropriately oriented and located (between, rather than within transcription units) short dispersed repeats seems to be a major factor in determining the prevalence of cpDNA inversions in land plants (Palmer, 1991). Similarly, recombination events between short dispersed repeats have also been proposed to account for the various rearrangements described in the Chlamydomonas chloroplast genome (discussed by Boudreau and Turmel, 1996). Moreover, the finding that trn-specific oligonucleotide probes hybridized near the endpoints of an inversion in C. pitschmannii cpDNA led Boudreau and Turmel (1995) to raise the possibility of intra- or intermolecular recombination events between duplicated tRNA genes being responsible for the observed inversion. In addition, in C. reinhardtii, the Wendy element is considered to have played a major role in the shuffling of chloroplast gene clusters in this lineage relative to other Chlamydomonas lineages; the fact that both copies of Wendy were found to be flanked by gene clusters that are contiguous in C. moewusii but are separated and inverted relative to each other in C. reinhardtii argues for such an involvement. The mechanisms involved in such rearrangements might have involved Wendy-dependent illegitimate homologous or site-specific recombination events, or both (Fan et al., 1995).

VI. Evolution of Mitochondrial and Chloroplast Gene Structure and Organization in Chlamydomonas

A. Intron-containing Coding Regions Both mitochondrial and chloroplast coding regions in chlamydomonadalean taxa are interrupted by introns. No introns of the group II type have been identified in the mitochondrial genes of the group, but a trans-spliced group II intron has been found in the chloroplast psaA gene of several Chlamydomonas taxa (see Turmel et al., 1995a for references). Introns of the group I family have been found in mitochondrial rRNA- and protein- but not tRNAcoding regions of Chlamydomonas. It is interesting, however, that the mitochondrial genome of C. rein­ hardtii is devoid of any intronic sequences. A group I

Aurora M. Nedelcu and Robert W. Lee intron has been also reported in the only other available mtDNA sequence of a chlorophycean taxon outside the chlamydomonadalean group, i.e., a partial sequence of mitochondrial rrnL from Scenedesmus obliquus (Kück et al., 1990). Because the insertion site of the rrnL group I intron in S. obliquus is four nucleotides downstream of the insertion site of one of the C. eugametos mitochondrial rrnL group I introns, it is difficult to make any suggestions as whether the most recent common ancestor of these two taxa, which belong to the two main evolutionary lineages within the chlorophycean group, namely the DO and CW lineages, had or did not have group I introns in its LSU rRNA gene. Nevertheless, two group I introns have been found in the mitochondrial rrnL of the trebouxiophycean (sensu Friedl, 1995) taxon Prototheca wickerhamii: one intron is present at the same position as the mitochondrial rrnL group I intron in S. obliquus (Wolff et al., 1993), where as the other intron shares an identical insertion site with one of the three mitochondrial rrnL group I introns in C. eugametos. This finding could argue for the presence of at least two introns in the mitochondrial LSU rRNA gene in the most recent common ancestor of the trebouxiophycean and chlorophycean lineages, followed by the loss of one of the introns as well as the acquisition of new introns at new insertion sites in each of the two lineages. It seems likely, therefore, that the most recent common ancestor of green algae had introns in its mitochondrial coding regions, and subsequently introns were independently lost or acquired in distinct evolutionary lineages. Interestingly, two of the three group I introns present in cox1 of Prototheca wickerhamii are located at positions identical to the sites of insertion of liverwort mitochondrial cox1 introns, and it has been suggested that they were already present in the common chlorophyte/embryophyte ancestor (Wolff et al., 1993). Moreover, both Scenedesmus obliquus and liverwort mtDNAs contain introns ofthe group II family, which represent the only type of intron found in the mitochondrial genes of angiosperms. It is interesting to note that the reverse transcriptase-like coding region present in the C. reinhardtii mtDNA seems to be in fact the intronic ORF of a degenerate group II intron (Nedelcu and Lee, 1998). These observations may suggest that the most recent common ancestor ofthe green algal/land plant group contained both group I and II introns in its mitochondrial coding regions and that a massive loss of all of the former, and most of the latter, has

Chapter 5

Mitochondrial and Chloroplast Genome Evolution

occurred in the tracheophyte (vascular plant) as well as C. reinhardtii lineage. In contrast, Chlamydomonas chloroplast genes contain both group I and II introns. The distribution of 39 group I introns representing 12 insertion sites is highly variable among 17 Chlamydomonas taxa and does not suggest the same phylogenetic relationships among Chlamydomonas lineages as do the chloroplast rDNA sequences (Turmel at al., 1993). Because the rrnL of cyanobacteria and of the Chlorella and land plant chloroplast lineages lack introns, it was suggested that all of the intron insertion positions in Chlamydomonas rrnL are of recent origin and that some of them might have arisen after the divergence ofthe two main Chlamydomonas lineages (Tunnel et al., 1993). Moreover, because the chloroplasts of both land plants and their closest relatives, the charophycean green algae, display overall a very small number of group I introns (Palmer, 1991), it has been suggested that the proliferation of group I introns occurred in the Chlamydomonas lineage after the charophycean divergence (Turmel et al., 1993). However, it was recently reported that the Chlorella vulgaris chloroplast rrnL is interrupted by a group I intron inserted at the same position as the single group I intron in the C. reinhardtii rrnL; intronic sequence comparisons as well as amino acid sequence similarities of their intronic ORFs suggested that the two closely related self-splicing rrnL group I introns in Chlorella vulgaris and C. reinhardtii descended from the same group I intron present in the most recent common ancestor of these two lineages (Kapoor et al., 1997). The only introns with features of the group II family identified to date in Chlamydomonas chloroplast genes are those found in psaA of several Chlamydomonas taxa (see Turmel et al., 1995a). While Chlamydomonas chloroplast genes have multiple introns in most of the genes possessing introns, land plant chloroplast genes contain single introns. Also, although land plant chloroplast tRNA genes harbor long single introns, no split tRNA genes have been found in algal chloroplasts. It is interesting that some of the land plant mitochondrial and chloroplast protein-coding genes consist of scattered exons flanked by 5´- or 3´segments ofgroup II introns; the exons are separately transcribed and spliced in trans (see Turmel et al., 1995a for references). In Chlamydomonas, such an organization has only been described among chloroplast genes. Chlamydomonas psaA coding

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regions are made up of three exons scattered around the genome and spliced in trans (Kück et al., 1987; Choquet et al., 1988; Turmel et al., 1991; Goldschmidt-Clermont et al., 1991; Turmel et al., 1995a). Although the location of the three exons in the genome is different between two very divergent taxa, C. moewusii and C. reinhardtii, the information contained is similar, indicating that the most recent common ancestor of these two lineages possessed a psaA coding region interrupted in a similar manner (Boudreau et al., 1994). In contrast, psaA is not trans-spliced in land plants or other genera of algae (with the exception of Euglena gracilis), but rps12 (which is uninterrupted in Chlamydomonas, Euglena and Cyanophora) is trans-spliced in all examined land plants (Sugiura, 1989). An additional feature contributing to variation in chloroplast gene structure in Chlamydomonas is the presence of translated large insertion sequences in the clpP gene of C. reinhardtii and C. eugametos (Huang et al., 1994) as well as chimeric RNA polymerase coding regions juxtaposed in-frame with long sequences of unknown origin in C. rein­ hardtii (Fong and Surzycki, 1992).

B. Fragmented Coding Regions In Chlamydomonas, both mitochondrial and chloroplast rRNA genes are fragmented into coding modules whose mature transcripts are not spliced together into covalently continuous rRNA molecules. However, the rRNA gene organization and expression as well as evolutionary origins are very different between the two organellar genomes. Mitochondrial rRNA genes in Chlamydomonas are not only highly fragmented but also scrambled (i.e., the gene pieces are interspersed with other coding regions and do not follow the 5'–3' transcriptional order of their counterparts in conventional continuous genes) (Fig. 2). The SSU and LSU rRNA-coding regions are fragmented into four and eight gene pieces, respectively, in C. reinhardtii (Boer and Gray, 1988a) and into three and six gene pieces, respectively, in C. eugametos (Denovan-Wright and Lee, 1994). The rRNA-coding modules are extensively interspersed with each other as well as with protein-coding and tRNA genes, and the rRNA fragments are most likely excised from longer multicistronic transcripts following precise endonucleolytic scissions (Boer and Gray, 1988a). Although the rRNA pieces are not spliced together,

84 they have the ability to interact through intermolecular base pairing, restore the conserved core of the rRNA secondary structure (Boer and Gray, 1988a; DenovanWright and Lee, 1994), and assemble into mito­ chondrial ribosomes (Denovan-Wright and Lee, 1995). Discontinuous mitochondrial rRNAs have been reported not only in other Chlamydomonas taxa (Denovan-Wright et al., 1996), but also in other chlorophycean taxa with a CW or DO flagellar configuration as well as in chlorococcalean taxa phylogenetiqally related to them (Nedelcu et al., 1996). A trend in the evolution of this trait, that is, a tendency towards an increase in the degree of discontinuity from continuous mitochondrial rRNAs to the highly fragmented mitochondrial rRNAs in C. eugametos and C. reinhardtii was suggested (Nedelcu et al., 1996; Nedelcu, 1997). Although mitochondrial rRNA genes are highly fragmented and scrambled in both C. reinhardtii and C. eugametos, the distribution of the coding in formation among their coding modules, as well the order of these modules within the genome, is different between the two species (Denovan-Wright and Lee, 1994). Calculations of the minimal number of transpositions required to convert hypothetical ancestral rRNA gene organizations to the arrange­ ments present in the two Chlamydomonas taxa, as well as a limited survey of the size of mitochondrial LSU rRNAs in other Chlamydomonas species, led Denovan-Wright et al. (1996) to propose that the last common ancestor of Chlamydomonas algae pos­ sessed fragmented mitochondrial rRNA genes whose coding modules were nearly co-linear with their counterparts in conventional continuous rRNA genes. The model presented by the authors predicted that in taxa basal to the Chlamydomonas group, mito­ chondrial rRNA genes would be fragmented but not scrambled. The presence of scrambled but not highly fragmented mitochondrial LSU rRNA coding regions in Scenedesmus obliquus, however, suggested that scrambling may have developed at an early stage in the evolution of discontinuous and scrambled rRNA genes within the chlorophycean green algal group, probably in parallel with the fragmentation events (Nedelcu, 1997). The mechanisms responsible for either the fragmentation or the scrambling ofthe mitochondrial rRNA coding regions in Chlamydomonas are not known yet, although several suggestions have been made. The GC-rich repeat clusters identified in

Aurora M. Nedelcu and Robert W. Lee C. reinhardtii mitochondrial DNA were suspected to have contributed to the extensive rRNA gene arrangements through a mechanism analogous to bacterial transposition (Boer and Gray, 1991). The absence of a reverse transcriptase-like open reading frame in C. eugametos mitochondrial DNA led Denovan-Wright and Lee (1994) to favor the view that the mitochondrial rRNA coding regions in Chlamydomonas became scrambled by recom­ bination between non-homologous regions of mtDNA molecules such as the dispersed repeated elements found in C. reinhardtii (Boer and Gray, 1991) and C. eugametos (Nedelcu and Lee, 1998) rather than by reverse transcription (Boer and Gray, 1988a). The authors further assumed that the unusual gene structure in Chlamydomonas mitochondria arose from conventional, continuous rRNA genes by two separate, consecutive processes: the introduction of processing signals and the scrambling of coding regions defined by these signals. Nedelcu (1997), however, proposed a recombination model that could disrupt and scramble a coding region in a single step. The proposal is an extension of the model presented by Fauron et al. (1995) for the evolution of the maize mitochondrial genome and involves an intramolecular homologous recombination event between two sets of two-copy inverted repeats. Generally, such an event would result in an interchange of the sequences situated between the two sets of inverted repeats (Fig. 4c). The way in which recombination events such as the one suggested above, as well as those proposed by Fauron et al. (1995), could have been involved in the evolution of chlorophycean mito­ chondrial rRNA genes is illustrated in Fig. 4d. Comparisons among the locations of the short repeated elements within the mitochondrial genome of C. reinhardtii, C. eugametos and the available DNA sequences of other chlorophycean green algae revealed similarities regarding the positions of these repeats relative to the rRNA-coding units within the respective genomes (Nedelcu, 1997; Nedelcu and Lee, 1998). It was suggested, therefore, that the fragmented and scrambled mitochondrial rRNA coding regions in the chlorophycean green algal group may have been generated through multiple recombination events triggered by the accumulation of short repeated sequences within the variable regions of the rRNA genes and the intergenic spacers of these mitochondrial genomes. To illustrate how recombination events similar to those proposed above could be entirely responsible for the extensive

Chapter 5 Mitochondrial and Chloroplast Genome Evolution mitochondrial rRNA gene rearrangements, a hypothetical pathway to gradually convert conven­ tional continuous mitochondrial rRNA genes to the rRNA gene arrangement described in C. eugametos was envisioned (Nedelcu, 1997). It is interesting that chloroplast LSU but not SSU rRNA-coding regions are also fragmented in Chlamydomonas. Three internal transcribed spacers (ITSs) located at the same position in the chloroplast rrnL of all 17 Chlamydomonas taxa investigated by Turmel et al. (1993) interrupt this gene into four gene pieces whose transcripts are not covalently linked after the removal of the ITSs from the primary transcript. Unlike the introns, but like the break points in the mitochondrial rRNA-coding regions, the ITSs are located within highly variable regions of primary and/or secondary structure. The ITSs in the Chlamydomonas chloroplast rrnL are usually less than 300 bp long and differ substantially in size and base composition. Although they are always excised post-transcriptionally from a precursor RNA to yield four mature rRNA species, no common sequence motif to account fora similar processing recognition signal has been identified, suggesting that either different recognition signals or specific threedimensional topology of the ribosome might be involved in the processing of ITSs (Tunnel et al., 1993). It is noteworthy that the size and base composition differences among corresponding ITSs in different Chlamydomonas taxa are not consistent with the phylogenetic relationships suggested by the LSU rRNA-coding sequences (Turmel et al., 1993). This feature of fragmented chloroplast LSU rRNAcoding regions is not confined to Chlamydomonas taxa. In Chlorella ellipsoidea, an insert that does not have the characteristics of an intron has been reported at the same position as ITS3 of Chlamydomonas (Yamada and Shimaji, 1987). Moreover, Nedelcu et al. (1996) showed that the chloroplast LSU rRNAs in green algal lineages from three green algal classes (sensu Mattox and Stewart, 1984), the Chlorophyceae, Pleurastrophyceae, and Micromonadophyceae, have fragmented chloroplast LSU rRNAs, in most cases the fragmentation pattern being similar to that described in Chlamydomonas. The distinct patterns observed in some lineages are most likely due to the absence or inability to process one of the ITSs. On the other hand, although three ITSs, one of which accounts for the 4.5S rRNA species, have been identified in the maize chloroplast LSU rRNA gene (Kössel et al., 1985), they are situated at different

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positions than in Chlamydomonas. It is interesting that the chloroplast rrnL of Chlamydomonas as well as the mitochondrial rrnL of higher plants are both missing the variable region in which the ITS that accounts for the 4.5S rRNA species is situated (see Turmel, 1993 for references). It was proposed that there is a direct evolutionary connection between variable regions and ITSs, in the sense that variable regions might have in fact evolved from the ITSs separating the rRNA coding modules in the progenote (Gray and Schnare, 1995). Although most of the ITSs in contemporary rRNA genes represent most likely derived rather than primitive traits, the acquisition of the processing sites responsible for the excision of the contemporary ITSs could be considered a ‘reversion to a primitive state’ (Gray and Schnare, 1995). Another unusual gene organization has been reported for the chloroplast RNA polymerase genes (rpo) of C. reinhardtii. The rpoB coding region is divided into two ORFs separated by a 616 bp spacer. Although evidence for the transcription of these ORFs is missing, if expressed, they most likely encode separate polypeptides (Fong and Surzycki, 1992).

VII. Evolution of Mitochondrial and Chloroplast DNA Sequences in Chlamydomonas In land plants, estimated rates of synonymous (silent) nucleotide substitution per site in mitochondrial and chloroplast protein-coding genes are lower than in nuclear genes (reviewed by Palmer, 1991; Bousquet et al., 1992; Laroche et al., 1997). This trend contrasts with the situation in mammals where synonymous substitution rates in mitochondrial genes are higher than in nuclear genes (discussed by Palmer, 1991). No extensive studies on point mutation levels in Chlamydomonas mitochondrial, chloroplast and nuclear genes have yet been published. However, K. J. Prendergast and R. W. Lee (unpublished) have noted that in Chlamydomonas, the number of synonymous substitutions per site, in contrast to land plants, is higher in mitochondrial and chloroplast protein-coding genes than in the one nuclear proteincoding gene (rbcS) for which such a value can be calculated at present. Moreover, in Chlamydomonas, the number of synonymous substitutions per site is higher in mitochondrial than in chloroplast genes, which is opposite to the situation observed in land

86 plants. Further support for such a trend in Chlamydomonas is provided by the mitochondrial and chloroplast LSU rRNA sequences (Turmel et al., 1993; Denovan-Wright et al., 1996). It is interesting that the number of substitutions in Chlamydomonas mitochondrial SSU and LSU rRNA sequences is several-fold higher than the accumulated substitutions in land plant mitochondrial counterparts (Denovan-Wright et al., 1996). Mitochondrial rRNA sequences of P. wickerhamii also seem to have a high rate of nucleotide substitution and, together with the Chlamydomonas, ciliate, fungal and yeast counter­ parts, constitute a rapidly evolving group (associated with long branches in phylogenetic analyses), in marked contrast to the slowly evolving land plant mitochondrial rRNA sequences. Although the number of transitional substitutions is probably saturated in the rapidly evolving mitochondrial rRNA sequences, Denovan-Wright et al. (1996) showed that the apparent affiliation of the Chlamydomonas sequences with ciliate/fungal/yeast counterparts, and therefore their separation from the land plant sequences, is not due to a ‘long-branch length attract’ artifact (i.e., the grouping of rapidly evolving sequences together, in spite of their true phylogenetic relatedness). Surprisingly, Chlamydomonas chloroplast rrnL sequences also display extensive sequence diver­ gence; the various Chlamydomonas lineages studied by Turmel et al. (1993) revealed at least twice the range of sequence variation seen in land plants. Moreover, within some Chlamydomonas lineages (including those leading to C. reinhardtii or C. eugametos) the level is greater than that found between the bryophyte Marchantia and the monocot Oryza (Turmel et al., 1993). Although the chloroplast genomes of the closely related C. pitschmannii and C. eugametos/C. moewusii are extremely similar in gene order they appear to be very divergent in DNA sequence, as deduced from differences in their cpDNA restriction patterns (Boudreau and Turmel, 1995). The ratio of point mutations to length mutations in Chlamydomonas cpDNA is, however, substantially lower than in angiosperm chloroplast DNA, due to the increased level of length mutations in the former (Palmer et al., 1985). The lack of knowledge as to the exact time of divergence of different Chlamydomonas lineages makes it difficult to assess absolute rates of nucleotide substitutions in Chlamydomonas organellar genomes and to compare their tempo of DNA sequence evolution with that of other counterparts. Although fossil evidence suggests that the chlamydo-

Aurora M. Nedelcu and Robert W. Lee monadalean group is at least 350 million years old, the first chlorophycean green algal fossils are around 900 million years old (Tappan, 1980). It seems reasonable to assume, therefore, that the C. reinhardtii/C. eugametos divergence could be as old as 900 million years or as recent as 400 million years. On the other hand, the bryophyte/tracheophyte divergence is believed to have occurred about 400 million years ago (Schopf, 1970; Shear, 1991). If nucleotide substitution levels in organelle DNAs of Chlamydomonas were roughly equal to or up to twice the level observed in their land plant counterparts, comparable rates of nucleotide substitutions in Chlamydomonas and land plants could be hypothesized. In contrast, levels of nucleotide substitution exceeding twice the level noted among land plant counterparts would suggest a higher rate of nucleotide substitution within Chlamydomonas. The observed levels of sequence divergence in organellar rRNA genes in Chlamy­ domonas, i.e., several-fold and at least two-fold higher in mitochondrial and chloroplast rRNA genes, respectively, indicate higher and slightly higher rates of nucleotide substitution in Chlamydomonas mitochondrial and chloroplast rRNA genes, respec­ tively, relative to their land plant counterparts. However, more knowledge about the levels of nucleotide substitution in protein-coding genes from all three genetic compartments from various lineages within the chlamydomonadalean group have to be available before the tempo of DNA sequence evolution in Chlamydomonas organellar genomes can be assessed with confidence. It is not fully understood why the DNA sequences in different genomes of a given lineage or among different groups have different evolutionary rates. Palmer (1990) suggested that error-free replication mechanisms, better postreplication repair systems or copy-correction mechanisms might explain the overall low substitution rates in plant organelle genomes. It is noteworthy that in contrast to land plants, the rDNA nucleotide substitution levels in both Chlamydomonas organellar genomes appealrelatively high. It is possible that this opposite trend could be determined by the same factors proposed for land plant organellar genomes but acting in an opposite direction: error-prone replication mechan­ isms, inefficient postreplication repair systems or copy-correction mechanisms. To explain the observation that mitochondrial and chloroplast genomes in land plants both have low rates of DNA sequence evolution, it was suggested that they might

Chapter 5

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be under common nuclear control (Palmer, 1990). Such a control can also be hypothesized for Chlamydomonas given that both organellar DNA sequences seem to have evolved at high rates.

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study group in attempts to elucidate the evolutionary processes that shape organellar genomes.

Acknowledgments VIII. Conclusions Land plant organellar genomes revealed ‘contrasting modes and tempos of genome evolution’ (Palmer, 1990). Our comparative analysis of evolutionary trends in the mitochondrial and chloroplast genomes of Chlamydomonas, however, allows us to suggest concerted modes and tempos of evolution of these two organellar genomes in this green algal lineage. While land plant mitochondrial genomes are more variable in size and organization but more stable in DNA sequence than their chloroplast counterparts, both organellar genomes in Chlamydomonas seem to have evolved at comparable rates with respect to genome size, organization as well as DNA sequence, as summarized in Table 1. The mechanisms and the selective pressures responsible for both the overall high rate of evolution in Chlamydomonas organellar genomes relative to the land plant counterparts, as well as their apparent concerted evolution are not fully understood (Nedelcu, 1998). The increasing amount of information on the biology, biochemistry, molecular biology, and genetics of these green flagellates (reviewed in this book), as well as the development of new technologies in manipulating their cellular genomes, make Chlamydomonas a good

We thank M. W. Gray for helpful discussion and critical reading of an earlier version of this paper and D. Bhattacharya, E. Boudreau, E. M. DenovanWright, and M. W. Gray for permission to use modified versions of figures previously published. A. M. N. was supported from a Natural Sciences and Engineering Research Council (N.S.E.R.C.) of Canada grant to R. W. L., a Dalhousie University Graduate Scholarship, and an Izaac Walton Killam Memorial Scholarship. Research in the laboratory of R. W. L. is supported by N.S.E.R.C. of Canada.

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Van de Peer Y, Neefs J-M and De Watcher R (1990) Small ribosomal subunit RNA sequences, evolutionary relationships among different life forms, and mitochondrial origins. J Mol Evol 30: 463–476 Waddle JA, Schuster AM, Lee KW and Meints RH (1990) The mitochondrial genome of an exosymbiotic Chlorella-like green alga. Plant Mol Biol 14: 187–195 Wolff G and Kück U (1990) The structural analysis of the mitochondrial S S U r R N A implies a close phylogenetic relationship between mitochondria from plants and from the heterotrophic alga Prototheca wickerhamii. Curr Genet 17: 347–351 Wolff G and Kück U (1993) Organization and coding capacity of mitochondrial genomes of algae. In: Brennicke A and Kück U (eds) Plant Mitochondria with Emphasis on RNA Editing and Cytoplasmic Male Sterility, pp 1 0 1 – 1 1 3 . VCH Verlagsgesellschaft,Weinheim Wolff G, Burger G, Lang BF and Kück U (1993) Mitochondrial genes in the colourless alga Prototheca wickerhamii resemble plant genes in their exons but fungal genes in their introns. Nucleic Acids Res 21: 719–726 Wolff G, Plante I, Lang BF, Kück U and Burger G (1994) Complete sequence of the mitochondrial DNA of the chlorophyte alga Prototheca wickerhamii. Gene content and genome organization. J Mol Biol 237: 75–86 Wolstenholme DR and Fauron CM-R (1995) Mitochondrial genome organization. In: Levings III CS and Vasil IK (eds) The Molecular Biology of Plant Mitochondria, pp 1–59. Kluwer Academic Publishers, Dordrecht Woodson SA and Cech TR (1989) Reverse self-splicing of the Tetrahymena group I intron: Implication for the directionality of splicing and for intron transposition. Cell 57:335–345 Yamada T (1991) Repetitive sequence-mediated rearrangements in Chlorella ellipsoidea chloroplast DNA: Completion of nucleotide sequence of the large inverted repeat. Curr Genet 19: 139–147 Yamada T and Shimaji M (1987) An intron in the 23S rRNA gene of the Chlorella chloroplasts: Complete nucleotide sequence of the 23S rRNA gene. Curr Genet 11:347–352 Zinn AR, Pohlman JK, Perlman PS and Butow RA (1988) In vivo double-strand breaks occur at recombinogenic GC-rich sequences in the yeast mitochondrial genome. Proc Natl Acad Sci USA 85: 2686–2690 Zurawski G, Clegg MT and Brown AHD (1984) The nature of nucleotide sequence divergence between barley and maize chloroplast DNA. Genetics 106: 735–749

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Chapter 6 Uniparental Inheritance of Chloroplast Genomes E. Virginia Armbrust University of Washington, Marine Molecular Biotechnology Laboratory, School of Oceanography, Box 357940, Seattle, WA 98195, U.S.A.

Summary I. Introduction II. Historical Overview of the Uniparental Inheritance of Chloroplast DNA A. Genetic Evidence for the Selective Elimination of Minus Chloroplast DNA in Early Zygotes B. Physical Evidence for the Selective Elimination of Minus Chloroplast DNA in Early Zygotes III. Mating-Type Control of Life Cycle Events A. Gamete Differentiation and Fusion B. Zygote Development C. Chloroplast DNA Inheritance IV. Protection of Plus Chloroplast DNA A. Evidence for a Specific Gene Required for Protection 1. Diploid Crosses 2. The mtl1 Mutation of C. monoica B. Is Plus Chloroplast DNA Protected by Methylation? V. Zygote Specific Elimination of Minus Chloroplast DNA Specific Factor A. Dependence on a UV Sensitive, but not Gametes Enhances Biparental Inheritance 1. Brief UV Irradiation of 2. The Ezy1 Gene 3. The uvsE1 Mutation 4. Involvement of RecA? Parent B. Dependence on the Chloroplast DNA Content of the 1. Treatment with an Inhibitor of Chloroplast DNA Replication, 5-fluorodeoxyuridine 2. The mat3 Mutation C. The Ezy2 Gene D. The sup1 Mutation VI. Regulation of Chloroplast DNA Inheritance A. Regulation of Protection B. Regulation of Destruction VI. Evolution of the Uniparental Inheritance of Organelle Genomes Acknowledgments References.

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Summary An intriguing feature of most eukaryotes is that chloroplast and mitochondrial genomes are inherited almost exclusively from one parent. This can be explained for those organisms that produce gametes of different sizes as chloroplasts and mitochondria are mostly excluded from the sperm or pollen. However, uniparental inheritance also typifies those organisms that produce gametes of identical sizes. In Chlamydomonas, the J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 93–113. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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uniparental inheritance of chloroplast genomes is achieved by a series of mating type-controlled events that culminate in the early zygote with the selective degradation of chloroplast DNA (but not the chloroplast) parent. Thus, only the chloroplast DNA from the mating-type plus contributed by the mating-type minus parent persists in the zygote to be transmitted to meiotic progeny. How Chlamydomonas selectively degrades a subset of organelle genomes has long fascinated researchers and is the subject of this chapter. The molecular mechanisms underlying this phenomenon remain elusive, but are hypothesized to entail two distinct events that occur during different stages of the life cycle: a ‘protection’ of plus chloroplast DNA, perhaps during gametogenesis, and a ‘destruction’ of unprotected minus chloroplast DNA during early zygote development. The gene(s) required for protection have yet to be isolated but all evidence to date indicates that they will be cells only. The degradation of unprotected minus chloroplast DNA appears to be accomplished, present in in part, by a zygote-specific nuclease targeted to the chloroplast during early zygote development. Interestingly, multiple genes have now been identified that all appear to play a role in accomplishing this degradation. As will be shown, the specific elimination of minus chloroplast DNA during early zygote development is a complex and carefully orchestrated phenomenon likely requiring the activity of several proteins. Potential mechanisms for regulating this process are highlighted.

I. Introduction Eukaryotes employ two fundamentally different mechanisms to transfer genetic information from parent to offspring. Nuclear genes are transmitted according to Mendel’s laws. The meiotic spindle ensures that nuclear alleles segregate from one another and genes located on nonhomologous chromosomes sort independently. The existence ofthese laws means that the inheritance patterns of nuclear genes are predictable. Organelle genomes, on the other hand, appear to defy Mendel’s laws and display their own mode of inheritance. In the vast majority of multicellular organisms, the egg and sperm or pollen are dramatically different in size such that organelles and their genomes are typically transmitted to offspring via the egg. Moreover, in mammals at least, any male mitochondria that do enter the zygote appear to be quickly destroyed (Yaneda et al., 1995). Thus most plants and animals inherit organelle genomes from the female parent only, in a process traditionally referred to as maternal inheritance. Conifers, the main exception to this rule, inherit organelle genomes from the male parent and thus display paternal inheritance (Whatley, 1982; Neale et al., 1989). In most unicellular organisms, organelle genomes are also inherited from only one parent even though the two gametes are generally the same Abbreviations: CsCl – cesium chloride; DAPI – 4´,6-diamidinophenylindole; ezy — early zygote; FdUr – 5-flurodeoxyuridine; – mating type minus; – mating type plus; mtl – mating type limited; RFLP – restriction fragment length polymorphism; sup – suppressor of uniparental inheritance

size and contribute equal numbers of organelles to the zygote. In these instances, uniparental inheritance is accomplished by selectively destroying one set of organelles (Braten, 1971, 1973) or organelle genomes (Harris, 1989; Kuroiwa, 1991; Meland et al., 1991) in the developing zygote. Although uniparental inheritance dominates in most eukaryotes, organelle DNA can also be inherited from both parents in a process known as biparental inheritance (Sears, 1980; reviewed in Birky, 1995). Biparental inheritance can occur when the mechan­ isms that typically exclude or destroy organelle genomes are either inefficient (Gyllensten et al., 1991), or else nonexistent (Sears, 1980). Because neither mitochondria nor chloroplasts possess a spindle, when organelle genomes of two types coexist within the zygote, they partition randomly during meiosis such that some progeny inherit the organelle DNA from one parent, some progeny inherit from the other parent, and some progeny inherit from both parents. The biparentally transmitted genomes further segregate during subsequent mitotic divisions such that eventually, the mitotic offspring are pure for the organelle genomes from one parent or the other. This mitotic segregation also results from a stochastic partitioning of genomes to progeny (Birky, 1983). And finally, perhaps the most intriguing version of organelle inheritance, known as doubly uniparental inheritance, is displayed by mussels: female mussels transmit mitochondrial DNA to both male and female offspring while male mussels selectively pass on to their male offspring the mitochondrial DNA they have inherited from their father (Hoeh et al., 1996).

Chapter 6

Chloroplast DNA Inheritance

Non-Mendelian transmission patterns and the mitotic segregation of alleles are now accepted as defining features of organelle inheritance. Beginning at the turn of the century, however, when seemingly aberrant inheritance patterns were first independently described by Correns and Bauer (reviewed in Birky, 1995) and continuing until the mid 1950s and 1960s, the possible existence of genetic information that did not obey Mendel’s laws was somewhat heretical (reviewed in Sager, 1965). DNA within the nucleus could be seen, but there was no notion that organelles could also contain their own genomes. Research with Chlamydomonas reinhardtii was instrumental in transforming organelle inheritance studies from a descriptive science to an experimental science and today C. reinhardtii remains a model organism for the study ofmechanisms underlying chloroplast DNA transmission (Rochaix, 1995). In Chlamydomonas spp., the basis of uniparental inheritance is a selective degradation in the early zygote of the chloroplast DNA (but not the chloroplast) from the mating-type parent, presumably due to the action of minus a zygote-specific nuclease. The chloroplast DNA from the mating-type plus parent is somehow protected from degradation and persists in the zygote to be transmitted to the meiotic progeny. Thus, meiotic progeny inherit chloroplast DNA uniparentally from parent only. the The molecular basis for the selective destruction of minus chloroplast DNA has been the subject of continual study since the phenomenon was first discovered in 1954 (Sager, 1954), but details of the underlying mechanisms remain elusive. What remains even more mysterious, however, is how the uniparental inheritance of organelle genomes evolved in the first place. A number of excellent reviews of chloroplast DNA inheritance have been published in recent years by Harris (1989), Gillham et al. (1991), Kuroiwa (1991), Gillham (1994), and Sears and VanWinkleSwift (1994). This chapter will focus on possible molecular mechanisms underlying the uniparental inheritance of chloroplast genomes. Unless otherwise stated, the work described involves C. reinhardtii since this has traditionally been the study organism of choice. However, it should be noted that all species of Chlamydomonas appear to destroy selectively the chloroplast DNA from one parent in the early zygote (McBride and McBride, 1975; Coleman and Maguire, 1983; VanWinkle-Swift and Aubert, 1983).

95 II. Historical Overview of the Uniparental Inheritance of Chloroplast DNA

A. Genetic Evidence for the Selective Elimination of Minus Chloroplast DNA in Early Zygotes The field of chloroplast genetics began in 1954 when Ruth Sager described the inheritance patterns of two UV-induced mutations, sr1 and sr2 (reviewed in Boynton et al., 1992). The sr1 mutation confers resistance to low levels ofthe antibiotic streptomycin and the sr2 mutation confers resistance to high levels of streptomycin (Sager, 1954; Table 1). Regardless or the parent, ofwhether sr1 is carried by the when a resistant strain is crossed to a sensitive strain, two of the meiotic progeny are resistant to low levels of streptomycin and two of the progeny are sensitive, just as predicted for a nuclear mutation inherited in a parent Mendelian fashion. In contrast, when a carrying the sr2 mutation is crossed to a sensitive parent, all the meiotic progeny are resistant to high levels of streptomycin; only rarely is a sensitive progeny obtained. When the reciprocal cross is conducted, and a resistant parent is crossed to a sensitive parent, the meiotic progeny are all streptomycin sensitive (Fig. 1). Sager (1954) performed a series of backcrosses with each type of sr2, sensitive, F1 progeny (– sr2, sensitive) to prove that these seemingly aberrant inheritance patterns were stable and that the resistance phenotype conferred by the sr2 mutation was transmitted to meiotic progeny only when carried by parent. It is now known that the sr2 mutation the is localized within the rps12 gene of the chloroplast genome (Liu et al., 1989) and that these inheritance patterns reflect the uniparental inheritance of parent. Those rare chloroplast DNA from the zygotes that transmit minus chloroplast DNA to meiotic progeny are referred to as exceptional since they represent the exception to the rule. Sager (1954) briefly raised the possibility that she might have discovered a Chlamydomonas version of maternal inheritance, but she ended this landmark paper by lamenting that ‘the nature of the physical carrier of this inheritance as well as the mechanism of transmission remains obscure.’ Over the next 10–15 years, additional uniparentally inherited mutations were isolated, all of which appeared genetically linked (Sager and Ramanis, 1963; Gillham, 1965; Sager and Ramanis, 1965; Gillham and Fifer, 1968), suggesting that the ‘physical

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carrier’ must be DNA (for discussion, see Gillham, 1969). During this early period, further evidence for the existence of chloroplast DNA came from work by Ris and Plaut (1962). Using a combination of electron and light microscopy, they described regions in the chloroplast stroma that stained with Feulgen and acridine orange, both indicative of the presence of DNA. Moreover, these DNA-staining regions, or nucleoids, contained 25Å microfibrils that were

E. Virginia Armbrust

eliminated by incubation with DNase. Just one year later, Chun et al. (1963) and Sager and Ishida (1963) described the isolation ofchloroplast DNA by cesium chloride(CsCl) densitygradient centrifugation.Thus, it seemed increasingly likely that chloroplasts con­ tained DNA and that it was this chloroplast DNA that was uniparentally transmitted to meiotic progeny.

Chapter 6

Chloroplast DNA Inheritance

B. Physical Evidence for the Selective Elimination of Minus Chloroplast DNA in Early Zygotes By the early 1970s it had been clearly documented that the two chloroplasts (one from each parent) fuse in their entirety in the early zygote (Cavalier-Smith, 1970) and yet mutations presumably located on the minus chloroplast genome were not transmitted to meiotic progeny. What happened to this minus chloroplast DNA in the zygote? To address this question, Sager and Lane (1972) labeled the DNA and gametes with either from and used CsCl density gradient centrifugation to monitor the fate of nuclear and chloroplast DNA in 6 h and 24 h zygotes. Six hours into zygote develop­ ment, the nuclear DNA contributed by both parents could be recovered. In contrast, the chloroplast DNA parent had disappeared from this same from the timepoint and the chloroplast DNA from the

97 parent had undergone a shift in density (Sager and Lane, 1972). Here, at last, was a physical explanation for the genetic results: the chloroplast DNA from the parent could not be transmitted to meiotic progeny because this DNA was somehow eliminated during early zygote development. But why was the gamete left chloroplast DNA contributed by the intact? Sager and Lane (1972) postulated that the observed density shift of the plus chloroplast DNA specific methylation event that was the result of a somehow protected this DNA from elimination in the zygote, a possibility discussed in more detail in a later section. Conclusive proof that minus chloroplast DNA was eliminated from zygotes came in the early 80s from two new sources—molecular biology and fluorescent microscopy. The first molecular evidence was provided when Grant et al. (1980) discovered that a C. reinhardtii mutant strain, ac-u-g-23 (Shepherd et al., 1979), carried two small deletions in its chloroplast DNA. The presence or absence ofthe deletions could be easily monitored by an analysis of chloroplast DNA restriction fragment length polymorphisms (RFLPs). Grant et al. (1980) found that both the deletions in the chloroplast DNA and the independent nonphotosynthetic phenotype were uniparentally inherited. A few years later, Dron et al. (1983) determined that the mutation, rcl-u-1-10-6C, which is in the gene encoding the large subunit of ribulose bisphosphate carboxylase (Spreitzer and Mets, 1980), was localized to chloroplast DNA. Mets and Geist (1983) showed that this gene was uniparentally inherited. The next year, Lemieux et al. (1984) found that RFLPs between the chloroplast genomes of the two interfertile species, C. eugametos and C. moe­ wusii, cosegregated with uniparentally inherited antibiotic resistance markers. Thus, both photo­ synthetic genes and antibiotic resistance mutations were physically localized to the chloroplast genome and it was this DNA that was uniparentally inherited. Beautifully convincing fluorescent microscopy data for the selective elimination of minus chloroplast DNA in the zygote was published during this same period by Kuroiwa et al. (1982). Chloroplast DNA in Chlamydomonas is localized into 8–10 nucleoids, easily distinguished from the much larger cell nucleus when stained with the DNA fluorochrome, 4, 6diamindio-2-phenylindole or DAPI (Fig. 2). Within about 40 min of zygote formation, each chloroplast parent begins to nucleoid contributed by the ‘dissolve’ from its periphery until the DAPI

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E. Virginia Armbrust III. Mating-Type Control of Life Cycle Events

fluorescence disappears completely while the parent remain chloroplast nucleoids from the intact. The two nuclei fuse within the next hour, followed 1–2 h later by the fusion of the two chloroplasts. By 10 h after zygote formation, the nucleoids have coalesced and migrated to form a single large nucleoid near the pyrenoid (Kuroiwa et al., 1982). The chloroplast DNA content ofthe zygote was shown later to remain roughly constant throughout zygote maturation (Coleman, 1984). A burst of chloroplast DNA synthesis occurs prior to meiosis so that each newly formed progeny will possess a full complement of chloroplast DNA (Coleman, 1984). Uchida et al. (1992b) have recently extended this visual analysis of minus chloroplast DNA degradation. By using electron microscopy to determine the distribution of immunogold labeled DNA-specific antibodies, they confirmed that the parent is completely chloroplast DNA from the eliminated and not simply dispersed during early zygote development. This then is the stepping-off point for this chapter. Yes, the chloroplast genome from the parent is specifically eliminated during early zygote develop­ ment, but how is this accomplished? What is known about underlying molecular mechanisms and how are these processes controlled? Many potential players in chldroplast inheritance have been identified, and as will be seen, an understanding of the roles played by these proteins is just emerging.

The simplest model for the uniparental inheritance of chloroplast DNA is that the process entails two distinct events that are likely restricted to different stages of the life cycle: a ‘protection’ of plus chloroplast DNA perhaps during gametogenesis and a degradation (‘destruction’) of unprotected minus chloroplast DNA during early zygote development (reviewed by Gillham et al., 1991; Gillham, 1994; Sears and VanWinkle-Swift, 1994). The genes required for the protection of plus chloroplast DNA and the destruction of minus chloroplast DNA are thought to be controlled by the mating-type locus, described in detail below. In fact, three complex lifecycle events — gamete differentiation, zygote maturation and organelle inheritance — are all governed by the mating-type locus, perhaps reflecting early steps in the evolution of sexual reproduction (reviewed by Goodenough et al., 1995). To understand the regulation of processes occurring during the uniparental inheritance of chloroplast DNA, the life cycle and its control by the mating-type locus will first be examined. Under nutrient-replete conditions, haploid vegetative cells reproduce mitotically and are unable to mate. In response to nitrogen starvation (Sager and Granick, 1954) and a blue light signal (reviewed by Beck and Haring, 1996), vegetative cells or differentiate into mating-competent gametes. When gametes of the opposite mating type are mixed, they rapidly fuse to form diploid zygotes. After an obligate period of maturation, zygotes undergo meiosis and germination to form four haploid progeny that can once again reproduce mitotically. Thus, those genes required for gamete differentiation are expressed in response to nitrogen starvation and those required for zygote maturation are expressed in response to gamete fusion (Goodenough and Ferris, 1987). Life-cycle events are governed by genes of two general types: ‘sex­ linked’ genes are physically located at only one mating-type locus and ‘sex-limited’ genes are unlinked to the mating-type locus but are nonetheless expressed in cells of only one mating type (reviewed in Galloway and Goodenough, 1985; Goodenough and Ferris, 1987). As will be seen, both sex-linked and sex-limited genes are likely involved in the uniparental inheritance of chloroplast DNA. The mating-type locus in C. reinhardtii is located

Chapter 6 Chloroplast DNA Inheritance on the left arm of linkage group VI (Ebersold et al., 1962) and in genetic crosses displays Mendelian and segregation of the two apparent alleles, (Smith and Regnery, 1950). In fact, however, the mating-type locus is a complex region of suppressed recombination that spans nearly 850 kb of DNA, almost 1% of the entire genome (Gillham, 1969; Ferris and Goodenough, 1994). Twelve genes, five required for life-cycle events and seven ‘house­ keeping’ genes, have to date been localized to the mating-type locus (Ferris and Goodenough, 1994); the physical locations of three housekeeping genes, Nic7, Thi10, and Ac29, have recently been described (Ferris, 1995). Our understanding ofthe mating-type control of life-cycle events has advanced rapidly in the past few years due primarily to the work of Ferris and Goodenough. In 1994, they published the results of a one megabase chromosome walk through the entire region of suppressed recombination (Ferris and Goodenough, 1994). They found that this region and cells except for a is homologous in central domain of about 190 kb, designated the R domain, in which chromosomal rearrangements have occurred and segments unique to one or the other mating type are located (Fig. 3). Genes required for mating type-controlled life-cycle events are hypothesized to reside within these unique segments. Presumably, the extensive rearrangements that have occurred within the R domain explain the lack of recombination around mating-type, an arrangement that may ensure that genes required for sexual differentiation are inherited as a unit (Ferris and Goodenough, 1994).

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A. Gamete Differentiation and Fusion The recent cloning oftwo mating type-specific genes, Fus1 and Mid, provides insight into the regulation of sexual differentiation and gamete fusion. Surprisingly, the differentiation circuitry appears to be even simpler than imagined originally (Goodenough and Ferris, 1987). Apparently, only the two sex-linked genes, Fus 1 and Mid, are necessary to determine whether a or a gamete, although a cell will mate as a third gene defined by the iso1 mutation and unlinked to mating type (Campbell et al., 1995), may be required for proper Mid function (Goodenough et al., 1995). The Fus 1 gene is found exclusively within locus (Fig. 3) and is the R domain at the hypothesized to encode a structural protein localized to the plus mating structure and necessary for gamete fusion (Ferris et al., 1996). If a cell possesses a defective Fus 1 gene, as occurs in imp1 strains, the gamete will recognize and agglutinate with imp1 a partner, but will fail to undergo cell fusion (Goodenough et al., 1976; Goodenough et al., 1982; Ferris et al., 1996). The Mid gene is found exclusively locus (Fig. 3) and is in the R domain at the hypothesized to encode a transcription factor that both turns on minus sexual differentiation and turns off plus sexual differentiation (Galloway and Goodenough, 1985; Ferris and Goodenough, 1997). cell possesses a defective Mid gene, as in the If a differentiation will not be turned imp 11 mutant, on and the cell will instead differentiate into a plus gamete. This ‘pseudo-plus’ gamete can do everything that a true gamete can do except it cannot fuse partner (Goodenough et al., 1982; Ferris with a

100 and Goodenough, 1997). It is now clear that the reason the imp11 cell can not fuse is that it lacks the necessary Fus1 gene found at the locus (Ferris et al., 1996). Remarkably, if the wild-type Fus1 gene is transformed into an imp 11 strain, this locus will differentiate into a func­ cell with a gamete to tional plus gamete that can fuse with a form viable ‘minus/minus’ zygotes (Fus1 imp11 able to undergo meiosis (Ferris et al., 1996). cell is transformed with a wildReciprocally, if a type Mid gene, this cell will differentiate into a fully functional minus gamete that can mate with a gamete to create ‘plus/plus’ zygotes (Mid that are also able to undergo meiosis (Ferris and Goodenough, 1997). As will be discussed in more detail later, this ability to change the mating type of any given cell and create sexual zygotes homozygous for mating type has revitalized our ability to examine the function of genes located exclusively within one or the other mating-type.

B. Zygote Development Less is know about the control of zygote-specific gene expression in Chlamydomonas. Fifteen genes that are expressed specifically during zygote differentiation have now been identified. Presumably, a cascade of expression is initiated by zygote formation since nine of the genes are transcribed almost immediately upon gamete fusion, three are transcribed about 1–3 h later (Ferris and Goodenough, 1987; Woessner and Goodenough, 1989; Matters and Goodenough, 1992; Uchida et al., 1993), and three are transcribed during late zygote maturation (Wegener and Beck, 1991). The almost immediate transcription of the early zygote genes occurs long before the two nuclei fuse and is independent ofnew protein synthesis (Ferris and Goodenough, 1987). Originally, it was hypothesized that zygote-specific gene expression was dependent upon an interaction in the zygote of mating type-specific regulatory gamete-specific, and M, proteins known as P, for for gamete-specific (Goodenough and Ferris, 1987). This hypothesis must be now modified to explain the zygote-specific expression that occurs in minus/minus zygotes. A P protein encoded exclu­ sively at the locus cannot exist in the minus/ locus. minus zygotes since they do not posses a parent in this cross Since the imp11 Fus1 differentiates as a plus gamete, however, the data are consistent with the existence ofa sex-limited gene P

E. Virginia Armbrust which is turned on during plus gamete differentiation (Goodenough et al., 1995).

C. Chloroplast DNA Inheritance The simplest models for the mating-type control of uniparental inheritance hypothesize that nuclear genes locus are required for located exclusively at the both the protection of plus chloroplast DNA and the degradation of minus chloroplast DNA (Gillham et al., 1991; Gillham, 1994). Until quite recently, all the data supported this simple hypothesis. As will be described in detail in the following sections, it now appears that genes located outside of the matingand the loci type locus as well as at both the are involved in controlling the uniparental inheritance of chloroplast DNA. Two points are clear, however, from the early studies. First, all the genes required for the selective elimination of minus chloroplast DNA are nucleusencoded; inhibitors of either the transcription of nuclear genes or the translation of nuclear gene products prevent destruction while comparable chloroplast-specific inhibitors have no affect (Kuroiwa et al., 1983a,b). Second, the elimination of minus chloroplast DNA occurs only in true zygotes, not in diploids generated by other means. For example, a low percentage (~5%) of sexually generated diploid cells fail to differentiate into zygotes and rather than undergo meiosis, they instead divide mitotically as stable vegetative diploids (Ebersold, 1967). Most (50–90%) of these vegetative diploids initially possess chloroplast DNA from both parents (Gillham, 1963). As the diploids divide mitotically, the two parental chloroplast genomes segregate to produce progeny that are eventually homoplasmic for one chloroplast genome or the other (Gillham, 1963; reviewed in Sears and VanWinkle-Swift, 1994). Vegetative diploids can also be generated artificially by incubating cells in the presence of polyethylene glycol (PEG) (Matagne et al., 1979). In these vegetative diploids, about one third of the fusion products are initially biparental, one third possess the chloroplast genome from one parent and one third possess the chloroplast genome from the other parent regardless cells, two of whether the fusion is between two and cell (Matagne and cells or a Hermesse, 1980; Matagne, 1981). Matagne (1980) suggests that the reduced frequency of biparental inheritance observed in the PEG fused diploids may occur because the first mitotic division is delayed in

Chapter 6

Chloroplast DNA Inheritance

these cells. Van Winkle- Swift (1978) had shown earlier that a delay in the timing of this first division somehow decreases the subsequent number of diploids that are initially biparental. Thus, a segregation of chloroplast alleles is observed in both types of vegetative diploids, but the directed elimination of the minus chloroplast genome occurs only in sexual zygotes.

IV. Protection of Plus Chloroplast DNA The degradation of minus chloroplast DNA in early zygotes is due presumably, at least in part, to the action of a nuclease (Kuroiwa et al., 1983a). The two most obvious ways that minus chloroplast DNA alone could be eliminated from the zygote are to specifically ‘mark’ the minus chloroplast DNA for elimination or else to specifically protect the plus chloroplast DNA from elimination. As will be shown, the most straightforward interpretation of the evidence is that the plus chloroplast DNA is specifically protected prior to zygote formation. But what exactly does it mean to be protected? It is still not known whether protection entails a covalent modification of plus chloroplast DNA that somehow protects the DNA from degradation (e.g., Sager and Ramanis, 1973; Sager and Kitchin, 1975) or whether protection instead entails the presence ofprotector proteins that somehow prevent the nuclease from gaining access to plus chloroplast DNA (e.g., Gillham et al., 1974; Kuroiwa, 1991). Regardless ofthe exact mechanisms underlying protection, the process must be reversible. Under conditions of strict uniparental inheritance, all the chloroplast DNA transmitted to meiotic progeny was once protected from elimination. At some point during the life cycle this protection must be erased so that the chloroplast DNA in the progeny will once again be susceptible to degradation during the next round of zygote formation.

A. Evidence for a

Specific Gene Required

for Protection

1. Diploid Crosses The earliest evidence that a specific gene is involved in protection came from crosses with a diploid gamete heterozygous for mating type. When heterozygous vegetative diploid is nitrogena gamete starved, the resulting gamete mates as a and thus is phenotypically minus (Ebersold, 1967).

101 In other words, is dominant to for sexual differentiation, a phenomenon known as minus dominance. As described in an earlier section, minus dominance is conferred by the Mid gene (Ferris and Goodenough, 1997). Over thirty years ago, Gillham (1963) showed that when a heterozygous gamete, the diploid is crossed to a haploid chloroplast genomes from both parents are bipar­ entally transmitted to meiotic progeny. Gillham must therefore be (1963) hypothesized that dominant to for uniparental inheritance. Matagne and Mathieu (1983) extended this analysis by using the PEG-fusion technique to create diploid gametes that were either homozygous or heterozygous for mating type. Uniparental inheritance of the plus chloroplast genome was observed in all crosses in which the minus parent was either haploid or homozygous for mating type: and In contrast, biparental inheritance was observed in all crosses in which the minus parent was heterozygous for mating type: In other words, if a phenotypically minus parent carries the locus, chloroplast genomes from both parents will persist in the zygote (Tsubo and Matsuda, 1984) and will be transmitted to the meiotic progeny. These data suggest locus carries a gene required for protection that the that is not repressed by Mid. Moreover, these results imply that protection occurs prior to zygote formation since the triploid zygotes resulting from the crosses and have identical genotypes but very different chloroplast transmission patterns (Matagne and Mathieu, 1983).

2. The mtl1 Mutation of C. monoica Thus far, the focus of this chapter has been on the transmission of chloroplast DNA by C. reinhardtii. In recent years, important information about chloroplast DNA inheritance has come from work with C. monoica conducted by Van Winkle-Swift and colleagues. Unlike C. reinhardtii, C. monoica is and cells homothallic which means that both are produced in a clonal population. Thus C. monoica can ‘self-mate’ (see for example, van den Ende and VanWinkle-Swift, 1994). A breakthrough in the genetic manipulation of C. monoica came when Van Winkle-Swift and Bauer (1982) and Van WinkleSwift and Burrascano (1983) generated self-sterile and zygote maturation-defective mutants. Although self-sterile, these strains can be outcrossed and thus

102 have greatly facilitated the analysis of interclonal crosses. It should be kept in mind, however, that even when an outcross is performed with C. monoica, the and gametes two parent strains form both and thus each mating includes reciprocal crosses. Once self-sterile mutants were available, an analysis of chloroplast genetics became feasible. When a C. monoica strain carrying a chloroplast mutation that confers resistance to erythromycin, ery-u-1, is outcrossed to an erythromycin-sensitive strain, two types of tetrads are obtained: those in which all four meiotic progeny are resistant to erythromycin and those in which all four meiotic progeny are sensitive to erythromycin (VanWinkle-Swift and Aubert, 1983). This version of chloroplast gene transmission has been termed bidirectional uniparental inheritance and is hypothesized to result from the fact that half gametes and half the the ery-u-1 cells mate as gametes. When the ery-u-1 ery-u-1 cells mate as parent, the resistancemutation is carried by the conferring mutation is transmitted to all the meiotic progeny, but when the ery-u-1 mutation is carried by parent, the chloroplast DNA from the sensitive the strain is transmitted to all the meiotic progeny (VanWinkle-Swift and Aubert, 1983). To identify mating type-specific genes required for chloroplast genome transmission, VanWinkleSwift and Hahn (1986) screened for mutants that create a shift from a bidirectional to a unidirectional transmission of the ery-u-1 mutation. The mtl1 mutation (for mating type limited) created just such a phenotype. If an mtl1 strain resistant to erythromycin is outcrossed to an erythromycin-sensitive strain, only erythromycin-sensitive progeny are obtained, mtl1 ery-u-1 parent is no longer able to as if the transmit chloroplast DNA. Moreover, 50% of the zygotes resulting from this outcross do not germinate. If instead, an mtl1 mutant is allowed to self-fertilize, none of the resulting zygotes germinate (VanWinkleSwift and Hahn, 1986). A careful analysis of the behavior of this mutant in both self-matings and outcrosses indicated that the mtl1 phenotype is parent. The expressed only when carried by the zygote-lethal phenotype was traced to the fact that parent carries the mtl1 mutation, all the when the chloroplast DNA in the resulting zygote is eliminated, leading VanWinkle-Swift and Salinger (1988) to hypothesize that the mtl1 mutant is unable to protect plus chloroplast DNA. Importantly, this degradation of all chloroplast DNA occurs relatively synchronously and long before the two chloroplasts

E. Virginia Armbrust fuse, indicating that the responsible nuclease is simultaneously present in both chloroplasts. Thus, a distantly related cousin of C. reinhardtii also appears to utilize a gene controlled by the locus to somehow protect plus chloroplast DNA from degradation in the zygote. No mtl1-like mutations have yet been identified in C. reinhardtii.

B. Is Plus Chloroplast DNA Protected by Methylation? This simple question reflects one of the more controversial topics in the field of Chlamydomonas chloroplast DNA inheritance. This topic has been reviewedrecently (Harris, 1989; Gillham et al., 1991; Kuroiwa, 1991; Gillham, 1994) and will only be summarized here. When Sager and Lane (1972) first specific methylation event postulated that a protected plus chloroplast DNA from destruction in the zygote, the restriction-modification system of bacteria had only recently been described. Restrictionmodification was an excitingnew means of ‘silencing’ DNA that might also explain chloroplast DNA inheritance in C. reinhardtii (e.g., Sager and Kitchin, 1975). Sager and colleagues rapidly accumulated convincing evidence that the chloroplast DNA of gametes contains a high percentage of 5-methyl gametes (Burton et al., 1979; cytosine relative to Royer and Sager, 1979; Sano et al., 1980), perhaps due to the action of a gamete-specific methyl transferase (Sano et al., 1 9 8 1 ) . Moreover, this methylation event was apparently reversible, as would be expected for protection (Sano et al., 1984). The problem was how to lest the effect of such a global event. Initially, Sager described a mutation mat1 that locus; the was believed to be tightly linked to the chloroplast DNA of the mat1 strain was methylated and the presence of the mutation resulted in a biparental transmission of chloroplast genomes (Sager and Ramanis, 1974; Sager et al., 1981). This ‘methylation mutant’ was later shown to be a diploid cell that had been generated inadvertently during a genetic cross (Sager and Grabowy, 1985). These data nonetheless suggest that in diploids, the chloroplast DNA is methylated and protected. A series of papers from Gillham and colleagues subsequently argued that methylation of plus chloroplast DNA could not explain protection adequately. Bolen et al. (1982) isolated a nuclear mutant me1 that constitutively methylales chloroplast and vegetative DNA to high levels in both

Chapter 6

Chloroplast DNA Inheritance

c e l l s . When e i t h e r or gametes w i t h hypermethylated chloroplast DNA were used for crosses, normal uniparental inheritance patterns were observed, prompting Bolen et al. (1982) to hypothesize that the methylation of chloroplast DNA described by Sager and colleagues could not adequately explain protection. Sager and Grabowy ( 1 9 8 3 ) countered this conclusion w i t h t h e i r observation that additional methylation occurred , but not cells during gametogenesis of carrying the me1 mutation. One year later, Feng and Chiang (1984) presented evidence that treatment of cells with the methylation inhibitors, L-Ethionine and 5-azacytidine, resulted in a hypomethylation of gamete chloroplast DNA. The hypomethylation of chloroplast DNA from either parent had no effect on the subsequent uniparental inheritance of plus chloroplast DNA, prompting Feng and Chiang (1984) to also conclude that protection was not conferred by methylation. In recent years, only one other paper has addressed the possible connection between methylation and protection. Kuroiwa and colleagues (reviewed in Kuroiwa, 1991) have used fluorescent microscopy to document that the elimination of minus nucleoids occurs rapidly and apparently in the absence of any nucleoid swelling, regardless of nucleoid size (Ikehara et al., 1996). Kuroiwa (1991) argues that this absence of swelling indicates that a nuclease similar to DNase I that can rapidly degrade DNA, rather than a restriction enzyme, eliminates the minus chloroplast DNA from early zygotes. The evidence accumulated over the years, then, argues against a global methylation event as a means of conferring protection to plus chloroplast DNA. However, it should be kept in mind that none of these studies eliminates the possibility that protection is conferred by a specific methylation event unaffected by either the me1 mutation or the methylation inhibitors.

V. Zygote Specific Elimination of Minus Chloroplast DNA The simplest explanation for the selective elimination of minus chloroplast DNA is that a nuclease is either activated (Matagne, 1987; Kuroiwa, 1991) or else expressed specifically during early zygote develop­ ment and subsequently degrades unprotected minus chloroplast DNA (Gillham et al., 1991; Gillham, 1994). Over the years, a number of endo- and exo­ nucleases, have been isolated from whole-cell extracts

103 of C. reinhardtii vegetative cells (Burton et al., 1977; Tait and Harris, 1977a,b; Frost and Small, 1984). Presumably, however, these enzymes play a role in DNA repair since each was isolated from vegetative cells (Small, 1987). In the mid 1980s, Kuroiwa and colleagues set out to identify nucleases specifically required for the uniparental inheritance of chloroplast DNA. Rather than use whole-cell extracts from vegetative cells as had been done before, Ogawa and Kuroiwa (1985b) began with 3 h zygotes. They -dependent nucleases that identified a class of displayed both endo- and exo-nucleolytic activity. The extracts were composed ofsix small polypeptides, and this preparation was collectively referred to as nuclease C (Ogawa and Kuroiwa, 1985a,b, 1986a,b, 1987). It had previously been shown (reviewed in Kuroiwa, 1985) that incubation of zygotes with the calcium chelator, EGTA, inhibited the specific elimination of minus chloroplast DNA. Thus, Ogawa and Kuroiwa (1985b) were hopeful initially that they had identified the critical factor. Unfortunately, nuclease C activity was found to be present throughout and the entire life cycle — in zygotes, vegetative cells, and and gametes. This finding prompted Kurowia (1991) to conclude that nuclease C is not specific to chloroplast DNA inheritance. Thus, the identification of a nucleus-encoded enzyme with a detectable nuclease activity that is directed to the chloroplast during early zygote formation still remains elusive. There are, however, two general ways to prevent zygote-specific elimination of minus chloroplast DNA in the gamete prior to zygote —UV irradiation of the mating and a reduction of the chloroplast DNA gamete. As more data are content of the accumulated through experiments involving the use of these two ‘inhibitors’, it appears increasingly likely that multiple gene products, and not just a single nuclease, are involved in the specific elimination of minus chloroplast DNA during early zygote development.

A. Dependence on a UV Sensitive, Specific Factor 1. Brief UV Irradiation of but not Gametes Enhances Biparental Inheritance Sager and Ramanis (1967) first reported and others have since verified (e.g., Gillham, 1969; Nakamura et al., 1988; Uchida et al., 1992a; Armbrust et al.,

104

1993) that brief UV-irradiation of gametes just prior to mating prevents the selective elimination of minus chloroplast DNA in the zygote and thus enhances the frequency of biparental inheritance. gametes prior to mating Comparable treatment of has no effect on the elimination of minus chloroplast DNA in the zygotes; as in wild-type crosses, the resulting meiotic progeny inherit chloroplast DNA parent only (Fig. 4). Furthermore, the from the number of zygotes that transmit minus chloroplast DNA to meiotic progeny is dependent on UV dose; gamete is exposed to UV, the the longer the greater the number of exceptional zygotes that are recovered. The UV effect is also photoreactivable; if gametes are allowed to recover in the irradiated

E. Virginia Armbrust

light prior to mating, the frequency of exceptional zygotes decreases, suggesting that at least one target of UV irradiation is DNA. Sager and Ramanis (1967) hypothesized that UV-irradiation blocked the specific gene product that is synthesis of a necessary for the elimination of minus chloroplast DNA in the zygote. Some twenty years later, Nakamura et al. (1988) presented evidence that the expression of at least six zygote polypeptides is inhibited when the , but not the parent is exposed to brief UV-irradiation just prior to mating. While it was not shown whether any of these UVsensitive gene products are directly involved in minus chloroplast DNA elimination, this work was the first gametes can clear evidence that UV treatment of

Chapter 6 Chloroplast DNA Inheritance inhibit specifically the expression of a subset of genes in the zygote.

2. The Ezy1 Gene The Ezy1 gene (for early zygote) was found in a differential screen designed to identify zygotespecific messages (Ferris and Goodenough, 1987). Ezy1 is tandemly repeated 7–8 times at both the loci (Ferris and Goodenough, 1987), and and the at least three copies of the gene are transcribed within minutes of zygote formation (Armbrust et al., 1993). The negatively charged Ezy1 protein localizes specifically to chloroplast nucleoids during the interval of minus chloroplast DNA degradation. Both uniparental inheritance of chloroplast DNA and transcription of Ezy1 are differentially inhibited by brief UV irradiation of gametes just prior to gametes prior mating; comparable treatment of to mating has no effect on either Ezy1 transcription or inheritance patterns. Importantly, transcription of a zygote-specific wall gene (Woessner and Goode­ nough, 1989) does not display differential sensitivity gametes. Thus, when Ezy1 to UV irradiation of is not expressed, minus chloroplast DNA is not destroyed. The Ezy1 gene has therefore been hypothesized to be involved in the selective elimination of minus chloroplast DNA (Armbrust et al., 1993). Detailed biochemical work with Ezy1 is just beginning so the role played by this protein is not yet known. gamete How could UV irradiation of the possibly affect transcription in the zygote? A common consequence of UV-irradiation is the generation of pyrimidine dimers in DNA; the presence ofunrepaired dimers can completely inhibit expression of a transciptional unit (reviewed in Britt, 1996). One possible explanation, discussed in Armbrust et al. (1993), is that UV exposure damages the irradiated nuclear DNA which in turn prevents subsequent transcription of Ezy1 in the zygote. This explanation copies of Ezy1 are never assumes that the transcribed. Preliminary data now indicates, however, and copies of Ezy1 are expressed that both the (Armbrust, unpublished). The unirradiated copies of Ezy1 should, therefore, be unharmed and available for transcription in the zygote. This result suggests that UV irradiation may affect non-DNA targets that are also necessary for Ezy1 transcription. Polypeptide 4, originally identified by Nakamura et al. (1988) as a zygote polypeptide whose expression is sensitive

105 parent, likely corresponds to UV irradiation of the to Ezy1. Thus, two different groups have indepen­ dently accumulated data that suggest that transcription of Ezy1 and perhaps additional genes may be specific UV-sensitive factor, a dependent on a dependence that may ensure that destruction ofminus chloroplast DNA is prevented when the plus chloroplast DNA is potentially UV-damaged. If this hypothesis is true, then it should be possible specific mutants that are unable to to isolate prevent transcription of zygote-specific genes when UV-irradiated prior to mating. The zygotes resulting mutant from the mating of such a UV-irradiated parent would be predicted to and a wild-type eliminate minus chloroplast DNA and thus would transmit the potentially damaged plus chloroplast DNA to meiotic progeny. In 1974, Sager and Ramanis described the isolation of a mutant with just such a phenotype. The mat2 mutation, which unfortunately has since been lost, was reportedly linked to the locus. When a parent carrying the mat2 mutation was exposed to relatively high levels ofUV irradiation prior to mating, the plus chloroplast genome was still inherited uniparentally (Sager and Ramanis, 1974), implying that destruction of the minus chloroplast genome had not been prevented in the mat2 zygote. cells and wild-type cells Interestingly, mat2 displayed identical UV survival curves, suggesting that the Mat2 gene was not required for DNA repair. Thus, the mat2 mutation may have identified a gene necessary for inactivating transcription of a subset of genes in the zygote when the plus parent is UVirradiated and its chloroplast DNA potentially damaged.

3. The uvsE1 Mutation The uvsE1 mutant was isolated by Rosen and Ebersold (1972) as a strain that is particularly sensitive to UV irradiation, perhaps due to a defect in either excision or recombination-mediated repair. The uvsE1 mutation is unlinked to mating type. Zygotes homozygous for uvsE1 display a reduced frequency of recombination of nuclear genes during meiosis (Rosen and Ebersold, 1972). Rosen et al. (1991) later examined whether the uvsE1 mutation also decreased the frequency of recombination of chloroplast genes as might be expected for a recombination defective mutant. Chloroplast DNA recombination was unaffected by uvsE1, but Rosen et al. (1991) found and the unexpectedly that when both the

E. Virginia Armbrust

106 parent carried uvsE1, as many as 20% of the zygotes displayed biparental inheritance, a dramatic increase from the negligible levels displayed by control crosses. It is provocative that a mutation in a gene that leads to UV-sensitivity, presumably due to a defect in an enzyme required for excision repair of nuclear genes, gamete prior to mating and UV irradiation of the both result in an increased number of exceptional zygotes. The connection between these two phenomena remains unclear, though. Obviously, however, the simple model of a single nuclease acting alone to accomplish the selective elimination of minus chloroplast DNA needs to be modified.

4. Involvement of RecA? In E. coli, the RecA protein is essential for homologous recombination and for a variety of responses to DNA damage. It now appears that a RecA-mediated recombination process in C. reinhardtii chloroplasts may be involved in the repair of chloroplast DNA when damaged by photooxidation, UV irradiation, or other environmental stresses (Cerutti et al., 1995). If a dominant negative mutant form of the E. coli RecA protein is expressed in C. reinhardtii chloroplasts, the transformed cells display reduced survival rates when exposed to a number of DNA damaging agents, and reduced repair and recombination of chloroplast DNA. The most likely explanation for this result is that Chlamydomonas chloroplasts contain a RecA homolog whose activity is inhibited by the dominant negative mutant form of the E. coli RecA protein (Cerutti et al., 1995). Thirty years ago, Sager and Ramanis (1967) hypothesized and Gillham et al. (1987) reiterated gametes prior to that the UV irradiation of mating might activate a RecA-mediated response that included inactivation of a factor necessary for the specific elimination of minus chloroplast DNA in the zygote. Whether this inactivation occurred in the chloroplast or the nucleus was not addressed. However, it should now be possible to determine if the RecA-like protein present in the chloroplast is involved in the uniparental inheritance of chloroplast DNA.

B. Dependence on the Chloroplast DNA Content of the Parent 1. Treatment with an Inhibitor of Chloroplast DNA Replication, 5-fluorodeoxyuridine

The chloroplast genome is present in about 80–90 copies per vegetative cell and accounts for approximately 10–15% of the total DNA (Gillham, 1978). During gametogenesis, the amount of chloroplast DNA per cell decreases to about half that of vegetative cells (Chiang and Sueoka, 1967). If vegetative cells are grown in the presence of 5fluorodeoxyuridine (FdUr), a specific inhibitor of chloroplast DNA replication, the amount of chloroplast DNA in the resulting gametes is reduced dramatically, perhaps as low as about 1% of the total DNA (Wurtz et al., 1977). When FdUr-treated gametes with a reduced chloroplast DNA content are gametes, the number of zygotes mated to untreated parent that transmit chloroplast DNA from the increases dramatically, suggesting that the specific elimination of minus chloroplast DNA has somehow been disrupted (Wurtz et al., 1977). In a manner reminiscent of UV-irradiation, comparable FdUrgametes has no effect on the treatment of subsequent uniparental inheritance of the plus chloroplast genome (Wurtz et al., 1977). Matagne and Beckers (1983) extended the FdUr treatments to include the generation of both diploid and triploid zygotes from parents with reduced chloroplast DNA contents. They too concluded that the relative amount gametes of chloroplast DNA contributed by the influences the resulting inheritance patterns. These early studies did not address how a reduction of plus chloroplast DNA could lead to an increased number of exceptional zygotes. Was the selective elimination of minus chloroplast DNA turned off in these zygotes or was the elimination process simply inefficient, with a decrease in the input of intact plus chloroplast genomes increasing the likelihood that a minus chloroplast DNA molecule would ‘slip through’ to be replicated prior to meiosis? Wurtz et al. (1977) found no increase in zygote lethality as might be expected if minus chloroplast DNA was mostly eliminated in a zygote with very little plus chloroplast DNA.

2. The mat3 Mutation New insight into how the chloroplast DNA content of the parent influences the zygote-specific elimination of minus chloroplast DNA has recently been obtained from an unexpected source. The mat3 mutation was isolated in a screen designed to identify mt+ specific genes required for the uniparental inheritance of the chloroplast genome (Gillham et

Chapter 6 Chloroplast DNA Inheritance al., 1987). The mat3 mutation is tightly linked to the locus and when present in the parent, prevents the elimination of minus chloroplast DNA in the zygote (Gillham et al., 1987; Munaut et al., 1990; Rosen et al., 1991). Thus, meiotic progeny inherit chloroplast genomes from both parents. Interestingly, the presence of mat3 has no effect on the inheritance of mitochondrial DNA (Gillham et al., 1987). It therefore seemed likely that wild-type Mat3 was specific and encoded either the zygote-specific nuclease itself or else a regulator of the nuclease required for minus chloroplast DNA elimination (Gillham et al., 1987). Recent work suggests, however, that Mat3 may play a more indirect role in chloroplast genome transmission (Armbrust et al., 1995). A recently described phenotype associated with the vegetative mat3 mutation is the generation of cells and gametes that are remarkably small (Fig. 5) and contain very little chloroplast DNA, comparable to those levels obtained with Fdur-treatment. The cell-size defect displayed by mat3 cells does not appear to be due to their reduced chloroplast DNA content. For example, a comparable reduction in the amount of chloroplast DNA by treatment with FdUr has no effect on cell size (Hosler et al., 1989; Armbrust et al., 1995). The primary defect of the mat3 mutation instead appears to be a disruption of cell-size control. An understanding of the relation between cell-size control and chloroplast DNA content awaits the cloning of Mat3. Similar to FdUr-treatment, the mat3 mutation is hypothesized to prevent the degradation of minus chloroplast DNA in the zygote as a secondary consequence of the reduced amount of chloroplast parent DNA contributed to the zygote by the (Armbrust et al., 1995). This suggests that Chlamy­ domonas may be able to count intact chloroplast genomes and inhibit minus chloroplast DNA elimination when the amount of chloroplast DNA parent is below a threshold contributed by the level. The fact that C. monoica destroys all chloroplast DNA in the zygote when plus chloroplast DNA is not protected (VanWinkle-Swift and Salinger, 1988) suggests that although zygotes can detect the amount parent, of chloroplast DNA contributed by the they may not be able to determine whether this DNA is protected. Moreover, since the biparental inheritance phenotype associated with mat3 appears to be variable (Gillham et al., 1987; Armbrust et al., 1995), it may be that the inhibition of destruction is also variable; the less chloroplast DNA contributed

107

by the parent, the more completely destruction is prevented in the zygote. Interestingly, when a mat3 zygote is also homozygous for uvsE1, an even higher level of biparental inheritance is observed (Rosen et al., 1991). How might the elimination of minus chloroplast DNA be prevented in zygotes with a reduced amount of plus chloroplast DNA? Ezy1 is expressed at wildtype levels in zygotes resulting from the mating of a and a wild-type parent (Armbrust et al., mat3 1995). Furthermore, the six UV-sensitive poly­ peptides identified by Nakamura et al. (1988) are expressed in zygotes resulting from a mating between gamete and an untreated an FdUr-treated gamete (Nakamura and Kuroiwa, 1989). Nonetheless, in both instances the selective destruction of minus chloroplast DNA is inhibited. These observations suggest that chloroplast DNA inheritance may be regulated by at least two different means: one that prevents the genes required for destruction from being transcribed, as occurs in UV irradiated cells, and one that somehow inactivates the already synthesized proteins involved in destruction, as may occur in strains that have either been treated with FdUr or else carry the mat3 mutation.

C. The Ezy2 Gene As described in an earlier section, the ability to and homozygous generate homozygous

E. Virginia Armbrust

108 sexual zygotes that are able to undergo zygote differentiation and meiosis now allows a direct analysis of the involvement of mating type-specific genes in chloroplast DNA inheritance. For example, despite the fact that ‘minus/minus’ zygotes are locus, their most missing all genes specific to the obvious phenotype is that chloroplast DNA from both parents is transmitted to meiotic progeny (Ferris et al., 1996). The simplest explanation for this biparental inheritance phenotype is that a gene specific locus, and thus absent from these zygotes, to the is required for minus chloroplast DNA elimination in the zygote. An obvious structural difference between the two mating-type loci is a 16 kb segment of DNA repeated locus and present at the 6–8 times at the locus as a single repeat, split in two (Fig. 3). A single zygote-specific gene of about 6 kb, now referred to as Ezy2 has been found within the repeat unit. The mt copy of Ezy2 appears to be a pseudogene as Ezy2 message is transcribed from the locus only within minutes of zygote formation. Preliminary sequence data suggests that the Ezy2 polypeptide has a basic pI and possesses a chloroplast transit peptide (Armbrust and Ferris, unpublished). The role of Ezy2 is still unknown, but it is intriguing that the gene product of a second, multicopy, zygote-specific locus may also be targeted gene located within the to the chloroplast during early zygote development.

a loss-of-function. VanWinkle-Swift et al. (1994) have hypothesized that the wild-type Sup1 gene encodes a nuclease expressed in the zygote from both parental genomes that is able to degrade unprotected chloroplast DNA. Thus, when the zygotespecific nuclease that would normally eliminate minus chloroplast DNA is disabled by the sup1 mutation, the presence or absence of protection of plus chloroplast DNA has no affect on subsequent inheritance patterns. These very simple hypotheses of a specific protector gene and a zygote-specific nuclease fit the original model presented for the uniparental inheritance of chloroplast DNA in C. reinhardtii. Why does uniparental inheritance in C. reinhardtii seem to be so complicated with multiple loci potentially involved, but so simple in C. monoica? Perhaps the apparent differences in the control of chloroplast genome inheritance simply reflect the different evolutionary histories of the two species (VanWinkle-Swift et al., 1994). C. monoica is in the same lineage as C. eugametos, and the C. eugametos lineage diverged from the C. reinhardtii lineage around 350 million years ago (Buchheim et al., 1990; Larson et al., 1992).

D. The sup1 Mutation

This chapter began with a simple model ofchloroplast DNA inheritance that entailed two distinct steps: protection of plus chloroplast DNA presumably due specific event occurring prior to zygote to a formation and destruction of unprotected minus chloroplast DNA during early zygote development. As more is learned about chloroplast DNA inheritance in C. reinhardtii, the number of steps and the number of genes potentially involved appears to keep increasing. I summarize here a slightly more complicated model for the regulation of chloroplast DNA inheritance in C. reinhardtii.

While the control of chloroplast DNA inheritance in C. reinhardtii seems to become more complicated with each newly collected piece of data, the control of uniparental inheritance in C. monoica appears to be remarkably simple. As described previously, VanWinkle-Swift and Salinger (1988) isolated the mtl1 mutant as a strain that is apparently unable to protect plus chloroplast DNA from elimination in the zygote, an inability that is lethal to the zygote. VanWinkle-Swift et al. (1994) have recently described the isolation of an extragenic nuclear mutation, sup1 (suppressor of uniparental inheritance) that sup­ presses this zygote lethality. The sup1 mutation appears to be recessive and unlinked to mtl1. When a zygote is homozygous for sup1, in the presence or absence of the mtl1 mutation, none of the chloroplast nucleoids are eliminated from the zygote and thus chloroplast genomes are transmitted biparentally. The recessive nature of sup1 suggests that it represents

VI. Regulation of Chloroplast DNA Inheritance

A. Regulation of Protection Although there is still nothing known about the molecular basis of protection, it has seemed clear since Gillham’s early studies (Gillham, 1963) that specific protection was due at least in part to a event that is not repressed by Mid. Furthermore, the evidence suggests that protection is reversible. A

Chapter 6

Chloroplast DNA Inheritance

simple way to accomplish this reversibility would be to initiate protection in response to the nitrogen starvation that accompanies gametogenesis. Pro­ tection could then be erased by dilution during the chloroplast DNA synthesis that occurs prior to meiosis and during vegetative growth. At this time, there is no direct way to test this hypothesis. The only potential protector mutation is in C. monoica. No comparable protector mutations exist in C. reinhardtii and there are no known treatments that prevent protection. This inability to disrupt protection has clearly hampered our ability to study its molecular basis. A more detailed understanding of protection must await the cloning of a protector gene.

B. Regulation of Destruction A number of potential components involved in destruction have now been identified. In C. monoica, the Sup1 gene is expressed in the zygote from both parental genomes. In C. reinhardtii, the UvsE1 gene is unlinked to mating type and is also expressed in the zygote from both genomes; the Ezy2 gene is present at the locus as a pseudo gene and at the locus as a functional gene and thus is expressed locus only; and the Ezy1 in the zygote from the and gene is both found at and expressed from the loci in the early zygote. Interestingly, Ezy1 is the and loci, while Ezy2 is multicopy at both the multicopy only at the locus; the Ezy2 pseudogene is present as a single copy. The evidence for the role played by any one of these polypeptides remains circumstantial. However, the activity of one or more enzymes able to completely destroy minus chloroplast DNA must obviously be tightly regulated. An unregulated nuclease that completely degrades minus chloroplast DNA even when the plus chloroplast DNA present in the zygote is damaged or otherwise deficient would likely be lethal. In fact, it appears that Chlamydomonas controls tightly those situations in which the elimination of minus chloroplast DNA is allowed to proceed: if the parent is either UV-irradiated or else possesses a reduced complement of chloroplast DNA, the zygotespecific degradation of minus chloroplast DNA is prevented. UV-irradiation of the parent prior to mating likely initiates a number of responses including the activation of DNA repair enzymes. The UvsE1 gene appears to encode an enzyme required perhaps for excision repair and recombination of the nuclear

109 genome, but is also somehow involved in minus chloroplast DNA elimination in the zygote (Rosen et al., 1991). It is not yet clear whether UvsE1 is actually localized to chloroplast DNA or whether the enzyme plays a more indirect role in uniparental inheritance. The chloroplast localized RecA-like protein also appears to be activated by UV irradiation (Cerutti et al., 1995), but the possible role of this protein in uniparental inheritance has not yet been examined directly. Chlamydomonas apparently also initiates additional responses to UV irradiation of gamete besides DNA repair, since the zygotethe specific expression of at least six polypeptides, including Ezy 1, is inhibited only when the parent is exposed to UV prior to mating (Nakamura et al., 1988; Armbrust et al., 1993). Chlamydomonas appears to respond to the potential damage of plus chloroplast DNA by UV by preventing the zygotespecific transcription of critical components that are necessary for minus chloroplast DNA elimination. Thus, meiotic progeny are assured of inheriting at least some undamaged chloroplast DNA originating parent. It is tempting to imagine that the from the mat2 mutation (Sager and Ramanis, 1974) identified a gene involved in the signaling pathway that leads to an inactivation of transcription in the zygote. The elimination of minus chloroplast DNA is also parent contains a reduced amount prevented if the of chloroplast DNA. However, this inhibition appears to occur after the presumed components of the destruction ‘machinery’ have already been synthe­ sized. Both Nakamura and Kuroiwa (1989) and Armbrust et al. (1995) found that the UV-sensitive zygote-specific polypeptides are expressed at wildtype levels when the elimination of the minus chloroplast DNA is inhibited due to a reduced parent. Since chloroplast DNA content of the minus chloroplast DNA elimination normally begins within the first hour of zygote development, these zygote-specific gene products must be quickly inactivated, perhaps due to a post-translational modification by pre-existing factors. It is intriguing to imagine that a system that may turn offdestruction once the elimination of minus chloroplast DNA is completed in wild-type zygotes, may also prevent destruction from ever beginning in zygotes whose parent either carries mat3 or else has been FdUr­ treated. If Chlamydomonas is truly able to measure the amount of chloroplast DNA contributed to the parent, then it should be possible to zygote by the identify mutants unable to monitor chloroplast DNA

E. Virginia Armbrust

110 content. These mutants would be predicted to destroy minus chloroplast DNA even when the amount of plus chloroplast DNA is below the threshold level, thereby creating a shift from biparental inheritance to either uniparental inheritance of plus chloroplast DNA or zygote lethality.

VI. Evolution of the Uniparental Inheritance of Organelle Genomes The uniparental inheritance of organelle genomes has evolved independently numerous times, and astonishingly diverse methods exist for accomplishing this feat. Yet the driving force for the evolution ofthis nearly universal phenomenon remains mysterious (reviewed in Birky, 1995). Some have hypothesized that the elimination of organelles or organelle genomes from one parent provides a means for preventing (or at least decreasing) the transmission of deleterious cytoplasmic factors (e.g. Coleman, 1982; Law and Huston, 1992). Others have hypothesized that the elaborate steps undertaken to accomplish uniparental inheritance represent a form of detente originally devised to prevent selfish organelle genomes from engaging in potentially lethal conflict for control of the cytoplasm (Cosmides and Tooby, 1981). This need to minimize ‘organelle warfare’ has even been suggested as a means for explaining why most eukaryotes possess only two sexes (Anderson, 1992; Hurst, 1992). Recently, Sears and VanWinkle-Swift (1994) have suggested that the selective elimination of minus chloroplast DNA in Chlamydomonas spp. may have evolved as a way of salvaging nucleotides from the multicopy organelle genome during the starvation that accompanies gametogenesis and zygote maturation. Birky (1995) suggests that, in fact, there may not be a single hypothesis that can adequately explain the myriad of patterns of organelle inheritance that have been described. But Birky also emphasizes the importance of identifying and evaluating the genes involved in uniparental inheritance, a process now well underway in Chlamydomonas.

Acknowledgments I thank Malcolm Campbell, Patrick Ferris, Ursula Goodenough, and Jim Umen for their many helpful comments on an earlier version of this chapter. I am particularly grateful to Ursula and Patrick for

introducing me to and helping me navigate my way through the wonderful world of the Chlamydomonas life cycle and its control by the mating-type locus.

References Anderson A (1992) The evolution of sexes. Science 257: 324– 326 Armbrust EV, Ferris PJ and Goodenough UW (1993) A mating type-linked gene cluster expressed in Chlamydomonas zygotes participates in the uniparental inheritance of the chloroplast genome. Cell 74: 801–811 Armbrust EV, Ibrahim A and Goodenough UW (1995) A mating type-linked mutation that disrupts the uniparental inheritance of the chloroplast genome also disrupts cell size control in Chlamydomonas. Mol Biol Cell 6: 1807–1818 Beck CF and Haring MA (1996) Gametic differentiation of Chlamydomonas. Int Rev Cytol 168: 259–302 Birky CW, Jr (1983) Relaxed cellular controls and organelle heredity. Science 222: 468–475 Birky CW, Jr (1995) Uniparental inheritance of mitochondrial and chloroplast genes: Mechanisms and evolution. Proc Natl Acad Sci USA 92: 1 1 3 3 1 – 1 1 3 3 8 Bolen PL, Grant DM, Swinton D, Boynton JE and Gillham NW (1982) Extensive methylation of chloroplast DNA by a nuclear gene mutation docs not affect chloroplast gene transmission in Chlamydomonas. Cell 28: 335–343 Boynton JE, Gillham NW, Newman SM and Harris EH (1991) Organelle genetics and transformation of Chlamydomonas. In: Herrmann RG (eds) Advances in Plant Gene Research: Cell Organelles, Vol 6, pp 3–64. Springer-Verlag, New York Braten T ( 1 9 7 1 ) The ultrastructure of fertilization and zygote formation in the green alga Ulva mutabilis. J Cell Sci 9: 621– 635 Braten T (1973) Autoradiographic evidence for the rapid disintegration of one chloroplast in the zygote of the green alga Ulva mutabilis. J Cell Sci 12: 385–389 Britt AB (1996) DNA damage and repair in plants. Ann Rev Plant Physiol Plant Mol Biol 47: 75–100 Buchheim MA, Tunnel M.Zimmer EA and Chapman RL(1990) Phylogeny of Chlamydomonas (Chlorophyta) based on cladistic analysis of nuclear 18S rRNA sequence data. J Phycol 26: 689–699 Burton WG, Roberts RJ, Myers PA and Sager R (1977) A sitespecific single-strand endonuclease from the eukaryote Chlamydomonas. Proc Natl Acad Sci USA 7: 2687–2691 Burton WG, Grabowy CT and Sager R (1979) Role ofmethylation in the modification and restriction of chloroplast DNA in Chlamydomonas. Proc Nat1 Acad Sci USA 76: 1390–1394 Campbell AM, Rayala HJ and Goodenough UW (1995) The isol gene of Chlamydomonas is involved in sex determination. Mol Biol Cell 6: 87–95 Cavalier-Smith T (1970) Electron microscopic evidence for chloroplast fusion in zygotes of Chlamydomonas reinhardi. Nature 228: 333–335 Cerutti H, Johnson AM, Boynton JE and Gillham NW (1995) Inhibition of chloroplast DNA recombination and repair by dominant negative mutants of Escherichia coli RecA. Mol Cell Biol 15: 3003–3011

Chapter 6

Chloroplast DNA Inheritance

Chiang K-S and Sueoka N (1967) Replication of chloroplast DNA in Chlamydomonas reinhardi during vegetative cell cycle: Its mode and regulation. Proc Natl Acad Sci USA 1506– 1513: Chun EHL, Vaughan MH, Jr. and Rich A (1963) The isolation and characterization of DNA associated with chloroplast preparations. J Mol Biol 7: 130–141 Coleman AW (1982) Sex is dangerous in a world of potential symbionts or the basis of selection of uniparental inheritance. J Theor Biol 97: 367–369 Coleman AW (1984) The fate of chloroplast DNA during cell fusion, zygote maturation, and zygote germination in Chlamydomonas reinhardi as revealed by DAPI staining. Exp Cell Res 152:528–540 Coleman AW and Maguire MJ (1983) Cytological detection of the basis of uniparental inheritance of plastid DNA in Chlamydomonas moewusii. Curr Genet 7: 211–218 Cosmides LM and Tooby J (1981) Cytoplasmic inheritance and intragenomic conflict. J Theor Biol 89: 83–129 Dron M, Rahire M, Rochaix J-D and Mets L (1983) First DNA sequence of a chloroplast mutation: A missense alteration in the ribulosebisphosphate carboxylase large subunit gene. Plasmid 9: 321–324 Ebersold WT (1967) Chlamydomonas reinhardi: Heterozygous diploid strains. Science 157: 447–449 Ebersold WT, Levine RP, Levine EE and Olmsted MA (1962) Linkage maps in Chlamydomonas reinhardi. Genetics 47: 531–543 Feng T–Y and Chiang K–S (1984) The persistence of maternal inheritance in Chlamydomonas despite hypomethylation of chloroplast DNA induced by inhibitors. Proc Natl Acad Sci USA 81: 3438–3442 Ferris PJ (1995) Localization of the nic-7, ac-29 and thi-10 genes within the mating-type locus of Chlamydomonas reinhardtii. Genetics 141: 543–549 Ferris PJ and Goodenough UW (1987) Transcription of novel genes, including a gene linked to the mating-type locus, induced by Chlamydomonas fertilization. Mol Cell Biol 7: 2360–2366 Ferris PJ and Goodenough UW (1994) The mating type locus of Chlamydomonas reinhardtii contains highly rearranged DNA sequences. Cell 76: 1135–1145 Ferris PJ and Goodenough UW (1997) Mating type in Chlamydomonas is specified by mid, the minus-dominance gene. Genetics 146:859–870 Ferris PJ, Woessner JP and Goodenough UW (1996) A sex recognition glycoprotein is encoded by the plus mating-type gene fus 1 of Chlamydomonas reinhardtii. Mol Biol Cell 7: 1235–1248 Frost BF and Small GD (1984) Partial purification and c h a r a c t e r i z a t i o n of the major AP endonuclease from Chlamydomonas reinhardi. Biochim Biophys Acta 782:170– 176 Galloway RE and Goodenough UW (1985) Genetic analysis of mating locus linked mutations in Chlamydomonas reinhardtii. Genetics 111:447–461 Gillham NW (1963) Transmission and segregation of a non­ chromosomal factor controlling streptomycin resistance in diploid Chlamydomonas. Nature 200: 294 G i l l h a m NW (1965) Induction of chromosomal and non­ chromosomal mutations in Chlamydomonas reinhardi with Nmethyl-N´-nitro-N-nirosoguanidine. Genetics 52: 529–537 Gillham NW (1969) Uniparental inheritance in Chlamydomonas

111 reinhardi. Am Nat 103: 355–388 Gillham NW (1978) Organelle Heredity. Raven Press, New York Gillham NW (1994) Organelle Genes and Genomes. Oxford University Press, New York Gillham NW and Fifer W (1968) Recombination of nonchromo­ somal mutations: A three-point cross in the green alga Chlamydomonas reinhardi. Science 162: 683–684 Gillham NW, Boynton JE and Lee RW (1974) Segregation and recombination of non-Mendelian genes in Chlamydomonas. Genetics 78: 439–457 Gillham NW, Boynton JE, Johnson AM and Burkhart BD (1987) Mating type linked mutations which disrupt the uniparental transmission of chloroplast genes in Chlamydomonas. Genetics 115: 677–684 Gillham NW, Boynton JE and Harris EH (1991) Transmission of plastid genes. In: Bogorad L and Vasil IK (eds) Cell Culture and Somatic Cell Genetics, Vol 7A, pp 55–92. Academic Press, Inc., New York Goodenough UW and Ferris PJ (1987) Genetic regulation of development in Chlamydomonas. I n : Loomis W (eds) Genetic Regulation of Development, pp 171–189. Alan R. Liss, Inc., New York Goodenough UW, Hwang C and Martin H (1976) Isolation and genetic analysis ofmutant strains of Chlamydomonas reinhardi defective in gametic differentiation. Genetics 82: 169–186 Goodenough UW, Detmers PA and Hwang C (1982) Activation for cell fusion in Chlamydomonas: Analysis of wild-type gametes and nonfusing mutants. J Cell Biol 92: 378–386 Goodenough UW, Armbrust EV, Campbell AM and Ferris PJ (1995) Molecular genetics of sexuality in Chlamydomonas. Ann Rev Plant Phys Plant Mol Biol 46: 21–44 Grant DM, Gillham NW and Boynton JE (1980) Inheritance of chloroplast DNA in Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 77: 6067–6071 Harris EH (1989) The Chlamydomonas Sourcebook. Academic Press, San Diego Harris EH, Burkhart BD, Gillham NW and Boynton JE (1989) Antibiotic resistance mutations in the chloroplast 16S and 23S rRNA genes of Chlamydomonas reinhardtii: correlation of genetic and physical maps of the chloroplast genome. Genetics 123:281–292 Hoeh WR, Sterwart DT, Sutherland BW and Zouros E (1996) Multiple origins of gender-associated mitochondrial DNA lineages in bivalves (Mollusca:Bivalvia). Evolution 50: 2276– 2286 Hosler JP, Wurtz EA, Harris EH, Gillham NW and Boynton JE (1989) Relationship between gene dosage and gene expression in the chloroplast of Chlamydomonas reinhardtii. Plant Physiol 91: 648–655 Hurst LD (1992) Intragenomic conflict as an evolutionary force. Proc R Soc Lond (B) 248: 135–140 Ikehara T, Uchida H, Suzuki L and Nakamura S (1996) Chloroplast nucleoids in large number and large DNA amount with regard to maternal inheritance in Chlamydomonas reinhardtii. Protoplasma 194: 1 1 – 1 7 Kuroiwa T (1985) Mechanisms of maternal inheritance of chloroplast DNA: An active digestion hypothesis. Microbiol Sci 2: 267–270 Kuroiwa T( 1991) The replication, differentiation, and inheritance of plastids with emphasis on the concept of organelle nuclei. Int Rev Cytol 128: 1–62

112 Kuroiwa T, Kawano S and Nishibayashi S (1982) Epifluorescent microscopic evidence for maternal inheritance of chloroplast DNA. Nature 298: 481–483 Kuroiwa T, Kawano S and Sato C (1983a) Mechanisms of maternal inheritance. I. Protein synthesis involved in preferential destruction of chloroplast DNA of male origin. Proc Jpn Acad (B) 59: 177–181 Kuroiwa T, Kawano S and Sato C (1983b) Mechanisms of maternal inheritance. II. RNA synthesis involved in preferential destruction of chloroplast DNA of male origin. Proc Jpn Acad ( B ) 59: 182–185 Larson A, Kirk MM and Kirk DL(1992) Molecular phylogeny of the Volvocine flagellates. Mol Biol Evol 9: 85–105 Law R and Huston V (1992) Intracellular symbionts and the evolution of uniparental cytoplasmic inheritance. Proc R Soc Lond B 248: 69–77 Lemieux C, Turmel M, Seligy VL and Lee RW (1984) Chloroplast DNA recombination in interspecific hybrids of Chlamy­ domonas: Linkage between a nonmendelian locus for streptomycin resistance and restricion fragments coding for 16S rRNA. Proc Natl Acad Sci USA 81: 1164–1168 L i u X–Q, Gillham NW and Boynton JE (1989) Chloroplast ribosomal protein gene rps12 of Chlamydomonas reinhardtii. J Biol Chem 264: 16100–16108 Matagne RF (1981) Transmission ofchloroplast alleles in somatic fuison products obtained from vegetative cells and/or ‘gametes’ of Chlamydomonas reinhardi. Curr Genet 3: 31–36 Matagne RF (1987) Chloroplast gene transmission in Chlamy­ domonas reinhardtii. A model for its control by the matingtype locus. Curr Genet 12: 251–256 Matagne RF and Beckers M-C (1983) Perturbation of chloroplast gene t r a n s m i s s i o n in d i p l o i d and t r i p l o i d zygotes of Chlamydomonas reinhardi by 5-fluorodeoxyuridine. Curr Genet 7: 335–338 Matagne RF and Hermesse M-P (1980) Chloroplast gene inheritance studied by somatic fusion in Chlamydomonas reinhardtii, Curr Genet 1: 127–131 Matagne RF and Mathieu D (1983) Transmission of chloroplast genes in triploid and tetraploid zygospores of Chlamydomonas reinhardtii: Roles of mating-type gene dosage and gametic chloroplast DNA content. Proc Natl Acad Sci USA 80: 4780– 4783 Matagne RF, Deltour R and Ledoux L (1979) Somatic fusion between cell wall mutants of Chlamydomonas reinhardi. Nature 278: 344–346 Matters GL and Goodenough UW (1992) A gene/pseudogene tandem duplication encodes a cysteine-rich protein expressed during zygote development in Chlamydomonas reinhardtii. Mol Gen Genet 232: 81–88 McBride AC and McBride JC (1975) Uniparental inheritance in Chlamydomonas eugametos (chlorophyceae). J Phycol 11: 343–344 Meland S, Johansen S, Johansen T, Haugli K and Haugli F (1991) Rapid disappearance of one mitochondrial genotype after isogamous mating in the myxomycete Physarum polycephalum. Curr Genet 19: 55–60 Mets LJ and Geist LJ (1983) Linkage of a known chloroplast gene mutation to the uniparental genome of Chlamydomonas reinhardii. Genetics 105: 559–579 M u n a u t C, Dombrowicz D and Matagne RF (1990) Detection of chloroplast DNA by using fluorescent monoclonal anti­ bromodeoxyuridine antibody and analysis of its fate during

E. Virginia Armbrust zygote formation in Chlamydomonas reinhardtii. Curr Genet 18: 259–263 Nakamura S and Kuroiwa T (1989) Selective elimination of chloroplast DNA by 5-fluorodeoxyuridine causing no effect on preferential digestion of male chloroplast nucleoids in Chlamydomonas. Eur J Cell Biol 48: 165–173 Nakamura S, Sato C and Kuroiwa T (1988) Polypeptides related to preferential digestion of male chloroplast nucleoids in Chlamydomonas. Plant Sci 56: 129–136 Neale DB, Marshall KA and Sederoff RR (1989) Chloroplast and mitochondrial DNA are paternally inherited in Sequoia sempervirens D. Don Endl. Proc Natl Acad Sci USA 86: 9347– 9349 Ogawa K and Kuroiwa T (1985a) Destruction of chloroplast nuclei of the male gamete by calcium and nuclease C in a cell model of Chlamydomonas reinhardtii. Plant Cell Physiol 26: 493–503 Ogawa K and Kuroiwa T (1985b) Nuclease C polymorphism of calcium-dependent nucleases in Chlamydomonas reinhardtii. Plant Cell Physiol 26: 481–491 Ogawa K and K u r o i w a T ( 1 9 8 6 a ) Base-specific endo­ exonucleolytic activity of Chlamydomonas nuclease C1 & 2. Plant Cell Physiol 27: 701–710 Ogawa K and Kuroiwa T (1986b) Purification of major isozymes of nuclease C and production of active fragments by trypsin. Plant Cell Physiol 26: 1473–1484 Ogawa K and Kuroiwa T (1987) Preferential resistance of phosphodiester bonds to deoxycytidine and 5´-adjacent bases to Chlamydomonas nuclease C. Plant Cell Physiol 28: 323– 332 Ris H and Plaut W (1962) Ultrastructurc of DNA-containing areas in the chloroplast of Chlamydomonas. J Cell Biol 13: 383–391 Rochaix J-D ( 1 9 9 5 ) Chlamydomonas reinhardtii as the photosynthetic yeast. Ann Rev Genet 29: 209–230 Rosen H and Ebersold WT (1972) Recombination in relation to ultraviolet sensitivity in Chlamydomonas reinhardi. Genetics 71: 247–253 Rosen H, Newman SM, Boynton JE and Gillham NW ( 1 9 9 1 ) A nuclear mutant of Chlamydomonas that exhibits increased sensitivity to UV irradiation, reduced recombination of nuclear genes, and altered transmission of chloroplast genes. Curr Genet 19: 35–41 Royer H-D and Sager R (1979) Methylation ofchloroplast DNAs in the life cycle of Chlamydomonas. Proc Natl Acad Sci USA 76: 5794–5798 Sager R (1954) Mendelian and non-Mendelian inheritance of streptomycin resistance in Chlamydomonas reinhardi. Proc Natl Acad Sci USA 40: 356–363 Sager R (1965) Mendelian and non-Mendelian heredity: A reappraisal. Proc Royal Soc Lond Ser B 164: 290–297 Sager R and Grabowy C (1983) Differential methylation of chloroplast DNA regulates maternal inheritance in a methylated mutant of Chlamydomonas. Proc Natl Acad Sci USA 80: 3025–3029 Sager R and Grabowy C (1985) Sex in Chlamydomonas: Sex and the single chloroplast. In: Halvorson HO and Monroy A (eds) The Origin and Evolution of Sex, pp 1 1 3 – 1 2 1 . Alan R. Liss, New York Sager R and Granick S (1954) Nutritional control of sexuality in Chlamydomonas reinhardi. J Gen Physiol 37: 729–742 Sager R and Ishida MR (1963) Chloroplast DNA in Chlamy­

Chapter 6 Chloroplast DNA Inheritance domonas. Proc Natl Acad Sci USA 50: 725–730 Sager R and Kitchin R (1975) Selective silencing of eukaryotic DNA. Science 189: 426–432 Sager R and Lane R (1972) Molecular basis of maternal inheritance. Proc Natl Acad Sci USA 69: 2410–2413 Sager R and Ramanis Z (1963) The particulate nature of nonchromosomal genes in Chlamydomonas. Proc Natl Acad Sci USA 50: 260–268 Sager R and Ramanis Z (1965) Recombination of nonchromo­ somal genes in Chlamydomonas. Proc Natl Acad Sci USA 53: 1053–1060 Sager R and Ramanis Z (1967) Biparental inheritance of nonchromosomal genes induced by ultraviolet irradiation. Proc Natl Acad Sci USA 58: 931–937 Sager R and Ramanis Z (1973) The mechanism of maternal inheritance in Chlamydomonas: Biochemical and genetic studies. Theoret Appl Genet 43: 101–108 Sager R and Ramanis Z (1974) Mutations that alter the transmission of chloroplast genes in Chlamydomonas. Proc Natl Acad Sci USA 71: 4698–4702 Sager R, Grabowy C and Sano H (1981) The mat-1 gene in Chlamydomonas regulates DNA methylation during gameto­ genesis. Cell 24: 41–47 Sano H, Grabowy C and Sager R (1981) Differential activity of DNA methyltransferase in the life cycle of Chlamydomonas reinhardi. Proc Natl Acad Sci USA 78: 3118–3122 Sano H, Grabowy C and Sager R (1984) Loss of chloroplast DNA methylation during dedifferentiation of Chlamydomonas reinhardi gametes. Mol Cell Biol 4: 2103–2108 Sano H, Royer H–D and Sager R (1980) Identification of 5­ methylcytosine in DNA fragments immobilized on nitro­ cellulose paper. Proc Natl Acad Sci USA 77: 3581–3585 Sears BA (1980) Elimination of plastids during spermatogenesis and fertilization in the plant kingdom. Plasmid 4: 233–255 Sears BB and VanWinkle-Swift KP (1994) The salvage/turnover/ repair (STOR) model for uniparental inheritance in Chlamydomonas: DNA as a source of sustenance. J Hered 85: 366–376 Shepherd HS, Boynton JE and Gillham NW (1979) Mutations in nine chloroplast loci of Chlamydomonas affecting different photosynthetic functions. Proc Natl Acad Sci USA 76: 1353– 1357 Small GD (1987) Repair systems for nuclear and chloroplast DNA in Chlamydomonas reinhardtii. Mut Res 181: 31–35 Smith GM and Regnery DC (1950) Inheritance of sexuality in Chlamydomonas reinhardi. Proc Natl Acad Sci USA 36: 246– 258 Spreitzer RJ and Mets LJ (1980) Non-mendelian mutation affecting ribulose-1,5-bisphosphate carboxylase structure and activity. Nature 285: 114–115 Tait GCL and Harris WJ (1977a) A deoxyribonuclease from Chlamydomonas reinhardii. 1. Purification and properties. Eur J Biochem 75: 357–364 Tait GCL and Harris WJ (1977b) A deoxyribonuclease from Chlamydomonas reinhardii. 2. Substrate specificity, mode of action and products. Eur J Biochem 75: 365–371 Tsubo Y and Matsuda Y (1984) Transmission of chloroplast genes in crosses between Chlamydomonas diploids: Correlation

113 with chloroplast nucleoid behavior in young zygotes. Curr Genet 8: 223–229 Uchida H, Kawano S, Nakamura S and Kuroiwa T (1992a) A study on the effects of UV on preferential digestion of chloroplast nuclei in young zygotes of Chlamydomonas reinhardtii. Cytologia 57: 395–399 Uchida H, Nozue F, Kuroiwa H, Osafune T, Sumida A, Ehara T and Kuroiwa T (1992b) Evidence for preferential digestion of male-derived chloroplast DNA in young zygotes of Chlamydomonas reinhardtii by histochemical immunogold electron microscopy. Cytologia 57: 463–470 Uchida H, Kawano S, Sato N and Kuroiwa T (1993) Isolation and characterization of novel genes which are expressed during the very early stage of zygote formation in Chlamydomonas reinhardtii. Curr Genet 24: 296–300 van den Ende H and VanWinkle–Swift KP (1994) Mating-type differentiation and mate selection in the homothallic Chlamydomonas monoica. Curr Genet 25: 209–216 VanWinkle-Swift KP (1978) Uniparental inheritance ispromoted by delayed division of the zygote in Chlamydomonas. Nature 275: 749–751 VanWinkle-Swift KP and Aubert B (1983) Uniparental inheritance in a homothallic alga. Nature 303: 167–169 VanWinkle-Swift KP and Bauer JC (1982) Self-sterile and maturation-defective mutants of the homothallic alga, Chlamydomonas monoica (Chlorophyceae). J Phycol 18: 312– 317 Van Winkle-Swift KP and Burrascano CG (1983) Comple­ mentation and preliminary linkage analysis ofzygote maturation mutants of the homothallic alga, Chlamydomonas monoica. Genetics 103: 429–445 VanWinkle-Swift KP and Halm J-H (1986) The search for mating-type-limited genes in the homothallic alga Chlamy­ domonas monoica. Genetics 113: 601–619 VanWinkle-Swift KP and Salinger AP (1988) Loss of mt+derived zygotic chloroplast DNA is associated with a lethal allele in Chlamydomonas monoica. Curr Genet 13: 331–337 VanWinkle-Swift K, Hoffman R, Shi L and Parker S (1994) A suppressor of a mating-type limited zygotic lethal allele also suppresses uniparental chloroplast gene transmission in Chlamydomonas monoica. Genetics 136: 867–877 Wegener D and Beck C (1991) Identification of novel genes specifically expressed in Chlamydomonas reinhardtii zygotes. Plant Mol Biol 16: 937–946 Whatley JM (1982) Ultrastructure of plastid inheritance: Green algae to angiosperms. Biol Rev 57: 527–569 Woessner JP and Goodenough UW (1989) Molecular charac­ terization of a zygote wall protein: An extensin-like molecule in Chlamydomonas reinhardtii. Plant Cell 1: 901–911 Wurtz EA, Boynton JE and Gillham NW (1977) Perturbation of chloroplast DNA amounts and chloroplast gene transmission in Chlamydomonas reinhardtii by 5-fluorodeoxyuridine. Proc Natl Acad Sci USA 74 (10): 4522–4556 Yaneda H, Hayashi J-I, Takahama S, Taya C, Lindahl K and Yonekawa H (1995) Elimination of paternal mitochondrial DNA in intraspecific crosses during early mouse embryo­ genesis. Proc Natl Acad Sci USA 92: 4542–4546

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Chapter 7 Replication, Recombination, and Repair in the Chloroplast Genetic System of Chlamydomonas Barbara B. Sears

Department of Botany & Plant Pathology, Michigan State University,

East Lansing, Michigan 48824-1312 U.S.A.

Summary I. Introduction II. Replication A. Replication of cpDNA during cell cycle B. Nucleoids as Indicators of cpDNA Abundance C. cpDNA Degradation and Nucleotide Recycling D. Origins of cpDNA Replication E. Enzymes of cpDNA Replication III. Recombination A. Genetic Analysis of Recombination 1. Assessment of Chloroplast Gene Recombination Frequencies in Meiotic Progeny 2. Assessment of Chloroplast Gene Recombination Frequencies Through the

Analysis of Biparental Zygospore Clones 3. Assessment of Chloroplast Gene Recombination Frequencies After Paternal

Marker Selection 4. Assessment of Recombination Frequencies in Vegetative Diploid Zygotes B. Factors that Affect Recombination Frequency C. Recombination Within the Inverted Repeat D. Recombination Hotspots E. Gene Conversion F. Intron Homing IV. Repair A. UV-Damage and Photoreactivation B. Specific Impact of FdUrd on cpDNA C. Other Mutagens D. The Interrelated Processes of Recombination/Repair V. Perspectives and Conclusions Acknowledgments References

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Summary Chloroplast DNA replicates throughout the Chlamydomonas cell cycle, with individual DNA molecules being randomly chosen for replication. Mosaic labeling of the cpDNA in density transfer experiments may indicate that replication is dispersive, rather than semi- conservative, but it also may result from multiple rounds of recombination between the highly polyploid cpDNA molecules. Repair of spontaneous DNA damage may also contribute to dispersive labeling, and cpDNA turnover and nucleotide recycling may comprise a further contributing factor. Two origins of replication have been identified on the cpDNA molecule, and their activity has been studied in vitro. Exposure of cells to a gyrase inhibitor initially inhibits replication, and then stimulates J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 115–138. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

Barbara B . Sears

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a cryptic origin, near a site identified as a recombination hot spot in other studies. Transformation experiments point to the possible existence of at least one additional origin. For cpDNA, recombination is not limited to the sexual cycle, but occurs during vegetative cell growth as well, probably because enzymes that function in recombination, including a RecA-like protein, play an essential role in the repair of DNA damage and hence are present throughout the cell cycle. In mitotic cells, copy correction and ‘flip-flop’ recombination between the two copies ofthe inverted repeat are a consequence ofthese activities. From sexual crosses, biparental zygotes have been recovered in which recombination and gene conversion have been examined using chloroplast genetic markers and/or restriction fragment length polymorphisms (RFLPs). These procedures, combined with chloroplast transformation, have identified a recombination hotspot in the inverted repeat of C. reinhardtii. Other hotspots of recombination have been characterized in crosses of C. eugametos and C. moewusii, as well as C. reinhardtii, where several DNA segments act as ‘selfish DNAs’, inserting themselves into the target site in the plastome contributed by the other parent, and thus proliferating among the progeny. Several of these segments contain homing introns, which encode endonucleases essential for their movement. Several aspects of repair have been characterized biochemically, including the direct reversal of UV-damage by photolyase, and the apparent absence of methyltransferases for reversal of alkylation damage. The removal of pyrimidine dimers in the dark may occur through excision repair, recombinational repair or a combination of the two. When cells are treated with 5-fluoro-deoxyuridine, the cpDNA of C. reinhardtii becomes reduced in abundance and is highly mutable. Thus, this thymidine analog is a potent and specific mutagen, and has been effectively combined with other mutagenic treatments for the induction of non-Mendelian mutations in C. reinhardtii. Future investigations doubtlessly will utilize the ability to transform both the nuclear and chloroplast genomes with the classical genetic approach of inducing and analyzing mutations, and biochemical procedures for in vitro analysis. Heterologous probes and E. coli mutants deficient in specific aspects of replication, recombination and repair will serve as useful resources for the identification ofgenes that affect those processes in the chloroplast genetic system of Chlamydomonas.

I. Introduction The chloroplast genetic system depends upon replication and repair for the perpetuation and preservation of its DNA. In prokaryotes and eukaryotes, some enzymes essential for replication and repair also are utilized for recombination, a complex process in the Chlamydomonas chloroplast, and a process that is not yet fully understood. This chapter reviews genetic, biochemical, and molecular investigations that have yielded insight into these aspects of chloroplast DNA metabolism. Instances are noted where features of replication, recom­ bination, and repair in the chloroplast genetic system of Chlamydomonas differ from higher plants. In this review, the recent literature has received more attention than the older literature because several thorough reviews from previous decades are available. Abbreviations: cpDNA – chloroplast DNA; DAPI – 4´,6diamidino-2-phenylindole; FdUrd – 5-fluoro-deoxyuridine; IPTG MMS – methylmeth­ anesulfonate; mt – mating-type; plastome – plastid genome; RFLP – restriction fragment length polymorphism

Of particular value are reviews that cover cpDNA replication (Keller and Ho, 1981) and repair in Chlamydomonas (Small, 1987), chloroplast recom­ bination, genetic mapping and mobile introns (Gillham, 1978, 1994), cytological observations of chloroplast nucleoids (Kuroiwa, 1991), and the Chlamydomonas Sourcebook (Harris, 1989), which encompasses all ofthese topics. This chapter will not consider the topic of chloroplast gene inheritance; for that, the reader is referred to the chapter by Armbrust.

II. Replication Studies of chloroplast DNA (cpDNA) replication in Chlamydomonas have been facilitated by two important traits that are not general features of higher plant chloroplasts. First, cpDNA of Chlamydomonas has a much higher A-T content than does nuclear DNA, allowing the two classes of DNA to be separated in CsCl buoyant density gradients, even without the addition of ethidium bromide or bisbenzimide (Sager

Chapter 7

Chloroplast Replication, Recombination and Repair

and Ishida, 1963). In those gradients, the major band while the second most represents nuclear or prominent band is composed of cpDNA, also referred to as the (Fig. 1). The cpDNA composes 8– 18% of the total cellular DNA in logarithmically growing cells, and about 3–8% in the gametes (Sager and Ishida, 1963; Chiang and Sueoka, 1967; Chiang, 1971; Sears et al., 1980; Turmel et al., 1980). The second useful trait is the fact that exogenous thymidine is incorporated only into cpDNA due to the presence of a chloroplast-specific thymidine kinase (Swinton and Hanawalt, 1972; Chiang et al., 1975). Since adenine will be incorporated into both nuclear and cpDNA, when cells are provided with two differ­ entially labeled nitrogenous bases (i.e., their differential incorporation and can be used to distinguish the replication patterns of nuclear and cpDNA.

A. Replication of cpDNA during cell cycle An overview of the cell cycle is shown in Fig. 2. When cells are grown under phototrophic conditions in liquid culture with alternating periods of 12 h light, 12 h dark, chloroplast and cell divisions occur

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fairly synchronously during the dark phase (reviewed by Harris, 1989). In the initial experiments designed to examine replication in C. reinhardtii, synchronized cells were grown in and then transferred just after cell division (Chiang and to Sueoka, 1967). When cells were harvested during the next 24 h, shifts in buoyant densities of the peak indicated that nuclear DNA underwent two rounds of semiconservative replication during the dark period, resulting in a band of hybrid density (labeled with ) and a less dense, more abundant band (labeled exclusively with ). Similarly, the peak showed density shifts that indicated cpDNA replicated in a semi-synchronized and semiconservative fashion during the 12-h light period. When the experiments were repeated, the timing of nuclear DNA replication was in agreement with the previous studies, but cpDNA was found to replicate in both the light and dark periods (Chiang, 1971; Lee and Jones, 1973; Turmel et al., 1980, 1981). Furthermore, after cells were transferred to the buoyant density of the broadened, with the peak shifting gradually, a result that was interpreted to be due to random selection of cpDNA molecules for replication (Turmel et al.,

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1980, 1981). Further analysis focused on isolated from cells after one 24-h period of growth in (Turmel et al., 1981). The cpDNA was denatured to examine the buoyant density distri­ butions of the single-strands, and no discrete heavy strand was apparent. This result indicates that nucleotides were incorporated into both strands, suggesting that cpDNA replication may be dispersive rather than semi-conservative. However, Turmel et al. (1981) pointed out that the dispersive labeling could have been caused by frequent recombination among the polyploid plastomes, some that had replicated, and some that had not yet replicated. Moreover, Woefle et al. (1993) have noted that recombination could act to initiate DNA replication and thus yield a cpDNA having a mosaicism of incorporated label. As described in more detail in

Barbara B . Sears

later sections, other studies with mitotically growing cells have demonstrated frequent intramolecular recombination between the inverted repeats (Aldrich et al., 1985) and intra- or intermolecular recom­ bination between duplications created through biolistic transformation (Cerutti et al., 1995), indicating that the enzymes for recombination are indeed present in vegetative cells.

B. Nucleoids as Indicators of cpDNA Abundance The abundance and distribution of cpDNA has been examined cytologically through the use of 4',6diamidino-2-phenylindole (DAPI) to visualize nucleoids, which represent aggregations of cpDNA molecules. The initial application by Coleman (1978) of this fluorochrome to study the Chlamydomonas

Chapter 7

Chloroplast Replication, Recombination and Repair

chloroplast was followed by Kuroiwa, Birky, and others to track cpDNA during the sexual cycle (e.g., Kuroiwa et al., 1982; Birky et al., 1984; Coleman, 1984; Nakamura et al., 1991). These important contributions are described in the chapter by Armbrust. Kuroiwa and coworkers examined changes in number and shape of nucleoids in synchronized liquid cultures of C. reinhardtii (Kuroiwa et al., 1981) and cells growing on solid media (Nakamura et al., 1986). They observed increases in nucleoid size and intensity of DNA staining as the chloroplasts and cells enlarged. Shortly before chloroplast division in the synchronous cultures, the nucleoids became dumbbell-shaped and divided semi-synchronously, with the daughter nucleoids segregating into the four daughter chloroplasts. A subsequent investigation by Ehara et al. (1990) refined the fixation procedures, and reinvestigated the changes in nucleoid shape and distribution during the cell cycle. They found that when cells are grown synchronously, early in the light phase, most of the cpDNA is clustered near the pyrenoid (Fig. 2). Just before the dark phase and prior to division, the cpDNA becomes dispersed throughout each chloroplast. The dispersed cpDNA was found to aggregate if high concentrations of glutaraldehyde (0.5%) were used during the fixation procedures. A functional basis for the dispersion of the cpDNA nucleoids during the dark phase is unclear, since the biochemical characterizations described in the previous section indicate that cpDNA replicates throughout the cell cycle. However, as discussed in Section I.D, the location of cpDNA near the pyrenoid may be meaningful.

C. cpDNA Degradation and Nucleotide Recycling Several observations indicate that cpDNA is not as stable nor as persistent as nuclear DNA. The dispersive labeling observed in cpDNA of vegetative cells was also reported for young zygotes shortly after gamete fusion and in meiotic progeny, and was interpreted to indicate extensive recombination had occurred between the and cpDNAs (Chiang, 1971). However, these experiments are open to criticism since the procedures required to break open the thick walled zygospores caused shearing of the DNA, and hence, the peaks were poorly resolved in the CsCl gradients. Furthermore, since chloroplast markers are inherited predominantly from the parent, and nucleoids disappear shortly after gamete fusion, cpDNA molecules are few doubt that most

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degraded soon after gamete fusion (reviewed in Chapter 6, Armbrust). A shift in the buoyant density of in gametes (see Fig. 1b) was interpreted by Ruth Sager and colleagues to indicate that the cpDNA becomes methylated during gametogenesis and hence protected from an endonuclease that was chloroplast hypothesized to be activated in the (Sager and Lane, 1972). Subsequent studies using methylation-sensitive restriction enzymes (Royer and Sager, 1979) and HPLC characterization ofthe DNA showed that cytosines become extensively methylated gametes (Burton et al., 1979). The cytidine in target was verified by Feng and Chiang (1984) who used 5-azacytidine and L-ethionine to inhibit methylation. A nuclear mutation was isolated that results in hyper-methylation of the cpDNA at all stages of the cell cycle, regardless of mating type (Bolen et al., 1982). For a discussion of the correlations between methylation and uniparental inheritance of chloroplast markers, the reader is referred to Chapter 6 (Armbrust). Sears and VanWinkle-Swift (1994) suggested that cpDNA could have a the degradation of the selective advantage, especially under the conditions of nitrogen starvation which induce gametogenesis and zygospore dormancy. The salvage/turnover/repair (STOR) model postulates that the high polyploidy of cpDNA allows it to be used as a cellular resource for carbon, nitrogen, and nucleotides, especially under conditions of nutrient limitation. The STOR model further suggests that if cpDNAs are damaged, they might be turned over rather than repaired, and that a balance between replication and degradation would thus exist in all stages of the cell cycle, including the vegetative growth phase. This model is consistent with the dispersive labeling of cpDNA that was reviewed previously, and the observations of Small and Greimann (1977), who were studying dark repair of UV-induced lesions. In their unirradiated C. reinhardtii control cultures, Small and Greimann found that when the cells were grown in and then incubated in the adenine and dark for 24 h, the nuclear DNA was stably maintained, but about half of the cpDNA was degraded during the dark period. Results from Cerutti et al. (1995) have shown that when the pathway is inhibited, both the integrity and abundance of cpDNA are diminished under conditions of extensive DNA damage. Their interpretation is that excessively damaged cpDNA molecules are probably degraded rather than repaired. On the other hand, since recombination-repair

120 involves partial DNA degradation and resynthesis, if that process is operating in vegetative cells and/or zygotes, it could contribute to the dispersive labeling patterns observed for cpDNA (H. Cerutti, personal communication). In C. moewusii, evidence for cpDNA turnover has density transfer experiments, been found in in which the density of the cpDNA decreased more rapidly than expected from the net amounts of DNA synthesized (M. Tunnel, personal communication). In addition, a linear 6-kb segment of cpDNA that is consistently recovered from this species (Tunnel et al., 1986) appears to represent a specifically-protected remnant of the degraded cpDNA.

D. Origins of cpDNA Replication Initial attempts to locate the origins of plastome replication identified sequences that were capable of

Barbara B . Sears replication after transformation into the yeast nucleus (Loppes and Denis, 1983;Valletetal., 1984). Another study fortuitously discovered that at least four cpDNA segments were capable of autonomous replication in the Chlamydomonas nucleus (Rochaix et al., 1984). Subsequent investigations (Vallet and Rochaix, 1985; Wu et al., 1986) indicated that those sites differed from the location of two replication bubbles or displacement loops (D-loops) on the native cpDNA molecule visualized with electron microscopy (Waddell et al., 1984; Wang et al., 1984), although several were intriguingly close. Two D-loops that were observed by Madeline Wu and colleagues were mapped to adjacent EcoRI fragments contained in the fifth largest Bam HI fragment on the cpDNA molecule (Bam 5, using the fragment nomenclature reviewed by Harris, 1989; Fig. 3). Two observations suggest that a single site probably serves to originate replication on each cpDNA molecule: 1) although

Chapter 7 Chloroplast Replication, Recombination and Repair the two D-loops map to the same Bam fragment, few (if any) Bam fragments contained two D-loops at the same time; 2) as depicted in Fig. 4, the two strands of each D-loop have double-stranded regions at opposite ends, indicating that individual origins initiate bidirectional replication. These observations suggest that initiation of replication in the Chlamydomonas chloroplast may differ from initiation in higher plants, where two origins are thought to be required in order to accomplish bidirectional replication (Kolodner and Tewari, 1975). The initial effort to define more precisely the location of oriA used Southern hybridizations and heteroduplex analyses, with cloned fragments of cpDNA from C. reinhardtii and strain WXM, with the reasoning that a locus as important as the origin ofreplication would be highly conserved among different strains of Chlamydomonas (Wang et al., 1984). However, the homologies thus indicated were probably due to the presence of ribosomal protein genes that were subsequently identified near the most frequent origin of replication, oriA (Wu et al., 1986; Lou et al., 1987), since conserved regions in eubacterial origins are composed of A-T rich motifs of only 11–16 bp (Kornberg and Baker, 1992). Although no further investigations were pursued with strain WXM, oriA of C. reinhardtii was characterized extensively by Wu and colleagues. Cloned DNA fragments containing oriA were three to eight times more active than the control vector sequences in directing incorporation of radioactivelylabeled dTTP when used for in vitro replication studies (Wu et al., 1986; Hsieh et al., 1991). Two ‘back-to-back’ prokaryotic promoters were identified and proposed to function as sites ofprimer synthesis for replication initiation (Wu et al., 1986; Lou et al., 1987; Nie et al., 1987). As Wu and coworkers dissected components involved in cpDNA replication, they found that a 500-bp fragment adjacent to the oriA promoters specifically bound polypeptides of 18, 24, and 26 kDa (Nie et al., 1987). The 18-kDa protein was deduced to be encoded by a cpDNA gene, since incubation of cells in chloramphenicol specifically reduced its abundance. The 18 kDa protein was isolated and purified by Wu et al. (1989) for Nterminal amino acid sequencing, with the surprising result that the protein appeared to be the product of the ndhI (frxB) gene, which encodes an iron-sulfur protein that is a component of a thylakoid-localized NADH dehydrogenase. The amino acid sequence information has enabled degenerate oligonucleotides

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to be constructed as probes, but the ndhI gene has not been located on the cpDNA of Chlamydomonas (J. D. Rochaix, personal communication), suggesting that the protein may be encoded in the nucleus, in spite of the chloramphenicol results. Wu and colleagues speculated that the protein might serve a dual function by responding to redox levels with a conformational change that would affect the protein complex at the origin and thus alter initiation of replication (Wu et al., 1989, 1993). Subsequent immunological analyses showed that the ndhI gene product was located in one or more membrane vesicles near the pyrenoid and closely apposed to a subset of chloroplast nucleoids (Zhang and Wu, 1993). Because a charge differential exists across the vesicular membrane (transmembrane potential), the vesicles probably contain other components of photosynthetic electron transfer. The abundance and distribution of the 18-kD protein generally paralleled that of the nucleoids: increasing in the light, decreasing in the dark, decreasing when cells were grown in the presence of 5-fluoro-deoxyuridine (FdUrd), and chloroplasts during becoming greatly reduced in gamete fusion. The one exception noted by Wu and Zhang was that gametes ofboth mating-types seemed to contain more of the protein than did vegetative cells, whereas cpDNA abundance and nucleoid number have been found to decrease in gametes (Chiang, 1971; Sears et al., 1980; Turmel et al., 1980). In vitro characterizations by Wu and coworkers delimited oriA on the Bam 5 fragment to a 224-bp segment that contains the two promoters and a portion ofthe fragment that shows protein-binding properties

122 (Hsieh et al., 1991). However, others found that the 224-bp oriA segment was unable to sustain independent replication of a small plasmid following biolistic transformation (Suzuki et al., 1997). In contrast, the Bam 10 fragment (which contains the atpB gene and neither oriA nor oriB) becomes highly amplified when a plasmid containing it is transformed into the chloroplast (Boynton et al., 1988; Kindle et al., 1994; Suzuki et al., 1997). The amplifications were found as extrachromosomal tandem arrays, but always in conjunction with rearrangements in the cpDNA, and with the apparent prerequisite that an integrated copy be present (Suzuki et al., 1997). These observations led Suzuki et al. (1997) to propose that the Bam 10 plasmid did not replicate autono­ mously, but rather that tandem arrays of the plasmid were generated after a single cross-over event during transformation created a third copy of the atpB gene and a portion of the inverted repeat in the same orientation as the neighboring segment. After initiation ofreplication of the cpDNA, recombination between the unreplicated third copy and a replicated original copy could occur, leading to rolling circle replication of the segment of the cpDNA delimited by these two copies. A second crossover event could terminate replication or result in the excision of the tandem repeats. Although complex, this model explains nicely the otherwise perplexing observations that the Bam 10 plasmid is recovered as an extra­ chromosomal amplicon in transformation experi­ ments. However, the model is not confirmed by the results of Dürrenberger et al. (1996), who used the Bam 10 fragment as the insertion site for a third copy of a portion of the 23S rRNA gene, in both direct and inverted orientations. As discussed in more detail in Section III.F, the presence of a third copy in direct orientation led to deletions between the two closest copies, without the appearance of extrachromosomal amplifications. This suggests that an attribute intrinsic to the Bam 10 fragment itself may be responsible for the generation of the extrachromosomal arrays. The possibility should be considered that additional replication origins exist on the cpDNA, including the 8-kb Bam 10 fragment, since the original D-loop studies of Waddell et al. (1984) reported that replication bubbles in Bam HI fragments of about 9 and 13-kb occurred at a similar frequency to those denoted as oriB in the 17-kb Bam 5 fragment. Moreover, Mosig and coworkers discovered that a third site becomes active in initiating cpDNA replication when the chloroplast gyrase is inhibited

Barbara B . Sears by novobiocin (Woefle et al., 1993). This cryptic ‘origin’ maps in or near a recombination hotspot characterized by Newman et al. (1992), and initiates unidirectional replication, possibly from recom­ bination intermediates.

E. Enzymes of cpDNA Replication Three research groups have undertaken the isolation and characterization of DNA polymerases from Chlamydomonas reinhardtii, and two of these have focused specifically on characterizing the chloroplast enzyme(s). The initial studies by Ross and Harris (1978a, 1978b) characterized three distinguishable DNA polymerase activities fractionated from sonicated cell extracts as complexes of about 100­ kDa and 200-kDa. Keller and Ho (1981) isolated chloroplasts from a cell wall-less strain, CW-15, and showed that radioactive deoxyribonucleotides were incorporated primarily into the DNA. From such a chloroplast fraction, a 180-kDa protein complex having DNA polymerase activity was isolated. The major component was composed of protein(s) of about 40-kDa, and minor components of 15- and 75-kDa were present. In the same study, another DNA polymerase activity was identified, which may represent the nuclear enzyme. Replication in the isolated chloroplasts was inhibited by low concentrations of ethidium bromide, which had been shown by Flechtner and Sager (1973) to inhibit cpDNA replication in Chlamydomonas, and by nalidixic acid, which inhibits prokaryotic gyrases. Keller and Ho concluded that the replication complex must be anchored in the thylakoid membrane since many chloroplasts were stripped of their envelopes during the isolation procedures. Their preliminary characterizations showed that a DNase activity was purified along with the DNA polymerase, and the 40­ kDa protein with exonuclease activity was proposed to provide proof-reading ability to the cpDNA polymerase. In contrast, Wu and coworkers isolated DNA polymerase activity after breaking open wild-type cells by sonication. Initial investigations used both a thylakoid membrane fraction and soluble proteins (Wu et al., 1986), but later investigations were limited to soluble proteins, since that fraction was the source of the polymerase activity (Wang et al., 1991). Extensive purification yielded a DNA polymerase activity composed of proteins of 116 and 80 kDa, sizes similar to proteins composing the animal

Chapter 7

Chloroplast Replication, Recombination and Repair

mitochondrial polymerase (Wang et al., 1991). Similar to enzymes from chloroplasts of higher plants, the Chlamydomonas DNA polymerase showed a preference for poly (dA).(dT) over calf thymus DNA, and was inhibited by N-ethylmaleimide. The ability to synthesize DNA was sensitive to ethidium bromide, and was not inhibited by aphidicolin, thus resembling the chloroplast enzyme from higher plants (Sala et al., 1980; McKown and Tewari, 1984). Since the DNA polymerase characterized by Wu and coworkers is clearly different from that characterized by Keller and colleagues from isolated chloroplasts, it is conceivable that one of those polymerases may actually be the mitochondrial enzyme, particularly since Chlamydomonas mitochondria are highly sensitive to ethidium bromide mutagenesis (Alex­ ander et al., 1974; Gillham et al., 1987a). Another possibility is that multiple DNA polymerases may function in cpDNA replication and repair, as is the case in eubacteria and mammalian mitochondria. Investigations by Mosig and colleagues have focused on enzymes that affect the superhelicity of cpDNA. In vitro studies have shown that both a gyrase activity (topoisomerase II) and a relaxing activity (topoisomerase I) can be isolated from Chlamydomonas cells, and in vivo studies have shown that the gyrase inhibitors, novobiocin and nalidixic acid, differentially affect synthesis of several chloroplast transcripts (Thompson and Mosig, 1985). Chloroplast DNA replication is also inhibited soon after the addition of novobiocin, which suggests that DNA gyrase activity is required for normal cpDNA synthesis (Woelfle et al., 1993). Other investigations assessed plastome structure in light- and dark-grown cells by measuring the ability of a psoralen compound to intercalate into cpDNA (Thompson and Mosig, 1990). Cells in the light phase yielded a cpDNA that bound less psoralen than did cpDNA from cells in the dark phase, indicating that in the light, the cpDNA either had less superhelicity or more proteins associated with it, which consequently limited its accessibility.

III. Recombination

A. Genetic Analysis of Recombination In sexual crosses of Chlamydomonas cellular fusion of the gametes is followed by nuclear and then chloroplast fusion (Cavalier-Smith, 1970). Thus, the

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cpDNAs have an opportunity to recombine, but this can occur only if cpDNAs from both parents are able to persist in the zygote (biparental zygotes). Consequently, the likelihood of recovering recom­ binant cpDNAs is highest if the frequency of biparental chloroplast gene transmission is high. In hybrid backcrosses following interspecific crosses of C. eugametos × C. moewusii, high frequencies of biparental inheritance of chloroplast markers are observed (Lemieux el al., 1980, 1981; Lee and Lemieux, 1986). The zygotes from the interspecific cross have low viability, producing only about 10% viable progeny (Gowans, 1963), however, the progeny from repeated backcrossing show improved zygote viability (Lee and Lemieux, 1990), and high levels of biparental transmission are still observed (Bussières et al., 1996). In C. reinhardtii, spontaneous recombinants are rare because the chloroplast genes are inherited predominantly from the maternal parent, but exogenous chemicals and gametes prior to irradiation can be applied to the mating to increase the frequency of biparental inheritance (reviewed by Harris, 1989; Sears and VanWinkle-Swift, 1994; Chapter 6, Armbrust).

1. Assessment of Chloroplast Gene Recombination Frequencies in Meiotic Progeny Sager and coworkers were the first to map chloroplast genetic distances by assessing recombination frequencies between markers that showed nonMendelian inheritance in C. reinhardtii (Sager and Ramanis, 1968, 1976). Laborious microdissections of tetrad and octad meiotic products were performed, and the progeny were allowed to produce colonies that were then scored for the presence of both nuclear and chloroplast markers. Because only the first three to four cytoplasmic divisions of the zygote were analyzed, a recombinant molecule had to sort out during the first few cell divisions of the zygote, if the recombination event was to be scored. The cumulative data allowed the markers to be mapped relative to each other, and the initial maps coincided reasonably well with results obtained subsequently by Gillham and Boynton and their coworkers (reviewed by Harris et al., 1989). However, Sager’s genetic data were subjected to a complex mathematical analysis to create a circular map (Singer et al., 1976), and a membrane attachment site was proposed as an anchor point for the genetic map and for the plastid chromosome. The circular diagram was an attractive

124 model because the cpDNA had been shown to be a circular molecule (Behn and Herrmann, 1977). However, as pointed out by Harris (1989), the antibiotic resistance markers, which span about half of the circle of Singer et al. (1976), are in fact located within a 21-kb stretch on the 200-kb cpDNA molecule. Their order on the circular map does not correspond well to the locations that subsequently have been shown to be the sites of mutation in the physical map of the cpDNA (Harris et al., 1989). When the zygotes fulfill the essential condition for recombination analysis of containing chloroplast markers from both parents, one might expect that most of the meiotic progeny would be heteroplasmic (containing a mixture of chloroplast alleles from both parents), and therefore be useless for analyzing recombination. Surprisingly, among the 8-cell progeny resulting from meiosis followed by a single mitotic division, the majority of cells are homoplasmic for at least one locus (Sager and Ramanis, 1968, 1976; Forster et al., 1980). The rapid segregation is probably due to many factors, including the reduction in cpDNA copy number in gametes and possibly in zygotes, destruction ofmost ofthe copies of the plastomes, internal chloroplast membrane structure acting as a barrier to plastome mixing (Mets and Geist, 1983) and the likelihood that nucleoids, rather than individual cpDNA molecules, are the functional genetic units (VanWinkle-Swift and Birky, 1978). Gene conversion may also act to reduce heteroplasmicity, with stochastic events resulting in genetic drift within the population of organelle DNA molecules (Birky and Skavaril, 1976).

2. Assessment of Chloroplast Gene Recom­ bination Frequencies Through the Analysis of Biparental Zygospore Clones In contrast to the analysis of meiotic progeny, the biparental zygospore clone analysis of Gillham, Boynton, Harris, and colleagues assesses chloroplast gene recombination long after it has actually occurred. The procedure is analogous to the assessment of recombination frequencies in phage ‘crosses’, where bacteria are subject to a mixed infection, and recombinant phage are recognized when the progeny phage are diluted for subsequent single infections (Gillham et al., 1974). For the Chlamydomonas analysis, zygospores are allowed to sporulate and form colonies, which are then replica-plated to distinguish those in which chloroplast genes were

Barbara B . Sears inherited from both parents. Fifty or more biparental zygote colonies are selected and taken through dilution-plating on non-selective media so that individual cells give rise to colonies. From each biparental zygote dilution plate, 64 subclones are then routinely transferred to an 8 × 8 grid, replicaplated and scored for chloroplast marker composition. To calculate recombination frequency between two markers, the denominator is taken as the number of total subclones tested, and the numerator is the sum of the number of subclones that displayed a nonparental combination of the two markers. Percentage recombination is then converted to map units. Figure 5 shows a genetic map that was composed from these data. As this figure shows, for the antibiotic-resistance markers in genes that encode components ofthe chloroplast ribosome, the physical map aligns well with the genetic map, wi th approximately 1% recombination per 1000 bases. However, when crosses include other markers, such as those in the nearby psbA gene, recombination occurs at a high frequency between them and the ribosomal antibiotic-resistance markers, making them appear unlinked, since they approach the theoretical limit of 25% recombination according to the phage model, when multiple rounds of recombination and replication occur prior to sampling (Harris et al., 1989). Subsequent studies have shown that a recombination hotspot lies between the psbA and 23S rRNA genes (Newman et al., 1992), as further described in Section III.D. A disadvantage of assessing recombination frequencies by subcloning biparental zygospore clones is that early recombination events are amplified in this scoring system. Although it is true that early events may be multiply counted, the likely occurrence of any recombination event should still be distancedependent. If large numbers of biparental zygospore clones are analyzed, sampling errors should diminish as the recovery ofearly and late recombination events balance each other. One other comment should be made about the analogy between this method of analysis and determination of recombination frequency in phage crosses. Phage crosses are performed with equal titers from the two phage to accomplish double infections, whereas in C. rein­ hardtii, a maternal bias in the contribution of chloroplast markers is usually reflected in the composition of the biparental zygospore colonies (reviewed by Harris et al., 1989).

Chapter 7

Chloroplast Replication, Recombination and Repair

3. Assessment of Chloroplast Gene Recom­ bination Frequencies After Paternal Marker Selection Mets recognized that zygospore clone analysis tended to overcount early recombination events, and proposed an alternative means of mapping chloroplast gene recombination frequencies (Mets and Geist, 1983). The paternal marker selection procedure begins with zygospore colonies, which are scored by replica-plating. From each biparental zygospore colony, a single subclone is initially chosen based on its growth on medium that selects for one of the paternal markers. That subclone is then scored for the presence of other markers. This procedure is modeled after techniques for mapping of bacterial genes by co-transformation or co-transduction, where the denominator is also defined by the selected marker. An additional goal of the procedure was to establish a method that required analysis of fewer progeny, so that it could be used for the assessment of RFLP inheritance in crosses. Since the initial selection for one paternal marker eliminates from consideration all of the non-recombinant maternal chloroplast molecules, recombination events compose a larger

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fraction of the progeny analyzed. Consequently, map distances are larger when assessed in this way. Because only a single product is scored from each biparental zygospore colony, a recombination event that occurs early in a single zygospore has a chance to be counted only once, rather than multiple times. However, recombinant molecules that have the maternal marker at the chosen paternal locus are excluded from consideration. Thus, although this procedure accomplishes its goal of removing a bias due to early recombination events, it has the disadvantage of excluding a subset of the recom­ bination events from consideration. For their analysis, Mets and Geist (1983) used erythromycin- and streptomycin-resistance markers that had been included in the analyses of others, a newly isolated rbcL mutation that results in acetatedependence (A( 10-6C)), a psbA mutation that results in a photosystem II deficiency, and a mutation in the psbA gene that confers resistance to DCMU. Using the paternal marker selection procedure in two crosses, the recombination frequencies did not give map distances that were additive for a linear map, and they also could not be assembled into a clear circular map. In comparing the disparate recom­

126 bination frequencies of their two crosses, Mets and Geist noted that one cross had a much lower recombination frequency than did the other cross. In the first cross, where maternal and paternal plastomes were more abundant among the progeny than were recombinants, Mets and Geist proposed that the normal, internal architecture of the chloroplasts in those strains could have limited the extent of mixing of the plastomes, and thus recombination may have been minimal. For the other cross, more recombinants were recovered than parental plastome types, and Mets and Geist suggested that the disruption of normal thylakoid stacking in the photosystem IIcells would have allowed more mixing deficient of the plastomes to occur in the zygotes. In the latter cross, the cpDNA appeared to have served as the resident DNA molecule, which incorporated segments of a fragmented cpDNA. It was further noted that if one or two of the genetic markers were located in one or both of the single copy regions of cpDNA, and one or more markers were present in the inverted repeats of the chloroplast DNA, it could be difficult to assemble a genetic map. In fact, subsequent cpDNA sequencing showed that those hesitations were well-founded, since rbcL gene was mapped to a single-copy region, and the other three markers were located within the inverted repeat.

4. Assessment of Recombination Frequencies in Vegetative Diploid Zygotes As summarized in the chapter by Armbrust, vegetative diploid zygotes can be selected using appropriate markers, and they transmit chloroplast markers from both parents at a high frequency (Gillham, 1963, 1969; VanWinkkle-Swift, 1978). An analysis of recombination frequencies in vegetative zygotes derived from three-marker crosses of C. reinhardtii showed no consistent correlation in the recovery of reciprocal recombinant progeny within a population of zygotes, nor within individual zygote clones (VanWinkle-Swift and Birky, 1978). In fact these observations provided the basis for an extensive comparison highlighting the non-reciprocality of organelle gene recombination in Chlamydomonas and yeast. In spite of these findings, Girard-Bascou found that she could use the production of wild-type recombinants in vegetative zygotes to estimate map distance between different mutations affecting photosynthesis (Girard-Bascou, 1987). The wildtype recombinants were recognized by their low

Barbara B . Sears fluorescence or by their ability to grow on minimal media. The recombination frequency between each pair of mutations was calculated to be twice the frequency of the wild-type recombinants, divided by a value representing the differential growth rate of the wild-type recombinants. Girard-Bascou used this procedure to distinguish five loci among the eight mutations analyzed (Girard-Bascou, 1987), with the two closest loci lying in one of the single copy regions of cpDNA, separated by 1–4% recombination and, 1910 bp (Girard-Bascou et al., 1987).

B. Factors that Affect Recombination Frequency The most important parameter for homologous recombination is the extent of sequence homology between the two DNAs. The transformation studies of Newman et al. (1990, 1992) and Suzuki et al. (1997) indicate that high levels of homologous recombination in the chloroplast of C. reinhardtii require at least 150–200-bp sequence homology. Most likely, DNAs with smaller segments of homology are also capable of recombination, but at lower frequencies, since eubacterial recombination can involve sites with as little as 25-bp homology (Allgood and Silhavy, 1988). Studies of recombination frequencies in crosses have shown that increasing levels of UV-irradiation applied to gametes raise the proportion of zygospores in which the progeny inherit chloroplast alleles from both parents, but numbers and types of recombinants recovered from each biparental zygospore do not change markedly (Gillham et al., 1974; Sager and Ramanis, 1976b). In contrast, when a single population of C. reinhardtii zygospores was sampled and induced to germinate after longer and longer periods of dormancy, recombination frequency and map distances increased over time (Sears, 1980a). During the extended dormancy, the overall frequency of biparental zygospores actually dropped, while the proportion of both maternal and paternal zygospores increased (Sears, 1980b; Rosen et al., 1991). These results suggest that at least part of the cpDNA can persist in the zygospore for reasonably long periods of time and eventually recombine. Further­ more, in terms of cpDNA metabolism, the zygospore is not in fact quiescent.

C. Recombination Within the Inverted Repeat As has been observed in higher plants, homologous

Chapter 7

Chloroplast Replication, Recombination and Repair

intramolecular recombination occurs between the two copies of the large inverted repeat, resulting in two isomers of cpDNA that have the two single copy regions in opposite orientations (Aldrich et al., 1985; Palmer et al., 1985). To determine if ‘flip-flop’ recombination is a frequent event, Aldrich et al. (1985) assessed the arrangement of the cpDNA isolated from three liquid cultures that had been derived from single vegetative cells. They reasoned that if flip-flop recombination of the cpDNA is infrequent, a single cell and the clonal line derived from it would have predominantly a single isomer. In fact, near-equal stoichiometry of the two arrange­ ments was demonstrated for all three cultures by Southern hybridizations. These studies cannot formally rule out the possibility that a sorting process exists that preserves equimolar amounts of the isomeric genomes from one cell generation to the next, but it is an unlikely explanation in light of other observations of vegetative segregation. Aldrich and colleagues also concluded that cpDNA recombination is not limited to the sexual cycle since their DNA was isolated from vegetative cell cultures. The fact that both copies of the inverted repeat are identical indicates that copy correction must occur between the two repeats. The existence of a copy correction process has been convincingly demon­ strated through the isolation of many symmetrical deletions in the two repeats (Myers et al., 1982), and the observation that transformational replacement within the inverted repeat is never recovered in a single copy (Boynton and Gillham, 1993). Myers et al. (1982) suggested that when the helix is broken within the inverted repeat, one copy of the inverted repeat will serve as the template for recombinational repair, with the end result that both copies will be identical. The occasional instances in which non­ identical copies of the inverted repeat have been recovered are those in which a large deletion extends into a single copy region ofcpDNA and/or an essential gene has been deleted from one copy of the inverted repeat (e.g., Myers et al., 1982; Dürrenberger et al., 1996). In their consideration of other recombination events involving the inverted repeat, Boynton et al. (1992) have posited that flip-flop recombination and copy correction can be viewed as ‘different manifestations of the same mechanism.’

D. Recombination Hotspots Interspecific crosses between C. moewusii and

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C. eugametos (and their or progeny) have shown that recombination occurs more frequently within the inverted repeat than between single copy regions of the cpDNA (Lemieux et al., 1990). The high recombination frequency of the inverted repeat is observed even when the homing introns that are discussed later are neutralized because they are present in the cpDNAs of both parents. The five-fold difference in recombination frequencies of the different segments of cpDNA was proposed to be due to the potential of the two copies of the inverted repeat to recombine with each other, after a donor parent recombinationally allele from the opposite replaces one copy of the allele present in the resident cpDNA. In C. reinhardtii, Newman et al. (1992) found that different intervals within the inverted repeat displayed markedly different levels of recombination. In their investigation, three-point crosses were performed with C. reinhardtii and C. smithii (UTEX strain 1062), which have extensive restriction fragment length polymorphisms (RFLPs), but may in fact be more appropriately classified as a single species (A. Coleman, personal communication.). Through biparental zygospore analysis of genetic recombination frequencies, a hotspot for recombination was identified near the 3´-end of the psbA gene. The hotspot was localized to a 500-bp segment by quantifying preferred sites of homologous recombination in transformation experiments. The hotspot contains an AT-rich segment, but does not house any of the short repeats (Schneider et al., 1985; Rochaix, 1978; Gelvin and Howell, 1979) that were previously hypothesized to play a role in recombination (Palmer et al., 1985; Boynton et al., 1992).

E. Gene Conversion Gene conversion events have been invoked to explain the observations that both copies of the large inverted repeat are always identical. As elaborated in Section III.C, even if alterations are introduced at a single site by mutation or transformation, a ‘copy correction’ process acts quickly to create a mirror image in the second copy of the inverted repeat. Assuming that strand exchange between the repeats results in a heteroduplex, mismatch repair is probably the process that acts to ‘restore’ perfect pairing, resulting in elimination or fixation of a new allele. Gene conversion between molecules could lead to genetic

Barbara B . Sears

128 drift in the population of cpDNA molecules within an individual chloroplast (Birky and Skavaril, 1976), while vegetative segregation would lead to homo­ plasmicity as the chloroplasts and cells divide. No matter what type of analysis has been used to measure frequencies of recombinant progeny from genetic crosses, a disparity has been found in the recovery of reciprocal recombination products (e.g., Gillham, 1965a; Gillham et al., 1974; Boynton et al., 1976; Sager and Ramanis, 1976a; VanWinkle-Swift and Birky, 1978; Forster et al., 1980; Mets and Geist, 1983; Harris et al., 1989). The production of non­ reciprocal recombination products is probably due to biased or directional gene conversion events (Birky and Skavaril, 1976). However, the likelihood that both reciprocal and nonreciprocal recombination events occur has been discussed by VanWinkle-Swift and Birky (1978), who pointed out that multiple exchange events occurring within the organelle may act to obscure the nature of the original event. This article should be consulted by anyone seriously interested in organelle recombination processes, because it considers all aspects that can complicate the recognition and scoring of recombinant progeny, including intercellular growth differences, sample size, and intracellular selection. In addition to these considerations, Galloway and Holden (1985) have pointed out that the occurrence of different recombination products within a single zygospore clone may result coincidentally because circum­ stances such as cytoplasmic mixing, which are necessary for the production of one recombinant type, may also lead to the occurrence of another. Interspecific crosses between C. moewusii and C. eugametos have provided much valuable information about the process of gene conversion. Since these species have many differences in cpDNA physical markers (summarized by Turmel et al., 1987), RFLPs spanning the entire plastome can be examined in recombinant progeny. A subset of progeny were chosen for analysis through paternal marker selection from reciprocal crosses. An initial study ofantibiotic­ resistance genes and physical markers in and near the rRNA operon showed that regardless of the direction of the cross, 100% of the progeny inherited an RFLP in the 23S rRNA of C. eugametos, which was subsequently shown to represent a 955-bp intron (Lemieux and Lee, 1987; Turmel et al., 1991). These observations were noted to resemble the preferential inheritance of the yeast omega factor, which is an intron in the mitochondrial large rRNA gene

(reviewed by Dujon, 1989), where adjacent markers also show a ‘polarity’ of preferential inheritance. Two additional sites also are inherited preferentially by the progeny, and represent insertions of 21-kb in the inverted repeat and 6-kb in one of the single-copy segments of the cpDNA of C. moewusii relative to C. eugametos, (Lemieux et al., 1988,1990;Bussières et al., 1996). As depicted in Fig. 6, most loci are inherited from both parents, but the three sites denoted as c, g, and r (23S rRNA intron, 21-kb segment, and 6-kb segment respectively) show unidirectional inher­ itance. In the case of the 23S rRNA intron and the 21­ kb insert, adjacent markers are preferentially coinherited, but show lesser degrees of polarity (Lemieux and Lee, 1987; Lemieux et al., 1990; Bussières et al., 1996).

F. Intron Homing As reviewed above, C. Lemieux, Turmel and colleagues had observed directional gene conversion involving the intron in the large subunit rRNA gene in progeny of interspecific crosses of C. moewusii and C. eugametos. Those results led them to hypothesize that intron transposition might be mediated by an endonuclease encoded by the 218­ codon open reading frame (ORF) found within the intron (Gauthier et al., 1991; Turmel et al., 1991). Expression of the ORF in E. coli showed that its polypeptide product was required to linearize a plasmid carrying the C. moewusii sequence that lacks the intron (Gauthier et al., 1991). Subsequent investigations characterized the recognition site of the endonuclease to a 15–19-bp non-symmetric, degenerate sequence, with cleavage producing a 4­ base staggered cut at the site of insertion (Marshall and Lemieux, 1991, 1992). These investigations showed that the C. eugametos intron was auto­ mobile, and should be classified as a ‘homing intron’, since its target is the exact site at which the intron inserts into the gene (Dujon et al., 1989). Following the convention for nomenclature (Dujon et al., 1989), the endonuclease encoded by the C. eugametos 23S rRNA intron has been named I-CeuI, for Intronencoded homing endonuclease from C. eugametos (Gauthier et al., 1991). Turmel, C. Lemieux and colleagues have performed similar characterizations on related mobile introns in the 23S rRNA gene of C. humicola (Côté et al., 1993) and C. pallido­ stigmatica (Turmel et al., 1995).

Chapter 7

Chloroplast Replication, Recombination and Repair

The 23S rRNA gene of Chlamydomonas reinhardtii also contains an intron that has an open reading frame. The 163-codon ORF has some resemblance to genes for mitochondrial maturases (Rochaix et al., 1985), however the polypeptide product has no intronprocessing activity (Thompson and Herrin, 1991). Consequently, Dürrenberger and Rochaix (1991) assessed whether the polypeptide product of the ORF was involved in intron mobility. By placing the ORF under the control of an IPTG-inducible promoter, they showed that when the protein is expressed in E. coli, it specifically cleaves a plasmid carrying a cDNA copy of about 620-bp of the 23S rRNA gene, including the splice junction that remains after intron removal from the RNA. The site of double-strand cleavage was at or near the exon-exon junction. This approximate location was more precisely defined by Thompson et al. (1992) and Dürrenberger and Rochaix (1993) to 19–20-bp spanning the exon junction. As is the case with ICeuI, endonucleolytic cleavage was found to leave 4-base staggered 3´-overhangs, with a 3´-OH and a (Thompson et al., 1992). The C. reinhardtii homing endonuclease is known as I-CreI. In their first investigation, Dürrenberger and Rochaix (1991) attempted to examine the endo­ nucleolytic targeting by transformation of C. rein­ hardtii chloroplasts with a copy of the 23S cDNA segment located ectopically on the cpDNA. However, all of the transformants that had comprehensible integration patterns contained copies of the cDNA segment in which the intron already had transposed from the resident 23S rRNA gene! A subsequent collaborative investigation between Rochaix, Herrin, and their coworkers overcame this obstacle through the construction of a strain in which the intron was present in the resident 23S rRNA gene, but the reading frame was disrupted (Dürrenberger et al., 1996). The deletion that disrupted the reading frame also reduced the size of the intron, enabling the copy to be distinguished from the wild-type copy that subsequently would be introduced. In the strain and its derivatives, the 620-bp cDNA segment of the 23S rRNA was introduced by transformation into a site in a single-copy region of the nearby Bam 10 fragment. I-CreI activity was constructs with brought in later by mating the wild-type strains. In those crosses, specific cleavage at the CreI recognition site in the 23S cDNA segment was seen 1.5–2 h after mating. The investigation of Dürrenberger et al. (1996) led

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to a surprising rinding when meiotic progeny were analyzed to assess the frequency of intron movement. In the crosses described above, different results were obtained depending on whether the cDNA copy was placed in an inverted or direct orientation relative to the intron-containing 23S rRNA gene. When the cDNA homing site was initially located in an inverted orientation, about half of the meiotic progeny analyzed (from four out of seven zygospores) intron in its original position contained the as well as in the cDNA site. Thus, the intron had successfully moved into the cDNA target site through the endonuclease activity provided in trans. In both this cross and a control cross between two parents, a similar fraction of the progeny (20–25%) contained cpDNA in which the ectopic cDNA copy was eliminated by replacement with the wild-type cpDNA, indicating that sizable segment from the cpDNA are able to survive gamete pieces of the fusion and be recombinationally active, even in the absence of the endonuclease. In contrast, when the cDNA homing site was initially located in a direct orientation, the only products recovered contained deletions that had occurred between the 23S rRNA intron and the nearby gene containing the cDNA of the 23S rRNA gene. Since the deletion removed the intervening 16S rRNA gene in that copy of the inverted repeat, the deletion product certainly would not have had a selective advantage, and yet all of the progeny contained the deletion. Thus, rather than mobilizing the intron, the double-stranded DNA cleavage caused by the CreI enzyme appeared to initiate recombination between the cut site and the closest region of homology. Several optional cpDNA elements are candidates to contain or be located next to homing introns, including the 21-kb and 6-kb segments of C. moewusii cpDNA represented as RFLPs ‘g’ and ‘r’ in Fig. 6 (Bussières et al., 1996), and the C. reinhardtii ‘Wendy’ element characterized by Mosig and coworkers (Fan et al., 1995). The 21-kb segment appears to be propelled into C. eugametos cpDNA due to its linkage to a homing intron in the nearby psbA gene of C. moewusii. The gene conversion events associated with the 6-kb segment have a less clear basis, although the recovery of cpDNA linearized at the right end of the 6-kb segment indicates that an endonucleolytic target is present (M. Turmel, personal commun­ ication). The Wendy element contains ORFs with some homology to transposases and homing endonucleases. Since it is absent from the cpDNA of

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C. smithii, an assessment of its presence in the progeny of reciprocal crosses should indicate whether it has the properties of a homing intron.

IV. Repair

A. UV-Damage and Photoreactivation Ultraviolet irradiation causes bonds to form between adjacent pyrimidines on the same strand of a DNA helix (Kornberg and Baker, 1992); these pyrimidine dimers are mutagenic because they cannot be ‘read’ by DNA polymerase. Analogous to other biological systems, the Chlamydomonas chloroplast genetic system can respond to UV-damage through both a light-dependent photolyase-mediated repair and a dark repair pathway (Small, 1987), although the enzymes have not yet been isolated. The existence of

Barbara B . Sears

a photolyase was initially suggested by the observation that the impact of UV on chloroplast gene inheritance is greatly reduced if irradiated gametes are exposed to light prior to mating (Sager and Ramanis, 1967). The photoreactivation process was studied in detail by Small and coworkers, as elaborated below. Initial biochemical characterizations failed to find evidence for thymine dimer excision in the dark (Swinton and Hanawalt, 1973a,b), but subsequent investigations by Small and colleagues documented the dark repair of thymine dimers in cpDNA by using lower fluences of ultraviolet radiation and by utilizing a dimer-specific endonuclease from Micrococcus luteus to quantify pyrimidine dimers in the cpDNA (reviewed by Small, 1987). Small and Greimann and exposed them (1977) grew cells in to UV followed by a dark or light period. The DNA was extracted and half of it was treated

Chapter 7

Chloroplast Replication, Recombination and Repair

with the dimer-specific nuclease, while the other untreated halfserved as a control. When more dimers were present, the nuclease cleaved the cpDNA into smaller fragments. The degree of fragmentation was assessed in alkaline sucrose gradients, where smaller molecules do not sediment as readily as do larger ones. The distribution of the tritium label in the gradients thus indicated the prevalence of thymine dimers at each time point. After the cells were exposed to 90 min of light, their cpDNA appeared to be completely normal, whereas little repair occurred in the dark during the corresponding amount of time. After 24 h in the dark, thymine dimers were diminished to 15% of their initial abundance. Although the evidence is not conclusive, it suggests the existence of nucleotide excision repair and/or recombinational repair in chloroplasts. Small and colleagues also characterized several UV-sensitive mutants of C. reinhardtii, but found that their repair defects were specific for nuclear DNA damage (summarized by Small, 1987). However, Rosen et al. (1991) found that one UVsensitive mutant, uvsE1, increased the frequency of transmission of paternal chloroplast markers in sexual crosses, but only if the allele was present in both the and parents. The uvs-E1 allele interacts synergistically with the mat-3 nuclear allele (Gillham et al., 1987b), in promoting transmission of chloroplast genes from the paternal parent. As a mutagenic agent, UV has been used by geneticists for the induction of nuclear mutations in Chlamydomonas (Gillham and Levine, 1962; Girard et al., 1980;VanWinkle-Swift and Burrascano, 1983; and others reviewed by Harris, 1989), but only a few plastome mutations have been isolated after UVirradiation in C. reinhardtii (Hudock et al., 1979) and C. monoica (VanWinkle-Swift and Aubert, 1983; Van Winkle-Swift and Thuerauf, 1986). The fact that cpDNA mutations are induced only rarely in UVmutagenesis experiments seems paradoxical to the finding that UV specifically affects chloroplast gene inheritance when gametes are irradiated (Chapter 6, Armbrust). However, as pointed out by Sager and Ramanis (1967), UV applied to gametes may have a minimal impact on nuclear genes because chrom­ osomal DNA replication and meiosis occur many days after the irradiation, with repair being possible during an extended period of time. Furthermore, if UV-damage to the cpDNA leads to recombinational repair, an effect might not be noticed in vegetative

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cultures, since each cell contains a chloroplast with many copies of the identical DNA molecule that can act as the repair template. In contrast, gametes would have fewer copies of the cpDNA, and in the young zygote, a subset ofthese would have been contributed parent. by the

B. Specific Impact of FdUrd on cpDNA The uridine analog 5-fluoro-deoxyuridine has been found to affect specifically the abundance and integrity of cpDNA in C. reinhardtii, reducing the fraction of cpDNA from 8–18% of the total cellular DNA to 1–2% (Wurtz et al., 1977), and also diminishing the number of chloroplast nucleoids (Matagne and Hermesse, 1981; Nakamura and Kuroiwa, 1989). In bacteria, FdUrd binds to and inhibits thymidylate synthetase, the enzyme that converts dUMP to dTMP (reviewed by Kornberg and Baker, 1992). Assuming that the analogous enzyme is its target in Chlamydomonas, it is noteworthy that FdUrd has little mutagenic impact on the plastome until C. reinhardtii cells enter stationary phase or gametogenesis (Wurtz et al., 1979). This observation suggests that the chloroplast thymidylate synthetase is more active in C. reinhardtii when the cells are starved than during logarithmic growth (Sears and VanWinkle-Swift, 1994). This deduction is consistent with activation of the nucleotide salvage pathway during the massive turnover of ribosomes and rRNA that occurs under conditions of nitrogen starvation (Siersma and Chiang, 1971; Martin et al., 1976). An alternative, and notmutually exclusive explanation, is that cpDNA repair is more efficient in exponentially growing cells. FdUrd has been combined effectively with ethylmethanesulfonate (Spreitzer and Mets, 1980), ICR-191 (Harris et al., 1982) and X-irradiation (Myers et al., 1982) in the production of nonMendelian mutations. In contrast, FdUrd induces nuclear mutations in C. monoica (VanWinkle-Swift and Hahn, 1986; VanWinkle-Swift and Thuerauf, 1991), and the nitrogenous base analog, 5-fluorouracyl, has been used to induce nuclear mutations in C. reinhardtii (Girard et al., 1980). As reviewed in the chapter by Armbrust, the impact of FdUrd on cpDNA has also been observed when cells exposed to the analog are crossed to untreated gametes. Those crosses show reduced trans­ mission of chloroplast markers from the maternal

132 parent, with a dramatically increased frequency of exclusively paternal transmission and moderate levels of biparental transmission (Wurtz et al., 1977, 1979). As reviewed in the chapter by Goldschmidt-Clermont, FdUrd has also been used to reduce cpDNA levels for chloroplast transformation experiments.

C. Other Mutagens In higher plants, cpDNA mutations are induced most effectively through the use of nitroso-methyl-urea and nitroso-guanidine (reviewed by Hagemann, 1976; Börner and Sears, 1986), but in Chlamydomonas these agents have been used primarily to target nuclear genes (reviewed by Harris, 1989). The nitroso­ compounds cause mutations because they alkylate nitrogenous bases, particularly guanine residues, which then mispair during replication (Dodson et al., 1982). In bacteria, the primary lesion, can be reversed by a methyltransferase which transfers the offending methyl group to one of its own cysteine residues (Yarosh, 1985). In higher plants, the nucleus most likely contains methyltransferases, while the chloroplast probably lacks these specific repair enzymes. In contrast, Frost and Small (1987) found that methyltransferase activity is completely absent from extracts of C. reinhardtii, suggesting that this particular repair pathway is not present in either the nucleus or chloroplast of Chlamydomonas. The susceptibility of the two genetic systems to alkylation damage is reflected by the induction of both nuclear and plastome mutations by nitrosoguanidine (Gillham, 1965b). Similarly, both 2-amino-3-phenylbutanoic acid and methylmethane­ sulfonate (MMS) have been used to induce nuclear as well as plastome mutations in several species of Chlamydomonas (McBride and McBride, 1975; Hawks and Lee, 1976; Lee and Lemieux, 1986).

D. The Interrelated Processes of Recombination/Repair In a study designed to investigate the molecular mechanisms of homologous recombination, Cerutti et al. (1995) transformed chloroplasts with constructs that introduced a copy of the wild-type E. coli recA gene into the chloroplast genetic system of C. reinhardtii. Cell growth and survival after treatment with DNA damaging agents appeared to be fairly normal in cells expressing the wild-type E. coli RecA, indicating that chloroplast repair processes

Barbara B . Sears were intact. Recombination frequencies were measured by assessing the deletion of a segment between direct repeats bracketing a selectable marker introduced as a disruption of the chlL gene. The chlL gene is required for light-independent chlorophyll synthesis: when the reading frame is intact, Chlamydomonas cells are green in the dark; when the gene is disrupted, cells are yellow in the dark. Dark-grown cells containing the E. coli RecA showed a larger number ofyellow colonies with green sectors than did control cells, indicating that presence of RecA increased the frequency of recombinational deletion of the chlL disruption. Parallel studies with a control construct having a 24-codon internal deletion that abolishes RecA function showed that expression of the functionless protein did not affect the chloroplast genetic system. A third type of construct had up to 42 codons deleted from the N-terminus, which in E. coli drastically reduces the ability of the protein to form oligomers and thus acts as a dominant negative mutation. The N-terminal truncation product reduced the frequency ofrecombination as measured through the chlL assay, reduced the ability of the cells to repair cpDNA damage incurred by methyl methanesulfonate, and decreased the survival ofcells exposed to FdUrd, MMS, or UV-light. The specific decrease in plastid DNA level caused by FdUrd was also more pronounced in transformants with the Nterminal deletion of RecA than in the other genotypes. Although antibodies for E. coli RecA cross-react only slightly with a protein from wild-type Chlamydomonas the functional assays indicate that a similar protein functions in the algal chloroplast system. The observation of active recombination in vegetatively-growing cells led Cerutti et al. (1995) to suggest that the primary biological role of recom­ bination in the plastids is for the repair of cpDNA damage. Although no experiments have directly addressed the question of whether mismatch repair occurs within the Chlamydomonas chloroplast, much of the evidence reviewed here points to its existence. An active mismatch repair system could provide justification for the extensive methylation of cpDNA observed during gametogenesis and in vegetative cells under certain conditions. In E. coli, hetero­ duplexes resulting from misincorporation during replication are corrected by the mismatch repair enzymes (Friedberg et al., 1995). Those enzymes recognize, cleave, and remove the unmethylated strand bracketing the mismatch. Since methylation

Chapter 7 Chloroplast Replication, Recombination and Repair is a post-replication modification, this repair strategy leaves the parental (template) strand intact, while removing the newly synthesized daughter strand which should contain the replicative error. In other biological systems, transformational replacement of alleles and gene conversion involve mismatch repair of heteroduplexes (Friedberg et al., 1995). Analogous genetic observations have been made with Chlamy­ domonas, where non-reciprocal events typify cpDNA recombination and copy correction occurs between the inverted repeats.

V. Perspectives and Conclusions Both structural and functional features of the chloroplast genetic system reflect its evolutionary derivation from an endosymbiotic, prokaryotic ancestor (reviewed by Gillham, 1994). Given this history, many components of the replication, repair, and recombination machinery are likely to resemble their eubacterial counterparts. Such similarities could allow the isolation of genes involved in cpDNA metabolism by screening of Chlamydomonas clone libraries with heterologous probes from cyano­ bacteria. A functional screen could also be undertaken to search for complementation of defined E. coli mutants using a cDNA expression library generated from Chlamydomonas RNA. This approach has been employed successfully for cloning genes involved in recombination/repair from an Arabidopsis expression library (e.g., Pang et al., 1992, 1993a,b). Reverse genetics could then be pursued to assess the functional contribution to cpDNA metabolism of the genes thus identified. Investigations into the processes of replication, recombination, and repair of cpDNA in Chlamy­ domonas have exploited to a very limited extent the ability to isolate mutations in genes that encode products important for these processes. The isolation and analysis ofmutators that target chloroplast genes will enable a genetic dissection of replication and repair, and will also provide a useful resource for biochemical characterizations. The ability to transform both the chloroplast and nuclear genetic compartments in this alga enables foreign genes to be introduced to antagonize or supplement the endogenous genes, and also provides the option of introducing a selectable target for studying recombination or repair. This technological advance in genetic engineering has enhanced the versatility

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of Chlamydomonas as a model experimental organism for studying chloroplast genetic processes.

Acknowledgments The author is grateful for the constructive critiques and suggestions provided by Heriberto Cerutti and Jean-David Rochaix, and the comments and information provided by Annette Coleman, Lib Harris, Karen Kindle, Gisela Mosig, MoniqueTurmel, Karen VanWinkle-Swift, and Madeline Wu during the preparation of this chapter. Contributors of the figures are also gratefully acknowledged.

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Chapter 8 Chloroplast Transformation and Reverse Genetics Michel Goldschmidt-Clermont Departments of Molecular Biology and of Plant Biology, University of Geneva, Sciences II, 30 quai E. Ansermet, 1211 Geneva 4, Switzerland

Summary I. Introduction II. Delivery of DNA to the Chloroplast III. Selectable Markers and Reporters A. Selectable Markers 1. Photosynthesis 2. Drug Resistance B. Reporters and Foreign Gene Expression IV. Fate of Transforming DNA A. Integration in the Chloroplast Genome B. Extrachromosomal DNA C. Homoplasmic and Heteroplasmic Transformants D. Co-Transformation E. Marker Recycling V. Reverse Genetics A. Gene Inactivation B. Site-Directed Mutagenesis VI. Conclusion and Perspective Acknowledgments References

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Summary For chloroplast transformation, the most efficient method to introduce DNA is particle bombardment. To then select the cells which harbor a transformed plastid, two classes of markers are available. With one class, selection is based on the rescue of a non-photosynthetic mutant with the wild-type chloroplast gene. With the other class, selection is based on a mutation or a foreign gene conferring resistance to an antibiotic or a herbicide. Transforming DNA is integrated by homologous recombination, and only in exceptional cases is it maintained extrachromosomally. The modified and wild-type copies of the highly polyploid plastid genome usually segregate rapidly, although in some circumstances a heteroplasmic mixture of genomes is retained. The available technology and markers readily allow chloroplast gene inactivation and site-directed mutagenesis. These possibilities are enhanced by strategies such as co-transformation or the repeated use of unstable markers.

J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria im Chlamydomonas, pp. 139–149. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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I. Introduction

II. Delivery of DNA to the Chloroplast

Stable transformation of the chloroplast involves three main steps: the introduction of DNA into the organelle, the expression of a marker for selection, and the replication of the introduced DNA. The major challenge has been the first step, the delivery of DNA across the cell wall and three membranes: the plasma membrane and the two chloroplast envelopes. The breakthrough for chloroplast transformation came with the development of microprojectile bombardment for DNA delivery (Klein et al., 1987; Boynton et al., 1988). The transforming DNA has to carry a marker that can be selected, or screened for, to identify the transformants. Wild-type copies of photosynthesis genes were used as the original selectable markers, with the corresponding photosynthesis mutants as hosts. Mutations conferring resistance to translation inhibitors or to photosynthesis inhibitors have also proven useful for selection, and engineered bacterial genes have provided further markers and reporters. The stable propagation of the introduced DNA results from its integration into the plastid genome, a step which is facilitated by the recombination machinery active in the chloroplast. The chloroplast genome is present in approximately eighty copies in the single chloroplast of C. reinhardtii. The high level of ploidy implies that in the initial stages after transformation, the plastid is heteroplasmic: it contains both wildtype and modified copies of the genome. This may hamper the expression of recessive markers, but the problem is alleviated by the rapid segregation that usually occurs. With the appropriate tools available, targeted gene inactivation and site-directed muta­ genesis of chloroplast genes have become routine experiments. In a very short time, an extraordinary amount of knowledge has been gained using this technology, mostly in the fields of photosynthesis and of chloroplast gene expression. The pioneering work on chloroplast transformation in Chlamy­ domonas has established strategies that have also been applied fruitfully for the transformation of tobacco plastids (Svab et al., 1990; Svab and Maliga, 1993).

Ten years after its original description, ‘biolistic’ bombardment with DNA-coated microprojectiles remains the method of choice for chloroplast transformation (Boynton et al., 1988). The alternative is vortexing in the presence of glass beads (Kindle et al., 1991), but this is less efficient and requires cellwall deficient algae, either following treatment with a preparation of autolysin (gamete lytic enzyme) to degrade the cell wall or due to a nuclear cw mutation in the host. Biolistic transformation involves the bombardment of cells with micron-sized microprojectiles carrying the transforming DNA. To reduce drag on the microprojectiles, the procedure is performed in a partly evacuated chamber. Acceler­ ation of the microprojectiles is commonly based on one of two principles. In the first, the microprojec­ tiles are deposited on a macroprojectile (a plastic bullet), or on a disk, which is accelerated and then stopped in flight by a perforated stopping plate or a mesh that allows the microprojectiles to continue their trajectory to the target cells. In the first generation of gene guns, the macroprojectile was accelerated by the explosion of a charge of gunpowder (Klein et al., 1987; Zumbrunn et al., 1989). More recent versions use a burst of compressed air or helium to propel the macroprojectile or a plastic membrane disk (reviewed by Sanford et al., 1993). The alternate principle is to accelerate the microprojectiles directly in a flow or a sudden burst of high-pressure gas, usually helium (Finer et al., 1992; Takeuchi et al., 1992).

Abbreviations: AAD – aminoglycoside 3´´-adenyl transferase ( or aminoglycoside 3´´-adenylyl transferase); GUS – -Glu­ curonidase; PCR – polymerase chain reaction; rDNA – ribosomal RNA gene cluster; RFLP – restriction fragment length poly­ morphism; rRNA – ribosomal RNA

III. Selectable Markers and Reporters

A. Selectable Markers 1. Photosynthesis Several decades of genetic research in C. reinhardtii have provided a rich collection ofchloroplast mutants with defects in photosynthesis. These mutants can be grown in the presence of acetate, but not on minimal medium in the light. Many of the corresponding chloroplast genes have been identified and cloned, providing a source of selectable markers for transformation of the corresponding mutant hosts: transformants can be selected for phototrophy on minimal medium. This is the strategy that was used in the original demonstration of chloroplast

Chapter 8

Chloroplast Transformation

transformation (Boynton et al., 1988). A mutant with a deletion of the atpB gene was rescued by bombardment with a plasmid containing a wild-type fragment covering the deletion, with segments of homology on either side allowing homologous recombination. An advantage of using a deletion mutant as a host is the lack of reversion. A similar strategy has also been used with mutations in a variety of other photosynthesis genes (for thorough reviews see Boynton and Gillham, 1993; Erickson, 1996). A variation on this theme is the use of the tscA gene as a marker in tscA deletion hosts (GoldschmidtClermont et al., 1991; Kindle et al., 1991, Choquet et al., 1992). The tscA RNA is required for transsplicing of the psaA RNA (Chapter 11, Herrin et al.), and thus indirectly for photosynthesis: the lack of tscA prevents the accumulation of PSI, so that selection for phototrophic growth can also be applied with this marker. However some mutants with deletions of the tscA locus, such as H13, have complex rearrangements of the chloroplast DNA (Choquet et al., 1992), and the marker is not simply integrated in replacement of the deletion when vectors such as rbcX or atpX are used (Goldschmidt-Clermont, 1991). Another variation is based on the light sensitivity of certain photosynthesis mutants, like those that are defective in Photosystem I. These mutants fail to grow at high light intensities even when acetate is provided in the medium. Transformants that have regained an active gene loose their light sensitivity and can be selected on acetate medium in the light (Redding et al., 1998; M. Fleischmann, N. Fischer and J. -D. Rochaix, personal communication). The two repeated copies of the psbA gene of C. reinhardtii contain four introns and span 6.5 kb, properties that complicate site-directed mutagenesis. However intron-less derivatives of the psbA gene can be used for transformation, and the resulting strains that lack all the introns have apparently normal phenotypes (Johanningmeier and Heiss, 1993; Minagawa and Crofts, 1994; Mayfield et al., 1994).

141 Kindle et al., 1991; Newman et al., 1991; Roffey et al., 1991). Likewise mutations of the psbA gene that confer resistance to a herbicide such as metribuzin or DCMU can be used for selection (Przibilla et al., 1991; Newman et al., 1992). The obvious advantage of drug resistance over selection for photosynthetic activity is that the host need not carry a mutation. The markers are integrated by homologous recom­ bination, replacing the wild-type (drug sensitive) copies of the corresponding genes. However herbicide resistance and also spectinomycin resistance mutations can have effects on photosynthetic activity that should be taken into account by using appropriate control strains in structure-function studies of the photosynthetic complexes (Lers et al., 1992; Monod et al., 1994; Heifetz et al., 1997; M. Fleischmann and J.-D. Rochaix, personal communication). Another strategy to obtain resistance is to express a gene product that can inactivate an antibiotic. This was achieved by transformation with the bacterial aadA gene, engineered with 5´ and 3´ sequences from Chlamydomonas chloroplast genes, to obtain expression of aminoglycoside 3´´-adenyl transferase (AAD) and thus resistance of the transformants to spectinomycin and streptomycin (GoldschmidtClermont, 1991). For the commonly used atpXAAD construct, the predicted protein product is a fusion ofAAD to the first 25 N-terminal amino acids of AtpA (Leu et al., 1992). In other constructs, the normal AAD protein is expressed by fusion at the level of the translation initiation codon. The aadA cassette provides a dominant marker which is portable since it can be integrated at virtually any site in the chloroplast genome if flanking regions of homology are provided for homologous recombination. For gene inactivation or site-directed mutagenesis, this has the advantage of allowing the marker to be linked to the gene of interest, while the use of resistance mutations in the rDNA generally calls for cotransformation approaches (Section IV.D). New versions of the aadA cassette that allow its excision permit repeated rounds of transformation with the same marker (‘recycling’) as discussed below (Section IV.E; Fischer et al., 1996).

2. Drug Resistance B. Reporters and Foreign Gene Expression Another approach has been to use mutations that confer resistance to various inhibitors. Mutations in the rRNA genes that bestow resistance to spectino­ mycin, streptomycin or erythromycin have been used to transform wild-type cells (Newman et al., 1990;

If regions of homology are provided for homologous recombination, foreign DNA can be stably integrated into the chloroplast genome (Blowers et al., 1989). Chimeric genes with alien DNA downstream of a

142 Chlamydomonas chloroplast promoter are trans­ cribed. The presence or addition of sequences from the 3´ end of a chloroplast gene can provide signals for the formation of a stable RNA with apparently homogeneous 3´ ends as assayed by Northern analysis (Chapter 10, Stern and Drager). Such chimeric transcripts with the sequences of bacterial genes like nptII, uidA, or aadA accumulate in the transformed cells (Blowers et al., 1989; 1990; GoldschmidtClermont, 1991; Klein et al., 1992). Chimeric RNAs with a mitochondrial intron from Scenedesmus obliquus or with the atpF gene of spinach are also expressed in Chlamydomonas chloroplasts (Herden­ berger et al., 1994; Deshpande et al., 1995). The S. obliquus intron is spliced in Chlamydomonas, but not the intron in the spinach atpF gene (Chapter 11, Herrin et al.). Transgenic expression of a foreign gene in the chloroplast to produce a functional protein was first demonstrated in Chlamydomonas with the aadA cassette by demonstrating AAD activity in crude extracts of transformants (Goldschmidt-Clermont, 1991). Using constructs with the uidA gene, glucuronidase (GUS) activity can also be measured in extracts from transformed cells (Sakamoto et al., 1993). These genes can thus be used as reporters of gene expression for the analysis of transcription, RNA processing, RNA stability and translation under the control of a variety of promoters as well as 5´and 3´-untranslated sequences (Sakamoto et al., 1993; Nickelsen et al., 1994; Zerges and Rochaix, 1994; Zerges et al., 1997; Stampacchia et al., 1997). With the aadA cassette, the level of resistance of the transformants to different concentrations of spectinomycin or streptomycin for growth on solid media can provide a rough indication of the expression level of the AAD reporter. The bacterial recA gene from E. coli has also been expressed in the Chlamydomonas chloroplast (Cerutti et al., 1995). Production of the wild type bacterial RecA protein leads to an increase in chloroplast recombination, but expression ofa dominant negative mutant RecA inhibits recombination. These results elegantly show that the transgenic RecA is functional in Chlamydomonas. It is also noteworthy that different truncated mutant forms of transgenic RecA accumulate to reduced levels compared to the wildtype, as assayed by immunoblotting. Because these proteins are expressed under the control of the same elements and the mRNA levels are similar, these

Michel Goldschmidt-Clermont differences in accumulation suggest that the truncated proteins have different degrees of susceptibility to proteolytic degradation in the chloroplast. Thus post­ translational events could also be important for achieving optimal transgenic expression, in addition to factors such as the rate of transcription, the processing and stability of the mRNA, and the efficiency of translation initiation and elongation. For genetic engineering it may be important to investigate the contribution of these different steps, since in many attempts to develop new markers, or to express genes in the chloroplast, the product was poorly expressed or undetectable.

IV. Fate of Transforming DNA

A. Integration in the Chloroplast Genome In most cases, the analysis of stable transformants shows that the introduced DNA has recombined with the chloroplast genome. This process requires homology between the transforming DNA and the recipient. Empirically, it seems that the length of homology has to be in the order of one kilobase for efficient transformation, but shorter segments are known to be sufficient for homologous recombination (Section IV.E). When the selectable marker or mutation is flanked by a region of homology on only one side, integration is the apparent result of a single crossover with a duplication ofthe region ofhomology (Fig. 1; Kindle et al., 1991). This type of integration is reversible and thus genetically unstable, since recombination between the direct repeats will lead to excision ofthe marker (Boynton and Gillham, 1993). When there is homology on both sides ofthe marker, DNA integration is formally the result of a double crossover. The actual mechanism is not known, and could involve exchanges on both sides of the marker or gene conversion. Most transformation experiments have been done with circular plasmid DNA. Transformation with linearized plasmid DNA can be more efficient when there is sequence homology on both sides of the marker (Blowers et al., 1989). Conversely linear DNA is not as effective for transformation when it has homology on only one side ofthe marker (Kindle et al., 1991). The sites of recombination have been mapped in transformation experiments using the genes for

Chapter 8

Chloroplast Transformation

ribosomal RNA marked with multiple drug resistance mutations and restriction fragment length poly­ morphisms (RFLP; Newman et al., 1990). Even though circular plasmid DNA is used for trans­ formation, a bias is observed for recombination events that occur near the junctions of the chloroplast DNA with the plasmid vector, and result in the integration of long segments of transforming DNA. The remaining exchange events are not randomly distributed along the region. Likewise, a segment where recombination occurs preferentially (‘hot­ spot’), both in sexual crosses and in transformation experiments, has been identified in the 3´ part of the psbA gene (Newman et al., 1992). Transformation of tobacco plastids also results in the incorporation of long segments of DNA (Staub and Maliga, 1992). The introduction of chimeric constructs that use chloroplast DNA fragments to direct gene expression generates duplications of these segments, which are usually already present in the host genome. Recombination between the introduced copy and the endogenous one can lead to rearrangements or deletions. A chimeric construct, with sequences of atpA driving the expression of aadA, can be inserted between psaB and rbcL. The introduced atpA segment can then recombine with the nearby atpA gene so that a deletion of around 2 kb is generated and

143

maintained as a minor heteroplasmic component (Bingham and Webber, 1994; heteroplasmic versus homoplasmic transformants are discussed in Section IV.C). Likewise, when an atpA-aadA cassette replaces the psbl gene, the atpA segment recombines with the nearby atpA gene to generate a stable, homoplasmic deletion of around 2kb (Künstner et al., 1995). A duplication of a sequence from psbC introduced at the psaB locus can recombine with the endogenous psbC gene more than 60 kb away, generating a minor population of fragmented genomes (Fischer et al., 1996). Similar alterations have also been described in tobacco chloroplast transformants (Svab and Maliga, 1993). Depending on the relative position and orientation of the duplicated sequences, the distance between them, and the effect of the rearrangements on chloroplast DNA maintenance and gene expression, there may or may not be selective pressure against the rearranged genomes. Thus in some cases a homoplasmic situation will be reached, while in others the altered genome will only be apparent as a component of a heteroplasmic mixture.

B. Extrachromosomal DNA A circular plasmid DNA with the atpB gene as a selectable marker can propagate in its free form after

144

transformation of an atpB deletion host (Boynton et al. 1988). However when the cells are transferred to solid medium, the free plasmid is lost, and only the integrated gene copy is retained. To my knowledge, attempts to create a stable replicating vector for the chloroplast of Chlamydomonas have not yet been successful (e.g., Suzuki et al., 1997). In tobacco plastids, following transformation with a segment from the rDNA locus, small extrachromosomal circular DNAs, constituted of monomers and multimers of a plastid DNA segment, have been observed (Staub and Maliga, 1994). This segment has been used as part of a shuttle vector that can be propagated in E. coli. After transformation of tobacco plastids with the shuttle vector, extrachromosomal plasmids and copies integrated in the genome are present, as well as non-transformed plastid genomes. It is therefore unclear whether the shuttle vector replicates autonomously, or is being continuously generated by excision from the transformed chloroplast genomes. An intriguing phenomenon of gene amplification has been observed in transformation experiments with deleted versions of atpB that lack part of the inverted repeat at the 3´ end of the mRNA (Kindle et al., 1994). Rescue of the photosynthetic defect of an atpB mutant host gene with some of these 3´ end truncations is only partial. The transformants show poor photoautotrophic growth which is attributed to the accumulation of reduced amounts of atpB transcripts which are unstable and ofvariable length, and hence of reduced levels of the subunit ofATP synthase (Chapter 10, Stern and Drager). However ‘robust’ transformants that have better photosyn­ thetic capability and wild-type levels ofthe subunit to In these arise at a frequency of transformants, tandem arrays of fifteen or more copies of the transforming plasmid increase the total copy number of the truncated atpB gene to 10–30 copies per chloroplast genome, and thus probably to thousands per chloroplast. Although these arrays appear to be episomal, a copy of the transforming plasmid is integrated (Suzuki et al., 1997). In addition, there are also complex rearrangements at the atpB locus in the robust transformants. The amplified DNA could be replicating autonomously, or could be continuously generated from the integrated copy. The transformed strains exhibit genomic instability in sexual crosses, and the amplified arrays are lost following new rounds of transformation. These two types of instability limit the potential of the amplified

Michel Goldschmidt-Clermont

units for genetic engineering aimed at the overexpression of gene products in the chloroplast.

C. Homoplasmic and Heteroplasmic Transformants In each C. reinhardtii cell, the genome of the single chloroplast is present in approximately eighty copies. This high degree of ploidy has important conse­ quences for transformation. Following delivery, the initial event is presumably integration of DNA into one copy of the chloroplast genome. Depending on its degree of dominance, a selectable marker will be expressed phenotypically if it is carried by a sufficient fraction of genome copies in the subsequent growth of the host cell or some of its descendants. The segregation process is stochastic but can be influenced by selection or screening of desired markers, it eventually leads to the appearance of segregants that have a uniform genetic content. In most cases, segregation of markers is rapid, suggesting that in addition to assortment of the genomes at each division, gene conversion is involved in the process. If the marker is introduced in the inverted repeat, both copies are identical in the transformants, demonstrating that an active mechanism of copy correction is operating (Newman et al., 1990). When all the copies ofthe chloroplast genome are identical, the transformant is termed homoplasmic. Transformation frequencies can be improved if the host cells are pretreated with FUdR (5­ fluorodeoxyuridine), an inhibitor of chloroplast DNA replication (Newman et al., 1990; Kindle, 1991; Boynton and Gillham, 1993). This treatment reduces the copy number of the chloroplast genomes, and thus probably facilitates the segregation and expression of recessive markers in the first stages of transformation. Because FudR is mutagenic, and because transformation frequencies are not limiting for most experiments, the treatment is not used widely. In some cases, both wild-type and recombinant copies are maintained in the transformants in spite of selection, and a heteroplasmic state persists. The heteroplasmic situation is genetically unstable, and the marker is lost if selective pressure is removed. Persistent heteroplasmy in gene disruption experi­ ments with the aadA cassette has been interpreted as revealing genes that have a vital function for the cell. Selection for spectinomycin resistance requires the maintenance of some copies of the chloroplast genome with the gene disruption and the aadA gene,

Chapter 8

Chloroplast Transformation

while the vital function requires wild-type copies. Persisting heteroplasmic states have been observed with a disruption of ORF472 (GoldschmidtClermont, 1991), but sequencing of the locus has recently shown that ORF 472 is actually a part of the 3119 bp rpoC2 gene (S. Nuotio and S. Purton, personal communication). Indeed disruptions of the genes encoding subunits of RNA polymerase, rpoB1, rpoB2 and rpoC2 remain heteroplasmic, as well as those of rps3 (a ribosomal protein), of clpP (a subunit of ClpP protease), and of an open reading frame (ORF 1995) of unknown function (Liu et al, 1993; Huang et al., 1994; Rochaix, 1995; Boudreau et al., 1997). Presumably these genes are important for the survival of the cell even on acetate-containing medium in the dark, growth conditions where disruptions of genes involved in photosynthesis, or in chlorophyll synthesis, readily become homoplasmic. An alternate explanation, that the gene disruptions remain heteroplasmic because they affect genes required for the expression of the aadA marker itself, appears plausible in some cases but less likely for clpP (Chapter 10, Stern and Drager). A somewhat different situation prevails in tobacco, where a disruption mutant of the rpoB gene becomes homoplamic and is viable in culture on sucrose-containing medium, although it is not competent for photosynthesis (Allison et al., 1996).

D. Co-Transformation When two different markers are introduced on separate vector molecules, there is a high frequency of events (up to 80%) where both are incorporated in the same transformant (reviewed by Boynton and Gillham, 1993; Webber et al., 1995; Erickson, 1996). This suggests that a limiting step for transformation is delivery of DNA to the chloroplasts rather than its integration and segregation. It is not known whether all the cells and their single chloroplasts in a population are competent for transformation. The high frequency of co-transformation is useful for directed mutagenesis of the chloroplast genome, since it allows strategies where the selectable marker and the mutation of interest are in different loci (Kindle et al., 1991; Newman et al., 1991; Roffey et al., 1991). Cells are bombarded with a mixture of two separate DNA molecules, one with a marker conferring resistance to an antibiotic and the other with the gene carrying the mutation of interest. The antibiotic resistance marker is usually a mutation in

145 the rDNA or the insertion of an aadA cassette in a genetically silent position of the genome. Transformants are selected on medium containing the antibiotic, and co-transformants which have also incorporated the second mutation in at least some copies of the genome are identified by testing their genotype (colony hybridization, Southern blotting, PCR), or when readily apparent, their phenotype. Subculturing of the co-transformants and testing then leads to the identification of strains that are homoplasmic for the unselected mutation.

E. Marker Recycling For some applications, it is useful to introduce several modifications in the chloroplast genome in successive rounds of transformation. When photosynthesis markers cannot be used because the genes under study are also involved in photosynthesis, there is currently a scarcity of suitable selectable markers: the most convenient rDNA marker confers resistance to spectinomycin, like the aadA cassette. This limitation can be overcome by using strategies where the selectable cassette is introduced in a genetically unstable form, and can thus be lost from the transformed strains before being re-used in the next round of transformation and selection (Fischer et al., 1996; Redding et al., 1998). Two such strategies, dubbed ‘marker recycling,’ have been developed. One relies on recombination between direct repeats, the other on heteroplasmic transformants. The first approach is to use an aadA cassette flanked by direct repeats for transformation, with selection for spectinomycin resistance (Fig. 2). When selective pressure is removed, recombination between the direct repeats leads to excision and loss of the marker, so that a spectinomycin sensitive strain can be obtained for the next step of transformation. While a 483 bp repeat (bacterial plasmid DNA) leads to efficient excision, a repeat of230 bp from the atpA promoter is not sufficient. A direct repeat of 216 bp flanking an aadA cassette inserted in the chlN gene does allow recombination and the appearance of green cells that have regained the ability to synthesize chlorophyll in the dark (Cerutti et al., 1995), so the precise length required may depend on the sequence used. Another factor which may influence how readily genomes with the excision are segregated and recovered is the recessive versus dominant nature of the marker being scored: spectinomycin sensitivity versus chlorophyll synthesis respectively in the

146

examples above. This is because spectinomycin sensitivity will not become phenotypically apparent until most cells are nearly homoplasmic, while chlorophyll synthesis in a sector of a colony can probably be scored even if the cells are still heteroplasmic. The second approach is to co-transform with the construct of interest and with the aadA marker inserted in a gene where it remains heteroplasmic (Fig. 3). Appropriate screening and subculturing leads to strains which are homoplasmic for the desired mutation, but heteroplasmic for the aadA cassette. On removal of selection for spectinomycin resistance, the cassette is rapidly lost, and a spectinomycin sensitive yet mutant strain can be obtained for the next round of transformation.

Michel Goldschmidt-Clermont

Chapter 8

Chloroplast Transformation

The recombination method is experimentally more direct: selection for the marker which is closely linked to the mutation of interest rapidly leads to strains that are homoplasmic for the mutation. However after excision, one copy of the direct repeat remains in the mutant genome (Fig. 2). The second method requires co-transformation, and thus calls for more laborious screening to obtain a homoplasmic mutant, but avoids the presence of any extraneous DNA in the mutant genome.

V. Reverse Genetics

147 advantages for subsequent site-directed mutagenesis: one is that there is no need to repeatedly subculture the transformants to obtain homoplasmic strains lacking any wild-type copy. The other is that there is no risk of obtaining recombinants that carry the selectable marker but not the desired mutation. If a selectable marker can be placed in close proximity to the gene of interest without affecting the phenotype, then the selection of the mutated strains is direct and efficient. Elegant vectors with engineered restriction sites, that facilitate mutagenesis and mutant analysis, and with the aadA cassette for selection have been constructed for psbA, psaA and psaB (Minagawa and Crofts, 1994; Redding et al., 1998).

A. Gene Inactivation The functions of the products from numerous chloroplast genes and open reading frames have been investigated by transformation-mediated disruption (see Rochaix, 1997, for a recent compilation). These disruptions are obtained with two main strategies: one is co-transformation of a plasmid carrying the altered gene, flanked by regions of homology for recombination on both sides, together with a plasmid carrying the selectable marker. The other is to insert within the gene a selectable aadA cassette, or to substitute the gene with a cassette. Variations ofboth ofthese strategies to allow recycling of the marker have been used to generate deletions of photosynthesis genes (Section IV.E). If the gene that is targeted has a non-essential or conditional phenotype, such as acetate requirement or lightsensitivity, homoplasmic transformants are obtained that allow a detailed analysis. However when the disruption remains heteroplasmic, little information can be derived other than inferring that the gene may have an essential function (Section IV.C).

B. Site-Directed Mutagenesis The development of efficient tools for chloroplast transformation allows one to obtain routinely sitedirected mutations in virtually any gene of interest. Structure-function analysis of many components of the photosynthetic complexes and of Rubisco are described in the corresponding chapters of this book. The most efficient strategy is probably to first generate a complete deletion of the gene of interest using the recyclable marker approach, unless, as is the case for psbA, a classical deletion mutant is available. The use of a deletion mutant as the host has two main

VI. Conclusion and Perspective With the available bombardment technology and selectable markers, chloroplast transformation frequencies are more than adequate for most experiments: hundreds of transformed colonies are usually recovered quite readily. Homologous recombination and gene conversion greatly enhance the usefulness of chloroplast transformation because they lead to predictable insertion of the introduced DNA at sites of homology, and because they contribute to the rapid segregation of transformed genomes. With these properties C. reinhardtii is a very useful organism to investigate the chloroplast genome, the expression of chloroplast genes, as well as the role of the proteins encoded and the relations of their structure and function. Indeed research projects that use C. reinhardtii chloroplast trans­ formation have flourished, and a large number of reports citing these approaches are in the published literature. Nevertheless, despite this bright perspective, there is still room for technical improvement. It would be very useful to develop an inducible or repressible gene expression system for the chloroplast. This would be of help to analyze the function of essential genes, since conditional mutants could be obtained. Homoplasmic cell lines would be viable under inducing conditions, but the gene could be turned off or repressed to investigate the resulting phenotype. There is also a need for additional selectable markers, even though marker recycling alleviates the problem. Ideally new markers should be unrelated to photosynthesis, dominant, and portable to any site in the plastid genome. There are still unresolved

148 problems with the production of foreign proteins in the chloroplast. This may be due to suboptimal gene expression, and perhaps also to post-translational events, including protein degradation. To date, only foreign proteins of bacterial origin (AAD, GUS and RecA) have been expressed successfully in the chloroplast of C. reinhardtii (Section III.B). To my knowledge, there are no reports of the introduction of nuclear genes in the chloroplast resulting in the accumulation of their protein products, although a chimeric protein consisting of plastocyanin fused to AAD did accumulate as assayed by immunoblotting (N. Rolland and J.-D. Rochaix, personal communi­ cation). A better understanding of chloroplast gene expression and post-translational events will be of prime interest in fundamental research, but also for applications in biotechnology. Transformation will be a basic tool in this endeavor, and in turn the technique will benefit from its results.

Acknowledgments I thank Nicolas Roggli for his expert help in preparing the figures. Experimental work was supported by the Swiss National Fund for Scientific Research (3134014.92).

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Chapter 8

Chloroplast Transformation

Lers A, Heifetz PB, Boynton JE, Gillham NW and Osmond, CB (1992) The carboxyl-terminal extension of the D1 protein of Photosystem II is not required for optimal photosynthetic performance under and light-saturated growth conditions. J Biol Chem 267: 17494–17497 Leu S, Schlesinger J, Michaels A and Shavit N (1992) Complete DNA sequence of the Chlamydomonas reinhardtii chloroplast atpA gene. Plant Mol Biol 18: 613–616 Liu X-Q, Huang C and Xu H (1993) The unusual rps3-like orf712 is functionally essential and structurally conserved in Chlamydomonas. FEBS Lett 336: 225–230 Mayfield SP, Cohen A, Danon A and Yohn CB (1994) Translation of the psbA mRNA of Chlamydomonas reinhardtii requires a structured RNA element contained within the 5´ untranslated region. J Cell Biol 127: 1537–1545 Minagawa J and Crofts AR (1994) A robust protocol for sitedirected mutagenesis of the D1 protein in Chlamydomonas reinhardtii: A PCR-spliced gene in a plasmid conferring spectinomycin resistance was introduced into a psbA deletion strain. Photosynth Res 42: 121–131 Monod C, Takahashi Y, Goldschmidt-Clermont M and Rochaix JD (1994) The chloroplast ycf8 open reading frame encodes a Photosystem II polypeptide which maintains photosynthetic activity under adverse growth conditions. EMBO J 13: 2747– 2754 Newman S, Boynton JE, Gillham NW, Randorph-Anderson BL, Johnson AM and Harris EH (1990) Transformation of chloroplast ribosomal RNA genes in Chlamydomonas: Molecular and genetic characterization of integration events. Genetics 126: 875–888 Newman S, Gillham NW, Harris EH, Johnson AM and Boynton JE (1991) Targeted disruption of chloroplast genes in Chlamydomonas reinhardtii. Mol Gen Genet 230: 65–74 Newman S, Harris EH, Johnson AM, Boynton JE and Gillham NW (1992) Nonrandom distribution of chloroplast recom­ bination events in Chlamydomonas reinhardtii: Evidence for a hotspot and an adjacent cold region. Genetics 132: 413–429 Nickelsen J, van Dillewijn J, Rahire M and Rochaix JD (1994) Determinants for stability of the chloroplast psbD RNA are located within its short leader region in Chlamydomonas reinhardtii. EMBO J 13: 3182–3191 Przibilla E, Heiss S, Johanningmeier U and Trebst A (1991) Sitespecific mutagenesis of the D1 subunit of Photosystem II in wild-type Chlamydomonas. Plant Cell 3: 169–174 Redding K, MacMillan F, Leibl W, Brettel K,Hanley J, Rutherford AW, Breton J and Rochaix J-D (1998) A systematic survey of conserved histidines in the core subunits of Photosystem I by site-directed mutagenesis reveals the likely axial ligands of P700. EMBO J 17: 50–60 Rochaix J-D (1995) Chlamydomonas reinhardtii as the

149 photosynthetic yeast. Annu Rev Genet 29: 209–230 Rochaix J-D (1997) Chloroplast reverse genetics; New insights into the function of plastid genes. Trends Plant Sci 2: 419–425 Roffey RA, Goldbeck JH, Hille CR and Sayre RT (1991) Photosynthetic electron transport in genetically altered Photosystem II reaction centers of chloroplasts. Proc Natl Acad Sci USA 88: 9122–9126 Sakamoto W, Kindle K and Stern DB (1993) In vivo analysis of Chlamydomonas chloroplast petD gene expression using stable transformation of beta-glucuronidase translational fusions. Proc Natl Acad Sci USA 90: 497–501 Sanford JC, Smith FD and Russell JA (1993) Optimizing the biolistic process for different biological applications. Methods in Enzymol 217: 483–509 Stampacchia O, Girard-Bascou J, Zanasco J-L, Zerges W, Bennoun P and Rochaix J-D( 1997) A nuclear-encoded function essential for translation of the chloroplast psaB mRNA in Chlamydomonas. Plant Cell 9: 773–782 Staub JM and Maliga P (1992) Long regions of homologous DNA are incorporated into the tobacco plastid genome by transformation. Plant Cell 4: 39–15 Staub JM and Maliga P (1994) Extrachromosomal elements in tobacco plastids. Proc Natl Acad Sci USA 91: 7468–7472 Suzuki H, Ingersoll J, Stern DB and Kindle KL (1997) Generation and maintenance of tandemly repeated extrachromosomal plasmid DNA in Chlamydomonas chloroplasts. Plant J 11: 635–648 Svab Z and Maliga P (1993) High-frequency plastid trans­ formation in tobacco by selection for a chimeric aadA gene. Proc Natl Acad Sci USA 90: 913–917 Svab Z, Hajdukiewicz P and Maliga P (1990) Stable transformation of plastids in higher plants. Proc Natl Acad Sci USA 87: 8526–8530 Takeuchi Y, Dotson M and Keen NT (1992) Plant transformation: A simple particle bombardment device based on flowing helium. Plant Mol Biol 18: 835–839 Webber AN, Bingham SE and Lee H (1995) Genetic engineering of thylakoid protein complexes by chloroplast transformation in Chlamydomonas reinhardtii. Photosynth Res 44: 191–205 Zerges W and Rochaix JD (1994) The 5´ leader of a chloroplast mRNA mediates the translational requirements for two nucleusencoded functions in Chlamydomonas reinhardtii. Mol Cell Biol 14: 5268–77 Zerges W, Girard-Bascou J and Rochaix J-D (1997) Translation of the chloroplast psbC mRNA is controlled by interactions between its 5´ leader and the nuclear loci TBC1 and TBC3 in Chlamydomonas reinhardtii. Mol Cell Biol 17: 3440–3448 Zumbrunn G, Schneider M and Rochaix J-D (1989) A simple particle gun for DNA-mediated cell transformation. Technique 1: 204–216

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Chapter 9 Chloroplast RNA Stability Jörg Nickelsen

Allgemeine Botanik, Ruhr-Universität Bochum,

Universitätsstr. 150, D-44780 Bochum, Germany

Summary I. Introduction II. Cell Cycle Dependent Regulation of Chloroplast RNA Stability III. Nuclear Mutants Affected in Chloroplast RNA Stability IV. Towards a Molecular Model of Chloroplast RNA Stabilization/Degradation A. Cis-acting Elements 1. The 3´ Untranslated Regions 2. The 5´ Untranslated Regions B. Trans-acting Factors C. Chloroplast Ribonucleases D. RNA Stability and Translation V. Conclusions and Perspectives Acknowledgments References

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Summary The potential importance of RNA stability to the regulation of chloroplast gene expression is documented by recent work on the molecular aspects of chloroplast biogenesis. In Chlamydomonas reinhardtii, two lines of evidence support the idea that gene-specific regulated RNA decay plays a crucial role in determining the different steady-state levels of single transcripts. First, during the cell cycle, a light-dependent and/or circadian control of RNA accumulation takes place that appears to involve both transcriptional and posttranscriptional mechanisms. Secondly, numerous well-characterized photosynthetic nuclear mutants exhibit defects in the stabilization of specific Chloroplast transcripts. This further substantiates the significance of gene-specific aspects of chloroplast RNA metabolism. The recently developed chloroplast and nuclear transformation techniques, combined with appropriate in vitro RNA degradation and RNA binding assays, now allow the identification of the cis-acting RNA elements involved and the associated trans-acting factors. Recent evidence suggests that both 3´ and 5´ untranslated regions contain determinants for mRNA stability. The cloning of nuclear genes affecting chloroplast RNA turn­ over has led to a molecular model of multisubunit complexes mediating RNA stabilization. Furthermore, this work provides the platform for molecular links to subsequent steps of gene expression, such as transcript 5´ processing and translation. The overall picture emerging is that of a complex molecular network in which a number of gene-specific regulatory mechanisms are involved.

J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 151–163. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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I. Introduction The coordinated expression of both nuclear and organellar genes results in the constitution of a functional chloroplast capable of performing photosynthesis. The different levels of gene expression that could serve as targets mediating this coordinate regulation are comprised oftranscriptional as well as posttranscriptional steps within the two cell compartments. In higher plants, the expression of nuclear genes for plastid proteins is mainly controlled at the transcriptional level (Kuhlemeier, 1992); however, recent evidence indicates that regulated RNA turn­ over also contributes to the establishment of particular RNA steady-state levels (Abler and Green, 1996). In contrast, within the plastid, regulation of gene expression has been shown to be mostly dependent on posttranscriptional processes like RNA stabili­ zation, translation or even posttranslational modifica­ tions (Gruissem and Tonkyn, 1993; Sugita and Sugiura, 1996). In Chlamydomonas reinhardtii, as compared to higher plants, relatively little is known concerning the control of nuclear gene expression. Thus, although transcriptional regulation has been demonstrated for Cab or RbcS gene expression for instance, posttrans­ criptional events also appear to contribute to overall regulation of gene expression, reminiscent of the situation found in higher plants (Gagne and Guertin, 1992; Hwang and Herrin, 1994; Jacobshagen et al., 1996). Furthermore, recent genetic evidence supports the idea that differential RNA stabilization might play a crucial role at least for the expression of some nuclear genes encoding chloroplast polypeptides (Hahn et al., 1996). Within the C. reinhardtii chloroplast there is a clear preference for posttranscriptional processes acting as control points for gene expression levels (Rochaix, 1996). The aim of this chapter is to present and discuss the genetic and molecular aspects of chloroplast RNA stabilization that have arisen to date. Accordingly, I will first summarize what is known about the significance of RNA stabilization throughout the cell cycle of C. reinhardtii. Then well-characterized nuclear mutants of chloroplast RNA metabolism will be presented that have led to Abbreviations: GUS – glucuronidase; TPR – tetratricopeptide repeat; UTR – untranslated region

Jörg Nickelsen recent molecular models of how RNA can be stabilized/degraded in a regulated fashion.

II. Cell Cycle Dependent Regulation of Chloroplast RNA Stability When C. reinhardtii cultures are grown photo­ autotrophically under 12 h light/12 h dark regimes, cells divide synchronously into two to eight daughter cells at the end of the dark phase giving rise to a stepwise two- to three-fold increase in cell number. The synthesis of photosynthetic components is maximal in the middle of the light phase indicating a tight control of gene expression by light and/or an endogenous circadian mechanism (Harris, 1989). In order to dissect the regulatory checkpoints where this control is mediated in the chloroplast of C. reinhardtii, thorough work has been done on analyzing the transcription rates, mRNA levels, and polypeptide synthesis rates during the cell cycle. To quantitate processes of RNA metabolism, in general, transcription rates of particular genes are compared to the steady-state levels of the corres­ ponding transcripts. While the RNA levels are determined by Northern analysis, transcription can be followed by in vitro run-on transcription assays. For these assays, cells are permeabilized by either toluene treatment (Guertin and Bellemare, 1979) or repeated freeze and thaw cycles (Gagne and Guertin, 1992) and, subsequently, preinitiated transcripts are elongated for a limited time in the presence of radioactively labeled ribonucleotides. Resulting RNAs are visualized by hybridization with filterimmobilized DNA fragments of the gene of interest. Alternatively, transcripts may be pulse-labeled in vivo by short incubations of phosphate-depleted cells with radioactive orthophosphate (Herrin et al., 1986). When these pulse-labeled cells are subsequently kept in light or darkness for defined times, the halflives of individual transcripts under different light conditions can be followed (Salvador et al., 1993). Table 1 summarizes some of these data. Despite the large differences with regard to particular values that might have been caused by the different techniques (transcription analysis in permeabilized cells versus in vivo pulse-labeling, Sieburth et al. 1991) or strains used, some important principles are revealed. The steady-state levels of RNAs can vary to some extent during the cell cycle. While the

Chapter 9

Chloroplast RNA Stability

maximum/minimum ratio of transcript levels for some highly expressed genes like psbA, psbD, rbcL, and rrnL range only from 1.3 to 2.3/2.8, the levels of transcripts corresponding to other genes fluctuate much more, with tufA RNA representing the most striking example. This suggests that gene-specific mechanisms regulate the abundance of individual transcripts within the chloroplast of C. reinhardtii. Recent work indicates that, in some cases, the different transcript levels are not related to the cell cycle directly, but rather, are light dependent (psaB, psbA, and rbcL) or follow an endogenous circadian control (atpA, atpB, and tufA; Salvador et al., 1993). When transcription rates of the these genes are analyzed in parallel, one finds that they are maximum in the very early light phase (20 min) with a significant decline toward the end of the light period (7 h) and a subsequent increase during the next late dark phase. For tufA, rrnL, atpA, and atpB, max/min ratios of transcription rates are quite high, pointing to an important role for transcriptional control of chloroplast gene expression. However, the corres­ ponding steady-state RNA levels do not fluctuate as much, suggesting that additional processes of differential RNA stabilization are superimposed on a general effect of transcriptional control. Overall, RNA half-lives vary remarkably between the genes analyzed but are generally greater in the dark. For instance, the most stable chloroplast transcript known is the 16S rRNA, which exhibits a half-life of roughly 7 h in the light and 30 h in the dark. In contrast, tufA transcripts have a half-life of just 30 min in the light-grown versus 1.3 h in dark-

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grown cells. For rbcL RNA a six-fold increase in mRNA stability (3.5 h in light as compared to 21 h in dark) was found which represents the highest light/ dark ratio reported. Thus, enhanced RNA stabilization during the dark phase of the cell cycle appears to compensate for the drop in transcriptional activity within this period thereby keeping the steady-state levels of some highly expressed genes like psbA, psbD, and rbcL relatively constant. Interestingly, inhibition of chloroplast transcription by rifampicin treatment at the beginning of the light phase results in the subsequent loss of synthesis of the psbA and psbD gene products D1 and D2, respectively, while translation of rbcL mRNA continues (Herrin et al., 1986). This suggests that residual rbcL transcripts from the preceding cycle remain competent for translation, in contrast to psbA and psbD mRNAs, which have to be newly synthesized. The molecular basis for this difference in gene expression is not clear, but it is possible that different 5´ modifications on these mRNAs might be involved in discrimination between translatable and non-translatable chloroplast messengers (Section IV.A). For other transcripts, like tufA, probably due to their short half-lives, a compensation of reduced transcription in the dark does not take place, resulting in a diurnal fluctuation of transcript levels during the cell cycle. In conclusion, evidence obtained to date suggests a complex regulatory network determining individual transcript levels involving processes of differential transcription as well as controlled RNA stabilization. These effects are apparently superimposed upon a general increase in the transcription rate in the early

154 light phase and enhanced RNA stability in the dark.

III. Nuclear Mutants Affected in Chloroplast RNA Stability As photosynthetic function is dispensable in C. reinhardtii when cells are grown in the presence of acetate, it has been fairly easy to isolate photosynthetic mutants of C. reinhardtii (Rochaix, 1995). Among these are a number of nuclear mutants that exhibit defects in the stability of single chloroplast transcripts (Table 2). None of these mutations appear to be analogous to the recently characterized Arabidopsis mutation hcf109, which reduces the RNA accumulation of particular segments from four different plastid polycistronic transcription units (Meurer et al., 1996). However, in C. reinhardtii, gene-specific defects seem to be a general phenom­ enon of nuclear mutations affecting not only RNA stabilization, but also other steps of chloroplast gene expression (see Chapter 10, Stern and Drager; Chapter 11, Herrin et al.; Chapter 12, Hauser et al.). When the phenotypes of the C. reinhardtii mutants are analyzed in more detail, subtle differences between them become apparent. While most of the mutants fail to accumulate any RNA from the affected chloroplast genes, mutants and ncc1 still accumulate 5 and 10% of wild-type levels of petB and atpA transcripts, respectively. Interestingly, is deficient in photosynthetic activity, but in ncc1 about half of the affected ATP synthase complex assembles allowing wild-type growth rates under phototrophic growth conditions (Drapier et al., 1992; Gumpel et al., 1996). Another difference concerns the effects on polycistronic chloroplast mRNAs. For instance, in the mutants 222E and GE2.10, besides psbB RNA, additional smaller transcripts—encoded further downstream of psbB and probably cotranscribed with it—are absent, pointing to a more pleiotropic effect in this case. In contrast, psaA exon 2 sequences that are cotranscribed with the upstream psbD gene are apparently not degraded in the psbD RNA stability mutant nac2-26, since normal accumulation of mature psaA message occurs. Interestingly, the same RNA is degraded when it remains connected to psbD sequences in double mutants carrying, in addition to nac2-26, a mutation which blocks psaA RNA maturation/splicing from this dicistronic transcript. This suggests that psbD sequences can target these splicing intermediates for

Jörg Nickelsen rapid degradation in the mutant nuclear background (Kuchka et al., 1989). All of the mutants listed in Table 2 have been shown to transcribe the affected genes to nearly wild-type levels by run-on and/or in vivo pulse labeling assays. One exception is a recently described mutant called 76-5EN that accumulates only very low amounts of rbcL mRNA. Apparently, in 76-5EN, the rbcL gene is only weakly transcribed in 10 min pulse-labeling experiments but the resulting mRNA appears to remain stable for at least 1 h (Hong and Spreitzer, 1994). Whereas this mutant would represent the first example of a nuclear mutation affecting the transcription of a chloroplast gene, these results are still a matter of debate (Rochaix, 1996). It remains possible that, in 76-5EN, the rbcL mRNA was rapidly degraded and thereby escaped detection during the relatively long 10 min pulse, as has been reported for psbB transcripts in the mutant 222E (Monod et al., 1992). Since the RNA stability assays of Hong and Spreitzer (1994) were not strand-specific, one cannot exclude the possibility that the stable low abundance transcripts that were detected were generated from the opposite DNA strand. A similar situation has been found for petD gene transcription (Sturm et al., 1994). To date, it is still unknown how many nuclear genes can be involved in the stabilization of a particular chloroplast transcript. It is unlikely that mutagenesis of the nuclear genome has been performed to saturation, suggesting that there might be additional loci to be discovered. However, for the nac2-26 mutation affecting psbD RNA accumulation, a second allele has been found by analyzing the (S. Purton, unpub­ photosynthetic mutant lished). The mutants GE2.10 and 222E both exhibit a defect in psbB transcript stabilization; it is unknown yet whether they represent different alleles of the same gene.

IV. Towards a Molecular Model of Chloroplast RNA Stabilization/Degradation The combined analysis of both nuclear mutants with defects in chloroplast RNA stability and chloroplast transformants carrying reporter genes fused to putative regulatory regions now represents a powerful approach for addressing questions of the molecular basis of RNA decay in the chloroplast of C. reinhardtii. First, the target sites within chloroplast

Chapter 9

Chloroplast RNA Stability

transcripts recognized by the nuclear factors can be identified and other cis-acting determinants of RNA stability can be tested in vivo. Secondly, the cloning of the mutated nuclear genes together with recently developed in vitro RNA stabilization and RNA binding assays provide the tools for detailed characterization of the factors involved. These data can help to elucidate putative molecular connections between processes of RNA stabilization and translation (Gillham et al., 1994; Nickelsen and Rochaix, 1994).

A. Cis-acting Elements The identification of RNA elements responsible for determining the half-lives of their corresponding transcripts has been one major goal of research in chloroplast molecular biology during the last years. To date, no common motifs have been identified, but accumulating evidence suggests that almost all regions of chloroplast transcripts—namely the 5´ and 3´ untranslated regions as well as coding sequences—can be involved in RNA stabilization/ degradation.

1. The 3´ Untranslated Regions Initial work on higher plants has suggested that stem-loop structures within the 3´ untranslated regions (3´ UTR) of plastid mRNAs serve as barriers against exonucleolytic degradation of upstream sequences in vitro (Stern and Gruissem, 1987) and in organello (Adams and Stern, 1990). Furthermore, they appear to mediate correct 3´ end formation, and thus, a close

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relationship between 3´ maturation and RNA stability has been proposed. With the availability of a biolistic chloroplast transformation system for C. reinhardtii (Boynton et al., 1988) this hypothesis became testable in vivo also. One of the best characterized examples is the atpB 3´ UTR (Stern et al., 1991). Successive deletion of the atpB stem-loop structure leads to a loss of up to 80% of the atpB mRNA, indicating a significant role of this secondary structure element for RNA stabilization. In addition, the residual transcripts are heterogenous with regard to the length of their 3´ ends, indicating a role for stem-loops in RNA maturation. Interestingly, a 3´ UTR from the spinach petD gene can substitute functionally for the C. reinhardtii 3´ region, raising the possibility that common structural motifs exist in the 3´ UTRs of higher plants and algae (Stern et al., 1991). Even more surprising is the finding that a sequence of 18 guanosine nucleotides replacing the atpB stem-loop allows wild-type levels of atpB mRNA to accumulate (Drager et al., 1996). This is reminiscent of the situation in Saccharomyces cerevisiae, where polyG tracts can impede 5´–3´ exonucleases. While a significant impact of the atpB 3´ UTR stem-loop on the level of its mRNA accumulation has been documented, results obtained with chimeric GUS genes containing different 3´ regions do not support the idea that 3´ UTRs, in general, are involved in determining RNA half-lives. In these experiments, constructs consisting of the atpB 5´ region and the bacterial uidA gene, were fused to either the rbcL or the psaB 3´ flanking regions and, subsequently, introduced into the chloroplast genome. Although it

156 could be shown that the 3´ UTRs are required for correct 3´ end formation of the chimeric transcripts, the measurement of RNA half-lives after pulse labeling of transformant cells with phosphate revealed no differences between constructs harboring a 3´ region or not (Blowers et al., 1993). Contradictory results, as far as the psaB 3´ UTR is concerned, have been obtained after removal of the 3´ stem-loop structure from the endogenous psaB gene, which resulted in a 75% reduction in the halflife of its mRNA (Lee et al., 1996). It is unknown what the reason for these different findings is but one might speculate that the coding region and/or the 5´ untranslated portion of psaB, and perhaps also of other genes, affect the function of the 3´ UTR by direct and/or indirect interactions. In other words, if the rate limiting step for RNA degradation, in the case of the psaB transcript, is mediated by the 5´ region and/or coding sequences, this can be compensated for by an intact 3´ stemloop, but not by a deleted version. In this way, the chimeric mRNA might not be dependent on an appropriate 3´ end for stabilization. A role for proteincoding sequences in RNA stabilization has been suggested for psbA transcripts in spinach (Klaff 1995) and also for the rbcL gene in C. reinhardtii (Salvador et al., 1993). Moreover, recent genetic and molecular evidence indicate that the 5´ UTRs play key roles in determining the stability of at least some chloroplast RNAs.

2. The 5´ Untranslated Regions Although mRNA stability has been shown to be affected by alterations of cis-elements within the 3´ UTR, a complete loss of RNA has never been observed. This might be due to the use of experimental approaches that are based on the selection of chloroplast transformants by photoautotrophic growth, thereby selecting for at least partial functional activity of the transcripts analyzed. Alternatively, incomplete RNA degradation might be an inherent feature of 3´ UTR-mediated RNA destabilization. This might be further supported by the finding that in the nuclear mutant ncc1, which still accumulates 10% of wild-type atpA mRNA amounts (Table 2), RNA degradation initiates from the 3´ end of the message (Drapier et al., 1992). However, most of the other mutants analyzed (Table 2) do not accumulate any transcripts from the affected genes and, consequently, cannot grow

Jörg Nickelsen

photoautotrophically. Among these the nuclear mutant nac2-26, which exhibits defects in psbD RNA stabilization, has been analyzed in detail (Fig. 1). Chimeric reporter genes containing either the 5´ or 3´ flanking regions of the psbD gene fused to the bacterial aadA gene (conferring spectinomycin resistance; Chapter 8, Goldschmidt-Clermont) were inserted into the chloroplast genome by biolistic transformation. After crossing of the resulting transformants with nac2-26 the analysis of tetrads demonstrated a cosegregation of Photo­ system II deficiency and spectinomycin sensitivity in case of 5´ psbD-aadA fusions. Subsequent analysis of RNA levels revealed that both psbD and the chimeric transcripts are destabilized in the mutant nuclear background. In contrast, an aadA-3´ psbD chimera remained stable independent of the genetic nuclear background. This indicates that the 5´ leader region but not the 3´ UTR of the psbD transcript is sufficient to mediate accelerated RNA degradation in the absence of NAC2 function. The degradation of psbD leader RNA has also been followed in lysed

Chapter 9 Chloroplast RNA Stability chloroplasts in vitro pointing to a pathway involving at least three endonucleolytic cuts (Nickelsen et al, 1994). This resembles the situation found in bacteria, where the significance of 5´ leader regions for RNA stabilization/degradation events is widely accepted and endonucleolytic 5´ degradation has been demonstrated to occur (Bouvet and Belasco, 1992; Hansen et al., 1994). Using a similar approach, evidence has been obtained recently that in mutant 222E, in which psbB transcript stability is affected, the psbB 5´ flanking region also contains the target site for the nucleusencoded factor (F. Vaistij, M. Goldschmidt-Clermont, and J.-D. Rochaix, unpublished). Interestingly, psbB transcripts are very similar to psbD RNAs with regard to their 5´ ends. In both cases two different 5´ ends can be detected that correspond to a longer less abundant form and a shorter predominant version (Nickelsen et al., 1994; J. Nickelsen, unpublished). Although it has not yet been demonstrated directly, preliminary indirect evidence (see below) suggests that, at least in case of the psbD gene, the shorter form represents a 5´ processed form (J. Nickelsen, unpublished results). As psbA RNA messages reveal similar 5´ heterogeneity (Nickelsen et al., 1994), these three mRNAs appear to represent a special group of chloroplast transcripts. The concerted synthesis of their products and the assembly of an intermediate complex structure comprising D1, D2, and P5 proteins has been observed during Photo­ system II biogenesis (De Vitry et al., 1989). Thus, a model has been proposed in which obligate mRNA 5´ end maturation reflects a coordinate step during gene expression of psbA, psbD, and psbB, respectively (Rochaix, 1996). In order to further test this hypothesis, chloroplast transformants have been generated which contain altered psbD 5´ UTRs. Preliminary data suggest that deletion of the region from position –74 (long mRNA 5´ end) to –47 (mature mRNA 5´ end), removing all sequences unique to the larger RNA, results in loss of psbD transcripts. The psbD transcription rate in this mutant is similar to wild-type indicating the absence of putative promoter elements, within the –74 to –47 region, that might have driven the transcription of the short psbD RNA (see above) and thus, the results confirm the significance of this region for RNA processing and stabilization. Further analyses narrow down the crucial stability element to the first 12 nt of the psbD leader (J. Nickelsen and J.-D. Rochaix, unpublished). This result is somewhat

157 surprising since the processed mature form starting at position –47, which does not contain this element, accumulates as the predominant transcript. Appar­ ently, during psbD gene expression, there is a strict requirement for a 5´ processing reaction, which is impaired in the mutated versions, that stabilizes the message. This might occur, for example, if the maturation reaction is closely coupled to subsequent steps of factor assembly on the mature mRNA. Mutations introduced into the 5´ part of the endogenous psbA gene, in general, affect RNA accumulation only slightly. Two exceptions are a deletion removing the putative Shine-Dalgarno sequence, GGAG, at position –28, which results in only 20% of wild-type RNA accumulation, and a substitution of nucleotides from position –56 to –60 by GCCTC, which then allows base pairing with the GGAG motif and results in only 2% of wild-type mRNA levels (Mayfield et al., 1994). It is not clear whether these decreases in mRNA steady-state levels are a consequence of reduced translatability of the molecules. The same also applies to the psbD mutants mentioned above and the petD 5´ deletions that have been tested in vivo (Sakamoto et al., 1994). The possible connection between translation and RNA stabilization will be discussed in Section IV.D. Additional data supporting the idea of 5´ regions playing a key role in RNA degradation have been obtained when 5´ rbcL-reporter gene fusions were analyzed in vivo. Reporter gene transcripts are degraded rapidly in cells grown in the light, but are stabilized after addition of 5´ sequences from the rbcL protein-coding region (Salvador et al., 1993), again pointing to long range interactions over the whole molecule. Finally, recent work based on the analysis of reporter gene constructs revealed that in the nuclear mutant F16 (Table 2) at least one determinant for petD mRNA stabilization is also located within its 5´ leader region (Drager et al., 1998). Taken together, recent reports about the ciselements mediating the regulation of RNA accumu­ lation suggest that almost all regions of chloroplast transcripts can contain positive or negative deter­ minants of RNA stabilization. One of the key issues for the future will be to elucidate those elements which control the rate limiting steps. The 5´ regions might include some of them since mutations in these regions can lead to a complete absence of the mRNA. Furthermore, in the nuclear mutants nac2-26, 222E and F16, that mediate their RNA destabilization

158 effects via chloroplast leader regions, no transcripts from the affected genes are detectable. From a functional point of view, degradation initiated from the 5´ end might result in an immediate inactivation of translation capacity avoiding the waste of energy for the synthesis of incomplete polypeptides. In addition, such incomplete polypeptides might be deleterious when they are assembled into inactive complexes because they could sequester other subunits. It is interesting to note that in higher plants, like in E. coli, polyadenylation of chloroplast transcripts appears to accelerate degradation of RNA molecules (Kudla et al. 1996). It is not clear yet whether similar processes also occur in C. reinhardtii, but it adds an exciting novel aspect to what is known to date about RNA stabilization in chloroplasts. However, in order to understand these processes more precisely, the trans-acting factors that interact with these RNA elements will also have to be characterized.

B. Trans-acting Factors To characterize and isolate the factors mediating the differential regulation of RNA stability in chloro­ plasts, biochemical as well as genetic approaches have been initiated. In higher plant plastids, a number of different RNA binding proteins have been identified that interact specifically with the 3´ flanking regions of various chloroplast transcripts in vitro. For some of these, functions have been assigned (for a review, see Sugita and Sugiura, 1996). While RNA protein interactions within the 3´ flanking regions of higher plant plastid transcripts have been studied intensively, so far, in vitro work on C. reinhardtii has mainly focused on analyzing 5´ leader regions in RNA binding experiments (Danon and Mayfield, 1991; Zerges and Rochaix, 1994; Hauser et al., 1996). The initial goal has been to identify protein factors involved in translational regulation of chloroplast mRNAs, but as mentioned above, these regions can also serve as determinants of RNA stability, at least in C. reinhardtii. Hence, additional genetic data and/or appropriate in vitro assays are required to distinguish whether a particular RNA binding activity is involved in either of these processes. Using wild-type chloroplast lysates, the psbD leader RNA, that serves as the target site of the nucleus-encoded factor affected in the mutant nac2-26 (Table 2), was shown to interact with a 47

Jörg Nickelsen kDa protein as monitored by UV-light mediated crosslinking of RNA-protein complexes (Nickelsen et al., 1994). Both the stability of the RNA probe and the binding activity were lower when chloroplast lysates from nac2-26 were analyzed. Interestingly, this factor only bound to a probe comprising the long leader from position –74 to +1. An in vitro transcript, that corresponds to the mature form (–47 to +1) did not form a stable complex indicating that the binding site of the 47 kDa protein is not within the short leader. Because deletion of positions –74 to –47 of the psbD leader resulted in RNA destabilization in vivo (compare Section IV.A.2), confirming the significance of this region for RNA stability, the 47 kDa protein represents a good candidate for a factor involved in RNA stabilization or subsequent steps of gene expression, such as 5´ processing or translation. Binding of proteins, with molecular weights in the range of 47 kDa, to 5´ regions of various chloroplast mRNAs in C. reinhardtii has been reported (Danon and Mayfield, 1991; Zerges and Rochaix, 1994; Hauser et al., 1996; Chapter 12, Hauser et al.) raising the question of whether these binding activities are transcript-specific or not. Although only the isolation of these factors and/or their respective genes will allow one to resolve this problem, the picture emerging from competition experiments is that numerous proteins in this size range are capable of interacting with RNA molecules. Depending on the sample preparation procedure used, one might enrich for particular proteins. For example, some of the methods involve whole cell extracts further purified on heparin-affinity columns (Danon and Mayfield, 1991; Hauser et al., 1996) while others start from purified chloroplasts lysed with (Nickelsen et al., 1994) or without (Zerges and Rochaix, 1994) solubilization of membrane proteins. In addition to a biochemical approach to identify factors regulating RNA stability, the isolation of nuclear genes from mutants listed in Table 2 is underway. The Nac2 cDNA has recently been cloned by using a strategy that involves the complementation of a photosynthetic mutant with a cosmid library and subsequent isolation of the introduced DNA by plasmid rescue (J. Nickelsen and J.-D. Rochaix, unpublished). The primary structure of the cDNA revealed a polypeptide with some interesting features. The main portion of the protein consists of 9 TPR characteristic, 34 amino-acid (tetratricopeptide) motif repeats (Goebl and Yanagida,

Chapter 9

Chloroplast RNA Stability

1991). TPR domains have been found in a variety of unrelated proteins and have been proposed to mediate protein-protein interactions (Lamb et al., 1995) suggesting that the Nac2 factor could be part of a multisubunit complex. The C-terminus contains a hydrophobic stretch capable of forming a single putative membrane spanning helix, and thus this factor is tentatively classified as a peripheral membrane protein. Based on the structure of the Nac2 factor as well as on the in vivo and in vitro data concerning the psbD leader analyses, the following model could be proposed on how the stabilization of psbD RNA might be achieved at the molecular level (Fig. 2). It is assumed that the nascent 5´ end of the psbD mRNA is bound by the 47 kDa protein, which could interact either directly or indirectly with the Nac2 factor and other proteins. As a consequence the whole RNAprotein complex would be anchored in a membrane system, e.g. the envelope or the thylakoids, by the hydrophobic tail of Nac2. Data obtained so far do not allow a precise localization of the Nac2 factor, but in view of information supporting the idea of a close relationship of these two major chloroplast membrane systems, with thylakoids emerging from the envelope membrane (Hoober et al., 1994; Chapter 19, Hoober et al.), both could potentially be involved in the biogenesis of Photosystem II. The RNA binding could result in targeting of the psbD transcript to its final destination for translation following the model of cotranslational membrane insertion of D1 protein in higher plants (Kim et al., 1991). Then 5´ processing and/or ribosomal loading would take place, with 5´ maturation possibly having the function of allowing correct binding of ribosomes that might otherwise be sterically blocked by the Nac2 complex. In addition, 5´ processing might fulfill the function of regenerating active Nac2 complex that is ready for binding to another psbD mRNA molecule. The idea of a membrane-associated stabilization complex is further supported by the observation that the RNA binding activity of the 47 kDa protein can only be detected in the presence of anionic detergents like Triton X-100 (Nickelsen et al., 1994). This hypothesis, which remains to be tested, takes into account the relationship between the different processes occurring on the 74 nt psbD leader, which have to follow a highly coordinated program in terms of spatial and temporal regulation. The ongoing isolation of additional genes involved in transcript-specific RNA stabilization in C. rein­

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hardtii will add more information on the molecular aspects of this process and will allow us to elucidate whether the mechanisms involved have common features. The psbB RNA stability mutant 222E (Table 2) has recently been complemented by using an indexed cosmid library (Zhang et al., 1994) and based on the genomic subclone a cDNA clone has been isolated that complements the mutation (F. Vaistij, M. Goldschmidt-Clermont and J.-D. Rochaix, personal communication). Also, the exhibiting a defect in petA insertional mutant mRNA accumulation, has been complemented by a 18 kb wild-type genomic fragment (Gumpel et al., 1995).

C. Chloroplast Ribonucleases The current models of differential mRNA stabili­ zation in chloroplasts predict distinct RNA elements that interact with trans-acting factors to protect the RNA against nucleolytic attack. Thus, an additional determinant of transcript decay that has to be considered is the chloroplast RNA degradation machinery. Work on higher plants has recently provided insights into these ribonucleases and their molecular organization. A 54 kDa protein in mustard and a 41 kDa from spinach have been shown to have intrinsic endonucleolytic activities that could be involved in 3´ end maturation and/or RNA degradation (Nickelsen and Link, 1993; Yang et al., 1996). Furthermore, a model has been proposed in which 3´ end maturation is mediated by a high molecular weight complex containing ancillary RNA binding proteins and endo- as well as 3´–5´ exonucleases resembling RNase E and polyribonucleotide phosphorylase from E. coli, respectively (Hayes et al., 1996). Much less is known about the presence and nature

160 of ribonucleases in chloroplasts of C. reinhardtii, although it has been shown that the 3´ end formation of atpB transcripts is mediated by an endonucleolytic cut downstream of the mature 3´ terminus near an essential stem-loop structure (Section IVA.1) and subsequent 3´–5´ exonucleolytic trimming of the last 10 nt (Stern and Kindle, 1993). The activity of one or more endonucleases in chloroplast lysates has also been detected when the pathway of psbD leader RNA degradation was analyzed in vitro (Section IVA.2). No 5´–3´ exonucleolytic activities have been detected in chloroplasts to date. But, recently, the in vivo analysis of reporter genes in C. reinhardtii demonstrated that poly G tracts—that are known to impede the activity of 5´–3´ exonucleases in yeast (Vreken and Raue, 1992; Section IVA.1)—can stabilize chloroplast transcripts when they are introduced into 5´ leader regions (Drager et al., 1998), This suggests that 5´–3´ exonucleases might also exist in chloroplasts, though other expanations for the observed effects are possible. Hence, identification and characterization of chloroplast ribonucleases in C. reinhardtii represents one of the challenges for future molecular research work.

D. RNA Stability and Translation Besides the putative existence of particular RNA stabilizing protein factors interacting with their cognate target sites on chloroplast transcripts, another aspect concerning RNA turn-over is the effect of ribosome assembly on the stability of translated mRNAs. A complex relationship between these two processes has been documented in both prokaryotes and eukaryotes (Petersen, 1993; Jacobson and Peltz, 1996). Recent data on this subject from C. reinhardtii are summarized below. Impairment of translation by mutations in the psbA (Mayfield et al., 1994) and petD (Sakamoto et al., 1994) 5´ untranslated regions was accompanied by a significant decrease in RNA levels. However, as already mentioned (IVA.2), it is possible that these cis-acting mutations affect RNA stability directly. Another example is the chloroplast mutant Fud47, which harbors a 42 bp direct repeat within the psbD gene (Erickson et al., 1986) and accumulates reduced levels of psbD mRNA, suggesting a stabilization effect of polysomal assembly. Also in this case, it is possible that the direct repeat in Fud47 forms a cryptic signal leading to enhanced degradation by

Jörg Nickelsen plastid ribonucleases. A second class of translational mutants exhibits no drastic changes in the levels of the transcripts. Among these are the nuclear mutants F35 and F34 affected in translation of psbA (Girard-Bascou et al., 1992) and psbC (Rochaix et al., 1989), respectively. Also psbD ATG-initiation codon mutants that have been introduced into the chloroplast genome (J.-D. Rochaix, unpublished results) accumulate wild-type levels of psbD mRNA. Likewise, only a slight decrease in petD RNA levels was observed after changing the start codon to AUU or AUC although this led to a five- to ten-fold reduction in the trans­ lation rate of subunit IV of the cytochrome complex (Chen et al., 1993). Finally, nuclear mutants deficient in atpA and psbD translation contain increased levels of the corresponding transcripts (Drapier et al., 1992; Kuchka et al., 1988). In addition, a psaB frame shift mutation also leads to a more than twofold increase in psaB mRNA, suggesting that reduced translation can stabilize chloroplast transcripts (Xu et al., 1993). A negative effect on psaB RNA accumulation was observed when the stop codon was moved further downstream, thereby extending the translated region of the psaB protein coding region. This supports the idea of cotranslational mRNA destabilization in chloroplasts of C. reinhardtii (Lee et al., 1996). To test this hypothesis, the impact of translation inhibitors like chloramphenicol and lincomycin on the accumulation of several chloroplast transcripts was examined. Interestingly, a significant stabilization effect of both chemicals was observed for psaA and psaB RNAs, while other RNAs, e.g. rbcL and atpB, remained unaffected. This points to gene-specific mechanisms of RNA degradation that can be linked to active translation in the case of the psaA and psaB transcripts. Secondly, the data show that in these two cases, polysomal assembly does not play an important role for RNA stabilization, since chloramphenicol inhibits peptidyl-transferase activity and blocks the ribosomes on the mRNA molecules. In contrast, lincomycin acts on the formation of the first peptide bond of newly initiated polypeptides leading to polysome run-off that forces the mRNA into a non­ polysomal state. Taken together, no general relationship between RNA stability and translation appears to exist. Nevertheless, gene-specific effects suggest that, in some cases, active translation can influence the halflife of the RNA. Assuming that defined cis-acting

Chapter 9

Chloroplast RNA Stability

RNA elements mark hotspots for the regulation of RNA stability by being exceptionally accessible or, alternatively, exceptionally protected, one might speculate that the ribosome moving across these sites can increase or decrease the degradation process. This could explain why particular transcripts behave differently when they are not translated and, depending on the location of the cis-element, blocking of translation at the level of initiation, elongation, or termination could result in different effects. Thus, gene-specific RNA stabilization after antibiotic treatment of cells could be a consequence of special RNA stability determinants within psaA and psaB mRNAs.

V. Conclusions and Perspectives The role of differential RNA synthesis and degradation as factors contributing to the process of regulated gene expression has been documented in almost all organisms examined to date. In chloroplasts as well as in mitochondria (Margossian and Butow, 1996) mRNA stabilization appears to play a key role in determining the levels of organellar transcripts. Analyses of nuclear and chloroplast mutants in C. reinhardtii have revealed that mRNA decay is not a random process in which non-specific nucleases degrade RNA based only on target size or ribosomal association. Instead, a picture of gene-specific stabilization effects is emerging in which defined cis-acting elements and their cognate trans-acting protein factors allow precise determination of particular RNA steady-state levels. Apparently, both 5´ and 3´ untranslated regions of chloroplast transcripts carry these cis-elements, and in the future, one of the most important questions will be to determine the rate limiting steps of RNA decay that cause the irreversible functional inactivation of an RNA molecule (Petersen 1992). As the 5´ leader mediates degradation of psbD, psbB and petD mRNAs in at least three nuclear mutants (nac2-26, 222E and F16, respectively), 5´ regions appear to play an important role in the control of these initial events. Consequently, the algal cell would benefit from avoiding partial translation of transcripts degraded from the 3´ end. However, it should be taken into account that a mutated nuclear or chloroplast background represents an artificial situation that might lead to misinterpretation of the results.

161 The available techniques for nuclear as well as chloroplast transformation will allow the identi­ fication of more trans-acting factors mediating genespecific RNA stabilization. The TPR domain on the Nac2 protein suggests that multisubunit complexes are involved, and possibly that these complexes are, at least in some cases, associated with the chloroplast membrane system. Together with the available in vitro systems for monitoring RNA degradation and RNA-protein complex formation, it should be feasible to characterize the molecular mechanics of RNA stability in more detail. One emphasis will be to elucidate the interrelationship ofprocesses involving 5´ leader regions; namely RNA stabilization, RNA targeting, 5´ processing, and translation. Furthermore, it will be exciting to test whether processes recently discovered in higher plants like RNA polyadenylation also occur in C. reinhardtii and to study their role in RNA metabolism. Finally, with the identification of factors involved in higher plant plastid RNA stabilization (Meurer et al., 1996), similarities and differences observed will provide ideas of how mechanisms of regulated RNA decay developed during evolution.

Acknowledgments I want to thank J.-D. Rochaix for his support and many stimulating discussions. In addition, I wish to thank those who kindly provided unpublished data of their work and E. Schmidt for critical reading of the manuscript. Furthermore, I would like to thank U. Kück for providing basic support and laboratory space. This work is supported by the Deutsche Forschungsgemeinschaft.

References Abler ML and Green PJ (1996) Control of mRNA stability in higher plants. Plant Molec Biol 32: 63–78 Blowers A, Klein U, Ellmore GS and Bogorad L (1993) Functional in vivo analyses of the 3´ flanking sequences of the Chlamydomonas chloroplast rbcL and psaB genes. Mol Gen Genet 238: 339–349 Bouvet P and Belasco JG (1992) Control of RNase E-mediated RNA degradation by 5´-terminal base pairing in E. coli. Nature 360: 488–491 Boynton JE, Gillham NW, Harris EH, Hosler JP, Johnson AR, Jones BL, Randolph-Anderson, Robertson TM, Klein KB, Shark B and Sanford J (1988) Chloroplast transformation in Chlamydomonas with high velocity microprojectiles. Science 240: 1534–1538

162 Chen X, Kindle K and Stern DB (1993) Initiation codon mutations in the Chlamydomonas chloroplast petD gene result in temperature-sensitive photosynthetic growth. EMBO J 12: 3627–3635 Danon A and Mayfield SP (1991) Light-regulated translational activators: identification of chloroplast gene-specific mRNA binding proteins. EMBO J 10: 3993–4002 De Vitry C, Olive J, Drapier D, Recouvreur M and Wollman F­ A (1989) Posttranslational events leading to the assembly of Photosystem II complex: a study using photosynthesis mutants from Chlamydomonas reinhardtii. J Cell Biol 109: 991–1006 Drager RG, Zeidler M, Simpson CL and Stern DB (1996) A chloroplast transcript lacking the 3´ inverted repeat is degraded by 3´–5´ exoribonuclease activity. RNA 2: 652–663 Drager RG, Girard-Bascou J, Choquet Y, Kindle KL and Stern DB (1997) In vivo evidence for 5´–3´ exoribonuclease degradation of an unstable chloroplast mRNA. Plant J 13: 85–96 Drapier D, Girard-Bascou J and Wollman F-A (1992) Evidence for nuclear control of the expression of the atpA and atpB chloroplast genes in Chlamydomonas. Plant Cell 4: 283–295 Gagne G and Guertin M (1992) The early genetic response to light in the green unicellular alga Chlamydomonas eugametos grown under light/dark cycles involves genes that represent direct responses to light and photosynthesis. Plant Mol Biol 18: 429–445 Gillham NW, Boynton JE and Hauser CR (1994) Translational regulation of gene expression in chloroplast and mitochondria. Annu Rev Genet 28: 71–93 Goebl M and Yanagida M (1991) The TPR snap helix: A novel protein repeat motif from mitosis to transcription. Trends Biochem Sci 16: 173–177 Gruissem W and Tonkyn JC (1993) Control mechanisms of plastid gene expression. Crit Rev Plant Sci 12:19–55 Guertin M and Bellemare G (1979) Synthesis of chloroplast ribonucleic acid in Chlamydomonas reinhardtii toluene-treated cells. Eur J Biochem 96: 125–129 Gumpel NJ, Ralley L, Girard-Bascou J, Wollman F-A, Nugent JHA and Purton S (1995) Nuclear mutants of Chlamydomonas reinhardtii defective in the biogenesis of the cytochrome complex. Plant Molec Biol 29: 921–932 Hahn D, Bennoun P and Kück U (1996) Altered expression of nuclear genes encoding chloroplast polypeptides in nonphotosynthetic mutants of Chlamydomonas reinhardtii: evidence for post-transcriptional regulation. Mol Gen Genet 252: 362–370 Hansen MJ, Chen L, Fejzo MLS and Belasco JG (1994) The ompA 5´ untranslated region impedes a major pathway for mRNA degradation in Escherichia coli. Molec Microbiol 12: 707–716 Harris EH (1989) The Chlamydomonas source book. Academic Press, San Diego Hauser CR, Gillham NW and Boynton JE (1996) Translational regulation of chloroplast genes: proteins binding to the 5´ UTRs of chloroplast mRNAs in Chlamydomonas reinhardtii. J Biol Chem 271: 1486–1497 Hayes R, Kudla J, Schuster G, Gabay L, Maliga P and Gruissem W (1996) Chloroplast mRNA 3´ end processing by a high molecular weight protein complex is regulated by nuclear encoded RNA binding proteins. EMBO J 15:1132–1141 Herrin DL, Michaels AS and Paul A-L (1986) Regulation of

Jörg Nickelsen genes encoding the large subunit of ribulose-1,5-bisphosphate carboxylase and the Photosystem II polypeptides D1 and D2 during the cell cycle of Chlamydomonas reinhardtii. J Cell Biol 103: 1837–1845 Hong S and Spreitzer RJ (1994) Nuclear mutation inhibits expression of the chloroplast gene that encodes the large subunit of ribulose-1,5-bisphosphate carboxylase/oxygenase. Plant Physiol 106: 673–678 Hoober JK, White RA, Marks DB and Gabriel JL (1994) Biogenesis of thylakoid membranes with emphasis on the process in Chlamydomonas. Photosynth Res 39: 15–31 Hwang S and Herrin DL (1994) Control of the gene transcription by the circadian clock in Chlamydomonas reinhardtii. Plant Molec Biol 26: 557–569 Jacobshagen S, Kindle KL and Johnson CM (1996) Transcription of CABII is regulated by the biological clock in Chlamydomonas reinhardtii. Plant Molec Biol 31: 1173–1184 Jacobson A and Peltz SW (1996) Interrelationships of the pathways of mRNA decay and translation in eukaryotic cells. Annu Rev Biochem 65: 693–739 Kim J, Gamble-Klein P and Mullet JE (1991) Ribosomes pause at specific sites during synthesis of membrane bound chloroplast reaction center protein D1. J Biol Chem 266: 14931–14938 Klaff P (1995) mRNA decay in spinach chloroplasts: psbA mRNA degradation is initiated by endonucleolytic cleavages within the coding region. Nucl Acids Res 23: 4885–4892 Kuchka MR, Goldschmidt-Clermont M, van Dillewijn J and Rochaix J-D (1989) Mutation at the Chlamydomonas nuclear NAC2 locus specifically affects stability of the chloroplast psbD transcript encoding polypeptide D2 of PS II. Cell 58: 869–876 Kudla J, Hayes R and Gruissem W (1996) Polyadenylation accelerates degradation of chloroplast mRNA. EMBO J 15: 7137–7146 Kuhlemeier C (1992) Transcriptional and post-transcriptional regulation of gene expression in higher plants. Plant Molec Biol 19: 1–14 Lamb JR, Tugendreich S and Hieter P(1995)Tetratrico peptide repeat interactions: to TPR or not to TPR. Trends Biochem Sci 20: 257–259 Lee H, Bingham SE and Webber AN (1996) Function of 3´ noncoding sequences and stop codon usage in expression of the chloroplast psaB gene in Chlamydomonas reinhardtii. Plant Mol Biol 31: 337–354 Leu S, White D and Michaels A (1990) Cell cycle-dependent transcriptional andpost-transcriptionalregulation ofchloroplast gene expression in Chlamydomonas reinhardtii. Biochem Biophys Acta 1049: 311–317 Margossian SP and Butow RA (1996) RNA turnover and the control of mitochondrial gene expression. Trends Biochem Sci 21: 392–396 Mayfield SP, Cohen A, Danon A and Yohn CB (1994) Translation of the psbA mRNA of Chlamydomonas reinhardtii requires a structured element contained within the 5´ untranslated region. J Cell Biol 127: 1537–1545 Meurer J, Berger A and Westhoff P (1996) A nuclear mutant of Arabidopsis with impaired stability on distinct transcripts of the plastid psbB , psbD/C, ndhH and ndhC operons. Plant Cell 8: 1193–1207 Monod C, Goldschmidt-Clermont M and Rochaix J-D (1992)

Chapter 9 Chloroplast RNA Stability Accumulation of chloroplast psbB RNA requires a nuclear factor in Chlamydomonas reinhardtii. Mol Gen Genet 231: 449–459 Nickelsen J and Link G (1993) The 54 kDa RNA binding protein from mustard chloroplasts mediates endonucleolytic transcript 3´ end formation in vitro. Plant J 3: 537–544 Nickelsen J and Rochaix J-D (1994) Regulation of the synthesis of D1 and D2 proteins of Photosystem II. In: Baker NR and Bowyer JR (eds) Photoinhibition of photosynthesis: From Molecular Mechanisms to the Field, pp 179–194. Information Press Ltd, Oxford Nickelsen J, van Dillewijn J, Rahire M and Rochaix J-D (1994) Determinants for stability of the chloroplast psbD RNA are located within its short leader region in Chlamydomonas reinhardtii. EM BO J 13: 3182–3191 Petersen C (1992) Control of functional mRNA stability in bacteria: Multiple mechanisms of nucleolytic and non­ nucleolytic inactivation. Mol Microbiol 6: 277–282 Purton S and Rochaix J-D (1994) Complementation of a Chlamydomonas reinhardtii mutant using a genomic cosmid library. Plant Mol Biol 24: 533–537 Rochaix J-D (1995) Chlamydomonas reinhardtii as the photosynthetic yeast. Annu Rev Genet 29: 209–230 Rochaix J-D (1996) Post-transcriptional regulation of chloroplast gene expression in Chlamydomonas reinhardtii. Plant Mol Biol 32: 327–341 Sakamoto W, Kindle KL and Stern DB (1993) In vivo analysis of Chlamydomonas chloroplast petD gene expression using stable transformation of beta-glucuronidase translational fusions. Proc Natl Sci USA 90: 497–501 Sakamoto W, Chen X, Kindle KL and Stern DB (1994) Function of the Chlamydomonas reinhardtii petD 5´ untranslated region in regulating the accumulation of subunit IV of the cytochrome complex. Plant J 6: 503–512 Salvador ML, Klein U and Bogorad L (1993a) Light regulated and endogenous fluctuations of chloroplast transcript levels in Chlamydomonas. Regulation by transcription and RNA degradation. Plant J 3: 213–219

163 Salvador ML, Klein U and Bogorad L (1993b) 5´ sequences are important positive and negative determinants of the longevity of Chlamydomonas chloroplast gene transcripts. Proc Natl Acad Sci USA 90: 1556–1560 Sieburth LE, Berry-Lowe S and Schmidt G W (1991) Chloroplast RNA stability in Chlamydomonas: rapid degradation of psbB and psbC transcripts in two nuclear mutants. Plant Cell 3: 175– 189 Stern DB and Kindle KL (1993) 3´ end maturation of the Chlamydomonas reinhardtii chloroplast atpB mRNA is a twostep process. Molec Cell Biol 13: 2277–2285 Stern DB, Radwanski ER and Kindle KL (1991) A 3´ stem/loop structure of the Chlamydomonas chloroplast atpB gene regulates mRNA accumulation in vivo. Plant Cell 3: 285–297 Sugita M and Sugiura M (1996) Regulation of gene expression in chloroplasts of higher plants. Plant Molec Biol 32: 315–326 Vreken P and Raue HA (1992) The rate limiting step in yeast PGK1 mRNA degradation is an endonucleolytic cleavage in the 3´-terminal part of the coding region. Mol Cell Biol 12: 2986–2996 Xu R, Bingham SE and Webber AN (1993) Increased mRNA accumulation in a psaB frame-shift mutant of Chlamydomonas reinhardtii suggests a role for translation in psaB mRNA stability. Plant Molec Biol 22: 465–474 Yang J, Schuster G and Stern DB (1996) CSP41, a sequencespecific chloroplast mRNA binding protein, is an endo­ ribonuclease. Plant Cell 8: 1409–1420 Zerges W and Rochaix J-D (1994) The 5´ leader of a chloroplast mRNA mediates the translational requirements for two nucleusencoded functions in Chlamydomonas reinhardtii. Mol Cell Biol 14: 5268–5277 Zhang H, Herman PL and Weeks DP (1994) Gene isolation through genomic complementation using an indexed library of Chlamydomonas reinhardtii DNA. Plant Mol Biol 24: 663– 672

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Chapter 10 Chloroplast RNA Synthesis and Processing David B. Stern and Robert G. Drager

Boyce Thompson Institute for Plant Research at Cornell University,

Tower Road, Ithaca, NY 14853, U.S.A.

Summary I. Transcription of Chloroplast Genes A. General Considerations B. RNA Polymerase 1. Plastid-Encoded rpo Genes 2. Nucleus-Encoded Factors 3. Evidence for a Nucleus-Encoded Chloroplast RNA Polymerase C. Promoters 1. Prokaryotic-Like –10/–35 Promoters 2. Other Promoter Types 3. Transcriptional Regulation by DNA Topology D. Transcription Termination 1. Inverted Repeats Are Inefficient Transcription Terminators 2. Consequences of Transcriptional Read-Through II. Processing of Chloroplast mRNAs A. General Considerations B. 5´-End Processing 1. The Extent of 5´-End Processing 2. Possible Mechanisms for 5´-End Processing 3. The Relationship Between 5´-End Processing and mRNA Accumulation C. 3´ End Processing 1. Examples of 3´-End Processed Transcripts 2. Mechanisms of 3´-End Processing 3. Nuclear Mutations Affecting Chloroplast mRNA 3´-End Formation 4. Does the 3´ UTR Regulate mRNA Stability? Acknowledgments References

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Summary Transcription and RNA maturation are two essential steps in gene expression. In chloroplasts, transcription is carried out by at least two biochemically and genetically separable activities, which may participate in establishing different basal expression rates for ribosomal RNAs, transfer RNAs and protein-coding genes. Because chloroplast RNA polymerases do not generally terminate transcription at sites corresponding to the 3´ termini of mature transcripts, these termini must be formed by RNA processing events. In Chlamydomonas reinhardtii chloroplasts, it appears that most or all transcript 5´-ends are also formed by RNA processing rather than by transcription initiation. Thus, RNA processing converts primary transcripts of generally unknown dimensions to the mature, accumulating transcripts. Molecular, genetic and biochemical approaches have been

J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 165–181. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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used to unravel the chloroplast transcription and RNA processing machinery, with the most information gained to date from the analysis of chimeric reporter genes introduced into chloroplasts by biolistic transformation. The picture painted by these data reveals both similarities and differences between these processes in Chlamydomonas and land plants. However, some perceived differences, particularly based on the phenotypes of nuclear mutants which affect chloroplast mRNA metabolism, may reflect selection or screening procedures and thus may mask an overall congruity between gene expression mechanisms in the chloroplasts of all organisms.

I. Transcription of Chloroplast Genes

B. RNA Polymerase

A. General Considerations

1. Plastid-Encoded rpo Genes

While most genes in land plant cpDNA are organized into clusters or operons, Chlamydomonas chloroplast genes are somewhat dispersed. These organizational differences are reflected in the patterns of accumu­ lating transcripts, which are often very complex in land plants, but usually simple in Chlamydomonas. However, Chlamydomonas cpDNA contains some conserved gene clusters, such as the ribosomal protein genes which resemble the E. coli S10 and spc operons (Boudreau et al, 1994), and co-transcription of nonconserved groups of genes does occur (Levy et al., 1997). As discussed in the second part of this chapter, rapid RNA processing can also lead to the illusion of independent transcription of two genes that are in fact co-transcribed. The enzymes that carry out chloroplast trans­ cription are poorly characterized in Chlamydomonas, with somewhat more information being available for land plants and Euglena. In this chapter, therefore, data for other organisms are also discussed. One reason for the lack of information regarding Chlamydomonas chloroplast RNA polymerases is that Chlamydomonas chloroplasts are somewhat difficult to isolate in large quantities, which has limited the amount of biochemical work that has been carried out to date. Also, because transcription initiation is not generally considered to be an important control point for chloroplast gene expression in Chlamydomonas, less experimental emphasis has been put on this area than on post­ transcriptional controls.

Like the chloroplast genomes of land plants (Ohyama et al., 1986; Shinozaki et al., 1986; Hiratsuka et al., 1989; Maier et al., 1995), Euglena gracilis (Hallick et al., 1993), and Porphyra purpurea (Reith and Munholland, 1993), Chlamydomonas spp. cpDNAs contains open reading frames encoding potential proteins with homology to subunits of E. coli RNA polymerase. While plants contain the genes rpoA, rpoB, rpoC1 and rpoC2, encoding putative subunits, in Chlamydomonas spp. cpDNA and only rpoB1, rpoB2 and rpoC2 have been located (Boudreau et al., 1994). In Chlamydomonas, the subunit has been split into two proteins, while only the C-terminal half of the E. subunit is encoded in cpDNA. Since the sequence of the Chlamydomonas chloroplast genome has not been completed, whether the remaining genes are present is unknown. Biochemical evidence from maize (Hu and Bogorad, 1990; Hu et al., 1991) and pea (Little and Hallick, 1988) suggests that the chloroplast-encoded RNA polymerase subunits are present in vivo and form part of the RNA polymerase. In addition, when the rpoB gene was deleted from tobacco cpDNA by biolistic transformation, transcription of many chloroplast genes was abolished (Allison et al., 1996). Supporting data were also obtained from a barley mutant, albostrians, which was reported to lack chloroplast ribosomes and also was deficient in transcription of a class of chloroplast genes (Hess et al., 1993). When similar deletions were attempted in Chlamydomonas chloroplasts, however, the cells remained heteroplasmic, suggesting that the rpoB1, rpoB2 and rpoC2 genes performed essential functions (Fischer et al., 1996). Since the deletions were performed by replacement of the coding regions with a selectable marker cassette which included the aadA coding region driven by the atpA promoter, it is possible that the rpo genes are not essential for cell

Abbreviations: cpDNA – chloroplast DNA; cpRNA – chloroplast RNA; IR – inverted repeat; LRP – light-regulated promoters; NEP – nucleus-encoded RNA polymerase; Rubisco – ribulosel,5-bisphosphate carboxylase/oxygenase; TAC – transcriptionally active chromosome; UTR – untranslated region

Chapter 10 Chloroplast Transcription viability per se, but rather for the expression of the selectable marker. In the case of the tobacco rpoB deletion, the aadA cassette was driven by a modified rRNA promoter; rRNA genes are thought to be transcribed by a nucleus-encoded chloroplast RNA polymerase (see below). The lingering hetero­ plasmicity of genomes containing the rpoB deletion in Chlamydomonas, however, argues strongly that these genes play an important role in chloroplast gene expression, most likely in transcription.

167 known gene-specific translational activators or RNA stabilizers, do in fact exist. If they do, then one would expect mutants to be in hand already; however, one cannot exclude the possibility that one or a very few genes are controlled in this way, much as psbD/C, rrn16, and petG are the only reported examples of gene-specific transcriptional regulation in land plants (reviewed in Stern et al., 1997).

3. Evidence for a Nucleus-Encoded Chloroplast RNA Polymerase

2. Nucleus-Encoded Factors E. coli RNA polymerase holoenzyme contains a sigma factor in addition to the core subunits; however, genes encoding putative factors have not been found in cpDNA. This suggests that might be nucleus-encoded in plants. Recently, a nuclear SigA gene was reported for the red alga Cyanidium caldarium, providing additional evidence for this possibility (Liu and Troxler, 1996; Tanaka et al., proteins 1996). The partial purification of from land plant (Lerbs et al., 1988; Tiller and Link, 1993) and Chlamydomonas (Surzycki and Shellen­ barger, 1976; Troxler et al., 1994) chloroplasts has been reported. The next several years should see the biochemical data meshing with the molecular genetic. Another line of evidence for nucleus-encoded RNA polymerase components would be the isolation of nuclear mutants defective in chloroplast transcription. However, in land plants such mutants have not been reported and in Chlamydomonas, only a single nuclear mutant strain of this class is known (76-5EN). In 76­ 5EN, transcription of the rbcL gene is reduced, based on a run-on assay (Hong and Spreitzer, 1994). However, the reduction in the transcription rate may not fully account for the Rubisco deficiency of these cells. The dearth of transcription defects among the many non-photosynthetic Chlamydomonas strains that have been isolated has, then, two possible explanations. One is simply that the target genes do not exist, and the other is that the target genes are essential for cell viability. The latter limitation can be overcome by the isolation of temperature-sensitive mutants, of which there are numerous examples for the RNA polymerases of yeast and E. coli. While it is highly likely that one or more nuclear genes encode a subunit of chloroplast RNA polymerase in Chlamydomonas, an additional question is whether gene-specific transcription factors, analogous to well-

Several lines of evidence suggest that chloroplasts contain both nucleus- and chloroplast-encoded RNA polymerase activities. First, the existence of two biochemically-separable polymerase activities has been reported in Euglena gracilis and in land plants (Greenberg et al., 1984; Pfannschmidt and Link, 1994). These activities are a membrane-associated DNA-protein complex called transcriptionally active chromosome (TAC), thought to transcribe principally rRNA genes, and a soluble RNA polymerase that recognizes canonical E. coli-like promoters. Since only the E. coli type of RNA polymerase gene is found in cpDNA, this suggests that TAC is encoded in the nucleus. A second line of evidence is the transcription of plastid genes in organisms that lack either plastid ribosomes (Hess et al., 1993), or functional plastid-encoded RNA polymerase (Wolfe et al., 1992; Allison et al., 1996). Additionally, the observation that chloroplasts contain a T7-like RNA polymerase activity (Lerbs-Mache, 1993), and the existence of multiple promoter types (see Section I.C) are evidence for multiple RNA polymerases or at least different transcription factors. While RNA polymerases have been purified from the chloroplasts of E. gracilis and several plants, relatively little has been done in the case of Chlamydomonas. Fractionation of total cell RNA polymerases revealed a rifampicin-sensitive, activity that had a preference for cpDNA (Surzycki and Shellenbarger, 1976). The activity that incorporates nucleoside triphosphates into cpRNA of toluene-treated cells has similar characteristics (Guertin and Bellemare, 1979), and these features are consistent with an E. coli-like RNA polymerase. However, when Chlamydomonas RNA polymerases were separated based on their the ability to transcribe trnE1, encoding and activity was insensitive to both rifampicin, but was sensitive to heparin (Jahn, 1992).

168 Although rifampicin insensitivity distinguishes this activity from the E. coli-like polymerase, heparin sensitivity is typical of soluble chloroplast RNA polymerase activities from land plants when used for in vitro transcription experiments (Gruissem et al., 1983). The trnE1 transcription activity might be nucleus-encoded, especially since it has RNA polymerase III characteristics based on promoter deletion analysis of trnE1 (Section I.C).

David B. Stern and Robert G. Drager promoters are present, but inactive. For example, the spinach rrn16 gene has two upstream –35/–10 promoters which are active in vitro but not in vivo. Further analysis revealed that a third promoter of another type was located between them, and that initiation at this promoter blocked access of RNA polymerase to the E. coli-like promoters (Iratni et al., 1994).

2. Other Promoter Types C. Promoters 1. Prokaryotic-Like –10/–35 Promoters Canonical E. coli promoters consist of –35 and –10 elements with the consensus sequences TTGACA and TATAAT. Sequences similar or identical to these were recognized when chloroplast genes were first isolated from plants, and these were later verified to be bona fide promoter elements using in vitro transcription systems, especially from spinach (Gruissem and Zurawski, 1985a,b). In addition, it has been possible to analyze plastid gene promoters in E. coli. For example, mutations in the maize atpB promoter had the same effects either in E. coli or in a chloroplast in vitro transcription system (Bradley and Gatenby, 1985). Several Chlamydomonas chloroplast promoters have been identified by their ability to direct transcription of reporter genes in transformed chloroplasts, and alignment of these and additional sequences have revealed E. coli-like promoters upstream of some Chlamydomonas chloroplast genes, for example rrn16 and rbcL (Klein et al., 1992). Thus, the –35/–10 promoter type is widespread in chloroplast genomes, and in most cases has sequence requirements indistinguishable from those of E. coli promoters. At least two variants of the scenario described in the preceding paragraph are also known. In one case, the –35 element is apparently absent, leading to a ‘–10’ type of promoter. A consensus sequence TATAATAT has been proposed for Chlamydomonas, based on deletion analysis of the atpB promoter and inspection of sequences upstream of known mRNA 5´ termini. However, because Chlamydomonas chloroplast intergenic regions are extremely rich in A and T, and because many 5´ termini may be generated by RNA processing rather than by transcription initiation (Section II. A), these sequence alignments should be viewed with caution. A second variant of –35/–10 promoters occurs when these

In land plants, at least three types of chloroplast promoters are known apart from the prokaryotic types (–35/–10 or–10 only). These are so-called NEP promoters, driven by a nucleus-encoded RNA polymerase, light-regulated promoters (LRPs), and internal tRNA promoters. NEP promoters are those that principally drive components of the gene expression apparatus, for example rRNAs, ribosomal proteins and RNA polymerase subunits. The NEP promoters of plastid 16S rRNA genes are highly conserved, with a G+C content of approximately 50% (Vera and Sugiura, 1995; Allison et al., 1996). NEP promoters are also found upstream of some photosynthetic genes such as atpB and atpI; however, these genes also have E. coli-like promoters (Hajdukiewicz et al., 1997; Kapoor et al., 1997). Based on continued initiation at NEP promoters in the absence ofchloroplast-encoded RNA polymerase (Hess et al., 1993; Allison et al., 1996; Hajdukiewicz et al., 1997), they are assumed to be transcribed by a nucleus-encoded enzyme. The best-characterized LRP drives blue lightinduced transcription of a major psbD/C transcript in barley (Sexton et al., 1990) and other land plant species (Christopher et al., 1992). Elements of this promoter type have been dissected using in vitro transcription (Kim and Mullet, 1995; Satoh et al., 1997) and in vivo using transcriptional fusions to the uidA reporter gene in transformed tobacco chloro­ plasts (Allison and Maliga, 1995). These experiments revealed that the LRP contains both classical –35/– 10 elements as well as enhancing and light-responsive elements not found in other promoters. While Chlamydomonas has not been demonstrated to have this type of response to high light levels, there is strong evidence for circadian control of transcript levels in this organism. The amplitude of this variation can be more than ten-fold, and is exhibited by a number of genes, including rrn16, tufA and genes for photosynthetic proteins (Hwang et al., 1996). That

Chapter 10 Chloroplast Transcription transcription rate oscillation is circadian rather than light-controlled was shown by leaving lightsynchronized cells in continuous light or darkness, and finding that the circadian rhythms persisted for two days or more. Similar results were obtained in another study, with the additional finding that the psaB transcription rate is controlled by light rather than by endogenous rhythm (Salvador et al., 1993b). The promoter elements that mediate this trans­ criptional regulation remain to be defined. In spinach chloroplasts, two classes of tRNA gene promoters have been found. For example, trnM2 has a –35/–10 promoter with standard functional elements (Gruissem and Zurawski, 1985b). The trnS1 and trnR1 genes, however, do not require upstream elements for in vitro transcription (Gruissem et al., 1986), and thus resemble nuclear tRNA genes transcribed by RNA polymerase III. This latter class is also represented in Chlamydomonas chloroplasts, as evidenced by results obtained with an in vitro transcription system and the trnE1 gene (Jahn, 1992). Deletion analysis showed that neither 5´ nor 3´ flanking sequences were required for full activity, but that deletions into the coding region abolished in vitro transcription. Thus, trnE1 has an internal promoter. Whether this can be generalized to other Chlamydomonas chloroplast tRNA genes will require additional in vitro transcription analysis or deletion analysis in vivo. The occurrence of different classes of chloroplast RNA polymerases and of different promoter types in Chlamydomonas and land plants is summarized in Table 1.

3. Transcriptional Regulation by DNA Topology It is well-known that accessibility of promoters to RNA polymerase can influence transcription initiation rates. Unlike E. coli RNA polymerase, land plant chloroplast RNA polymerases have typically exhibited a preference for supercoiled over linear templates in vitro (Lam and Chua, 1987; Zaitlin et al., 1989), suggesting that DNA topology may play a role in establishing basal transcription rates of chloroplast genes. While similar in vitro transcription experiments have not been carried out for Chlamy­ domonas, treatment of cells with novobiocin, an inhibitor of DNA gyrase, has substantial effects on transcript accumulation patterns. It can be shown in vitro that Chlamydomonas chloroplasts have an ATPdependent supercoiling activity (Thompson and Mosig, 1985), and that the relative accumulation of

169

several transcripts is altered when this activity is inhibited by novobiocin (Thompson and Mosig, 1987), resulting in either increases or decreases for individual genes. These experiments suggested that torsional stress in DNA was being altered, which was subsequently examined by measuring the ability of an intercalating dye to bind to different regions ofthe Chlamydomonas chloroplast genome. It was found that DNA structure changed in response to light (Thompson and Mosig, 1990), consistent with a chromosome structure-mediated transcriptional response to light conditions. The light- and circadian rhythm-directed changes in chloroplast transcription rates that have been characterized (see previous section) may well result from, or require these alterations in chloroplast chromosome structure.

D. Transcription Termination 1. Inverted Repeats Are Inefficient Transcription Terminators The 3´-ends of most chloroplast protein-coding regions are flanked by inverted repeat sequences that can potentially form stem-loop structures. While these stem-loops superficially resemble bacterial terminators, they often lack the Urich tract immediately downstream that is required for efficient termination activity (Platt, 1986).

170 Measurements of termination using in vitro transcription systems from spinach showed that while terminators, for example the known E. coli trp attenuator and the bacteriophage T7 early terminator, could be recognized by RNA polymerase, chloroplast 3´ stem-loops were very inefficient terminators (Stern and Gruissem, 1987; Chen et al., 1990). A sole exception was trnS1, which had strong termination activity either within the tRNA coding region or downstream (Stem and Gruissem, 1987). The subsequent discovery that chloroplasts contain a T7-like RNA polymerase activity (Lerbs-Mache, 1993) raises the interesting question of whether the observed termination was due to a nuclear or chloroplast-encoded RNA polymerase activity. In Chlamydomonas chloroplasts, the termination efficiencies of 3´ UTRs have been measured by pulse-labeling RNA, and quantifying incorporation into sequences before and after possible of termination signals (Stern and Kindle, 1993; Rott et al., 1996). In these studies, no significant termination activity was seen (the atpB 3´ stem-loop imparted the highest level of transcription termination at ), even for known terminators. These results are ostensibly at odds with observations of polar effects of 3´ stem-loops on the transcription of downstream genes in transformed Chlamydomonas chloroplasts. For example, read-through of the rbcL 3´ stem-loop does not appear to occur when a reporter gene is flanked by three consecutive iterations ofthis sequence, because all accumulating transcripts end at the first stem-loop (Blowers et al., 1993). However, this result can also be explained by efficient processing at the first stem-loop, followed by degradation of downstream sequences. This is in fact what appears to occur at the 3´-end of the atpB gene (Stern and Kindle, 1993). Thus, what appears at first glance to be efficient termination may in fact represent efficient processing followed by or concurrent with polar RNA degradation. At an extreme, it can be envisioned that transcription termination simply does not occur in Chlamydomonas chloroplasts, at least not at a level sufficient to be a primary mechanism for generating 3´ termini. This would be akin to transcription of metazoan mitochondrial genomes, where the entire 16 kb genome is co-transcribed and subsequently processed to yield individual mRNAs and tRNAs (see Tracy and Stern, 1995 and references therein), although termination does occur by a special mechanism downstream of the rRNA genes (Daga et

David B. Stern and Robert G. Drager al., 1993). Another possibility is that transcription in chloroplasts is punctuated by tRNA genes capable of directing efficient termination (Stern and Gruissem, 1987), although this phenomenon has not yet been demonstrated in Chlamydomonas. Finally, termin­ ation may be stochastic, with the polymerase relying on the relative instability of RNA-DNA hybrids in the extremely A+T-rich intergenic regions in the chloroplast genome. Further studies in Chlamy­ domonas will probably have to await purification of the RNA polymerase or development of an in vitro transcription system.

2. Consequences of Transcriptional ReadThrough A lack of efficient transcription termination has several consequences for gene expression. First, rapid and precise RNA processing is required to generate functional transcripts. The known mechanisms of chloroplast mRNA processing are discussed in Section II, while chloroplast tRNA processing and rRNA processing have been described elsewhere (Gruissem et al., 1983; Keus et al., 1984; MarionPoll et al., 1988; Wang et al., 1988; Barkan, 1993). A second and related consequence is that the chloroplast must be able to distinguish, and degrade, the many untranslatable RNA segments that would accumulate initially as part of primary transcripts. Evidence for rapid degradation of non-coding RNA has been obtained by comparing transcript accumulation levels and run-on transcription rates in both Chlamy­ domonas (Stern and Kindle, 1993) and spinach (Deng et al., 1987) chloroplasts, and in maize mitochondria (Finnegan and Brown, 1990). The way in which these transcripts are recognized as non-coding is not known; however, it may be that the default pathway is degradation, and mature mRNAs are protected from degradation by specific RNA-binding proteins and/or secondary structures. A third consequence of transcriptional readthrough is the potential synthesis of antisense RNA, which could be detrimental to gene expression, especially in a genome with tightly clustered genes on both strands. Antisense RNA synthesis has been observed in run-on transcription experiments (Deng et al., 1987), but this RNA may be rapidly degraded as discussed above. In addition, because the chloroplast genome is polyploid, symmetric transcription does not necessarily pose topological problems. It may be possible to address the transcriptional activities of different genomes

Chapter 10 Chloroplast Transcription within a single plastid by using in situ methods, since cpDNA is organized as discrete bodies, called nucleoids(Fujie et al., 1994; Nakamura et al., 1994). Finally, in the absence of efficient termination, but with the existence of efficient RNA processing, the majority of chloroplast promoters are in effect obsolete. In the case of the Chlamydomonas petD gene, deletion of its promoter had no consequences for gene expression, since the upstream petA gene provided sufficient transcriptional readthrough (Sturm et al., 1994). Thus, the Chlamydomonas chloroplast may be evolving to contain fewer promoters, in keeping with a general trend of post­ transcriptional regulatory mechanisms.

II. Processing of Chloroplast mRNAs

A. General Considerations Messenger RNA processing in chloroplasts encom­ passes intron removal and steps that form the 5´ and 3´ transcript termini. Other forms of mRNA processing include polyadenylation, which in chloroplasts is related to RNA degradation (Kudla et al., 1996; Lisitsky et al., 1996), and RNA splicing and decay (Chapter 9, Nickelsen; Chapter 11, Herrin et al.). As discussed above, transcription termination is inefficient or absent in Chlamydomonas chloroplasts, requiring that 3´ termini be formed by processing. At the same time, researchers have been unable to detect primary transcripts in Chlamydomonas chloroplasts using vaccinia virus guanylyl transferase to ligate GTP to di- or triphosphate 5´ termini (Johnson and Schmidt, 1993; N. Sturm and D. B. Stem, unpub­ lished). In contrast, in vitro capping of land plant chloroplast transcripts is readily accomplished (Woodbury et al., 1989), and at least some Euglena gracilis chloroplast transcripts can be capped (Drager, 1993). This raises the possibility that most or all 5´ termini of Chlamydomonas chloroplast transcripts are also generated by RNA processing, rather than directly by transcription initiation. These considerations underscore the importance of RNA processing in the generation of translatable RNAs in Chlamydomonas chloroplasts. It is possible that the mechanisms of RNA processing are linked to those governing translation and RNA stability, which are subjects ofChapters 12 (Hauser et al.) and 9 (Nickelsen), respectively. These connections will

171 be revealed by continued biochemical and molecular genetic analysis of mRNA metabolism.

B. 5´-End Processing 1. The Extent of 5´-End Processing The determination that a mRNA 5´ terminus results from processing rather than initiation can be inferred from its inability to be capped in vitro, or from in vitro processing experiments using synthetic RNAs, In land plants, it is generally the case that the longest or primary transcript of a gene cluster can be capped, while a larger number of processed transcripts resulting from intercistronic cleavages accumulate to similar levels (e.g. Westhoff and Herrmann, 1988). In simpler cases, such as rbcL transcription, only a monocistronic mRNA accumulates, but two to three closely-spaced 5´ termini are found; one primary and the others processed (e.g. Erion, 1985; HanleyBowdoin et al., 1985; Mullet et al., 1985). In at least one case, this processing may have a function in regulating accumulation of the protein product (Reinbothe et al., 1993). In other cases, such as the spinach atpB transcripts, most multiple 5´ termini result from initiation and not processing (Chen et al., 1990). The situation in Chlamydomonas chloroplasts with respect to 5´-end formation is somewhat different, largely because many gene-specific probes hybridize with only a single mRNA, rather than the as many as 10–15 identified by the same probes in land plants. It has therefore been assumed that most Chlamy­ domonas genes include a promoter just upstream of the coding region which directs transcription of a monocistronic mRNA. However, several instances have now been documented where genes can be cotranscribed. Examples of co-transcribed Chlamy­ domonas reinhardtii chloroplast genes include psbD with psaA exon2 (Choquet et al., 1988), tscA-chlN (reviewed in Rochaix, 1996), psbB-psbT (Monod et al., 1992; Johnson and Schmidt, 1993), atpA-psbIcemA-atpH (Levy et al., 1997), atpE-rps7 (Robertson et al., 1990) and ycf9-psbM (D.C. Higgs, K.L. Kindle and D.B. Stern, unpublished observation). In addition, a single transcript can have multiple 5´ termini, as exemplified by petA (Matsumoto et al., 1991). The supposition that the 5´-ends of Chlamy­ domonas chloroplast transcripts are generated directly by transcript initiation is also supported by the identification of promoters immediately upstream of

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the mature 5´-ends of atpB and petD transcripts (see section I.C and Sakamoto et al., 1994). However, efforts to cap petD or other mRNAs in vitro have been unsuccessful, suggesting that Chlamydomonas chloroplast mRNAs may be processed at their 5´ends (although it cannot be ruled out that the 5´ di- or triphosphate is converted to a monophosphate without removal of nucleotides). In the only confirmed example of 5´ processing of a co-transcript emanating from an upstream promoter, it was found that the petD promoter could be deleted without substantially reducing the amount of monocistronic petD mRNA accumulating in vivo (Sturm et al., 1994). Whether the case for petD is typical for Chlamydomonas chloroplast mRNAs can only be determined by precisely deleting promoters for other mRNAs, and asking whether monocistronic mRNAs can still be produced by transcriptional readthrough ofupstream genes (the upstream gene is petA in the case of petD) followed by endonucleolytic processing. This type of experiment can also be performed in an ectopic context: for example, a promoterless petD 5´ UTR­ uidA fusion placed in the petA-petD intergenic region produced monocistronic uidA mRNA (Sakamoto et al., 1994).

2. Possible Mechanisms for 5´-End Processing Processed 5´-ends can in principle be generated by three post-transcriptional mechanisms which are illustrated in Fig. 1. In the first, an endonucleolytic cleavage generates directly the mature 5´ terminus; exonucleolytic processing in the second, trims the precursor to give the mature 5´-end; and in the third mechanism, endonucleolytic cleavage is exonucleolytic processing. In followed by the absence of in vitro RNA processing systems, it is difficult to distinguish between these possibilities. However, one approach is to use mutagenesis to define processing sites in vivo, and then determine whether a new upstream and presumably unprocessed 5´-end is produced, which could potentially be capped in vitro. In the case of petD, as described above, the promoter could be deleted without affecting 5´-end formation substantially (Sturm et al., 1994). In this sequences from –2 to –81 with deletion, termed respect to the mature 5´-end were removed. This suggests that the processing signal lies entirely within the 5´ UTR, a hypothesis that was confirmed by placing this promoterless 5´ UTR upstream of the uidA gene and showing that monocistronic uidA

mRNA could accumulate (Sakamoto et al., 1994). In genes, both the case of and the chimeric however, some unprocessed mRNA accumulated as co-transcripts with the upstream petA mRNA. This indicates that sequences upstream of–2 can contribute to the efficiency of mRNA processing.

3. The Relationship Between 5´-End Processing and mRNA Accumulation Post-transcriptional processes often affect RNA halflife and accumulation. For example, in maize nuclear mutants deficient in chloroplast RNA splicing, there is no increase in the accumulation of unspliced precursors, suggesting that the inability to remove introns may increase transcript susceptibility to nucleases (Jenkins et al., 1997). This does not appear

Chapter 10

Chloroplast Transcription

to be the case for Chlamydomonas nuclear mutants defective in psaA splicing, because unspliced precursors accumulate to increased levels (Choquet et al., 1988; Goldschmidt-Clermont et al., 1990). In other cases, nuclear or chloroplast mutations resulting in a translational defect have been shown to lead to increased degradation of the affected chloroplast transcripts, although in some cases there is no effect on transcript accumulation (reviewed in Rochaix, 1996). There are at least three land plant nuclear mutants that affect chloroplast mRNA processing. One is the crp1 mutant of maize, which is defective in the accumulation of monocistronic petB and petD mRNAs (Barkan et al., 1994), the 5´ termini of which are formed by intercistronic processing (Rock et al., 1987). The accumulation of 17 other transcripts from this complex gene cluster is unaffected, including the putative precursors to monocistronic petB and petD. This could be explained by a block in processing of the precursors coupled to some type of feedback control of precursor accumulation at the transcriptional level such that the precursor level remains constant, or by destabilization of the monocistronic products. Other possible processing mutants are Arabidopsis thaliana hcf5 and hcf109. The h c f 5 mutant accumulates a very low level of rbcL mRNA, and also has altered transcript patterns for the psbB gene cluster (Dinkins et al., 1997), whereas hcf109 fails to accumulate a subset of transcripts from the psbB, ndhC and ndhH gene clusters (Meurer et al., 1996). Again, in these cases it is difficult to say whether RNA processing or RNA stability is the primary defect, but the results nevertheless raise the possibility of a link between the two. In Chlamydomonas there are no confirmed nuclear mutants with defects in mRNA 5´ processing, although as for land plant mutants, nuclear mutants affecting mRNA half-lives could have primary defects in RNA processing. These RNA stability mutants are discussed in detail in Chapter 9 (Nickelsen). Two of them, nac2-26 (Kuchka et al., 1989) and F16 (Drager et al., 1998) were shown to destabilize psbD or petD mRNAs, respectively, via the 5´ UTR (Nickelsen et al., 1994). Thus, a defect in 5´ processing could be the primary lesion leading to increased RNA degradation, if processing were required to permit the binding of an RNA stabilizing protein, such as the 47 kD protein that binds to the psbD 5´ UTR in vitro (Nickelsen et al., 1994).

173 Other hints at the relationship between 5´ processing and RNA stability in Chlamydomonas chloroplasts come from deletions that remove processing signals. For example, in the chloroplast mutant FUD6 (Lemaire et al., 1986), the cytochrome complex fails to accumulate due to a 236-bp deletion that encompasses part of the petD 5´ UTR and also the upstream promoter region (Sturm et al., 1994). As a result, a petA-petD co-transcript accumulates instead of monocistronic petD mRNA. However, the level of this co-transcript is quite low, suggesting that it is unstable. This instability could be due to a lack of processing, its inability to be translated, and/or an altered RNA structure due to the deletion, that creates RNAse-hypersensitive sites. strain The petA-petD co-transcript of the related (section II.B.2.) also accumulates to a low level, and as noted above, there are several possible ways to account for the low abundance of this transcript. Overall, the discussion above emphasizes the interdependence ofpost-transcriptional processes in chloroplasts, and how a defect in one process can have pleiotropic effects. 5´ RNA processing is likely to interact with other mechanisms that stabilize transcripts and facilitate the initiation of protein synthesis. The isolation of appropriate mutants and the development of in vitro systems will help to define the interrelationships between these essential steps of chloroplast gene expression.

C. 3´ End Processing 1. Examples of 3´-End Processed Transcripts Like 5´ termini, mRNA 3´-ends can be formed by a direct mechanism, transcription termination, or indirectly through RNA processing. While some chloroplast RNA polymerases can recognize bacterial terminators (Stern and Gruissem, 1987; Chen and Orozco, 1988), there is little evidence for efficient termination of transcription at chloroplast 3´ UTR stem-loops (Stern and Gruissem, 1987; Stern and Kindle, 1993; Rott et al., 1996). However, these stem-loop structures define the ends of, and are required for accumulation of discrete transcripts (Stern et al., 1991; Lee et al., 1996). Thus, most or all chloroplast mRNA 3´-ends are probably formed by RNA processing. The strongest evidence for the existence of such mechanisms is the patterns of transcription from land plant, Euglena gracilis and Chlamydomonas complex operons or gene clusters,

174 where a primary transcript is processed to yield multiple smaller mRNAs (e.g. Rock et al., 1987; Westhoff and Herrmann, 1988; Stevenson and Hallick, 1994; Stollar and Hollingsworth, 1994; Hong et al., 1995;Levy et al., 1997). However, the existence of these transcripts does not prove a precursor-product relationship, i.e. that intercistronic termination does not occur in spite of the in vivo and in vitro results on transcription termination cited above. That the enzymatic machinery for processing exists in chloroplasts is proven adequately by a variety of in vitro results (Stern and Gruissem, 1987; Nickelsen and Link, 1993; Stern and Kindle, 1993; Hayes et al., 1996). Thus, the lack of termination and the presence of appropriate enzymatic activities processing is nearly support the notion that universal for chloroplast transcripts both in land plants and in Chlamydomonas. One confounding observation, however, has been that certain sequences appear to act as terminators in chimeric gene constructs introduced into Chlamydomonas (Blowers et al., 1993; Stern and Kindle, 1993; Sakamoto et al., 1994), whereas analogous constructs in transformed tobacco chloroplasts clearly allow readthrough of UTR, yielding polycistronic the chimeric gene transcripts (Staub and Maliga, 1994). RNA processing followed by degradation of downstream sequences, however, would give the same amount ofaccumulated RNA as transcription termination. This was inferred to be the case for the Chlamydomonas atpB mRNA (Stern and Kindle, 1993), and may explain the similar results cited above for the Chlamydomonas rbcL UTR (Blowers et al., 1993).

2. Mechanisms of 3´-End Processing As illustrated in Fig. 2, three mechanisms are possible for maturation, which in most cases results in immediately downstream of a stema mature loop or other stable secondary structure. In the first, is formed directly by a site-specific the mature endonucleolytic cleavage. In the second, the endonucleolytic cleavage occurs downstream of the and exonucleolytic resection is mature required. In the third mechanism, transcription termination occurs downstream of the mature either at a specific site or in a stochastic manner, exonucleolytic degradation continues and until the nuclease encounters a stable secondary structure, which thereby defines the transcript terminus.

David B. Stern and Robert G. Drager

In vitro evidence for each ofthe above mechanisms has been published for chloroplast transcripts, although for Chlamydomonas, only the maturation of the atpB has been studied; this maturation occurs by the two-step endonuclease-exonuclease mechanism (Stern and Kindle, 1993). Direct was reported endonucleolytic formation ofthe for spinach petD mRNA (Hayes et al., 1996); however, could be formed it was also reported that this by exonuclease alone (Stern and Gruissem, 1987, 1989). This inconsistency may merely reflect the fact that different protein fractions were used in the assays. Endonuclease activity was also implicated in the formation of the mustard trnK and rps16 mRNA (Nickelsen and Link, 1993). In the spinach petD study (Hayes et al., 1996), a partial cDNA sequence encoding a protein resembling E. coli polynucleotide phosphorylase was reported, as well as a protein which cross-reacted with an antibody raised against E. coli RNAse E. Poly­ exonuclease nucleotide phosphorylase is a which can process the 3´-end of petD mRNA correctly in vitro (Stern and Gruissem, 1989); RNAse E is an

Chapter 10 Chloroplast Transcription endonuclease involved in certain types of RNA maturation in E. coli (e.g. Hajnsdorf et al, 1994). Once the plant genes are cloned, reverse genetics may lead to an understanding of the roles of these enzymes in chloroplast mRNA metabolism. Other chloroplast ribonuclease activities have been reported (Chen and Stern, 1991; Yang and Stern, 1997); however, it is not clear whether the primary roles of these activities is in RNA processing or RNA degradation.

3. Nuclear Mutations Affecting Chloroplast mRNA Formation In the case of nuclear mutants affecting the accumulation of multiple RNAs from complex operons or gene clusters, such as maize crp1 or Arabidopsis hcf5 and hcf109 which were discussed above, one cannot rule out a primary defect in formation. For example, crp1 could be defective in petB formation and/or petD formation, since the data show only that the petD monocistronic transcript does not accumulate. In Chlamydomonas, two nuclear mutants clearly have defects in chloroplast processing and/or mRNA stability mediated by UTR elements. In the mutant ncc1, ATP synthase subunit accumulates to 50% of the wild-type level, and monocistronic atpA mRNA accumulates to 10% of the wild-type level (Drapier et al., 1992). However, other mRNAs emanating from the atpA promoter, for example the dicistronic atpA-psbI transcript, are unaffected. This suggests that ncc1 is defective in formation between atpA and psbI. The phenotype of ncc1 is wild-type, since the remaining ATP synthase is sufficient to support photoautotrophic growth. which carries a recessive The nuclear mutant mutation in the Crp3 (chloroplast RNA processing) formation within the gene, is also defective in atpA atpA gene cluster (Levy et al., 1997). In monocistronic mRNA accumulates to wild-type levels, whereas the dicistronic atpA-psbI transcript accumulates to 50% of the wild-type level, and the tricistronic atpA-psbI-ycf10 transcript fails to accumulate (the tetracistronic atpA-psbI-ycf10-atpH transcript is unaffected). These results indicate that the products of at least two nuclear genes participate maturation or stabilization of transcripts of in has at least the atpA gene cluster. Interestingly, two other phenotypes. First, it was identified by robust phototrophic growth on minimal medium

175 resulting from high levels of accumulation of a nuclease-susceptible atpB transcript that lacks the stem-loop structure (see section 4, below). Secondly, processing presumptive intermediates in petD accumulate in but not in its wild-type progenitor. Although these intermediates accumulate to only a low level, these results suggest that the Crp3 gene product functions in the maturation of multiple chloroplast mRNAs. Since screens for non-photosynthetic mutants of Chlamydomonas have yielded several that act on the level of RNA stability, one might naturally expect that a certain percentage would have defects in stabilization. However, as first shown for the psbD stability mutant nac2-26 (Nickelsen et al., 1994) and subsequently for the atpB stability mutant thm24 (Drapier et al., 1992), the petA stability mutant MØ11 (Gumpel et al., 1995; D. B. Stern and R. G. Drager, unpublished), and the petD stability mutant F16 (Drager et al., 1997), these nuclear gene products stabilize their cognate mRNAs by acting on the UTR. This raises the question of why more stabilitydeficient mutants have not been isolated. The two main possibilities are that such mutants would not have a detectable phenotype with the screen used, or that such mutants would be lethal. The fact that both ncc1 and crp3 (which we have shown to be unlinked loci by genetic crosses) are able to grow photo­ autotrophically under normal conditions supports the first possibility. In this case, it will be necessary to design screens for mutants defective in processing processing, for example by inserting elements ectopically into essential genes or selectable markers. If stability mutants are lethal, then temperature-sensitive mutant screens might get around this problem. In E. coli, however, mutations that inactivate enzymes such as RNAse II, RNAse III or polynucleotide phosphorylase are not lethal; only multiple mutations cause lethality (Donovan and Kushner, 1986; Babitzke et al., 1993). This again supports the notion that there is redundancy in the formation machinery, or that RNA secondary structure alone allows sufficient accumulation of transcripts to support photoautotrophic growth under the conditions most often used for mutant screening— that is, total inability to grow without added acetate (see Stern et al., 1991; Drager et al., 1996).

4. Does the

UTR Regulate mRNA Stability?

In both land plants and Chlamydomonas, chloroplast

176 mRNAs exhibit a wide range of half-lives, which can vary between developmental stages, within the cell cycle, or in a circadian manner (Deng and Gruissem, 1987; Mullet and Klein, 1987; Leu et al., 1990; Kim et al., 1993; Salvador et al., 1993a,b; Hwang et al., 1996). To what degree is the 3´ UTR a determinant of chloroplast mRNA half-life? As noted above, most chloroplast mRNAs have inverted repeats in their 3´ UTRs; these repeats pair to form stem-loop structures, at least in vitro (Stern et al., 1989), and such structures impede the progress of exoribonucleases (McLaren et al., 1991; Drager et al., 1996). Thus, RNAs lacking these structures are unstable in vitro in chloroplast protein extracts (Stem and Gruissem, 1987; Stern and Kindle, 1993). In Chlamydomonas cells altered by chloroplast transformation, an atpB gene lacking the IR yielded unstable and heterogeneous mRNA (Stern et al., 1991), while either removal of the psaB stem-loop or extension of the psaB coding region into the stemloop also destabilized the transcript (Lee et al., 1996). In contrast, measurement of RNA half-life by slot blots did not reveal an effect of stem-loop removal (Blowers et al., 1993). However, these experiments did not distinguish between intact and partially degraded or heterodisperse transcripts. Overall, it appears that the IR has an important role in defining the 3´-end of the transcript, and that in its absence, transcripts are less stable and more heterogeneous. On the other hand, mRNAs lacking 3´ IRs appear to be translatable, and no absolute requirement for a 3´ stem-loop to support photoautotrophic growth has been demonstrated. Although the atpB 3´ IR deletion referenced above did not lead to obligate heterotrophic growth, the cells accumulated only 20% of the wild-type level of the ATP synthase, and grew slowly on minimal medium, and were sensitive to high light or elevated (35°C) temperature. These phenotypes allowed selection or screening for phenotypic suppressors, of which two types were isolated. In the first type, a gene amplification event had occurred, resulting in a 20-fold increase in the number of atpB genes still carrying the IR deletion (Kindle et al., 1994). These amplification events were associated with complex rearrangements of the chloroplast genome, and were unstable when the suppressed cells were supertransformed with an atpB gene containing the IR, in which case the copy number of atpB returned to one per genome (Suzuki et al., 1997). One possible explanation for this would be that very high levels of

David B. Stern and Robert G. Drager atpB mRNA titrate a factor required for the expression ofother genes. The other type of atpB suppressor was a nuclear mutation that was impeded in the degradation of the normally unstable transcript (Levy et al., 1997). As discussed in section 3, this suppressor mutation was in the gene Crp3, which appears to mRNA degrada­ define a factor required for tion. This demonstrates that nuclear factors can, at least in certain cases, determine chloroplast RNA half-life by interacting with the 3´ UTR. The above discussion still leaves open the question of what regulates chloroplast mRNA half-lives that vary either intrinsically or according to cellular signals. In the case of rbcL, there is evidence for light control of mRNA stability via the 5´ UTR (Salvador et al., 1993a), and similar conclusions have been reached for psbA mRNA in tobacco (Staub and Maliga, 1994). Since in the case of psbA the same sequences also control light-regulated translation (Staub and Maliga, 1994), this raises the more general question of whether light-, developmentally-, or cell cycle-regulated changes in RNA stability are secondary consequences of translational control. The existence of a variety of Chlamydomonas nuclear mutants defective in chloroplast translation (Chapter 12, Hauser et al.) offers an opportunity to test this possibility. The intrinsic differences in chloroplast RNA half-lives, which vary from about 30 min to several days and include both protein-coding and structural RNAs, cannot be explained solely by translational differences. In addition, there is no clear correlation between transcript stability and translatability in Chlamydomonas (Chapter 9, Nickelsen). Rather, multiple determinants within the untranslated regions and coding regions may confer differential susceptibility to nucleases. Recently, a new pathway of land plant chloroplast mRNA degradation, which can involve the 3´ UTR, has been described. Two reports have shown that sequences within the 3´ UTR, as well as sequences within the coding region, can serve as sites for polyadenylate addition (Kudla et al., 1996; Lisitsky et al., 1996). Know sites of polyadenylate addition and aproposed degration mechanism (Lisitsky et al., 1997) are illustrated in Fig. 3. The resulting polyadenylated transcripts are particularly susceptible to exonuclease digestion. Similarpolyadenylation of transcripts also occurs in E. coli, where it also leads to transcript degradation (Cohen, 1995; HaugelNielsen et al., 1996). It is not yet clear whether polyadenylation is the rate-limiting step in land plant

Chapter 10 Chloroplast Transcription

chloroplast mRNA decay, nor is it known whether polyadenylation also occurs in Chlamydomonas chloroplasts. This should therefore prove a fertile area for the study of chloroplast mRNA 3´ UTR function over the next several years.

Acknowledgments Work on chloroplast RNA processing and stability in the Stern laboratory was supported by grants from the Department of Energy and the National Science Foundation. R. G. D. was supported by a fellowship from the National Institutes of Health.

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David B. Stern and Robert G. Drager Stern DB and Gruissem W (1989) Chloroplast mRNA 3´ end maturation is biochemically distinct from prokaryotic mRNA processing. Plant Mol Biol 13: 615–625 Stern DB and Kindle KL (1993) 3´ end maturation of the Chlamydomonas reinhardtii chloroplast atpB mRNA is a twostep process. Mol Cell Biol 13: 2277–2285 Stern DB, Jones H and Gruissem W (1989) Function of plastid mRNA 3´ inverted repeats: RNA stabilization and gene-specific protein binding. J Biol Chem 264: 18742–18750 Stern DB, Radwanski ER and Kindle KL (1991) A 3´ stem/loop structure of the Chlamydomonas chloroplast atpB gene regulates mRNA accumulation in vivo. Plant Cell 3: 285–297 Stern DB, Higgs DC and Yang J (1997) Transcriptional and translational activities of chloroplasts. Trends Plant Sci 2: 308–316 Stevenson JK and Hallick RB (1994) The psaA operon prc­ mRNA of the Euglena gracilis chloroplast is processed into photosystem I and II mRNAs that accumulate differentially depending on the conditions of cell growth. Plant J 5:247–260 Stollar NE and Hollingsworth MJ (1994) Expression of the large ATP synthase gene cluster from spinach chloroplasts. J Plant Physiol 144: 141–149 Sturm N, Kuras R, Büschlen S, Sakamoto W, Kindle KL, Stern DB and Wollman FA (1994)The petD gene is transcribed by f u n c t i o n a l l y redundant promoters in Chlamydomonas reinhardtii chloroplasts. Mol Cell Biol 14: 6171–6179 Surzycki SJ and Shellenbarger DL (1976) Purification and characterization of a putative sigma factor from Chlamy­ domonas reinhardii. Proc Natl Acad Sci USA 73: 3961–3965 Suzuki H, Ingersoll J, Stern DB and Kindle KL (1997) Generation and maintenance of tandemly repeated extrachromosomal plasmid DNA in Chlamydomonas chloroplasts. Plant J 11: 635–648 Tanaka K, Oikawa K, Ohta N, Kuroiwa H, Kuroiwa T and Takahashi H (1996) Nuclear encoding of a chloroplast RNA polymerase sigma subunit in a red alga. Science 272: 1932– 1935 Thompson RJ and Mosig G (1985) An ATP-dependent supercoiling topoisomerase of Chlamydomonas reinhardtii affects accumulation of specific chloroplast transcripts. Nucleic Acids Res 13: 873–891 Thompson RJ and Mosig G (1987) Stimulation of a Chlamy­ domonas chloroplast promoter by novobiocin in situ and in E. coli implies regulation by torsional stress in the chloroplast DNA. Cell 48: 281–287 Thompson RJ and Mosig G (1990) Light affects the structure of Chlamydomonas chloroplast chromosomes. Nucleic Acids Res 18:2625–2631 Tiller K and Link G (1993) Sigma-likc transcription factors from mustard Sinapis alba L. etioplasts arc similar in size to, but functionally distinct from, their chloroplast counterparts. Plant Mol Biol 21: 503–513 Tracy RL and Stern DB (1995) Mitochondrial transcription initiation: promoter structures and RNA polymerases. Curr Genet 28: 205–216 Troxler RF, Zhang F, Hu J and Bogorad L (1994) Evidence that sigma factors are components of chloroplast RNA polymerase. Plant Physiol 104: 753–759 Vera A and Sugiura M (1995) Chloroplast rRNA transcription from structurally different tandem promoters: An additional novel-type promoter. Curr Genet 27: 280–284

Chapter 10 Chloroplast Transcription Wang MJ, Davis NW and Gegenheimer P (1988) Novel mechanisms for maturation of chloroplast transfer RNA precursors. EMBO J 7: 1567–1574 Westhoff P and Herrmann RG (1988) Complex RNA maturation in chloroplasts: the psbB operon from spinach. Eur J Biochem 171:551–564 Wolfe KH, Morden CW and Palmer JD (1992) Function and evolution of a minimal plastid genome from a nonphotosynthetic parasitic plant. Proc Natl Acad Sci USA 89: 10648–10652

181 Woodbury NW, Dobres M and Thompson WF (1989) The identification and localization of 33 pea chloroplast transcription initiation sites. Curr Genet 16: 433–446 Yang J and Stern DB (1997) The spinach chloroplast endoribonuclease CSP41 cleaves the 3´ untranslated region of petD mRNA primarily within its terminal stem-loop structure. J Biol Chem 272: 12874–12880 Zaitlin D, Hu J and Bogorad L (1989) Binding and transcription of relaxed DNA templates by fractions of maize chloroplast extracts. Proc Natl Acad Sci USA 86: 876–880

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Chapter 11 RNA Splicing in the Chloroplast David L. Herrin, Tai-Chih Kuo Department of Botany and Institute of Cellular and Molecular Biology, University of Texas at Austin, Austin, TX 78713, U.S.A.

Michel Goldschmidt-Clermont Department of Molecular Biology and Department of Plant Biology, University of Geneva, Sciences II, 30 quai E. Ansermet, CH-1211 Genève 4, Switzerland

Summary I. Introduction II. Group I Introns A. Group I Introns: Splicing Mechanism and Structure B. Group I Introns in Chlamydomonas spp. Chloroplast Genes 1. Distribution and Classification 2. Identification of Self-Splicing Group I Introns 3. Biochemical Characterization of Two Self-Splicing Group I Introns C. Protein Factors Involved in Group I Splicing D. Regulation of Group I Splicing III. Group II Introns and Trans-Splicing A. Group II Introns: Splicing Mechanism and Structure B. Trans-Splicing of the psaA Introns 1. Structure of the psaA Gene 2. Trans-Splicing Mutants 3. The tscA RNA 4. Evolution of Trans-splicing C. Splicing of Heterologous Group II Introns IV. Perspective Acknowledgments References

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Summary The chloroplast genomes of Chlamydomonas spp. contain introns that belong to the two major classes found in organelles: group I and group II. Some of the members of both classes are ribozymes capable of self-splicing in vitro and, indeed, most of the group I introns studied in Chlamydomonas spp. are autocatalyic. The biochemical characterization of these introns suggests that, in vivo, their splicing must be dependent on protein factors. It is particularly interesting in this respect that light regulates the splicing of the four group I introns in the psbA gene of C. reinhardtii. The two group II introns of the psaA gene are split into two or more pieces which are transcribed separately from different chloroplast loci. The mature psaA RNA is assembled from the separate precursors in two steps of intermolecular trans-splicing. There are many nuclear mutants defective in psaA mRNA maturation, which identify a large number of protein factors that are required, directly or indirectly, for trans-splicing. J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 183–195. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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I. Introduction Some chloroplast genes contain intervening sequences, or introns as they are more often called, that interrupt the coding sequences (i.e. exons). Two main types of introns have been found in Chlamy­ domonas spp. chloroplast genes, group I and group II. These designations were originally given to the two structurally different families of introns that were found mainly, but not exclusively, in organelles, and was based primarily on conserved primary and secondary structure elements (Michel and Dujon, 1983). Subsequently, it was shown that these two intron groups also differ in their mechanism of splicing (see Cech, 1990; Michel and Ferat, 1995, for reviews). However, some members of both groups are self-splicing introns, i.e. introns that splice in vitro in the absence of proteins. Those remarkable findings indicated that the pre-RNA itself catalyzes the splicing reactions. Consequently, these molecules are often referred to as catalytic RNAs, or ‘ribozymes’. In vivo, however, proteins probably facilitate splicing of most group I and group II introns (Saldanha et al., 1993), and recent evidence indicates that, at least in the case of group I introns, some of the proteins act by promoting the correct folding of these large catalytic RNAs (Mohr et al., 1992; Shaw and Lewin, 1995; Weeks and Cech, 1995). Group I introns are more common than group II in Chlamydomonas spp. chloroplasts, a situation that differs from land plant chloroplasts, where group II introns are more common (Plant and Gray, 1988). The only group II introns described so far in Chlamydomonas spp. chloroplasts are the two introns in the psaA gene (Kück et al., 1987; Turmel et al., 1995a). Interestingly, both of these introns are spliced in trans: the psaA exons, with flanking intronic sequences, are found at widely separated sites in the chloroplast genome (Kück et al., 1987), and are apparently transcribed independently (Choquet et al., 1988; Herrin and Schmidt, 1988). In addition, an internal segment ofintron 1 is encoded at yet another site in the genome, the tscA locus, which is transcribed to produce a small RNA (Goldschmidt-Clermont et al., 1991). Thus, at least three RNAs must be brought together for the splicing of psaA intron 1. This mode of splicing shows a strong dependence on protein Abbreviations: LSU – large subunit ribosomal RNA; ORF – open reading frame

factors, since self-splicing has not been reported for any trans-spliced intron. It is also interesting that trans-splicing introns have only been found so far among group II, but not group I introns. Research with C. reinhardtii has played an important role in our current understanding of RNA splicing in green plant chloroplasts. The first selfsplicing RNAs from chloroplasts were identified using C. reinhardtii (Herrin et al., 1990, 1991), as were the first mutants blocked in splicing of chloroplast introns (Choquet et al., 1988; Herrin and Schmidt, 1988). In fact, mutants affected in chloroplast biogenesis played key roles in both those studies. In addition, chloroplast transformation in C. reinhardtii has greatly facilitated the study ofcis­ and trans-acting expression elements encoded in the chloroplast genome (Rochaix, 1996). Finally, Chlamydomonas is proving to be an excellent genus for studying the evolution of introns, due in part to the large number of related species, and the fact that most, if not all, contain these classes of introns (e.g. Turmel et al., 1993a, 1995a). The purpose of this chapter is to summarize our current state ofknowledge concerning RNA splicing in Chlamydomonas spp. chloroplasts.

II. Group I Introns

A. Group I Introns: Splicing Mechanism and Structure Figure 1 shows the splicing mechanism for group I introns (reviewed in Cech, 1990). The first step involves a free guanosine nucleotide (probably GTP), which makes a nucleophilic attack at the 5´ splicesite, breaking the pre-RNA chain, and becoming covalently attached to the 5´ end of the intron-exon molecule. In the second step, the 3´ OH of the 5´ exon attacks the 3´ splice-site, effectively displacing the intron and ligating the exons. The liberated intron often undergoes cyclization reactions via the conserved 3´ terminal G, which attacks one or more phosphodiester bonds near the 5´ end of the intron. Intron cyclization is commonly seen in vitro, but has also been seen in vivo (Daros and Flores, 1996). Because circular RNAs do not migrate strictly according to their size in electrophoresis gels, the principal fate ofthe free intron in vivo is not clear for most systems. We will briefly review the structure of group I

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185 5´ splice-site, which is also part of the P1 helix, is usually the sequence GU (U being the last nucleotide of the exon), with the U being involved in a wobble base-pairing interaction with a G in the intron. The 3´ terminal nucleotide of the intron is nearly always a G, and it binds to the guanosine-binding site in P7 during the second splicing step, exon ligation. A short sequence near the 5´ end of the intron, known as the internal guide sequence (IGS), base-pairs with the end of the 5´ exon, and during the second step, the beginning of the 3´ exon. Thus, the IGS plays a role in both steps by helping to align the splice-sites relative to the catalytic core. These conserved features make up only a fraction of the sequence for many group I introns, which range in size from ~250 to ~2000 bp. Most of the larger group I introns contain Open Reading Frames (ORFs), many of which have been shown to encode site-specific DNA endonucleases that promote intron mobility (reviewed in Dujon, 1989). Other less conserved domains in group I introns have also been found to play important roles in the ability of the respective introns to self-splice, apparently by contributing to the stable formation of the catalytic core (Jaeger et al., 1996). The presence or absence of these so-called peripheral domains have been used to classify Group I introns into four major classes (A,B,C, and D), and eleven subclasses in total (Michel andWesthof, 1990).

B. Group I Introns in Chlamydomonas spp. Chloroplast Genes introns; for a more detailed and recent review the reader is referred to Jaeger et al. (1996). Group I introns are identified according to their secondary structure, which includes a conserved set of basepaired regions (called P1, P2, etc.), but only a few highly conserved nucleotides. Figure 2 shows the secondary structure of the single intron in the 23S rRNA gene of C. reinhardtii (Rochaix et al., 1985), depicted in the ‘original’ style (Fig. 2A), and in the newer style (Fig. 2B); the latter is more representative of the intron’s three-dimensional structure in that the P4-P6, and the P8, P3 and P7 helices are stacked colinearly (Cech et al., 1994). The latter form also facilitates depiction of some of the tertiary interactions. Together, the paired regions P3-P8 constitute the catalytic core, and P7 contains the guanosine binding site. The ends of the intron, together with a few nucleotides into each adjacent exon, are important for splice-site recognition. The

1. Distribution and Classification Table 1 provides a list of published group I introns from Chlamydomonas spp. chloroplast genes; most of them fall into the A or B classes, but with only one B subclass, IB4, represented so far. A number of other Group I introns have been found in the rrnL gene of different Chlamydomonas species, although the sequences of these have not yet been published (Turmel et al., 1993a). The introns reported to date have been found mainly in rRNA genes (both rrnL and rrnS), and in genes coding for proteins involved in photosynthesis. It should be noted, however, that not all of the chloroplast genes in these species, including C. reinhardtii, have been sequenced. Thus, it is possible that additional Group I introns will be found in these and other species of Chlamydomonas. The irregular, but phylogenetically extensive,

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distribution of group I introns suggests that they have migrated during evolution. This migration could have occurred at the DNA level (see Dujon, 1989 for a discussion thereof), or at the RNA level via reverse splicing (Woodson and Cech, 1989), followed by reverse transcription of the RNA into cDNA, and then recombination with genomic DNA. One characteristic shared by the chloroplast genes that contain group I introns is that they encode abundant RNAs (see Table 1). The presence of abundant ‘substrate’ would certainly promote reverse splicing, although the successful completion of all steps is probably a rare event (Thompson and Herrin, 1994). Group I intron migration at the RNA level during evolution is also strongly supported by the preferential location of introns in regions of the rrnL gene that correspond to sites in the mature RNA that are exposed on the surface of the ribosome (Turmel et al., 1993a).

2. Identification of Self-Splicing Group I Introns Because not all group I (or group II) introns selfsplice in vitro, searching for self-splicing introns by subcloning each one (with some flanking exon sequences), synthesizing pre-RNA in vitro, and testing it for self-splicing activity could be a frustrating endeavor. However, an alternative approach is to take advantage of the group I splicing mechanism, which involves the covalent addition of a guanosine nucleotide to the intron (Fig. 1), to rapidly look for self-splicing introns in total cellular RNA. Of course, this approach requires the presence of at least some precursor RNA in the cells. Using RNA from the ac20 mutant of C. reinhardtii (which, as it turns out, contains increased levels ofunspliced RNAs from the rrnL and psbA genes), it was straightforward to identify self-splicing group I in the introns by incubating total RNA with

Chapter 11 Chloroplast RNA Splicing

presence of and then hybridizing the GTPlabeled RNA to total chloroplast DNA, or to cloned DNA fragments (Herrin et al., 1990, 1991). In vitro synthesis of pre-RNAs for each intron was used to confirm that they self-splice in vitro. The single intron in the rrnL gene, and all 4 introns in the psbA gene, proved to be catalytic RNAs (Herrin et al., 1990, 1991). Moreover, these 5 introns constitute all the known group I introns in C. reinhardtii. Using RNA synthesized in vitro, nine other Chlamydomonas chloroplast group I introns were tested for self-splicing activity, and eight of them were positive (Côté and Turmel, 1995; Table 1). Thus, of the group I introns from Chlamydomonas that have been tested, the vast majority have been found to be capable of self-splicing in vitro. It is also interesting to note that all of the self-splicing introns are in the A class, while the non-catalytic intron is from the B class (Table 1).

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3. Biochemical Characterization of Two SelfSplicing Group I Introns Since the RNA plays such an important role in group I splicing, acting both as catalyst, and providing specificity to the splicing reactions, it was of interest to carry out some biochemical analyses of selfsplicing reactions catalyzed by chloroplast group I introns. Table 2 shows the conditions that were necessary for efficient self-splicing of the CrLSU and CrpsbA2 introns; these data were gathered using synthetic RNAs produced by in vitro transcription of linearized plasmid with phage RNA polymerases. Both introns were highly specific for guanosine nucleotides, with only a weak reaction with ATP (at least 100-fold less than with GTP); there was no was strictly reaction detected with CTP or UTP. required for activity with either intron, and the optimal concentration range was 15–25 mM (in the presence

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of monovalent salt). The temperature optimum for self-splicing was quite high (45–47 °C), and splicing at 23 °C was very slow. Spermidine, which can promote RNA folding, stimulated the self-splicing of only the CrpsbA2 intron. Kinetic analyses of selfsplicing by both introns were also performed. The and values were similar to those of the Tetrahymena self-splicing intron (Bass and Cech, 1984), which has been the most studied group I intron (Cech, 1990); they were and for CrLSU, and and for CrpsbA2, respectively (Bao, 1993; T-C. Kuo and D.L. Herrin, unpublished). The rate values for self-splicing by these introns are also comparable to our estimated rates for splicing of these introns in vivo (Fig. 3), although, for the in vivo studies, the cells were grown and analyzed at 23 °C. Thus, from this perspective, these introns appear to splice more readily in vivo than in vitro. It is also apparent that these two introns are functionally related, which may not be surprising, since they are also in the same subclass, 1 A3 (Table 1). Finally, these studies showed that chloroplast group I introns self-splice using the same overall mechanism as the Tetrahymena intron.

C. Protein Factors Involved in Group I Splicing The splicing of most group I introns in vivo is probably facilitated by trans-acting factors that are proteins. This suggestion is supported by the following: (1) some group I introns are not capable of self-splicing in vitro, (2) of those that can, the conditions needed for efficient self-splicing are often non-physiological, and (3) fungal nuclear mutants have been isolated that are deficient in splicing mitochondrial group I introns in vivo, including introns that are capable of self-splicing in vitro. Proteins that promote splicing of mitochondrial group I introns in Saccharomyces cerevisiae and

Neurospora crassa were initially identified using genetic approaches; the proteins are encoded in the nuclear genome and within the introns themselves (Saldanha et al., 1993). The intron-encoded proteins are known only through genetics; they are usually found in-frame with the preceding exon, and they promote splicing of mainly the intron where they reside. However, two ofthe nucleus-encoded splicing factors have been biochemically characterized, these are products ofthe CYT18 and CBP2 genes. Cyt18 is a bifunctional protein that can promote splicing of many group I introns, including the non self-splicing NcLSU intron; its other function is in tyrosyl-tRNA charging (Guo and Lambowitz, 1992). In contrast, Cbp2 specifically promotes splicing of the Sc.cob.5 intron, which requires non-physiological (i.e. high) concentrations to self-splice in vitro (Gampel and Tzagoloff, 1987; Gampel et al., 1989). Both of these proteins seem to act by promoting folding of the pre-RNAs into their catalytically active forms (Mohr et al., 1992; Shaw and Lewin, 1995; Weeks and Cech, 1995). The conditions required for efficient self-splicing by the C. reinhardtii chloroplast group I introns, CrLSU and CrpsbA2 (see Table 2 and above), suggest that proteins probably facilitate splicing of these introns in vivo. However, splicing factors for chloroplast group I introns have not been clearly identified. Most of the intron-encoded ORFs described to date are free-standing rather than inframe like the mitochondrial maturases (although CrpsbA3 contains an ~18 kDa ORF that is in-frame with the upstream exon; N. Deshpande, unpublished). Moreover, several ofthese ORFs have been shown to encode DNA endonucleases that function in intron mobility (see Table 1), although that by itself does not preclude them from also having RNA maturase activity. However, it should be noted that deletion of most of the ORF from the CrLSU intron did not

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189 affect splicing of that intron in vivo (Thompson and Herrin, 1991). Previously, it was suggested that the ac20 mutant of C. reinhardtii, which is deficient in functional chloroplast ribosomes, might be a splicing mutant, since it over-accumulates unspliced CrLSU (i.e. pre­ 23S) RNA (Herrin et al., 1990). However, deletion of the CrLSU intron from ac20 did not restore the wildtype phenotype, which would have been expected if the primary defect in ac20 was a deficiency in CrLSU splicing (S. Holloway and D.L. Herrin, unpublished). Although this does not preclude a role for the ac20 gene product in group I splicing, it does indicate that the apparent splicing deficiency in this mutant is not the primary cause of its phenotype.

D. Regulation of Group I Splicing The fact that most group I introns probably require proteins for efficient splicing in vivo, suggests that splicing could be regulated. However, until recently, evidence for regulated splicing of group I introns was generally lacking. Under light-dark cycles, C. reinhardtii cell growth and division become synchronized, thereby providing a homogeneous population of cells (Harris, 1989). Moreover, under these conditions, there are extensive changes in gene expression (Herrin and Michaels, 1984). By focusing on the introns of the psbA gene of C. reinhardtii, a gene known to be strongly light-regulated (e.g. Herrin et al. 1986), Deshpande et al. (1997) obtained evidence that light promotes splicing of all four of the CrpsbA introns coordinately. These authors observed that unspliced psbA pre-RNAs accumulate in the dark, apparently due to a slow rate of splicing, and then disappear quickly upon illumination. This effect was observed despite a simultaneous, lightstimulated increase in transcription of the psbA gene (Deshpande et al., 1997). By blocking transcription with an inhibitor, the rate of splicing of each intron was estimated as shown in Fig. 3. The results indicate that light stimulates splicing ofpsbA introns six- to ten-fold, and that the introns splice with a half-life of ~12–15 min in the light. By using specific inhibitors, mutants, and chloroplast transformation, Deshpande et al. (1997) also showed that the light-regulated splicing of psbA introns requires photosynthetic electron transport, but not ATP synthesis. Thus, this study provided the first evidence that RNA splicing in chloroplasts can be regulated by light.

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III. Group II Introns and Trans-Splicing

A. Group II Introns: Splicing Mechanism and Structure Some members ofgroup II have the ability to undergo self-splicing in vitro under appropriate conditions of elevated temperature and high concentrations of salt and magnesium, which are clearly not physiological. This suggests that the reaction in vivo is facilitated by protein factors. Figure 4 shows the splicing mechanisms for group II introns (reviewed by Saldanha et al., 1993; Michel and Ferat, 1995; Jacquier, 1996). While the prevalent pathway involves two trans-esterification reactions, under certain conditions in vitro the first step can proceed via hydrolysis. In the first trans-esterification reaction, the 2´ OH of the A nucleotide at the branch site attacks the 5´ exon-intronjunction to produce a lariat intermediate (lariat intron- 3´ exon) and the separated 5´ exon. In the second step, the free 3´ OH of the 5´ exon attacks the 3´ intron-exonjunction, yielding the spliced exons and releasing the intron. Both transesterification reactions are reversible, so that a free intron can insert by reverse-splicing into the appropriate exon-exon junction. The excised intron is a versatile ribozyme and can also catalyze other reactions, such as reverse-splicing into DNA, or polymerization of RNA or DNA in a 3´– 5´direction (Möril et al., 1992; Mueller et al., 1993;Zimmerley et al., 1995 ;Hetzer et al., 1997). Group II introns share conserved secondary structure and tertiary interactions, but relatively few strictly conserved nucleotides in the primary sequence (Michel and Ferat, 1995; Jacquier, 1996; Costa etal., 1997). The secondary structure is usually represented as six partly helical domains surrounding a central wheel (see Section B.3 and Fig. 6). Domain V, which has a very conserved structure, and domain I are the most important for catalysis, while domain VI contains the bulging A at the branch point which is involved in the first trans-esterification reaction. An important tertiary interaction entails base-pairing of a sequence in domain I of the intron (exon binding site 1, EBS 1) with the last bases of the upstream exon (intron binding site 1, IBS1). The nucleotide just 5´ to EBS 1 can interact with the first nucleotide of the downstream exon and thus may act as a guide in the second trans-esterification. Another part of domain I (EBS2) can pair with nucleotides in the upstream exon (IBS2) adjacent to IBS 1. There are many further

tertiary interactions between different parts of the structure. Domain IV sometimes encodes a poly­ peptide which in some cases is required for splicing in vivo (maturase) and in other cases is involved in intron homing and transposition, with a domain related to reverse transcriptases (reviewed by Grivell, 1996; Curcio and Belfort, 1996).

B. Trans-Splicing of the psaA Introns 1. Structure of the psaA Gene In 1987, Kück and coworkers discovered that in C. reinhardtii, the psaA gene is composed of three exons encoded at widely separate loci of the chloroplast genome, and that sequences adjacent to

Chapter 11 Chloroplast RNA Splicing

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these exons share some of the conserved primary structure features of group II introns. This suggested that the psaA exons may be assembled via a process of splicing in trans. The first intron of the rps12 gene in land plants is similarly fragmented, and numerous split introns also occur in land plant mitochondria (reviewed by Bonen, 1993). The psaA gene in land plant plastids and the rps12 gene in C. reinhardtii are not fragmented (Liu et al., 1989). In other Chlamydomonas taxa, the psaA gene is also composed of three widely separated exons (Boudreau et al., 1994; Turmel et al., 1995). Even in the distantly related C. moewusii, the three exons are colinear with those of C. reinhardtii, indicating that the split structure of the psaA intron was present in a common ancestor of these Chlamydomonas taxa (Boudreau et al., 1994; Turmel et al., 1995). To our knowledge, efficient in vitro splicing has not been demonstrated for any of the fragmented group II introns, suggesting that splicing of these introns is strictly dependent on trans-acting factors.

2. Trans-Splicing Mutants The analysis of C. reinhardtii mutants defective in photosystem I activity revealed a large number of mutations in both the chloroplast and the nuclear genome that affect the maturation of the psaA mRNA (Choquet et al., 1988; Herrin and Schmidt, 1988; Roitgrund and Mets, 1990). The mutants accumulate unspliced precursors and partly spliced transcripts which are presumably intermediates in the psaA mRNA maturation pathway. Their analysis indicates that the three exons of psaA are transcribed independently, and that the mature psaA mRNA is normally assembled in two steps of splicing in trans (Fig. 5). Some of the precursors are polycistronic: exon 2 is transcribed together with the psbD gene upstream, and exon 1 is most likely transcribed together with trnI downstream (Choquet et al., 1988; Turmel et al., 1995). The mutants can be assigned to three classes, according to the precursors and intermediates that they contain. In one class ofmutants (class C), trans-splicing of exons 1 and 2 is blocked, but exons 2 and 3 are assembled normally. Conversely in a second class (class A), trans-splicing of exons 1 and 2 can proceed, but not of exons 2 and 3. In the third class (class B), neither trans-splicing reaction occurs. The mutants in each class belong to multiple nuclear loci, as determined by complementation tests: five in class A, two in class B and seven in class C

(Goldschmidt-Clermont et al., 1990). It is likely that there are many more than fourteen nuclear genes required for trans-splicing of psaA mRNA, because most loci in classes A and C are represented by only one allele. It is striking that the products of the numerous loci in classes A and C are specifically required, directly or indirectly, in the splicing ofjust one of the two split introns. The loci in class B may encode factors more generally involved in splicing of group II introns, since the two split introns of psaA are the only group II introns described so far in the C. reinhardtii chloroplast genome. Three of the nuclear genes required for trans-splicing have been cloned by gene-tagging or by complementation (M.

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Goldschmidt-Clermont, K. Perron, C. Rivier and J.D. Rochaix, unpublished; see Chapter 4, Kindle). The analysis of these and other genes should provide insight into the function of factors involved in group II intron splicing and trans-splicing. There are a few mutants of Saccharomyces cerevisiae which are affected in splicing ofmitochondrial group II introns (reviewed by Saldanha et al., 1993). Two nuclear mutants of maize (crs1 and crs2) are deficient in splicing of different subsets ofthe group II introns in the plastid (Jenkins et al., 1997). It will be interesting to determine whether the affected genes bear any relation to those affected in the splicing mutants of C. reinhardtii.

3. The tscA RNA There is also a chloroplast gene (tscA, trans-splicing in the chloroplast) which is required for trans-splicing of exons 1 and 2 (Roitgrund and Mets, 1990; Goldschmidt-Clermont et al., 1990). The product of tscA is a small RNA devoid of coding sequence which is probably involved in forming the conserved structure of group II introns (Goldschmidt-Clermont et al., 1991). In the proposed model (Fig. 6), the tscA RNA forms a helix with the precursor of exon 1 (domain I of the intron), contributes domains II and III, and forms a helix with the precursor of exon 2 (domain IV). The first intron of psaA is thus composed of (at least) three separate transcripts. A comparative sequence analysis of the first split intron in the related species C. gelatinosa and C. zebra supports this model (Turmel et al., 1995a). A puzzling feature is that conserved elements of domain I are absent, and in particular the EBS1 region. Whether the missing structures and tertiary interactions are replaced by protein factors, whether additional RNAs are involved, or both, remains to be determined.

4. Evolution of Trans-splicing From an evolutionary point of view, it is interesting that group II introns share structural and mechanistic features with nuclear introns, but their phylogenetic relation is still a matter of debate (reviewed by Jacquier, 1996). The split structure of the first intron of psaA, with part of the intron on a small separate transcript, can be taken as a simple example of how some of the structural or catalytic domains of nuclear introns may have been transferred to the snRNAs (Goldschmidt-Clermont et al., 1991; Sharp, 1991). Another issue is whether split introns are archaic

remnants of an ancestral situation, or alternatively, whether they have appeared more recently during evolution as the result of genome rearrangements, perhaps with the recruitment of novel factors to mediate intermolecular intron assembly (Laird, 1989). Group II introns are split in different genes in the lineages of land plants (rps12) compared to Chlamydomonas spp (psaA), but this difference does not allow a distinction between the two alternatives. The split introns may have appeared after the separation of the two lineages, or may have been present in both genes in a common ancestor of algae and land plants and subsequently differentially lost. It is possible that the mechanism of trans-splicing is ancestral, and that it has repeatedly allowed the appearance of novel split introns during evolution.

C. Splicing of Heterologous Group II Introns Group II introns from organelles of other organisms have been introduced into the C. reinhardtii chloroplast genome by transformation. The rI1 intron from the mitochondrial rDNA of Scenedesmus obliquus was found to excise accurately when it was inserted, together with the flanking EBS1 sequence, within the tscA RNA (Herdenberger et al., 1994). In contrast, when the chloroplast gene of spinach,

Chapter 11 Chloroplast RNA Splicing containing a group II intron, was introduced in the C. reinhardtii chloroplast genome in an expression vector, the chimeric atpF transcript accumulated to high levels but the heterologous intron was not spliced (Deshpande et al., 1995). According to details of their structure, group II introns can be subdivided in two classes, IIA and IIB (Michel et al., 1989). To understand the different fates ofthe two heterologous introns, it may be significant that the rI1 intron is a member of subgroup IIB, that the atpF intron belongs to subgroup IIA, and that the two split introns of C. reinhardtii, although untypical, are more closely related to subgroup IIB. This could indicate that splicing factors in the C. reinhardtii chloroplast are specific for introns from subgroup IIB. The analysis of maize mutants deficient in chloroplast splicing also suggests that different splicing factors are specific to one or other of the subgroups (Jenkins et al., 1997). However it should also be noted that the rll intron is capable of self-splicing in vitro (Kück et al, 1990), while the atpF intron is not: thus the requirements for splicing factors may be more stringent for the latter than for the former.

IV. Perspective An important goal for the future will be to study the factors that promote splicing of group I and group II introns in vivo, and this will be facilitated by the isolation of the corresponding genes. Mutations identify numerous nuclear loci which are required for trans-splicing of the psaA group II introns, some of which have recently been cloned (Section III.B.2). The putative proteins that promote splicing of the psbA group I introns should also be particularly interesting, since splicing of these introns is regulated by light via a redox pathway. It would also be important to determine whether splicing of any of the other introns in photosynthesis-related genes is light-regulated. Finally, the development of proteindependent assays for splicing in vitro, or in organello, would advance studies of the factors that mediate splicing and its regulation.

Acknowledgments We thank Drs. S. Holloway and H.-H. Kim for communicating results prior to publication, and N. Roggli for preparing some of the figures. DLH and TCK were supported by grants from the US Dept. of

193 Agriculture (96-35301-3420), and the Robert A. Welch Foundation (F-1164) during preparation of the manuscript. MGC was supported by a grant from the Swiss National Fund for Scientific Research (31 34014.92).

References Bao Y (1993) Studies of a self-splicing group I ribozyme in the chloroplast psbA gene of Chlamydomonas reinhardtii. PhD Dissertation, University of Texas at Austin Bao Y and Herrin DL (1993) Nucleotide sequence and secondary structure of the chloroplast group I intron Cr.psbA-2: Novel features of this self-splicing ribozyme. Nucleic Acids Res 21: 1667 Bass B and Cech TR (1984) Specific interaction between the selfsplicing RNA of Tetrahymena and its guanosine substrate: Implications for biological catalysis by RNA. Nature 308: 820–826 Bonen L(1993) Trans-splicing of pre-mRNA in plants, animals and protists. FASEB J 7:40–46 Cech TR (1990) Self-splicing of group I introns. Annu Rev Biochem 59: 543–568 Cech TR, Damberger SH and Gutell RR (1994) Representation of the secondary and tertiary structure of group I introns. Nature Struct Biol 1: 273–280 Choquet Y, Goldschmidt-Clermont M, Girard-Bascou J, Kück U, Bennoun P and Rochaix J-D (1988) Mutant phenotypes support a trans-splicing mechanism for the expression of the tripartite psaA gene in the C. reinhardtii chloroplast. Cell 52: 903–913 Costa M, Dème E, Jacquier A and Michel F (1997) Multiple tertiary interactions involving domain II of group II selfsplicing introns. J Mol Biol 267: 520–536 Côté M-J and Tunnel, M (1995) In vitro self-splicing reactions of chloroplast and mitochondrial group-1 introns in Chlamy­ domonas eugametos and Chlamydomonas moewusii. Curr Genet 27: 177–183. Côté V, Mercier J-P, Lemieux C and Turmel M (1993) The single group I intron in the chloroplast rrnL gene of Chlamydomonas humicola encodes a site-specific DNA endonuclease. Gene 129: 69–76 Curcio MJ and Belfort M (1996) Retrohoming: cDNA-mediated mobility ofgroup II introns requires a catalytic RNA. Cell 84: 9–12 Daròs J and Flores R (1996) A group I plant intron accumulates as circular RNA forms with extensive 5´ deletions in vivo. RNA 2: 928–936 Deshpande NN, Hollingsworth M and Herrin DL (1995) The atpF group-I I intron-containing gene from spinach chloroplasts is not spliced in transgenic Chlamydomonas chloroplasts. Curr Genet 28: 122–127 Deshpande NN, Bao Y and Herrin DL (1997) Evidence for light/ redox regulated splicing of the psbA introns in Chlamydomonas chloroplasts. RNA 3: 37–48 Dujon B (1989) Group I introns as mobile genetic elements: Facts and mechanistic speculations—a review. Gene 82: 91– 114 Durocher V, Gauthier A, Bellemare G and Lemieux C (1989) An

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optional group I intron between the chloroplast small subunit rRNA genes of Chlamydomonas moewusii and C. eugametos. Curr Genet 15: 277–82 Dürrenberger F and Rochaix J-D (1991) Chloroplast ribosomal intron of Chlamydomonas reinhardtii: In vitro self-splicing, DNA endonuclease activity and in vivo mobility. EMBO J 10: 3495–3501 Gampel A and Tzagoloff A (1987) In vitro splicing of the terminal intervening sequence of Saccharomyces cerevisiae cytochrome b pre-mRNA. Mol Cell Biol 7: 2545–2551 Gampel A, Nishikimi M and Tzagoloff A (1989) CBP2 protein promotes in vitro excision of a yeast mitochondrial group I intron. Mol Cell Biol 9: 5424–5433 Gauthier A, Turmel M and Lemieux C (1991) A Group I intron in the chloroplast large subunit rRNA gene of Chlamydomonas eugametos encodes a double-strand endonuclease that cleaves the homing site of this intron. Curr Genet 19: 43–47 Goldschmidt Clermont M, Girard Bascou J, Choquet Y and Rochaix JD (1990) Trans-splicing mutants of Chlamydomonas reinhardtii. Mol Gen Genet 223: 417–25 Goldschmidt-Clermont M, Choquet Y, Girard-Bascou J, Michel F, Schirmer-Rahire M and Rochaix J-D (1991) A small chloroplast RNA may be required for trans-splicing in Chlamydomonas reinhardtii. Cell 65: 135–143 Grivell LA (1996) Transposition: Mobile introns get into line. Curr Biol 6: 48–51 Guo Q and Lambowitz A (1992) A tyrosyl-tRNA synthetase binds specifically to the Group I intron catalytic core. Genes Dev 6: 1357–1372 Harris EH (1989) The Chlamydomonas Sourcebook: A Comprehensive Guide to Biology and Laboratory Use. Academic Press, San Diego Harris EH, Boynton JE and Gillham NW (1974) Chloroplast ribosome biogenesis in Chlamydomonas. Selection and characterization of mutants blocked in ribosome formation. J Cell Biol 63: 160–179 Herdenberger F, Holländer B and Kück U (1994) Correct in vivo RNA splicing of a mitochondrial intron in algal chloroplasts. Nucleic Acids Res 22: 2869–2875 Herrin D and Michaels A (1984) Gene Expression during the Cell Cycle of Chlamydomonas reinhardtii. In: Stein G and Stein J (eds) Recombinant DNA and Cell Proliferation, pp 87– 106. Academic Press, Orlando Herrin DL and Schmidt GW (1988) trans-Splicing of transcripts for the chloroplast psaA gene. In vivo requirement for nuclear gene products. J Biol Chem 263: 14601–14604 Herrin DL, Michaels AS and Paul A-P (1986) Regulation of genes encoding the large subunit of ribulose-1,5-bisphosphate carboxylase and the photosystem II polypeptides D-1 and D-2 during the cell cycle of Chlamydomomas reinhardtii. J Cell Biol 103: 1837–1845 Herrin DL, Chen Y-F and Schmidt GW (1990) RNA splicing in Chlamydomonas chloroplasts: Self-splicing of 23S preRNA. J Biol Chem 265: 21134–21140 Herrin DL, Bao Y, Thompson AJ and Chen Y-F (1991) Selfsplicing of the Chlamydomonas chloroplast psbA introns. Plant Cell 3: 1095–1107 Hetzer M, Schweyen RJ and Mueller MW (1997) DNA polymerization catalysed by a group II intron RNA in vivo. Nucleic Acids Res 25: 825–1829 Jaeger L, Michel F and Westhof E (1996) The structure of group

I ribozymes. In: Eckstein F and Lilley DMJ (eds) Nucleic Acids and Molecular Biology, Vol 10, pp 33–52. SpringerVerlag, Berlin Jacquier A (1996) Group II introns: Elaborate ribozymes. Biochimie 78: 474–487 Jenkins BD, Kulhanek DJ and Barkan AB (1997) Nuclear mutations that block group II RNA splicing in maize chloroplasts reveal several intron classes with distinct requirements for splicing factors. Plant Cell 9: 283–296 Kück U, Choquet Y, Schneider M, Dron M and Bennoun P (1987) Structural and transcriptional analysis of two homologous genes for the P700 chlorophyll a apoproteins in Chlamydomonas reinhardtii: Evidence for trans-splicing. EMBO J 6: 2185–2195 Kück U, Godehardt I and Schmidt U (1990) A self-splicing group II intron in the mitochondrial large subunit rRNA (LSUrRNA) gene of the eukaryotic alga Scenedesmus obliquus. Nucleic Acids Res 18: 2691–2697 Laird PW (1989) Trans-splicing in trypanosomes—archaism or adaptation? Trends Genet 5: 204–216 Liu X-Q, Gillham NW and Boynton JE (1989) Chloroplast ribosomal protein gene rps12 of Chlamydomonas reinhardtii. J Biol Chem 27: 16100–16108 Michel F and Dujon B (1983) Conservation ofsecondary structures in two intron families including mitochondrial-, chloroplastand nuclear-encoded members. EMBO J 2: 33–38 Michel F and Ferat J-L (1995) Structure and activities of group II introns. Annu Rev Biochem 64: 435–461 Michel F and Westhof E (1990) Modelling of the threedimensional architecture of Group I catalytic introns based on comparative sequence analysis. J Mol Biol 216: 585–610 Michel F, Umesono K and Ozeki H (1989) Comparative and functional anatomy of group II catalytic introns—a review. Gene 82: 5–30 Mohr G, Zhang A, Gianelos J, Belfort M and Lambowitz A (1992) The Neurospora cyt-18 protein suppresses defects in the phage T4 td intron by stabilizing the catalytically active structure of the intron core. Cell 69: 483–494 Mueller MW, Hetzer M and Schweyen RJ (1993) Group II intron RNA catalysis of progressive nucleotide insertion: A model for RNA editing. Science 261: 1035–1038 Plant AL and Gray JC (1988) Introns in chloroplast proteincoding genes of land plants. Photosynth Res 16: 23–39 Rochaix J-D (1996) Post-transcriptional regulation of chloroplast gene expression in Chlamydomonas reinhardtii. Plant Mol Biol 32: 327–341 Rochaix J-D, Rahire M and Michel F (1985) The chloroplast ribosomal intron of Chlamydomonas reinhardtii codes for a polypeptide related to mitochondrial maturases. Nucleic Acids Res 13: 975–984 Roitgrund C and Mets LJ (1990) Localization of two novel chloroplast genome functions: Trans-splicing of RNA and protochlorophyllide reduction. Curr Genet 17: 147–153 Saldanha R, Mohr G, Belfort M and Lambowitz A (1993) Group I and Group II introns. FASEB J 7: 15–24 Shaw LC and Lewin AS (1995) Protein induced folding of a Group 1 intron in Cytochrome b pre-mRN A. J Biol Chem 270: 21552–21562 Thompson AJ and Herrin DL (1991) In vitro self-splicing reactions of the chloroplast Group I intron Cr.LSU from Chlamydomonas reinhardtii and in vivo manipulation via gene replacement.

Chapter 11 Chloroplast RNA Splicing Nucleic Acids Res 19: 6611–6618 Thompson AJ and Herrin DL (1994) A chloroplast group I intron undergoes the first step of reverse splicing into host 5.8S rRNA: Implications for intron-mediated RNA recombination, intron transposition and 5.8S rRNA structure. J Mol Biol 236: 455–468 Thompson AJ, Yuan X, Kudlicki W and Herrin DL (1992) Cleavage and recognition pattern of a double-strand-specific endonuclease (l-Crel) encoded by the chloroplast 23S rRNA intron of Chlamydomonas reinhardtii. Gene 119: 247–251 Tunnel M, Boulanger J and Lemieux C (1989) Two group I introns with long internal open reading frames in the chloroplast psbA gene of Chlamydomonas moewusii. Nucleic Acids Res 17: 3875–87 Turmel M, Boulanger J, Schnare MN, Gray MW and Lemieux C (1991) Six group I introns and three internal transcribed spacers in the chloroplast large subunit ribosomal RNA gene of the green alga Chlamydomonas eugametos. J Mol Biol 218: 293– 311 Turmel M, Gutell RR, Mercier JP, Otis C and Lemieux C (1993a) Analysis of the chloroplast large subunit ribosomal RNA gene from 17 Chlamydomonas taxa. Three internal transcribed spacers and 12 group I intron insertion sites. J Mol Biol 232: 446–467 Turmel M, Mercier J-P and Côté M-J (1993b) Group I introns interrupt the chloroplast psaB and psbC genes in Chlamy­ domonas. Nucleic Acids Res 21: 5242–5250

195 Turmel M, Choquet Y, Goldschmidt-Clermont, Rochaix J-D, Otis C and Lemieux C (1995a) The trans-spliced intron 1 in the psaA gene of the Chlamydomonas chloroplast: A comparative analysis. Curr Genet 27: 270–279 Turmel M, Côté V, Otis C, Mercier J-P, Gray MW, Lonergan KM, and Lemieux C (1995b) Evolutionary transfer of ORFcontaining group I introns between different subcellular compartments (chloroplast and mitochondria). Mol Biol Evol 12: 533–45 Turmel M, Mercier J-P, Côté V, Otis C and Lemieux C (1995c) The site-specific DNA endonuclease encoded by a group I intron in the Chlamydomonas pallidostigmatica chloroplast small subunit rRNA gene introduces a single-strand break at low concentrations of Nucleic Acids Res 23: 2519– 2525 Weeks KM and Cech TR (1995) Protein facilitation of group I intron splicing by assembly of the catalytic core and the 5´ splice site domain. Cell 82: 221–230 Woodson SA and Cech TR (1989) Reverse self-splicing of the Tetrahymena group I intron: Implication for the directionality of splicing and for intron transposition. Cell 57: 335–345. Zimmerly S, Guo H, Eskes R, Yang J, Perlman PS and Lambowitz AM (1995) A group II intron RNA is a catalytic component of a DNA endonuclease involved in intron mobility. Cell 83: 529–538

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Chapter 12

Regulation of Chloroplast Translation Charles R. Hauser, Nicholas W. Gillham and John E. Boynton DCMB Group, Departments of Botany and Zoology, Box 91000, Duke University, Durham, NC 27708-1000, U.S.A. Summary I. Introduction II. The Role of Physiological and Environmental Factors in Translational Control III. Current Biochemical and Genetic Approaches to Dissect Mechanisms of Translational Regulation IV. Cis-acting Sequences Involved in Translation Initiation V. Translational Regulation Involves Interactions between cis-Acting Sequences and trans-Acting Factors A. Regulation of Individual Photosynthetic Proteins B. Coordinate Regulation of Photosynthetic versus Ribosomal Proteins VI. Ribosomes, Membranes and Tethers VII. Translational Regulation of Complex Assembly A. Chloroplast Thylakoid Membranes B. Chloroplast Ribosomes VIII. How are the Regulatory Proteins Regulated? IX. Is there Hierarchical Control of Chloroplast mRNA Translation? Acknowledgments References

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Summary During the evolution of chloroplasts from cyanobacterial endosymbionts, many genes encoding components necessary for protein synthesis and photosynthesis have been transferred to the nucleus. Assembly of the machinery for both processes now relies on the concerted expression of genes in the nuclear and plastid genomes. Evidence accumulating in C. reinhardtii indicates that the expression and regulation of chloroplast genes probably differs in certain respects from the prokaryotic (E. coli) paradigm. Both mutational analysis of putative Shine-Dalgarno (SD) sequences, and creation of canonical SD sequences (–9 to –5) reveals that initiation is not mediated by SD-16S rRNA interactions for the majority of chloroplast-encoded mRNAs. Recent evidence in cyanobacteria, the most likely ancestors of the Chlamydomonas chloroplast, indicates that most genes lack SD sequences at the classical position. Interactions between cis-acting sequences in the 5´ untranslated regions (UTRs) and specific trans-acting nuclear gene products appear to mediate translation of chloroplast-encoded mRNAs for specific genes. Both UV crosslinking and gel mobility shift assays with several chloroplast leaders show that multiple proteins interact with these sequences. Some of these appear to be involved in gene-specific regulation, whereas others may be core proteins of a general ribonucleoprotein (RNP) complex. Inverted repeat sequences present in most chloroplast leaders predict higher order structures within the 5´ UTR of mRNAs that could serve as scaffolding for the formation of the RNP complex. One model predicts that the RNP complex directly enhances ribosome binding and translation initiation. A second model predicts that the RNP complex plays a role in making topological distinctions in the sites of synthesis of chloroplast mRNAs. A challenge in the coming years will be to determine which, if any, of these models are correct. J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 197–217. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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I. Introduction Posttranscriptional control plays a major role in regulating the expression of chloroplast and mitochondrial genes. This chapter specifically focuses on translational control mechanisms, whereas posttranscriptional control mechanisms including mRNA stabilization and degradation, processing, and splicing are reviewed elsewhere in this volume (Chapter 9, Nickelsen; Chapter 10, Stern and Drager; Chapter 11, Herrin et al.). Before considering trans­ lational control mechanisms in the chloroplasts of C. reinhardtii and land plants, we will briefly review comparable mechanisms operative in prokaryotes and eukaryotes. This is relevant because chloroplasts and mitochondria almost certainly arose from cyanobacteria and purple bacteria respectively by endosymbiosis (Gray, 1992) and many essential organelle genes were subsequently transferred to the eukaryotic nucleus (Gillham, 1994). Although the chloroplast translational apparatus is obviously of prokaryotic origin and resembles that of bacteria in many respects (Harris et al., 1994; Mache, 1990), it may have acquired certain eukaryotic characteristics over time. About two thirds of chloroplast ribosomal proteins and all the chloroplast tRNA synthetases in land plants and most algae are known to be encoded in the nucleus, synthesized in the cytosol and imported into the organelle (Gillham, 1994; Harris et al., 1994). Certain initiation and elongation factors appear to be chloroplast encoded (e.g. IF-1 in land plants, IF-2 in red algae and EF-Tu in C. reinhardtii and Euglena gracilis). However, analysis of several completely sequenced plastid genomes indicates that genes encoding the majority of the proteins involved in mediating translation on chloroplast ribosomes are absent from the chloroplast genomes of land plants and they are therefore assumed to be encoded by the nuclear genome of these organisms (Reith, 1995). We will begin by examining translational regulation in prokaryotes and then compare this to what is Abbreviations: 5´UTR – 5´ untranslated region; AAD – aminoglycoside adenine transferase; ASD-anti Shine-Dalgarno sequence; DBMIB – 2,5-dibromo-3-methyl-6-isopropyl-pbenzoquinone; DCMU – 3-(3,4 dichlorophenyl)-1,1 -dimethyl urea; DTNB–dithionitrobenzoicacid; PAR–photosynthetically active radiation; PQ – plastoquinone; p-protein – photosynthetic protein; RNP-ribonucleoprotein complex; r-protein–ribosomal protein; Rubisco – ribulose-1,5 bisphosphate carboxylase/ oxygenase; SD – Shine-Dalgarno sequence (GGAGG)

known in eukaryotes. In prokaryotes, control of translation initiation is dependent upon several factors (McCarthy and Brimacombe, 1994;Voorma, 1996). The pool of free 30S subunits available to begin translation is determined by the binding of initiation factors IF1 and IF3. These shift the equilibrium of the 70S ribosome monomers and their subunits towards dissociation into the 30S and 50S subunits. The 30S subunit, carrying IF 1, IF3, and probably IF2 as well, interacts with mRNA, and GTP through a series of intermediates giving rise to the 30S initiation complex. Recognition is governed largely by the extent of secondary structure in the mRNA and by two RNA-RNA interactions: 1) the codon-anticodon interaction between and the initiation codon (usually AUG), and 2) base pairing between a conserved purine rich ShineDalgarno (SD) sequence (GGAGG) 7 ± 2 nucleotides upstream of the mRNA initiator AUG codon and a complementary pyrimidine rich anti-SD (ASD) sequence near the 3´ end of the 16S rRNA in the small subunit. These two RNA-RNA interactions stabilize the 30S preinitiation complex in the proper reading frame. Other mRNA interactions may involve sites on the 30S subunit that might be regarded as providing an enlarged ‘catchment area’ capable of binding mRNA molecules that subsequently move into the major mRNA track (McCarthy and Brimacombe, 1994) Ribosomal protein S1 also plays a role in mRNA recognition in E. coli (Voorma, 1996). The amino terminus of this protein is associated with the 30S subunit by protein-protein interactions, but its long and flexible carboxyl terminal domain has a high affinity for pyrimidine sequences. Many E. coli mRNAs have pyrimidine rich sequences upstream of the SD sequence and the RNAs of phages f 2 and have U rich sequences upstream of the coat protein open reading frame which may represent S1 binding sequence sites. Site-directed mutagenesis of a upstream of rnd, an E. coli gene encoding RNase D, revealed that alteration of two to five U residues within this sequence had no effect on mRNA levels, but decreased RNase D protein and activity by as much as 95 percent. Furthermore, E. coli 30S subunits will recognize translational start sites of mRNAs lacking SD sequences such as alfalfa mosaic virus RNA4 and tobacco mosaic virus RNA only if S1 and IF3 are present. In fact, a recent scheme for initiation complex formation starts with S1-dependent formation of a 30S-mRNA binary complex followed

Chapter 12

Chloroplast Translation

by the SD-ASD interaction. Two major patterns of negative regulation have been identified in prokaryotes (Gold, 1988; McCarthy and Gualerzi, 1990;Voorma, 1996). The first involves repression of translation by trans-acting proteins that block formation of competent initiation complexes. Thus, the ribosomal (r-) protein operons are negatively auto-regulated by specific r-proteins whose target is usually the translational initiation region of the first gene in the operon. The second mechanism modulates mRNA secondary structure in order to control availability ofthe ribosome binding site for initiation of complex formation. In addition, nascent peptides have been found to play a role in translational regulation of specific genes via mechanisms as diverse as translational attenuation, ribosome hopping, and auto-regulation of mRNA half-life (Lovett, 1994). Translational regulation in eukaryotes has been reviewed by Standart and Jackson (1994) and in a recent volume edited by Hershey et al. (1996). In eukaryotic mRNAs the 5´ cap, the ‘context’ of the sequence flanking the initiator AUG, enhancer sequences (Kozak, 1983; Yamamoto et al., 1995; Schmitz et al., 1996), and the presence of upstream open reading frames (uORFs) in the 5´ UTR (Mathews et al., 1996) all affect the general efficiency of translation. Translational regulation of uORF­ containing mRNAs depends on many factors including the amino acid sequence encoded by the uORF, the length of intercistronic regions, and the sequence context of the termination codon of the uORF. In addition, the translation of arg-2 mRNA is negatively regulated in vitro by increasing concen­ trations of arginine (Wang and Sachs, 1997). Specific cis-elements and trans-acting factors are also important in regulation of translation. A wellstudied example of a eukaryotic translational repressor acting on the 5´ UTR is the iron-binding proteins (IRP1 and 2) which interact with a stemloop structure (IRE) in the 5' UTR of ferritin mRNA to turn off translation. In the absence of IRP, the IRE acts as a positive translational enhancer and causes preferential binding of initiation factors to ferritin mRNA (Standart and Jackson, 1994; Kim et al., 1995). Eukaryotic initiation factors (eIF) and the poly(A) tail at the 3´ end of eukaryotic mRNAs acting in synergy with the 5´ cap structure of the mRNA are also involved in regulating expression of eukaryotic mRNAs. Formation of the eukaryotic pre-initiation

199 complex with the 40S subunit is an elaborate process which requires not only the eIFs, but also novel properties of the poly(A) binding protein (Hentze, 1995, 1996; Proweller and Butler, 1996). The cap is bound by eIF-4E which plugs into the N-terminal portion of an adapter protein, eIF-4G whose central portion is associated with eIF-3, a multimeric complex bound to the 40S subunit. The multifunctional eIF­ 4G protein is also the site of interaction of the poly (A) binding protein which helps to recruit 40S subunits. Since the polyadenylation process itself depends on a whole array of proteins (Proudfoot, 1996), each of these is a potential target for translational regulation. Translational regulation in chloroplasts and mitochondria has been the subject of numerous reviews (Gillham et al., 1994; Mayfield et al., 1995; Fox, 1996a,b; Rochaix, 1996; Sugita and Sugiura, 1996; Cohen and Mayfield, 1997). Most of the data for mitochondria and chloroplasts derive from experi­ ments with yeast and C. reinhardtii respectively. In both organelles control is frequently exerted via trans-acting factors which bind to the 5´ untranslated regions (5´UTRs) of organellar mRNAs and affect message half-life or translatability. Fox (1996a,b) calls attention to the fact that translational activators binding to the 5´UTRs of specific mitochondrial mRNAs also interact with the small subunit of the mitochondrial ribosome, quite probably with specific ribosomal proteins. Although polyadenylation was considered to be a unique property of nucleus-encoded mRNA in eukaryotic cells, polyadenylation of chloroplastencoded mRNAs has recently been reported (Kudla et al., 1996, Lisitsky et al., 1996). In vivo, endonucleolytic cleavage of the petD mRNA is followed by polyadenylation ofthe cleavage products which in turn substantially increases their rate of degradation (Kudla et al., 1996). Similarly, mRNA can be targeted for rapid degradation in vitro by the addition of a poly(A)-rich sequence to an endo­ nucleolytic cleavage product (Lisitsky et al., 1996). Following transcription, chloroplast precursor mRNA may follow one of two paths: 1) undergo 3´ end formation to generate a stable product for translation, or 2) endonucleolytic cleavage and addition of poly(A)-rich sequences leading to rapid degradation. This branch point in mRNA metabolism is regulated in vitro by ATP concentration. However, Lisitsky et al. (1996) state that in vivo this branch point may be regulated by the chloroplast redox potential,

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photosynthetic electron flow or specific trans-acting proteins. Clearly, regulation of the enzymes responsible for chloroplast mRNA polyadenylation must now be considered among the factors determining which transcripts become available for translation. In addition, translation of chloroplast mRNAs may be regulated by other diverse mechan­ isms such as alternative processing of mRNA 5´ ends (Reinbothe et al., 1993, Shapira et al., 1997) or codon bias optimized for translational efficiency (Morton, 1996). These observations, taken together with the findings reviewed above relating to translational control mechanisms in eukaryotes and prokaryotes, should alert investigators studying translational control mechanisms in chloroplasts. The roles of specific chloroplast ribosomal proteins, initiation factors and other diverse factors (e.g. nascent peptides) will probably need to be taken into account in trying to reach a complete depiction of the events that accompany activation of chloroplast messages for translation.

II. The Role of Physiological and Environmental Factors in Translational Control Chloroplast development in response to light exposure in most land plants and green algae is controlled at the transcriptional, post-transcriptional and transla­ tional level. Light mediated reduction ofprotochloro­ phyllide accumulated in the dark to chlorophyllide triggers reorganization of the paracrystalline prolamellar body of the etioplast, synthesis of new chlorophyll pigment and photosynthetic proteins, formation of photosynthetic thylakoid membranes and their organization into grana stacks (von Wettstein et al., 1995). Clear evidence exists that transcription of many nuclear genes encoding photosynthetic proteins is activated by light (Gilmartin et al., 1990; Schindler and Cashmore, 1990). Concomitantly, light mediates chloroplast gene expression primarily at the translational level, although transcription and stability of chloroplast encoded mRNAs are also enhanced by light induced greening (Gruissem and Tonkyn, 1993; Gillham et al., 1994; Mayfield et al., 1995). In contrast to angiosperms, wild-type C. reinhardtii, like gymnosperms and liverworts (Gillham, 1994), has a light-independent protochloro­ phyllide reductase. Therefore, this alga reduces protochlorophyllide to chlorophyllide enzymatically

in the dark and maintains highly differentiated chloroplasts under heterotrophic growth conditions (Harris, 1989). Light has been shown to regulate translation of psbA mRNA encoding the D1 protein of Photo­ system II, which is degraded and re-synthesized more rapidly than any other chloroplast protein (Fig. 1). Synthesis of this protein is enhanced in wild-type C. reinhardtii grown phototrophically versus hetero­ trophically (Danon and Mayfield, 1991; Hauser et al., 1996). Pulse-labeling experiments indicate that D1 is not synthesized in heterotrophically grown cells of y1 and other mutants blocked in the enzymatic reduction of protochlorophyllide, but synthesis of D1 is activated by exposure of such mutants to light (Malnoë et al., 1988; Danon and Mayfield, 1991). However, a fast turn-over of D1 in these mutants cannot be ruled out. Binding of 47 and 60 kDa proteins to a 36 nt stem-loop structure upstream of the ribosome binding site in the 5´UTR of the psbA mRNA has been reported to correlate with light stimulated D1 synthesis in wild-type cells (Danon and Mayfield, 1991) and to be controlled by the redox potential in the chloroplast (Danon and Mayfield, 1994). However, a 47 kDa protein was also shown to crosslink to the leaders of the chloroplast rbcL and atpB mRNAs encoding photosynthetic proteins and the chloroplast rps7 and rps12 mRNAs encoding ribosomal proteins when extracts from heterotrophic and phototrophic cells were compared (Hauser et al., 1996; Rochaix, 1996). Whether the same or distinct 47 kDa proteins are involved in these binding reactions remains to be ascertained (Gillham et al., 1994; Hauser et al., 1996; Rochaix, 1996). Silk and Wu (1993) found that illumination of dark grown y1 cells is also accompanied by a five­ fold increase in the accumulation of tufA mRNA encoding the elongation factor EF-Tu due to an increased half-life of this mRNA in the light. While the significance of changes in tufA transcript levels is unknown, the authors speculate that EF-Tu might be essential for the accumulation ofchloroplast encoded proteins during greening of y1 cells. Transfer of low light adapted C. reinhardtii (CC­ PAR, 125) cells to high light (70 and respectively) has a dramatic transient effect on the differential translation of psbA and rbcL mRNAs (Shapira et al., 1997; Fig. 1). Synthesis of the D1 protein increases ten-fold during the first six hours whereas synthesis of ribulose-1,5 bisphosphate carboxylase/oxygenase large subunit (Rubisco LSU)

Chapter 12 Chloroplast Translation

drops dramatically within 15 minutes and only gradually resumes at the end of the six hour period. Changes in D1 and LSU synthesis cannot be explained by shifts in the levels of accumulated psbA or rbcL mRNA or by the temporary decrease observed in the ratio of the short to long form of the rbcL transcripts. These several distinct effects of temporary light stress are correlated with a rapid, sustained increase a transient decline in in the reduction state of photosynthetic activity, a less rapid drop in total chlorophyll content and a delay in cell division. Whether these changes result in differential binding of translational activator and repressor proteins to the psbA and rbcL mRNA leaders in response to changes in the cell’s redox potential or to another regulatory mechanism involving the generation of free radicals, remains to be determined.

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Most studies of expression of nuclear and chloroplast genes affecting photosynthetic function in C. reinhardtii are carried out on cells grown mixotrophically on medium containing acetate as a reduced carbon source. The presence of reduced carbon sources represses expression of nuclear RbcS and Cab genes encoding the Rubisco small subunit and the chlorophyll a/b binding protein in C. rein­ hardtii (Gibbs et al., 1986; Kindle, 1987) and strongly inhibits the expression of nuclear genes encoding seven different photosynthetic proteins in maize tissue culture cells (Sheen, 1990). Much less has been published about the effect of carbon source on chloroplast gene expression. In C. reinhardtii cells grown in the light on acetate, the level of D1 protein increased and the level of the LSU protein was reduced, but no differences were detected in extracts

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from these cells with respect to the binding of six proteins that crosslink to the 5´UTRs of mRNAs for three genes encoding photosynthetic proteins and two genes encoding ribosomal proteins (Hauser et al., 1996). Internal concentration may also affect chloroplast gene expression at the translational level (Winder et al., 1992). Transfer of phototrophically grown cells of C. reinhardtii from elevated to limiting resulted in transient declines in synthesis of both the chloroplast-encoded large subunit and the nucleus-encoded small subunit of Rubisco with no changes in levels of the corresponding mRNAs. Nitrogen starvation in C. reinhardtii is known to effect a significant reduction in the levels of both chloroplast and cytoplasmic ribosomes as vegetative cells differentiate into sexual gametes (Harris, 1989). During a 10–12 h period following restoration of nitrogen, gametes begin to regenerate into vegetative cells, resynthesize rRNAs and ribosomal proteins and assemble the components into the normal complements of chloroplast and cytoplasmic ribosomes without cell division (Myers et al., 1984). Whether preferential translation of chloroplast mRNAs encoding chloroplast ribosomal proteins observed under conditions of reduced chloroplast protein synthesis (see below) allows C. reinhardtii cells to respond to fluctuations in nitrogen content in their environment remains to be determined.

III. Current Biochemical and Genetic Approaches to Dissect Mechanisms of Translational Regulation Development of techniques for stable transformation of chloroplast genes in C. reinhardtii (Boynton et al., 1988; Boynton and Gillham, 1993) and tobacco (Svab et al., 1990; Maliga, 1993; Svab & Maliga, 1993) has permitted the direct demonstration that the 5´UTR regions of chloroplast mRNAs are essential for their translation (Mayfield et al., 1994; Staub and Maliga, 1994; Zerges and Rochaix, 1994; Zerges et al., 1997). Recent efforts to identify cis-acting elements required for mRNA translation and stability have utilized the dominant eubacterial aadA gene (conferring spectinomycin resistance) in chimeric reporter constructs as a rapid means of accessing the functionality of putative cis regulatory elements in chloroplasts of C. reinhardtii (GoldschmidtClermont, 1991; Nickelsen et al., 1994; Zerges and

Rochaix, 1994; Fargo et al., 1997). Furthermore, analysis of AAD expression from reporter constructs in the progeny ofcrosses involving photosynthetic or ribosomal protein mutant strains and the wild type has allowed direct demonstration of the requirement for 5' UTR sequences in translalional regulation (Nickelsen et al., 1994; Zerges and Rochaix, 1994; C. R. Hauser, A. M. Johnson, N. W. Gillham and J. E. Boynton, unpublished). Chloroplast transformants containing 5´ UTR: aadA: 3´UTR reporter constructs should also be valuable for use in isolating cis-acting (5´ and/or 3´UTR) mutants in the chloroplast sequences and trans-acting nuclear mutations that suppress loss of function mutations in the cis-acting sequences. Gel mobility shifts (GMS), UV crosslinking and RNA affinity chromatography (Leibold and Munro, 1988; Meerovitch et al., 1989) have been utilized to identify the proteins mediating 5´UTR-driven selective translation in C. reinhardtii. Heparinenriched extracts from whole cells or chloroplast fractions isolated and analyzed in the presence of reductants have been used to identify 5´ UTR-binding proteins. Since the reduction state of certain 5´UTRbinding proteins has been shown to affect RNP complex formation in vitro (Danon and Mayfield, 1994b), caution should be exercised in extrapolating these in vitro results to in vivo regulation. GMS identifies a ribonucleoprotein complex (RNP) by the retarded migration of an RNA in a non-denaturing gel system, whereas UV crosslinking identifies the proteins in the complex which are in direct contact with labeled nucleotides in the RNA. Second dimension analysis by SDS PAGE of the proteins binding to an RNA identified as a gel mobility shifted band in a GMS assay, can identify additional proteins in the complex, regardless of whether the proteins participate via protein-RNA or protein-protein interactions. RNA affinity chromatography utilizing specific RNA ligands immobilized on matrices is a powerful approach to identify the trans-acting factors which bind to the chloroplast leader sequences (Rouault et al., 1989; Danon and Mayfield, 1991, Prokipcak et al., 1994). However, this method has so far seen only limited application in the identification of trans-acting proteins that bind to chloroplast 5' UTRs (Danon and Mayfield, 1991). The widely used yeast 2-and 3-hybrid systems designed to identify protein:protein or protein:RNA interactions required for gene expression (Fields and Sternglanz, 1994; Putz et al, 1996; SenGupta et al., 1996) should

Chapter 12

Chloroplast Translation

also be useful for the identification of trans-acting proteins that are part of the chloroplast 5´UTR complexes, particularly for those proteins that do not bind directly to RNA. With improved nuclear transformation efficiencies in C. reinhardtii and the development of dominant nuclear selectable markers (Stevens et al., 1996; Cerutti et al., 1997a,b), genomic complementation (Purton and Rochaix, 1994; Zhang et al. 1994; Vashishtha et al., 1996; Funke et al., 1997) and tagging of nuclear genes (Tam and Lefebvre, 1993; Gumpel and Purton, 1994) are viable routes for the identification of genes encoding trans-acting proteins which regulate the localization and/or translation of mRNAs.

IV. Cis-acting Sequences Involved in Translation Initiation Although chloroplast ribosomes share many structural and functional properties with prokaryotic ribosomes, questions have arisen as to whether translation initiation involves different mechanisms in the two systems. Conserved anti-SD sequences occur near the 3´ end of the highly conserved 16S rRNA molecule from chloroplast ribosomes of higher plants, green algae and cyanobacteria (Harris et al., 1994; Kaneko et al., 1996). However, complementary SD sequences are missing from the leaders of certain chloroplast and cyanobacterial genes, and when present, are hyper-variable in position and nucleotide composition (Gillham et al., 1994; Harris et al., 1994; Kaneko et al., 1996; Fargo et al., 1997). Furthermore, tracts and other prokaryotic UTR sequences reported to enhance translation are absent from the 5´UTRs of chloroplast mRNAs examined to date (Fargo et al., 1997). Experiments have been carried out in tobacco and C. reinhardtii to test whether SD-like sequences present in the 5´UTRs of chloroplast mRNAs function in translation as they do in E. coli. In tobacco, addition of a canonical SD sequence (GGAGG, –11 to –7) to the strong constitutive rRNA operon promoter (Prrn) fused to the uidA gene encoding the (GUS) reporter enzyme, leads to the efficient translation of nidA mRNA in chloroplast transformants (Staub and Maliga, 1994). In C. reinhardtii transformants, deletion of the SD-like sequence (GGAG) located 27 nt upstream of the ATG in the psbA mRNA reduced both half-life

203 and translation of this message greatly (Mayfield et al., 1994). Two dimensional modeling studies of the mutant leader using the M-fold algorithm (Zuker, 1994) suggest that the four nucleotide deletion is sufficient to alter the predicted secondary structure of the leader (Fargo et al., 1997). Strains containing a mutation (CUCC) within the putative 36 nt stemloop of the leader which sequesters the SD sequence (GGAG) by base pairing and extends the stem also blocked psbA expression and severely reduced mRNA stability (Mayfield et al., 1994). Mutant leaders containing a 32 nt deletion immediately upstream of the SD sequence translate psbA mRNA at high rates (Mayfield et al., 1994). This deletion, which eliminates 65% of the putative stem-loop sequence encompassing presumed binding sites for the 47 and 60 kDa proteins (Danon and Mayfield, 1991), is modeled to have an unstructured SD sequence with a short stem loop at the immediate 5´ end of the message. This stem loop may serve as a negative translational attenuator (Mayfield et al., 1994). Whether the observed reduction in psbA expression in these 5' UTR mutants resulted directly from a reduction in mRNA translation, or from a reduction in mRNA half-life, cannot be ascertained from the published data (Rochaix, 1996). Interpretation is further complicated by evidence that the half-life of chloroplast mRNAs can be enhanced or diminished by their association with ribosomes (Sakamoto et al., 1994; Yohn et al., 1996; Chapter 9, Nickelsen). Using an in vitro translation system derived from tobacco chloroplasts and a series of mutant psbA 5´UTRs, Hirose and Sugiura (1996) identified four elements within the psbA 5´UTR which are required for translation in vitro. Three of these 5´UTR elements appear to be putative ribosome binding sites (RBSs), RBS1 (AAG [–9]), RBS2 (UGAUGAU [–22]) and RBS3 (GGAG [–33]) based on complementarity to the 3´ terminus ofthe tobacco chloroplast 16S rRNA. Replacement mutagenesis of RBS1 or RBS2 reduced translation to 33% and 38% of wild type respectively, while mutagenesis of the canonical SD sequence, RBS3, had little effect (12% decrease) on translation. Disruption of both RBS1 and RBS2 resulted in a drastic decrease (92%) in translation whereas the mRNAs from two other double mutants (RBS1&3 and RBS2&3) were translated at levels similar to the single mutants RBS1 or RBS2. Deletion of a fourth element (UAAAUAAA [–17]) abolished translation and the authors suggest this sequence may be a target for potential trans-acting translation factor(s).

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Deletion of ca. 70% of the chloroplast petD 5´ UTR from a translational fusion with the uidA reporter gene reduced mRNA half-life four- to six-fold and glucuronidase activity 8 fold in chloroplast transformants of C. reinhardtii while the transcription rate of the uidA gene was unaltered (Sakamoto et al., 1993). In contrast, replacement mutagenesis of a trinucleotide SD-like sequence located 10 nt upstream of the ATG codon had no effect on the accumulation of the petD gene product subunit IV (Sakamoto et al., 1994a). Similarly, a change in the psbD SD sequence from GGAG to AAAG did not affect phototrophic growth or photosynthetic efficiency (unpublished data cited in Rochaix, 1996). The putative SD-like sequences in A-U rich 5´UTRs of two r-protein mRNAs (rps4 [SD at–149] and rps 7 [SD at –116]) and two photosynthetic protein mRNAs (atpB [SD at –85] and atpE [SD at –22]) were altered by replacement mutagenesis to destroy complementarity to the ASD in the 16S rRNA (Fargo et al., 1997). Transformants homoplasmic for a chloroplast expression cassette carrying the four upstream regulatory regions with either the wildtype or mutated SD-like sequences fused to the bacterial aadA reporter gene showed identical levels of spectinomycin resistance in vivo and identical levels of aminoglycoside adenine transferase (AAD) activity in vitro. Similar results were obtained when the uidA sequence encoding was substituted for the aadA coding sequence in the rps4 and atpB leader constructs. Based on the observation that certain chloroplast genes can be transcribed and their mRNAs translated in E. coli (Liu et al., 1989a), expression of these 5´UTR reporter constructs carried on pUC18 plasmids was also examined in E. coli. Whether wild-type or mutant SD-like sequences were present, all four chloroplast promoter/leader regions coupled to aadA resulted in comparable levels of spectino­ mycin resistance and AAD activity in the bacterial transformants (Fargo et al., 1997). Similar results were obtained in the case of E. coli transformants containing the uidA gene under the control of rps4 and atpB regulatory regions with the wild-type and mutant SD-like sequences. Creation of a canonical SD sequence (GGAGG) by replacement mutagenesis at positions –9 to –5 in the leaders of these constructs resulted in a slight elevation in reporter gene expression in E. coli and no detectable increase in C. reinhardtii. The use of two independent and

functionally unrelated reporter genes with the different chloroplast leaders greatly diminishes the possibility that internal coding sequences could be promoting formation of the initiation complexes in the absence of the SD-like sequences. In contrast, expression of both the aadA and uidA reporter genes in E. coli was also shown to require the presence of an active SD sequence when these genes were fused to the 5´UTR of mutant and wild-type leaders of the phage fI gene VII. Native gene VII mRNA contains a defective initiation site and is inactive in assays of independent initiation (Ivey-Hoyle and Steege, 1992). The chloroplast mRNA leaders are highly A-U rich (70–84%) and show no significant primary sequence identity to one another. However, analysis of these sequences using energy minimization algorithms (Zuker, 1994) predicts highly folded structures. Folding of the naked RNA depends on kinetic barriers to achieving the thermodynamically most stable structure. Thus the leader may assume a multitude of local energy minima conformations on a path to the most thermodynamically stable structure (Konings and Gutell, 1995). The structure(s) acquired by RNA in vivo are dependent also on their interaction(s) with trans-acting proteins, which once bound, may regulate accessibility to sequences required for translation. While certain 5´ UTR binding proteins have been reported to protect leader regions from RNase digestion (Danon and Mayfield, 1991; Hauser et al., 1996) there are no published accounts examining the effect(s) of protein binding to C. reinhardtii leader sequences on the higher order structure of the RNA. That the leaders of at least some chloroplast mRNAs undergo conformational changes prior to translation is implicit in a model proposed by Rochaix (1996) for PS II mRNAs. An RNP complex formed between the 5´ ends of the UTRs and trans-acting proteins is proposed to target the mRNA to a membrane site, at which point the structured 5´ end of the leader is clipped off, leaving a shorter and unstructured mRNA poised for binding to the 30S subunit and translation initiation. As in the case of certain E. coli genes (Zhang and Deutscher, 1992), changes in the initiation codon can affect translation of particular chloroplast mRNAs (Chen et al., 1993,1995). Mutating the AUG initiation codon of the chloroplast petD mRNA encoding complex, to AUU subunit IV of the cytochrome or AUC results in a temperature sensitive nonphoto­ synthetic phenotype with a 50% reduction in mRNA

Chapter 12

Chloroplast Translation

accumulation and a 80–90% reduction in both synthesis and accumulation of subunit IV under permissive photosynthetic conditions (Chen et al., 1993). A dominant nuclear mutation (sim30-1d ) appears to increase the translation initiation rate of the petD mRNA containing the mutant AUU codon (Chen et al., 1997). Changing the initiation codon of the chloroplast petA gene from AUG to AUU, ACG, ACC, ACU or UUC had variable effects on synthesis of cytochrome f and photosynthetic competence. All mutants except the one carrying the UUC mutation, accumulated detectable levels of cytochrome f, but only the AUU codon mutant grew well at 24 °C. Introduction of in-frame UAA stop codon mutations immediately at, or immediately downstream of, the initiation codon prevented accumulation of cyto­ chrome f, suggesting that neither the second codon nor codons downstream ofthe second codon serve as the initiation codon in vivo. In addition, strains containing a stop codon inserted immediately upstream of the AUG codon accumulated wild-type levels of cytochrome f, suggesting that initiation upstream of the AUG codon does not contribute significantly to cytochrome f accumulation. The authors suggest from these data that in each initiation codon mutant strain that accumulates cytochrome f, the mutant codon is used as the initiation codon. Therefore, the petA AUG codon is not required to specify the site oftranslation initiation in chloroplasts, but the strength of the initiation codon-anticodon interaction may determine the rate of translation initiation.

V.Translational Regulation Involves Interactions between cis-Acting Sequences and trans-Acting Factors

A. Regulation of Individual Photosynthetic Proteins Interactions between cis-acting sequences in the 5´UTRs of chloroplast mRNAs and specific trans­ acting nuclear gene products appear to be essential for the expression of the several chloroplast genes so far examined (Fig. 1). This gene-for-gene relationship has been deduced from characterization of nuclear mutants that either block translation of particular chloroplast mRNAs (Table 1) or perturb their halflife (Gillham et al., 1994; Mayfield et al., 1995; Rochaix, 1996). While there are many mutations

205 affecting the expression of genes encoding photo­ synthetic proteins, nuclear mutants in which the synthesis of a specific ribosomal protein is affected have not been identified. Nuclear gene products are required for the stabilization ofthe atpA and atpB transcripts encoding the and subunits of the chloroplast ATP synthase, respectively, as well as the translation of the atpA transcript (Stern et al., 1991, Drapier et al., 1992). The nuclear mutant F54 shows a four-fold increase in atpA mRNA accumulation over wild type, yet it fails to synthesize the subunit. These data suggest that the expression of the atpA gene is blocked at the level of translation in mutant F54, and that the translational block is accompanied by an increased half-life of the transcript. In contrast to the impaired subunit, the F54 mutant also synthesis of the displays a stimulation in the synthesis the subunit, while maintaining wild-type levels of atpB transcript. Together these data point to the interlinked expression of the chloroplast-encoded ATP synthase subunits. The molecular basis for the increased translation of the subunit in F54 is postulated to result from competition between atpA and atpB 5´UTRs for regulatory proteins which are present in limiting conditions in the wild type and which have a higher affinity for the atpA transcript. Under these conditions, a larger proportion of the regulatory proteins would be available for the translation of the atpB transcripts in mutant F54, where atpA transcripts would not associate with polysomes (Drapier et al., 1992). Deletion analysis carried out by Mayfield et al. (1994) has suggested that the putative stem-loop structure in the psbA 5´UTR located upstream of the SD-like sequence may function to modulate expression of the D1 protein. Mutants lacking the loop sequences or altering pairing within the stem accumulate only ~20% of the normal levels of D1 protein, so the authors speculate that this region continues to function as a translational attenuatorbut that its activity can no longer be overcome by binding of trans-acting activator proteins. The nuclear mutant F35 that blocks D1 synthesis (Girard-Bascou et al., 1992) has been studied in detail by Yohn et al. (1996). They report that F35 reduces association of psbA mRNA with chloroplast ribosomes and decreases the half-life of this transcript. Analysis of psbA mRNA distribution on polysomes suggests that the F35 mutant is impaired in translation initiation. The reduction in thepsbA mRNA loading onto polysomes in the F35 mutant correlates with a decrease in the

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formation of a psbA RNP complex compared to wild type as monitored by gel mobility shift analysis. However, in contrast to the results of experiments in which psbA expression was studied in illuminated versus dark adapted cells where formation of a RNP complex correlates with levels of translation (Danon and Mayfield, 1991), the total block of psbA translation in F35 cells is not paralleled by a complete loss of the psbA RNP complex. The authors explain the pleiotropic effects of the F35 mutant in terms of the modest reduction in formation of psbA mRNA initiation complexes causing a larger decrease in translation initiation of mRNA encoding the D1 protein. Preliminary results suggest that a 55 kDa protein may be absent from the complex in extracts from the F35 mutant. Similarly, deletion analysis of the 362 nt 5´UTR of the chloroplast petD gene denned two domains (+153 to 302 and +312 to 330) required for synthesis of subunit IV (Sakamoto et al., 1994). Isolation of nuclear suppressors that restore translation of the petD mRNA (Sakamoto et al., 1994) again points to the involvement of nucleus-encoded trans-acting

proteins in translational regulation of chloroplast mRNAs. A light-dependent psbA RNP complex has been identified and characterized (Danon and Mayfield 1991). The protein complex interacts with the aforementioned 36 base stem-loop RNA structure within the 5´untranslated region (UTR) of the psbA mRNA immediately 5´ to the putative ribosome binding site (GGAG). Binding of the protein complex to the RNA was detected using RNA gel mobility shifts, while its constituents were identified by UV crosslinking and RNA affinity chromatography. A minimum complex consists of proteins of 47,75,60, 55 and 38 kDa and possibly two additional proteins of 30 and 33 kDa (Danon and Mayfield, 1991,1994a). The 47 kDa protein was further shown to resolve into four species by two dimensional gel electrophoresis. Direct contact of the protein complex with the RNA appears to be mediated by the 47 kDa protein(s) whereas the 60 kDa protein may interact with the RNA through contacts with the 47 kDa protein. A correlation was found between RNP complex formation and the level of translation of the psbA

Chapter 12

Chloroplast Translation

mRNA under mixotrophic (light) and heterotrophic (dark) growth conditions as measured by RNA gel mobility shifts (Danon and Mayfield, 1991). Illuminated cells, which rapidly synthesize D1 protein, contained high levels of psbA RNP, whereas dark-grown cells, which accumulate low levels of Dl , showed only minimal levels of RNP formation. Dark-grown cells of the y1 mutant, which lack photosynthetic membranes and do not translate psbA mRNA, were completely deficient in RNP complex formation. However, only minor differences in the amounts of the 47 and 60 kDa proteins present in extracts from light- and dark-grown wild-type cells were detected as monitored by UV crosslinking. From these data, the RNP complex is hypothesized to regulatepsbA mRNA translation either by targeting the message to the thylakoid membranes for translation (Rochaix, 1996) or by directly enhancing ribosome binding and translation initiation (Danon and Mayfield, 1991, Mayfield et al., 1994). The D2 protein encoded by the chloroplast psbD gene in Chlamydomonas is also an integral thylakoid membrane constituent of the PS II reaction center. It is synthesized on thylakoid membrane associated ribosomes and is cotranslationally inserted into the membrane (see Rochaix, 1996). Synthesis of the D2 protein is dependent upon at least three nuclear genes: NAC1 and AC115 whose products promote D2 translation (Kuchka et al., 1988), and NAC2. The NAC2 product appears to be essential for protecting psbD mRNA from degradation (Kuchka et al., 1989) and also affects the ability of a 47 kDa protein (presumably also nucleus-encoded) to bind to the 74 nt psbD 5´UTR sequence (Nickelsen et al., 1994). Mutations at the nuclear NAC1 and AC115 loci that specifically reduce synthesis of the D2 protein by affecting translation of psbD mRNA (Kuchka et al., 1988) can be suppressed by a dominant nuclear mutation in strain sup4b (Wu and Kuchka, 1995). While the molecular basis for this suppression is not known, the hypothesis has been put forth that the sup4b mutation may bypass the requirement for NAC1 and AC115 gene products in psbD gene expression (Wu and Kuchka, 1995). Two recessive nuclear mutants (F34, F64) marking the TBC1 and TBC2 loci, respectively and a chloroplast mutant (FUD34) acting through the 5´UTR specifically block translation of psbC mRNA encoding PS II subunit P6 (Rochaix et al., 1989; Zerges and Rochaix, 1994, Zerges et al., 1997). Chloroplast transformants carrying a chimeric

207 construct in which the psbC 5´UTR is fused to the aadA reporter gene require the presence of both the TBC1 and TBC2 wild-type gene products for expression (Zerges and Rochaix, 1994). This confirms the specificity of cis-acting sequences in the psbC 5' UTR for the two nucleus-encoded, trans-acting proteins. Subsequent deletion analysis in which an inverted repeat sequence specifying a putative stemloop in the middle of the psbC 5´UTR was removed showed that deletion of this binding site for the TBC1 gene product reduced expression of the aadA reporter considerably (Zerges et al., 1997). The interaction of the wild-type TBC1 gene product with the stem-loop region in thepsbC 5´UTR is supported by the finding of a cis-acting chloroplast suppressor of the F34 mutation (psbC-F34suI) which contains a base change in the psbC 5´UTR (Rochaix et al., 1989). This suppressor mutation presumably diminishes the stability of the stem, partially restoring translation of the psbC mRNA. The nonphotosynthetic chloroplast mutant, FUD34, also alters the stability of the stem of a putative stem loop structure in the 5´UTR of the psbC mRNA (Rochaix et al., 1989). Insertions of two adjacent T residues and deletion of a C residue five nucleotides away are thought to increase the thermodynamic stability of the stem structure. Thus, the sequence alterations in F34suI and FUD34 affect two complementary sequences within the stem structure and have opposite effects on the trans­ latability of psbC mRNA which correlate with their ability to destabilize and over-stabilize the stem loop in the 5´UTR of psbC, respectively. A spontaneous partial phenotypic revertant of FUD34, tbc3-rb1, has also been isolated. This dominant nuclear mutation (tbc3-rb1) can restore expression ofthe aadA reporter gene from the 5´UTR bearing the psbC -FUD34 mutation or the deletion of the entire stem-loop (Zerges et al., 1997). The ability of tbc3-rb1 to suppress the deletion of the stem loop demonstrates that 5´UTR sequences outside of this region are sufficient for restoration of translation. That tbc3-rb1 also suppresses tbc1-F34 shows that the mutation alleviates the requirement for the functional interaction between the TBC1 gene product and the putative stem-loop region. From these data, Zerges and Rochaix (1997) propose two alternative models. A TBC1 -dependent protein interacts with the putative stem-loop to inhibit a TBC3-dependent translational repressor from acting on a site outside this region. In the absence of either

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the stem-loop or wild-type TBC1 function, the TBC3­ dependent repressor would block translation of the psbC mRNA. The tbc3-rb1 mutation acting as a dominant negative mutation would restore psbC translation. Alternatively, the TBC1 and TBC3 gene products activate psbC mRNA translation by binding to the stem loop. In the absence of TBC1 or the stemloop, wild-type TBC3 would be insufficient to promote translation. The dominant tbc3-rb1 mutation might alter the TBC3 gene product in such a way that it is sufficient to promote psbC mRNA translation in the absence of TBC1 or the stem-loop. Clearly, the stem-loop region of the psbC mRNA and the TBC1 and TBC3 gene products interact functionally to control translation of the psbC mRNA. This conclusion is tempered by the observation that in three of four strains carrying the tbc3-rb1 mutation analyzed, synthesis but not accumulation of Rubisco large subunit was apparently reduced relative to the wild type (Zerges et al., 1997). This may indicate that, in contrast to all other characterized mutations affecting chloroplast or mitochondrial translation, TBC3 function may not be gene specific, but required for the translation of other mRNAs as well. Molecular analysis of trans-acting proteins capable of interacting with chloroplast encoded 5´UTRs revealed that extracts of wild-type, F34 and F64 cells contained proteins of 95, 65, 40 and 33 kDa which crosslink to the 5´UTRs of psbC, psbA and psbD mRNAs, although the 33 kDa protein was not detected in extracts from purified chloroplasts (Zerges and Rochaix, 1994). As these proteins are present in all extracts, there is no reason to believe that they are related to the products of the wild-type TBC1 and TBC2 genes. However, a protein of 46 kDa (RBP46) specific to the extract from the F64 mutant was crosslinked to the psbC 5´UTR, suggesting that this protein may be responsible for the effect of the F64 mutation on psbC translation (Zerges and Rochaix, 1994).] RBP46 may well have other functions as this protein also crosslinks to the leaders of psbA and psbD (Zerges and Rochaix, 1994). Binding of RBP46 was inhibited by incubation of extract with poly(A) or poly(U) showing that the protein either has an affinity for AU-rich sequences, or has little sequence specificity. A mutation in the nuclear TAB1 gene, defined by strain tab1-F15, specifically interferes with the synthesis of the two reaction center subunits of Photosystem I (PS I), PsaA and PsaB encoded by the chloroplast psaA and psaB genes respectively

(Stampacchia et al., 1997). The tab1-F15 mutation was shown to affect either the preinitiation or initiation of psaB mRNA translation, but not that of psaA by examining the expression of aadA reporter constructs driven by the promoters and 5´UTRs of psaB and psaA. A photoautotrophic revertant of the tab1-F15 mutant, suF15, was isolated and shown to contain a G to A transition adjacent to a putative SD sequence 17 nucleotides upstream of the AUG translation initiation site of psaB. The loss of PsaA synthesis in the tab1-F15 mutant is a consequence of the absence of PsaB synthesis and not the presence of the tab1­ F15 mutation as demonstrated by the fact that the suF15 mutation in the psaB 5´UTR is capable of restoring the synthesis of both proteins. The authors speculate that TAB1 function is required for opening the stem structure to present an unstructured SD sequence analogous to the mom/com paradigm in bacteriophage Mu (Wulczyn et al., 1989).

B. Coordinate Regulation of Photosynthetic versus Ribosomal Proteins A syndrome ofphotosynthetic defects has been found in all chloroplast ribosome deficient mutants so far characterized (Harris, 1989; Harris et al., 1994). Experiments with the nuclear double mutant ac20 cr1 which is severely deficient in chloroplast ribosomes showed that synthesis and accumulation of chloroplast encoded photosynthetic proteins was greatly reduced while chloroplast-encoded ribosomal proteins (r-proteins) continue to be synthesized and accumulate (Liu et al., 1989b). A spectinomycin resistant mutant (spr-u-1-27-3) which carries a mutation in the chloroplast 16S rRNA, exhibits the same symptoms of reduced synthesis of photo­ synthetic proteins when grown in the presence of spectinomycin, but synthesizes and accumulates normal levels of r-proteins (Liu et al., 1989b). These results suggest that when the capacity for chloroplast protein synthesis is reduced, photosynthetic and rprotein mRNAs are distinguished in a class specific manner and a hierarchy of translation is established which favors expression of r-protein mRNAs (Gillham et al., 1994). Analysis of heparin-actigel purified whole-cell extracts fromwild-type cells grown phototrophically, mixotrophically or heterotrophically and extracts of mutants deficient in chloroplast protein synthesis demonstrated that proteins of 81, 62, 56, 47, 38, 36 and 15 kDa UV crosslink to the leaders of several

Chapter 12 Chloroplast Translation chloroplast, but not nucleus-encoded 5´UTRs (Hauser et al., 1996; Fig. 1). In extracts from mutant cells with reduced chloroplast protein synthesis (ac20 cr1 and spr-u-1-27-3) binding of the 36 kDa protein present in wild-type cells is undetectable (Hauser et al., 1996). This suggests that the 36 kDa protein may be required for the translation of photosynthetic, but not r-protein mRNAs. It is also possible that the 36 kDa protein is absent because it is synthesized on chloroplast ribosomes. The observation that six of the RNA binding proteins bind to five distinct chloroplast leaders under a variety of environmental conditions in conjunction with data cited above (Danon and Mayfield, 1991, 1994b; Zerges and Rochaix, 1994) suggest that at least some of these proteins are general leader binding proteins. Of particular interest is the 47 kDa protein family, whose members migrate as doublets when they are crosslinked to various leader sequences by UV irradiation (Hauser et al., 1996). The RBP46 protein identified in extracts of the F64 strain, in which psbC translation is inhibited (Zerges and Rochaix, 1994), and the 47 kDa protein that is involved in the lightactivated translation of psbA mRNA (Danon and Mayfield, 1991, 1994b) may either be genetically unrelated, or members of a related family of proteins. Together, these results suggest that members of the of 47 kDa RNA binding protein family may be either class-, message- or environment-specific, while others may bind ubiquitously to all chloroplast leaders. Proteins that UV crosslink directly to the 5´UTRs of chloroplast mRNAs may represent only a subset of the proteins constituting these RNP complexes. Other proteins which interact in the complex via protein-protein interactions will not be detected using this assay. Thus the aforementioned six to eight trans-acting proteins binding to the 5´UTRs of chloroplast mRNAs may be an underestimate of the number of the participants in the translational regulatory complex. Therefore, failure to observe differences in the pattern of proteins that U V crosslink to the 5´UTRs of chloroplast mRNAs in extracts from cells exposed to different environmental conditions may indicate that the key regulatory proteins do not bind directly to the mRNA, but rather to other proteins in the translation complex. Alternatively, since all of these extracts were prepared and analyzed under reducing conditions, differences in their state of reduction in vivo that might be related to their binding in response to environmental stimuli could be obscured. The 60 kDa protein

209 reported to be part of the complex on the psbA leader in C. reinhardtii (Danon and Mayfield, 1991) appears to be a regulatory protein of this type. Proteinprotein crosslinking experiments, use of the yeast two-hybrid system and/or isolation ofan intact 5´UTR binding complex will be necessary to determine whether additional translational regulatory proteins exist that are not in direct contact with the mRNA.

VI. Ribosomes, Membranes and Tethers A model has emerged, principally from studies of COX3 message translation in the yeast mitochondrion, which assumes that an inner-membrane-bound translational activator complex recognizes both the mRNA 5´UTR and the mitochondrial ribosome (Fox, 1996a,b). The possible application of this model to translational control in chloroplasts has already been recognized (Gillham et al., 1994; Rochaix, 1996). The model derives its appeal for chloroplasts from the fact that the COX3 protein, like many other proteins encoded in chloroplasts and mitochondria, is very hydrophobic. Coupling of the translation complex to the inner mitochondrial membrane would ensure that the nascent, hydrophobic polypeptide is synthesized close to its insertion site in the inner membrane. Translation of COX3 mRNA requires three proteins encoded by the nuclear genes PET54, PET122 and PET494 (Fox, 1996a,b). Several lines of evidence suggest that these three proteins form a complex. The PET54 product (Pet54p) is found in equal amounts as a soluble polypeptide and a peripheral membrane protein whereas the other two polypeptides are probably integral membrane proteins. Since the Pet54p plays a role in splicing of a mitochondrial intron in COX1 pre-mRNA in some, but not all, yeast strains, Fox speculates that the soluble form of the protein is responsible for intron splicing while the membrane-bound form is part of the COX3 mRNA translation complex. Other evidence suggests that Pet54p may participate in COX3 mRNA recruitment to the inner membrane while the Pet122p makes contact with the small subunit of the mitochondrial ribosome. Loss of function caused by carboxy­ terminal truncation of the Pet 122p through the use of deletion or nonsense alleles can be suppressed by mutations in the genes encoding three different proteins of the small subunit of the mitochondrial ribosome. All three proteins are required for

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mitochondrial protein synthesis in yeast, but none displays sequence similarity with ribosomal proteins from other organisms. Fox (1996a,b) believes that his model is consistent with data available for translation of other mitochondrial genes in yeast although they are less complete, and he further speculates that these proteins may play a role in making topological distinctions in the sites of

synthesis of each mitochondrion-encoded protein. Such specific inferences concerning the role of translational activator proteins in membrane binding and the recruitment ofmRNA and ribosomes are not yet possible to make in the case of the chloroplast of Chlamydomonas (Fig. 2). However, there is much evidence that many chloroplast encoded proteins are synthesized on thylakoid membrane-bound poly­

Chapter 12 Chloroplast Translation somes (see Gillham, 1994 and Harris et al., 1994 for reviews). During greening of etiolated barley, membrane-bound chloroplast ribosomes engaged in translation of the D1 protein pause at specific sites during the translation process (Kim et al., 1991). Six different pause sites have been identified which correspond roughly to the insertion of the five of D1 into the thylakoid hydrophobic membrane. There seems to be a correlation between pausing and the presence of the hydrophobic amino acids of these helices in the ribosome tunnel. Since the ribosome tunnel is itself somewhat hydrophobic, pausing may involve the interaction of the newly synthesized hydrophobic amino acids and those present in the ribosome tunnel. In addition, D1 translation intermediates at the third and fourth pauses may bind phaeophytin while those at the fifth pause may bind chlorophyll. Interestingly, nonphoto­ synthetic D1 mutants of C. reinhardtii with amino to Gln, Glu, His, and acid substitutions from Asp synthesize two forms of D1 one of which is slightly larger (33–34 kDa) and the other smaller (24–25 kDa) than the mature protein (Lardans et al., 1997). While the 33–34 kDa protein does not correspond to the precursor and may result from structural modification ofthe mature 32 kDa protein, the 24–25 kDa form is likely to be a translation intermediate. This polypeptide may result from abortion of translation in the C-terminal portion of IV and V during the loop connecting ribosome pausing at the next to last or last pause site. In summary, much evidence exists to support the idea that many chloroplast mRNAs program translation on thylakoid-bound polysomes and that some of the protein products are very hydrophobic. This would be consistent with the scheme envisioned by Fox (1996a,b) for yeast mitochondrial mRNAs in that these chloroplast mRNAs might bind close to the site on the membrane where their protein product is to be inserted. However, there is still no evidence that the nucleus-encoded proteins that bind to the 5´UTRs of chloroplast mRNAs either function to localize these mRNAs to the thylakoid membranes or that they make contact with the chloroplast ribosome. Recently, a nucleus-encoded homolog of E. coli ribosomal protein S1 which is targeted to the chloroplast has been identified (Franzetti et al., 1992; Harris et al., 1994). This protein could play a role in mRNA recognition as it does in E. coli (see above Section I).

211 VIl.Translational Regulation of Complex Assembly

A. Chloroplast Thylakoid Membranes The numerous nuclear mutants of Chlamydomonas deficient in photosynthesis or chloroplast protein synthesis also afford tools for the dissection of the mechanisms regulating assembly of these multimeric protein complexes (Fig. 1, Table 1). Nuclear mutants identified to date that affect the synthesis of chloroplast encoded photosynthetic proteins appear to function in a gene specific manner (Gillham et al., 1994; Rochaix 1995, 1996). A possible exception is the nuclear tbc3 mutant recently described by Zerges et al., (1997) that appears to affect expression of both psbC and rbcL mRNAs. Translational regulation of the synthesis of chloroplast-encoded proteins appears to mediate two distinct phenomena: (1) the coordinated assembly of photosynthetic and translational complexes, and (2) a response to specific environmental signals, such as light (reviews: Gillham et al., 1994; Rochaix, 1996). The precise role(s) these nucleus-encoded proteins play in regulating chloroplast mRNA translation is still being resolved, however intriguing models with parallels to the yeast mitochondrial translation system have been invoked (Gillham et al, 1994; Rochaix, 1996). The Photosystem II core complex is composed of proteins D1, D2, P5 and P6 encoded by psbA,psbD, psbB and psbC mRNAs. Previous work has revealed the concerted expression of the psbA,psbD and psbB genes (reviewed by Rochaix, 1996). During the assembly of the PS II complex the D1, D2 and P5 proteins form an intermediate complex prior to association with P6. Primer extension studies suggest the possibility that the mRNAs encoding these three hydrophobic proteins are synthesized as precursors and cleaved within their 5´UTRs to generate mature mRNAs. Analogous to the yeast mitochondrial system, discussed above, a protein complex by binding to the 5´ end of the chloroplast leader might serve to target and/or dock the mRNA to the thyla­ koid membrane (Fig. 2). Cleavage of the leader is hypothesized to induce a conformational change allowing ribosome binding and initiation of translation (Rochaix, 1996). The psbC product P6 is integrated in a subsequent step to form the PS II core complex. Assembly of the C. reinhardtii cytochrome complex appears to be regulated by two distinct

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mechanisms: 1) proteolytic degradation of unassem­ bled subunits and, 2) a process in which the pro­ duction of cytochrome f depends on its immediate interaction with cytochrome and subunit IV (Kuras and Wollman, 1994). Using insertional mutagenesis, mutants affected in the half-life of cytochrome f and have recently been isolated (Gumpel et al., 1995). Whether the tagged genes are required for constitutive expression of the subunits, or if they coordinate the synthesis of the subunits and thus assembly of the complex is unknown. Rubisco holoenzyme plays a pivotal role in photosynthesis because it catalyzes the carboxylation and oxygenation of RuBP (reviewed by Spreitzer, 1993; Chapter 27, Spreitzer). While most Rubisco­ deficient mutants result from alterations in the chloroplast encoded rbcL gene encoding LSU, which perturb its enzymatic functions, nuclear mutants not linked to rbcS encoding SSU, have been isolated which affect Rubisco at transcriptional, posttrans­ criptional and cotranslational steps (see Hong and Spreitzer, 1994). In addition, data from transgenic tobacco plants expressing antisense rbcS mRNA suggest that SSU protein abundance specifically contributes to the regulation of LSU protein accumulation at the level ofrbcL translation initiation (Rodermel et al., 1996). Sequences near the 5´ end of the rbcL leader were found to destabilize chimeric mRNAs in dark grown cells following illumination (Salvador et al., 1993). This effect is eliminated by the addition of sequences from the adjacent coding region of the rbcL gene, and these are probably responsible for stabilizing endogenous rbcL transcripts (Salvador et al., 1993).

B. Chloroplast Ribosomes The chloroplast translational apparatus is composed of five rRNA species and approximately 64 r-proteins (Harris et al., 1994). While all of the rRNAs are chloroplast encoded, only a third of the r-proteins are specified by the chloroplast genome with the remainder presumably being nuclear gene products (Gillham, 1994, Harris et al., 1994). To assemble such a molecular machine efficiently, the cell must provide equimolar amounts of each of the com­ ponents. While the mechanisms by which this is achieved in E. coli and the cytoplasm of eukaryotic cells have been extensively studied (see reviews by Nomura et al., 1984; Amaldi et al., 1989; Woolford and Warner, 1991), they remain to be determined in

chloroplasts. However, the identification of cis-acting sequences and trans-acting factors responsible for modulating the differential translation of chloroplastencoded photosynthetic and ribosomal protein subunits has begun (Section V).

VIII. How are the Regulatory Proteins Regulated? Except in the case of the psbA mRNA of C. reinhardtii, little is currently known about regulation of the nucleus-encoded trans-acting factors that govern expression of chloroplast mRNAs (Fig. 1). As discussed earlier, the 5´UTR of the psbA mRNA contains a 36 nt inverted repeat sequence thermo­ dynamically capable of forming a stem-loop which is crosslinked by UV to a complex of four closelyrelated 47-kDa proteins (Danon and Mayfield, 1991). A 60-kDa protein found to be associated with the isolated complex is not crosslinked to the 5´UTR by UV (Danon and Mayfield, 1994a). Although illuminated wild-type cells rapidly synthesize the D1 protein while dark-grown cells do not, there is less than a two-fold difference in the abundance of the 47-kDa proteins in light-and dark-grown cells. A serine/threonine phosphotransferase associated with the mRNA binding complex (Danon and Mayfield, 1994a) has been identified that utilizes the of ADP to phosphorylate the 60-kDa protein in vitro. Although phosphorylation of this protein inhibits the mRNA binding activity of the complex, the high levels ofADP required in vitro are only attained in vivo in the chloroplasts of plants grown in the dark. Phosphorylation of the 60 kDa protein in vivo appears to be sufficient to overcome any effect of reduction of the 47 kDa protein in these in vitro assays. Thus, the activator complex would bind well to the psbA 5´UTR in light- but not darkgrown cells. Synthesis of the 47-kDa protein appears to require the presence of chlorophyll and/or a fully developed chloroplast, since the 47-kDa protein is not detected in dark-grown cells of the y1 mutant which, unlike wild type, does not synthesize chlorophyll, chloroplast membranes or D1 (Danon and Mayfield, 1991). Light-modulation of psbA mRNA expression via redox potential has been proposed as a translational regulatory mechanism (Danon and Mayfield, 1994b). Once again the evidence is based mostly on in vitro experiments and shows that oxidation of the

Chapter 12 Chloroplast Translation regulatory protein complex with dithionitrobenzoic acid (DTNB) blocks binding ofthe complex topsbA­ mRNA. This effect is reversed by incubation of the complex with reducing agents like dithiothreitol (DTT). Furthermore, an even greater restoration of RNA-binding activity of the complex was observed when reduced thioredoxin was added to the DTNBoxidized complex suggesting that thioredoxin was the agent responsible for reducing the protein complex in vivo. In chloroplasts, reduction of thioredoxin by ferredoxin is stimulated by photosynthetic electron transfer in response to light (Buchanan et al., 1994; Chapter 26, Jacquot et al.). Reduced ferredoxin activates three of the four major regulatory enzymes of the carbon cycle. This led Danon and Mayfield (1994b) to postulate that reduced ferredoxin can also activate the protein complex that binds to the 5´UTR of psbA mRNA and stimulates translation of D1. The model was tested in vivo using a Photosystem I-deficient mutant (ac-u-g-2-3) which can no longer mediate reduction of thioredoxin by ferredoxin (Danon and Mayfield, 1994b). The mutant, as predicted, contained much less of the psbA mRNA protein binding complex and much less D1 than wild type although psbA mRNA accumulation was similar in the mutant and wild type. Synthesis of three other chloroplast-encoded Photosystem II proteins (D2, CP43 and CP47) was also reduced in ac-u-g-2-3. Rochaix (1996), states that these results are at variance with earlier pulse-labeling studies with other Photosystem I-deficient mutants of C. reinhardtii which did not reveal a diminution in the synthesis of the core Photosystem II polypeptides including D1. Obviously this discrepancy will have to be resolved before the redox model for D1 regulation can be accepted as explaining light-regulation of psbA mRNA translation in vivo. Redox regulation, possibly via the thioredoxin-ferredoxin pathway, has also been implicated recently in splicing of group I introns from the psbA pre-mRNA in C. reinhardtii (Desh­ pande et al., 1997) so the mechanisms by which redox regulation of chloroplast gene expression is exerted are potentially far-reaching. In addition, experiments with Dunaliella tertio­ lecta, a relative of Chlamydomonas, reveal that the redox state of plastoquinone may modulate tran­ scription rates of the nuclear Cab genes encoding the chlorophyll a/b proteins via a phosphorylation cascade (Escoubas et al., 1995). The reduction state of the plastoquinone (PQ) pool can be manipulated using the electron transport inhibitors DCMU [3-

213 (3,4 dichlorophenyl)-1,1 -dimethyl urea) and DBMIB [2,5-dibromo-3-methyl-6-isopropyl-p-benzoquinone] to block the reduction and oxidation of the PQ pool respectively. DCMU induced a doubling of Cab mRNA abundance mimicking the effect of shifting cells from high to low light intensity while DBMIB induced a 75% decrease in Cab mRNA, mimicking the effect of a shift to higher irradiance. Gel shift assays identified a protein-DNA complex in extracts from cells grown at high, but not low light. Phosphatase inhibitors blocked derepression of Cab gene expression in low light. These results have been interpreted in terms of a model in which reduced PQ is sensed by a chloroplast kinase which phos­ phorylates a chloroplast phosphoprotein. This phosphoprotein then directly or indirectly activates a cytosolic kinase which phosphorylates a repressor which can bind in the Cab gene promoter region in its phosphorylated state and block transcription. Whether similar models will prove applicable to nucleus-encoded regulators of chloroplast mRNA translation remains to be seen.

IX. Is there Hierarchical Control of Chloroplast mRNA Translation? Specific nuclear gene products are clearly essential for expression of individual chloroplast genes in C. reinhardtii at the transcriptional, posttrans­ criptional or translational levels as discussed in this and earlier reviews (Gillham et al., 1994; Mayfield et al., 1995;Rochaix, 1995,1996; Cohen and Mayfield, 1997). The notion that translation initiation is modulated by a complex of proteins bound to the 5´UTRs of chloroplast mRNAs is supported by both genetic and biochemical evidence. Such translational activation complexes are likely to contain both genespecific proteins discussed earlier and ubiquitous proteins required for the translation of all chloroplast mRNAs (Hauser et al., 1996). Binding of specific proteins to the translational complexes may also effect concordant regulation of classes of chloroplast genes with related functions or in response to particular environmental stimuli (Gillham et al., 1994). Binding of gene-specific, class-specific or environment-specific proteins to the complex could exert either a negative or positive effect on translation initiation. This might occur directly by altering the ability of the mRNA to associate with the small subunit of the chloroplast ribosome to form an

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initiation complex. Alternatively, it could occur indirectly, by preventing binding of certain of the ubiquitous proteins essential for initiation to occur. In addition, these proteins may play a role in targeting the messages to specific sites within the organelle to facilitate coordinated translation of members of multisubunit complexes (Fox 1996b, Rochaix, 1996, Gillham et al., 1994). Clear evidence exists for classspecific translational regulation of ribosomal protein genes versus photosynthetic protein genes under conditions of reduced chloroplast protein synthesis in the C. reinhardtii chloroplast (Gillham et al., 1994; Liu et al., 1989b). The precise mechanism(s) regulating the entry of a mRNA into a translationally competent state in a gene, class or environment specific manner remains to be determined.

Acknowledgments This research was supported by NIH Grant GM­ 19427 to JEB and NWG. We thank D. Stern and J.-D. Rochaix for providing unpublished results.

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215 Kim HY, Klausner RD and Rouault TA (1995) Translational represser activity is equivalent and is quantitatively predicted by in vitro RNA binding for two iron-responsive elementbinding proteins, IRP1 and IRP2.J Biol. Chem 10:4983–4986 Kindle K (1987) Expression of a gene for a light-harvesting chlorophyll a/b-binding protein in Chlamydomonas reinhardtii: Effect of light and acetate. Plant Mol Biol 9: 547–563 Kozak M (1983) Comparison of initiation of protein synthesis in procaryotes, eucaryotes and organelles. Microbiol Rev 47: 1– 45 Kuchka M, Mayfield SP and Rochaix J-D (1988) Nuclear mutations specifically affect the synthesis and/or degradation of the chloroplast-encoded D2 polypeptide of Photosystem II in Chlamydomonas reinhardtii. EMBO J 7: 319–324 Kuchka M, Goldschmidt-Clermont M, van Dillewijn J and Rochaix J-D (1989) Mutation at the Chlamydomonas nuclear NAC2 locus specifically affects stability of the chloroplast psbD transcript encoding polypeptide D2 of PS II. Cell 58: 869–876 Kudla J, Hayes R and Gruissem W (1996) Polyadenylation accelerates degradation of chloroplast mRNA. EMBO J 15: 7137–7146 Kuras R, Wollman F-A (1994) The assembly of cytochrome b6/ f complexes: An approach using genetic transformation of the green alga Chlamydomonas reinhardtii. EMBO J 13: 1019– 1027 Lardans A, Gillham NW and Boynton JE (1997) Site-directed mutations at residue 251 of the Photosystem II Dl protein of Chlamydomonas that result in a nonphotosynthetic phenotype and impair D1 synthesis and accumulation. J Biol Chem 272: 210–216 Leibold EA and Munro HN (1988) Cytoplasmic protein binds in vitro to a highly conserved sequence in the 5´ untranslated region of ferritin heavy- and light-subunit mRNAs. Proc Natl AcadSci USA 85: 2171–2175 Lisitsky I, Klaff P and Schuster G (1996) Addition ofdestabilizing poly(A)-rich sequences to endonuclease cleavage sites during the degradation of chloroplast mRNA. Proc Natl Acad Sci USA 93: 13398–13403 Liu X-Q, Gillham NW and Boynton JE (1989a) Chloroplast ribosomal protein gene rps12 of Chlamydomonas reinhardtii. J Biol Chem 264: 16100-16108 Liu X-Q, Hosler JP, Boynton JE and Gillham NW (1989b) mRNAs for two ribosomal proteins are preferentially translated in the chloroplast of Chlamydomonas reinhardtii under conditions of reduced protein synthesis. Plant Mol Biol 12: 385–394 Lovett P (1994) Nascent peptide regulation of translation. J Bact 176: 6415–6417 McCarthy JE and Brimacombc R (1994) Prokaryotic translation: The interactive pathway leading to initiation. Trends Genet 10: 402–407 McCarthy JE and Gualerzi C (1990) Translational control of prokaryotic gene expression. Trends Genet 6: 78–85 Mache R (1990) Chloroplast ribosomal proteins and their genes. Plant Sci 72: 1–2 Maliga P (1993) Towards plastid transformation in flowering plants. Trends Biotech 11: 101–107 Malnoë P, Mayfield SP and Rochaix J-D (1988) Comparative analysis of the biogenesis of Photosystem 11 in the wildtype and y-1 mutant of Chlamydomonas reinhardtii. J Cell Biol

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106:609–616 Mathews MB, Sonenberg N and Hershey JWB (1996) Origins and targets of translational control. In: Hershey JWB, Mathews MB and Sonenberg N (eds), Translational Control, pp 1–29. Cold Spring Harbor Laboratory Press, Plainview Mayfield SP, Cohen A, Danon A and Yohn CB (1994) Translation of the psbA mRNA of Chlamydomonas reinhardtii requires a structured RNA element contained within the 5´ untranslated region. J Cell Biol 127: 1537–1545 Mayfield S, Yohn CB, Cohen A and Danon A (1995) Regulation of chloroplast gene expression. Annu Rev Plant Physiol Plant Mol Biol 46: 147–166 Meerovitch K, Pelletier J and Sonenberg N (1989) A cellular protein that binds to the 5´-noncoding region of poliovirus RNA: Implications for internal translation initiation. Genes Dev3: 1026–1034 Morton BR (1996) Selection on the codon bias of Chlamydomonas reinhardtii chloroplast genes and the plant psbA gene. J Mol Evol 43: 28–31 Myers AM, Harris EH, Gillham NW and Boynton JE (1984) Mutations in a nuclear gene of Chlamydomonas cause the loss of two chloroplast ribosomal proteins, one synthesized in the chloroplast and the other in the cytoplasm. Curr Genet 8: 369– 378 Nickelsen J, van Dillewijn J, Rahire M and Rochaix J-D (1994) Determinants for stability of the chloroplast psbD RNA are located within its short leader region in Chlamydomonas reinhardtii. EM BO J 13: 3182–3191 Nomura M, Gourse R and Baughman G (1984) Regulation of the synthesis of ribosomes and ribosomal components. Ann Rev Biochem 53: 75–117 Prokipcak RD, Herrick DJ and Ross J (1994) Purification and properties of a protein that binds to the C-terminal coding region of human c-myc mRNA. J Biol Chem 269: 9261–9269 Proudfoot N (1996) Ending the message is not so simple. Cell 87: 779–781 Proweller A and Butler JS (1996) Ribosomal association of poly(A)-binding protein in poly(A)-deficient Saccharomyces cerevisiae. J Biol Chem 271: 10859–10865 Purton S and Rochaix J-D (1994) Complementation of a Chlamydomonas reinhardtii mutant using a genomic cosmid library. Plant Mol Biol 24: 533–537 Putz U, Skehel P and Kuhl D (1996) A tri-hybrid system for the analysis and detection of RNA-protein interactions. Nucl Acids Res 24: 4838–4840 Reinbothe S, Reinbothe C, Heintzen C, Seidenbecher C and Parthier B (1993) A methyl jasmonate-induced shift in the length of the 5´ untranslated region impairs translation of the plastid rbcL transcript in barley. EMBO J 12: 1505–1512 Reith M (1995) Molecular biology of rhodophyte and chromophyte plastids. Annu Rev Plant Physiol. Plant Mol. Biol. 46: 549–575 Rochaix J-D (1995) Chlamydomonas reinhardtii as the photosynthetic yeast. Annu Rev Genet 29: 209–230 Rochaix J-D (1996) Post-transcriptional regulation ofchloroplast gene expression in Chlamydomonas reinhardtii. Plant Mol Biol 32: 327–341 Rochaix J-D, Kuchka M, Mayfield S, Schirmer-Rahire M, GirardBascou J and Bennoun P (1989) Nuclear and chloroplast mutations affect the synthesis or stability of the choroplast psbC gene product in Chlamydomonas reinhardtii. EMBO J 8:

1013–1021 Rodermel S, Haley J, Jiang, C-Z, Tsai, C-H and Bogorad L (1996) A mechanism for intergenomic integration: Abundance of ribulose bisphosphate carboxylase small-subunit protein influences the translation of the large-subunit mRNA. Proc Natl Acad Sci USA 93: 3881–3885 Rouault TA, Hentze MW, Haile, DJ and Harford JB (1989) The iron-responsive element binding protein: A method for the affinity purification of a regulatory RNA-binding protein. Proc Natl Acad Sci USA 86: 5768–5772 Sakamoto W, Kindle KL and Stern DB (1993) In vivo analysis of Chlamydomonas chloroplast petD gene expression using stable transformation of beta-glucuronidase translational fusions. Proc Natl Acad Sci USA 90: 497–501 Sakamoto W, Chen X, Kindle KL and Stern DB (1994) Function of the Chlamydomonas reinhardtii petD 5´ untranslated region in regulating the accumulation of subunit IVof the cytochrome complex. Plant J 6: 503–512 Salvador ML, Klein U and Bogorad L (1993) 5´ sequences are important positive and negative determinants of the longevity of Chlamydomonas chloroplast gene transcripts. Proc Natl Acad Sci USA 90: 1556–1560 Schindler U and Cashmore AR (1990) Photoregulated gene expression may involve ubiquitous DNA binding proteins. EMBO J 9: 3415–3427 Schmitz J, Prüfer D, Rohde W and Tacke E (1996) Non-canonical translation mechanisms in plants: Efficient in vitro and in planta initiation at AUU codons of the tobacco mosaic virus enhancer sequence. Nucl Acids Res 24: 257–263 SenGupta DJ, Zhang B, Kraemer B, Pochart P, Fields S and Wickens M (1996) A three-hybrid system to detect RNAprotein interactions in vivo. Proc Natl Acad Sci USA 93: 8496–8501 Shapira M, Lers A, Heifetz PB, Irihimovitz V, Osmond CB, Gillham NW and Boynton JE (1997) Differential regulation of chloroplast gene expression in Chlamydomonas reinhardtii during photoacclimation: Light stress transiently suppresses synthesis of the Rubisco LSU protein while enhancing synthesis of the PS II D1 protein. Plant Mol Biol 33: 1001–1011. Sheen J (1990) Metabolic repression of transcription in higher plants. Plant Cell 2: 1027–1038 Silk GW and Wu M (1993) Posttranscriptional accumulation of chloroplast tufA (elongation factor gene) m R N A during chloroplast development in Chlamydomonas reinhardtii. Plant Mol Biol 23: 87–96 Spreitzer RJ (1993) Genetic dissection of Rubisco structure and function. Annu Rev Plant Physiol Plant Mol Biol 44: 411–434 StampacchiaO, Girard-Bascou J,ZanascoJL, Zerges W, Bennoun P and Rochaix JD (1997) A nuclear-encoded function essential for the translation of the chloroplast psaB mRNA in Chlamydomonas. Plant Cell 9:773–782 Standait N and Jackson RJ (1994) Regulation of translation by specific protein/mRNA interactions. Biochimie 76: 867–879 Staub JM and Maliga P (1994) Translation of psbA mRNA is regulated by light via the 5´-untranslated region in tobacco plastids. Plant J 6: 547–553 Stern D, Radwanski ER, Kindle KL (1991) A 3´ stem/loop structure of the Chlamydomonas chloroplast atpB gene regulates mRNA accumulation in vivo. The Plant Cell 3: 285–297 Stevens DR, Rochaix JD and Purton S (1996) The bacterial phleomycin resistance gene ble as a dominant selectable marker

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217 Cold Spring Harbor Press, Cold Spring Harbor Wu HY and Kuchka MR (1995) A nuclear suppressor overcomes defects in the synthesis of the chloroplast psbD gene product caused by mutations in two distinct nuclear genes of Chlamydomonas. Curr Genet 27: 263–269 Wulczyn FG, Bolker M and Kahman R (1989) Translation of the bacteriophage MU mom gene is positively regulated by the phage com gene product. Cell 57:1201–1210 Yamamoto Y, Tsuji H and Ogokata J (1995) 5´-Leader of a Photosystem I gene in Nicotiana sylvestris, psaDb contains a translational enhancer. J Biol Chem 270: 12466–12470 Yohn CB, Cohen A, Danon A and Mayfield SP (1996) Altered mRNA binding activity and decreased translational initiation in a nuclear mutant lacking translation of the chloroplast psbA mRNA. Mol Cell Biol 16: 3560–3566 Zerges W and Rochaix JD (1994) The 5´ leader of a chloroplast mRNA mediates the translational requirements for two nucleusencoded functions in Chlamydomonas reinhardtii. Mol Cell Biol 14: 5268–5277 Zerges W, Girard-Bascou J and Rochaix J-D (1997) Translation of the chloroplast psbC mRNA is controlled by interactions between its 5´ leader and the nuclear loci TBC1 and TBC3 in Chlamydomonas. Mol Cell Biol 17: 3440–3448 Zhang J and Detitscher MP (1992) A uridine-rich sequence required for translation of prokaryotic mRNA. Proc Natl Acad Sci USA 89: 2605–2609 Zhang H, Herman PL and Weeks DP (1994) Gene insolation through genomic complementation using an indexed library of Chlamydomonas reinhardtii DNA. Plant Mol Biol 24: 663– 672 Zuker M (1994) Prediction of RNA secondary structure by energy minimization. Methods Mol Biol 25: 267–294

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Chapter 13

Chloroplast Protein Translocation Mireille C. Perret, Karen K. Bernd and Bruce D. Kohorn Developmental Cell and Molecular Biology, Box 91000, LSRC, Duke University, Durham, NC 27708, U.S.A.

Summary I. Introduction II. Chloroplast Import A. Signal Sequences B. The Import Apparatus III. Sorting of Proteins Within the Chloroplast IV. Thylakoid Translocation V. Mutations Affecting Translocation A. Chloroplast Suppressors B. Nuclear Suppressors VI. Perspectives Acknowledgments References

219 220 220 220 221 222 223 226 226 227 228 229 229

Summary The signal hypothesis, first put forth by G. Blobel and B. Dobberstein over 20 years ago, suggested that all proteins had as part of their primary amino acid sequence an amino-terminal region that defined the destination of proteins carrying that signal. This hypothesis has stood the test of numerous experiments and appears to be pertinent to most cellular compartments and proteins within eukaryotes and prokaryotes, including the chloroplast. Studies with Chlamydomonas reinhardtii, the subject of this chapter, have tested these concepts in live cells and have provided both new findings and the confirmation of in vitro results derived from vascular plants. Proteins synthesized outside the plastid carry chloroplast specific signals, often at their amino terminus. This signal can be removed upon entry of the protein into the chloroplast, but those destined for internal compartments within the plastid also contain additional signals. This sub-organelle targeting information can be removed once the protein is correctly localized. Thus chloroplast sorting is determined by multifunctional signal sequences. Experiments with isolated organelles have defined a number ofdifferent energetic parameters that are required for the translocation of proteins with distinct types of signal sequences, and models suggest that multiple mechanisms exist within the chloroplast. C. reinhardtii can be grown either heterotrophically or photoautotrophically thereby allowing for the selection and propagation of mutations that affect the biogenesis of chloroplasts, and thus some that affect protein translocation directly. The genetic analysis of thylakoid protein translocation in C. reinhardtii has revealed at least six loci whose products are involved in the process, and has provided genetic means to dissect those paths in vivo that have been described in isolated organelle studies. While there are likely to be a variety of requirements for the targeting and translocation of proteins to and within the chloroplast, these mechanisms may well share components between apparatuses, and between diverse groups of organisms. J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 219–231. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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I. Introduction Yellow green shapes (Greek: chloros plastos) containing sac-like forms (Greek: thytakos) were detected in the early days of microscopy and were named by Schimper in the 1880’s (Reed, 1942). Since this initial observation, we have come to realize the importance of this organelle in a variety of roles, including amino acid biosynthesis, lipid production and photosynthesis (Gillham, 1994). While chloro­ plasts and other plastids have been studied in a variety of photosynthetic species, C. reinhardtii has provided a unique contribution. The ability of C. reinhardtii to survive on acetate supplemented media without photosynthesis has allowed the creation and propagation of mutants impaired in chloroplast function. Moreover, the rapidity and ease of radiolabeling proteins in live C. reinhardtii has led to important conclusions about processes in vivo. The import of proteins into chloroplasts and the translocation ofproteins into and across the thylakoid membrane has been recently reviewed extensively (Cline and Henry, 1996; Kouranov and Schnell, 1996), and it would be redundant to provide another detailed account. Rather, we will highlight important findings that describe the mechanisms and, where appropriate, indicate how studies with C. reinhardtii have contributed. In most cases, work with C. reinhardtii has provided a basis for in vivo mechanisms, while studies with vascular plants have been restricted to either in vitro studies or gene identification. While many have touted one or another organism as ‘model,’ and C. reinhardtii is no exception, it is perhaps more useful to realize that we will discover something interesting wherever we look and that it is easier to do certain experiments with particular organisms. The internal structure of C. reinhardtii chloroplasts appears ultrastructurally and compo­ sitionally extremely similar to that of higher plants, and thus what we find true for events within the C. reinhardtii chloroplast is also likely true for other algae and angiosperms. Differences, such as reduced amounts of granal stacks in the algal chloroplasts Abbreviations: Cyt f – cytochrome f; DHFR – dihydrofolate reductase; LHCP– light harvesting chlorophyll a/b protein; OEC – oxygen evolving complex; OEC33, OEC23, OEC17 – 33 kDa, 23 kDA and 17 kDa subunits of the oxygen evolving complex; PC – plastocyanin; Rubisco SSU –small subunit of ribulose-1,5bisphosphate carboxylase oxygenase; SRP54 – 54 kDa subunit of the signal recognition particle; Tip – thylakoid insertion protein

(Harris, 1989) and the presence ofalternative electron (Wood, 1978; acceptors such as cytochrome Merchant and Bogorad, 1987), may reflect adaptive strategies. There are distinct differences between the organization of a chloroplast within a C. reinhardtii cell and within a vascular plant cell. The most obvious is that C. reinhardtii hasjust one chloroplast that fills most of the cell and is usually oriented in the same way within the cell. Vascular plant cells are normally characterized as having multiple chloroplasts that can stream with the cytoplasm or reside in numerous cellular locations. Thus, a priori, one might expect to find that there are unique features in C. reinhardtii versus vascular plants in mechanisms that target proteins from the cytoplasm to the chloroplast surface.

II. Chloroplast Import

A. Signal Sequences The signal hypothesis, first put forth by Blobel and coworkers (Blobel and Sabatini, 1971; Blobel and Dobberstein, 1975) over 20 years ago, suggested that all proteins had, as part of their primary amino acid sequence, an amino-terminal region that defined the destination of proteins carrying that signal. This concept has dominated and driven the field of protein targeting, and most experiments are consistent with the hypothesis. Not long after putting forth their signal hypothesis, which was based on studies with the endoplasmic reticulum, Blobel and Dobberstein (Blobel and Dobberstein, 1977) detected a ‘putative precursor’ of C. reinhardtii Rubisco small subunit (SSU) that had been translated in vitro, was precipitated by anti-SSU antibody, and could be processed to the size of SSU. They proposed that the extra sequence included chloroplast targeting information. Indeed, chloroplast proteins whose translation is initiated in the cytosol usually contain amino-terminal signals that are necessary and sufficient for targeting to the chloroplast envelope. These signals are often removed by one or more stromal peptidases upon entry into the chloroplast (Su and Boschetti, 1993, 1994; Rufenacht and Boschetti, 1995). While there are a number of predicted signal sequences in C. reinhardtii (von Heijne et al., 1991), in only a few cases have these sequences been shown to be functional and removed upon import (Yu et al., 1988; Schnarrenberger et al., 1994; Funke et al., 1997). The definition of a signal

Chapter 13 Chloroplast Protein Translocation

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sequence has often assumed that only the portion that is removed constitutes the signal. However, it has been observed that regions of the mature protein of two vascular plant proteins can influence the import reaction in vitro, suggesting that some signals may extend beyond the processing site into the sequence of the mature protein (Kohorn et al., 1986; Wan et al., 1995). Comparisons of amino-terminal signals from a variety of vascular plant proteins reveal little or no sequence conservation, although the region is rich in hydroxylated amino acids and alanine (von Heijne et al., 1989). The amino-terminal regions of nucleusencoded C. reinhardtii chloroplast proteins also appear to have no conserved primary structure, are rich in hydroxylated amino acids, and particularly rich in alanine. In solution, the signal has no apparent secondary structure, although some have suggested that signals can form helices in the presence of lipids (Horniak et al., 1993; see Chapter 26, Jacquot et al.). A structural study of the chloroplast signal sequence of the C. reinhardtii gamma subunit of the chloroplast ATP synthase showed that this region has a random coil conformation in a nonpolar environment (Theg and Geske, 1992). C. reinhardtii chloroplast signals resemble mitochondrial targeting signals in that they contain amphipathic, positively charged regions (Franzen et al., 1990; Franzen and Falk, 1992). Indeed, the amino-terminal region of the Rubisco SSU from C. reinhardtii can direct a fusion protein of CoxIV to the mitochondrion of coxIV deficient yeast (Hurt et al., 1986) and an SSU-DHFR fusion can be imported into isolated yeast mito­ chondria (Hurt et al., 1986). These experiments demonstrate that the presequence can function as a mitochondrial targeting signal if the cell requires such targeting for survival or if the signal is only presented with isolated mitochondria. The C. reinhardtii nucleus-encoded chloroplast protein PsaF can be imported in vitro into isolated mitochondria from spinach even when its presequence is removed (Hugosson et al., 1995) suggesting that a cryptic signal may be revealed under in vitro conditions. However, it is not yet clear whether in live C. reinhardtii the sequence functions as both a mitochondrial and a chloroplast signal. In C. rein­ hardtii neither SSU nor PsaF are found in the mitochondrion, suggesting that if mistargeting occurs, then the proteins are rapidly degraded. Alternatively, the chloroplast signal may be dominant to the cryptic mitochondrial one, or, more likely, additional

regulatory mechanisms exist in the cytosol that differentiate between chloroplast and mitochondrial proteins in vivo as is the case for angiosperms (Whelan and Glaser, 1997). In summary, C. reinhardtii chloroplasts can be distinguished from those in vascular plants by their mitochondrial-like chloroplast targeting signals and by their dominating presence within a single cell. Whether these two unique characteristics are related remains to be tested. There are many reports of molecular dissection of chloroplast targeting signals. The studies provide conflicting results depending on whether the signals are assayed in vitro or by transformation into a plant. However, all studies do conclude that the region is necessary and sufficient for chloroplast import. For example, deletion studies (Hageman et al., 1990) have shown that the plastocyanin (PC) transit sequence is necessary for import into the chloroplast, and fusion protein studies have shown that the PC transit sequence is sufficient to transfer dihydrofolate reductase (DHFR) across the chloroplast envelope. Also, the pea SSU transit sequence can direct the bacterial protein neomycin phosphotransferase into pea chloroplasts (Van den Broeck et al., 1985). A systematic deletion analysis ofthe C. reinhardtii PC import signal (K. Kindle, personal commun­ ication) reveals that the region is required for import, but that the in vitro reactions are more sensitive to signal sequence perturbations than are live cells. Overall, it is clear that small deletions or substitutions can be accommodated in vivo and this may indicate a plasticity in the targeting and import apparatus. The absence of a phenotype provided by signal sequence mutations is not conducive to a genetic suppressor screen to identify proteins that interact with the signal sequence. Moreover, mutations in this import process may be lethal (Smith and Kohorn, 1994; Schnell, 1995), and together, these restrictions may explain the lack of reports utilizing C. reinhardtii or any vascular plant for a genetic analysis of chloroplast import. C. reinhardtii may provide the best genetic system for this analysis as at least problems from pleiotropic effects on photosynthesis could be avoided by growth on acetate-containing medium.

B. The Import Apparatus All ofthe studies that have characterized chloroplast envelope proteins involved in import have been performed with isolated vascular plant chloroplasts,

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and therefore will not be reviewed extensively. Briefly (as reviewed by Kouranov and Schnell, 1996), the chloroplast envelope contains two lipid bilayers and translocation occurs as a result of the coordinated action of protein translocation machinery in each bilayer at specialized regions called contact zones where the envelope membranes meet. Outer envelope proteins include the translocation-intermediate associating IAP34, IAP75, and IAP86 integral membrane protein complex and two Hsp70-like proteins, COM70 on the cytoplasmic side of the outer envelope and Hsp70-IAP on the inner side of the outer envelope. The Hsp70 proteins and the GTPbinding domain-containing IAP34 and IAP86 are considered to be components of a two-step translocation mechanism where initial precursor binding occurs with ATP and GTP hydrolysis and is then followed by translocation that requires higher amounts of ATP hydrolysis. IAP75 and IAP86 have been cross-linked to the transit sequences in vitro, as have the inner envelope proteins IAP21 (integral) and IAP25 (localization unknown). Inner envelope proteins that associate with import intermediates in vitro have also been found (IAP100, IAP36). It has also been suggested that the targeting of precursor proteins to the outer envelope machinery is aided by the unique lipid composition of this bilayer (van’t Hof et al., 1993). Interaction between the signal sequence and receptor components may be predicated by lipid-precursor protein contact that induces a more translocation-competent secondary structure in the signal sequence. Many of the conclusions concerning chloroplast targeting have been based upon the assumption that import is post-translational. This assumption is derived from the early observation that completed protein chains can be imported into chloroplasts (Chua and Schmidt, 1978; Mishkind et al., 1985) and many have used this post-translational assay to further our understanding of the process. It is quite difficult to determine whether this is indeed what occurs in vivo for all precursors. Estimations that have been made suggest that the in vivo rate of import exceeds that seen in vitro by orders of magnitude (Hoober et al., 1994). What has been lacking in the literature is a thorough analysis of the possible co-translational insertion of at least some proteins into the chloroplast. This has been shown to be a distinct possibility with yeast mitochondria, and ribosomes can be detected on the surface of mitochondria (Fujiki and Verner, 1991, 1993; Verner, 1993). C. reinhardtii would be

well suited for such studies as pulse labeling and ultrastructural observations could be coupled with in vitro analysis.

III. Sorting of Proteins Within the Chloroplast There are six identifiable compartments within the chloroplast: the outer and inner membranes of the envelope, the space between these bilayers, the stroma, the thylakoid membrane, and the thylakoid lumen. In 1986, Smeekens et al. proposed that some chloroplast signals were multifunctional, a concept that has also been supported in mitochondrial import studies (Smeekens et al., 1986; Schatz and Dobberstein, 1996). According to this hypothesis, the signal is bipartite; the amino-terminal region mediates import across the chloroplast envelope and the remaining portion ofthe signal specifies sub-organellar targeting (Fig. 1). Their experiments and subsequent analysis in other labs addressed the targeting of thylakoid lumen proteins and provided evidence suggesting that upon entry into the chloroplast, the chloroplast import region was removed by a stromal peptidase, and the remaining amino-terminal region then directed the protein to the thylakoid membrane where it mediated translocation into the lumen. Many in vitro experiments with vascular plant chloroplasts demonstrate that newly imported proteins destined for the thylakoid vesicle indeed pass through the stroma (Cline and Henry, 1996). The fact that insertion into and passage through the thylakoid membrane can be performed posttranslationally in vitro with isolated thylakoid membranes is indeed consistent with the passage of precursors through the stroma in vivo. In most cases, the thylakoid transfer signal is required for the in vitro translocation into the lumen, but the envelope signal is not. Despite their rigor and clarity, none of these experiments address directly the possible transfer of some proteins directly from the envelope to the thylakoid. While many have confirmed this sorting model for chloroplast proteins destined for the thylakoid membrane or lumen by using in vitro experiments, there is only one publication supporting this view for events that occur in vivo, and this was carried out with C. reinhardtii. Howe and Merchant (1993) were able to detect the intermediate form of lumen-destined plastocyanin (PC), and the 33 kDa cytochrome oxygen-evolving complex protein (OEC33) in which

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the envelope targeting signal had been removed, but the thylakoid signal remained. Some of this intermediate was detected in a soluble fraction which supported the idea that it was stromal. Pulse chase experiments showing the concomitant disappearance of the intermediate and increase in the amount of mature protein supported the identity of the larger form as a true intermediate. These data stand alone as the only in vivo evidence of stromal intermediates in sub-chloroplast targeting. However, some of the intermediate was insoluble, and since its location remains unknown, the issue requires more attention.

triphosphates (NTPs), and the precursor of light harvesting chlorophyll a/b binding protein (LHCP) (Kirwin requires a stromal protein and GTP and a et al., 1989; Mould and Robinson, 1991; Cline et al., 1992; Hulford et al., 1994; Nielsen et al., 1994; Yuan and Cline, 1994; Kouranov and Schnell, 1996). The stromal protein required for pre-LHCP import is likely to be CP54, a homologue of the SRP 54-kD protein, as its depletion from stroma prevents insertion (Li et al., 1995), although it has not been reported that CP54 can be added back to restore depleted extracts. Studies of the integral membrane protein found that nucleoside triphosphates and stromal extracts are not required for its integration into the membrane, and that a only slightly enhances its integration (Michl et al., 1994). The energetic requirements for the translocation of precursors of thylakoid lumen proteins appear to be dependent upon the thylakoid transfer signal as exchange of signals also exchanges the energetic requirement. A chimeric construct consisting of the pea OEC23 thylakoid transit sequence fused to the mature region of PC does not require stromal extract or NTPs for thylakoid translocation, and the translocation of both this chimera and an OEC17 transit/mature PC fusion protein is inhibited by across the nigericin, which dissipates the

IV. Thylakoid Translocation A number of detailed studies with isolated pea chloroplasts show that differentproteins have varying energetic requirements for successful translocation into and across the thylakoid membrane. All of these precursors require thylakoid membranes and thylakoid proteins, but in addition, pre-PC and pre­ OEC33 (the 33-kD oxygen-evolving complex protein) require SecA and ATP, the precursors of the oxygen-evolving complex 23- and 17-kD proteins (pre-OEC23 and pre-OEC 17) and the photosystem I but no nucleoside N-subunit protein require a

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thylakoid membrane (Robinson et al., 1994). Another study (Henry et al., 1994) showed that the translocation of an OEC23/PC chimera is not sensitive to sodium azide, an inhibitor of the SecA ATPase, while the translocation of an OEC33/OEC17 chimera is inhibited by azide. From these experiments, it appears that the energetic requirements of a chimeric protein for translocation are determined by the thylakoid transfer signal. In order to determine the region within the thylakoid signal sequence that specifies the required energetics, chimeras were made within this region (Henry et al., 1997). A protein construct consisting of the charged region of the OEC23 lumen targeting domain followed by the hydrophobic core of the PC lumen targeting domain and the mature PC sequence was able to cross the thylakoid membrane efficiently, andthis translocation and competitively was dependent upon ATP and inhibited by an OEC23 translocation intermediate. Thus, this ‘dual-targeting’ signal sequence resulted in a combination of energetic requirements. However, the flexibility and function of chimeric thylakoid signal sequences is limited: a chimera consisting of the charged region of the PC signal fused to the hydrophobic core of OEC23 and the mature sequence of PC was not translocated, and the ‘dual-targeting’ signal used for PC allowed only dependent transport when OEC23 and OEC17 were used as passenger proteins. In contrast, pre-LHCP does not have a simple cleaved thylakoid membrane-targeting signal sequence (Auchincloss et al., 1992, Huang et al. 1992). Some information resides in the mature protein because correct insertion requires multiple trans­ membrane helices. Collectively, the energetics data have been interpreted to indicate the presence of multiple pathways of translocation across the thylakoid membrane. However, the results are also consistent with there being a variety of requirements necessary for different precursors to interact with the components of just one pathway. The truth may lie somewhere between these two extremes and in the identification of the transport apparatus. Homologues of both the bacterial SecA and SecY proteins that mediate translocation across the bacterial inner membrane (Schatz and Dobberstein, 1996) have been identified in vascular plant and algal chloroplasts and cyanobacteria (Nakai et al., 1992, 1994; Laidler et al., 1995). A SecA requirement for translocation in vitro has been demonstrated for PC, cytochrome f(Cyt f) and OEC33 (Nakai et al., 1994;

Yuan et al., 1994; Nohara et al., 1996). As SecY of bacterial membranes and its mammalian homologue, Sec61p of the ER, are protein translocases, it is assumed that chloroplast SecY has a similar function. One series of studies so far has established that indeed SecA has a role in the thylakoid translocation apparatus in vivo in vascular plants. Several nuclear mutants ofmaize are impaired in the translocation of a number of thylakoid proteins in a way that is entirely consistent with the models supported by the in vitro experiments with pea chloroplasts. The tha1 mutant has a transposon-disruption of the chloroplast SecA, and it has reduced levels of properly localized PC, OEC33, PSI-F, and Cyt f while the translocation of OEC23 and OEC17 is unaffected. These results suggest that SecA can function in the translocation of some proteins across the thylakoid membrane in vivo, but that it is not strictly required for the translocation of these proteins (Voelker et al., 1997). The hcf106 mutant has the opposite translocation phenotype of tha1. It has reduced levels of properly localized OEC23 and OEC17, while the levels of PC, PSI-F, and Cyt f are reported to be unaffected (Voelker and Barkan, 1995). However, in some cases Cyt f is reduced in the hcf106 mutant (R. Martienssen, personal communication), suggesting that Cyt f pathway. translocation is influenced by the Recent crosslinking experiments suggest that CP54 can associate with LHCP, Cyt f and the Rieske FeS protein (High et al., 1997), and these were proteins thought to be on separate pathways defined by energetics. Thus, the multiple pathways may indeed involve overlapping sets of components. Consistent with the view that different precursors may interact with similar translocation apparatuses are the results obtained from in vivo studies of the translocation of C. reinhardtii Cyt f into the thyla– koid lumen (Smith and Kohorn, 1994). Mutations within the 31 amino acid signal sequence of chloroplast encoded pre-apo Cyt f (Fig. 2) reduce or eliminate its translocation into the thylakoid membrane, causing the cells to become nonphotosynthetic. Point mutations (A15E and V16D) and deletions (Fig. 2) in the predicted hydrophobic core of the signal sequence had the most pronounced effects and other mutations delineated the boundaries of this essential region (Smith and Kohorn, 1994; Baillet and Kohorn, 1996) The signal sequence mutation A15 E inhibits not only the insertion ofpreCyt f but also decreases the accumulation of integral thylakoid membrane proteins LHCP and D1

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(Fig. 3A). The accumulation of PC and OEC 33, which are lumenal, is not affected (Fig. 3 A). OEC 17 and 23, lumenal proteins having different energetic requirements for in vitro translocation, are increased in the A15E strain (Fig. 3A). As expected, pre-Cyt f A15E also reduces the translocation of wild type pre-

Cyt f (Smith and Kohorn, 1994). Since the Cyt f A15E mutant is found as a precursor in the stroma and thylakoid membrane surface, it may be jamming the translocation apparatus. Thus, precursors using the same path would also be jammed while those on alternative routes would not be affected (we call this

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the Los Angeles effect). This is diagrammed in Fig. 3B. The decrease in LHCP levels suggests that Cyt f A15E blocks a common step, while the increase in OEC 23 is consistent with the view that it enters the pathway after the pre-Cyt f A15E induced block. This implies that components of the translocation apparatus may be limiting. The steady state level of PC is not affected by Cyt f A15E, and thus PC may have different requirements than do Cyt f and LHCP. Some studies show that Cyt f and PC are assisted by SecA (Voelker and Barkan, 1995; Voelker et al., 1997), but if SecA is not limiting, then Cyt f A15E may not be expected to affect PC. These data collectively suggest that integral thylakoid proteins and lumenal proteins use distinct translocation apparatuses, but these pathways may have common components (Smith and Kohorn, 1994). The changes in steady state levels of other thylakoid proteins may indicate that those precursors utilize the same apparatus, but small changes in other protein levels may go undetected, and thus, this analysis is not comprehensive. The experiments on the translocation of pre-Cyt f in C. reinhardtii are consistent with the early microscopic observations of ribosomes on the thylakoid surface which suggests that translocation of some thylakoid proteins is co-translational (Falk, 1969; Margulies and Michaels, 1974). Processing of Cyt f occurs in the thylakoid lumen and only those proteins being translocated will be processed. Therefore, those that are stromal and not processed are necessarily blocked from entry into the lumen. Precursor Cyt f is never detected in wild type cells, and could only be detected in mutants defective in translocation (Smith and Kohorn, 1994). These data are consistent with co-translational translocation of Cyt f.

ability to grow photosynthetically (Smith and Kohorn, 1994; see above, Section IV). Mutations that suppressed the Cyt f signal sequence alterations were selected for their ability to restore photosynthetic growth (Fig. 4). Suppressor mutations were identified as these were thought to represent gain of function alleles, and the generation of null mutations in the translocation process might be lethal. This scheme was based on a genetic analysis of protein translocation in E. coli (Emr et al., 1981). It was expected that some ofthe suppressors would reveal proteins that were involved in the translocation of Cyt f and perhaps interact with the signal sequence. The suppressor mutations map to six nuclear loci and therefore represent extragenic suppressors of the signal sequence mutations in Cyt f (Smith and Kohorn, 1994; Bernd and Kohorn, 1998). These loci defined by the suppressor mutations have been termed TIP for thylakoid insertion proteins. Two suppressors that lie within the Cyt f signal were also isolated (Smith and Kohorn, 1994; Baillet and Kohorn, 1996).

A. Chloroplast Suppressors V. Mutations Affecting Translocation To gain an understanding of thylakoid translocation in vivo, and to produce a selection scheme aimed at identifying proteins that are directly involved in the process, C. reinhardtii was used for a systematic characterization of the Cyt f thylakoid signal se­ quence and genetic analysis of the translocation mechanism of this protein. Strains carrying point and deletion mutations of the Cyt f signal sequence were unable to translocate pre-Cyt f into the thyla­ koid membrane, and this resulted in an impaired

The two chloroplast suppressor mutations result in the removal ofthe positively charged amino acid that borders the amino-terminus of the signal sequence hydrophobic core, and replaces this arginine with either a cysteine or a leucine (Fig. 5). The original mutations at A15E and V16D are unchanged. The occurrence of this type of suppressor mutation suggests that the hydrophobic core can be shifted in position within the signal sequence. One of the two isolates of the V16D suppressor strain was used for site-directed mutagenesis to create a triple mutant, in which the hydrophobic core of the signal sequence

Chapter 13 Chloroplast Protein Translocation

was further disrupted by a charged residue, A12E, which, when alone, has no phenotype. The R10L substitution is no longer capable of restoring photosynthetic growth in this triple mutant in a context of insufficient stretches of hydrophobic core regions (Fig. 5; Baillet and Kohorn, 1996). Thus, the signal that mediates translocation into the thylakoid membrane is characterized by a hydrophobic region whose exact amino acid content is not critical, but must be a minimum length of residues. Furthermore, the hydrophobic region of the signal sequence does not have to be flanked on its amino terminus by a charged residue.

B. Nuclear Suppressors Genetic analysis reveals six unlinked loci that were selected for their suppression of the Cyt f A15E mutation. The tip suppressors allow translocation of both wild-type thylakoid proteins and Cyt f A15E, indicating that they are mutations that make the translocation machinery more permissive. Thus it is not surprising that the tip mutants show no detectable change in photosynthetic growth in a wild-type Cyt f background, relative to the A15E Cyt f strain (Bernd and Kohorn, 1998). Wild-type Cyt f is processed from a 3 5-kD stromal precursor to the mature 32-kD thylakoid membrane form by a lumenal peptidase (reviewed in Gray, 1992). Pulse-chase experiments that detect Cyt f processing revealed that the kinetics of Cyt f maturation in the tip strains ranges from rates indistinguishable from wild type to rates where the precursor of Cyt f accumulates and is detected only after relatively long chase periods (Bernd and Kohorn, 1998). As processing is a measure of translocation,

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these data confirm that the tip suppressors restore the translocation of mutant Cyt f signal sequences but show that the strength of suppression varies between tip alleles. The six tip alleles were selected for their ability to suppress the A15E mutation, but crosses with C. reinhardtii strains carrying other Cyt f signal sequence mutations show that the tip mutants can also suppress these, albeit to differing degrees (Table 1). Thus the suppressor mutations are not allele specific. The original expectation of this selection scheme was that a given suppressor would compensate for a specific signal mutation. However, as the tip suppressors can also suppress the phenotype of strains carrying a deletion of the hydrophobic core of the signal, the data suggests that the suppressors render the thylakoid more permissive to translocation of proteins without a signal or a less than ideal signal. This same observation is made for similar selection schemes in bacteria (Emr et al., 1981). Significantly, improperly localized proteins are not found in the thylakoid membrane nor in bacterial membranes in these permissive mutants. Double tip mutant analysis indicates that a tip4/ tip5 strain is not photosynthetic under stringent conditions of selection, while all other combinations of tip mutations have no affect larger than the single alleles. Thus the proteins encoded by TIP4 and TIP5 may interact. The tip5 mutation, and to a lesser degree tip2, can suppress V16D and and the V16D and mutations block the signal sequence from binding the thylakoids (Smith and Kohorn, 1994). Thus we propose that Tip4, TipS and Tip2 act early in the translocation pathway and this is shown in Fig. 6. Mutations in Tip 1, 3 and 6 might not

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suppress V16D as this altered signal sequence may never reach the later steps mediated by these proteins. The A15E mutation blocks the Cyt f precursor at the membrane and mutations in Tip1, 3, and 6 can therefore suppress this mutation.The tip suppressor alleles are being cloned by transformation of the Cyt f A15E strain with a library, and characterization of these loci should clarify some of these issues. The TIP loci may well describe SecA and SecY homologues, but the identity of the other four is unknown. The thylakoid membrane contains a number of lipids unique to the chloroplast, but their role in protein translocation has yet to be studied (Kouranov and Schnell, 1996). These lipids help to determine the shape of the flattened thylakoid structure (Janero and Barrnett, 1981), and perhaps an alteration in the structure of the lipids by a translocation protein apparatus could prepare the bilayer for the translocating protein. This model is still consistent with the various energetic requirements that are dependent upon the type of signal rather than the mature sequence (Robinson et al., 1994), and with the signal sequence hypothesis that states that only the signal is of importance. Chlamydomonas can also be used to address the question of whether the in vitro energetic experiments truly reflect multiple independent translocation mechanisms as suggested (Robinson and Klosgen, 1994; Cline and Henry, 1996) as Cyt f, PC and OEC23 appear to have different energetic require­ ments for translocation. Null mutations of C. rein­ hardtii PC, OEC33 and OEC23 (ac208, Fud44 and Fud39, respectively) have been isolated (Gorman and Levine, 1966; Mayfield et al., 1987a; de Vitry et al., 1989) and the genes cloned (Mayfield et al., 1987b; Mayfield et al., 1989; Quinn et al., 1993), making the construction and analysis of signal sequence mutants that inhibit their translocation

possible. If there is interaction or sharing between translocation components used by Cyt f and PC or OEC23, then at least some of the tip mutants should suppress inhibitory mutations within those signals. Different tip mutants might be expected to suppress different signal types if there are several pathways that overlap at different points (as proposed in Fig. 3). If the tip suppressor alleles only suppress Cyt f mutations, then new tip mutants specific to PC or OEC33 or OEC23 can be isolated to describe participants in those import processes.

VI. Perspectives It is assumed that the ultimate goal of many researchers is to create a thylakoid translocation system that has been reconstituted from isolated components identified through in vitro or genetic approaches. This would enable a detailed analysis of mechanism. However, these experiments will still leave the question of what occurs in vivo. Hoober has proposed that proteins transversing the envelope do

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not pass through stromal intermediates, and perhaps those intermediates that are detected are rare escapes (Hoober et al., 1994). Those PC, and OEC 33 soluble intermediates detected in C. reinhardtii (Howe and Merchant, 1993) may be species that escape from the general path, or are in vesicles that fractionate with soluble components; the insoluble intermediates remain to be characterized. This alternate view is not generally accepted but is consistent with the large amounts of vesicular traffic and contacts between thylakoid and envelopes observed in chloroplasts active in thylakoid biogenesis (Hoober et al., 1994; Chapter 19, Hoober et al.). Direct transfer oflipophilic proteins such as the 22-kD thylakoid heat-shock protein and LHCP, which do not contain simple thylakoid signal sequences (Grimm et al., 1989; Auchincloss et al., 1992), from the envelope to the thylakoid membrane would obviate the need in vivo for stromal proteins such as CP54. Alternatively, CP54 may mediate envelope to thylakoid transfer. While most in vitro experiments are clear and concise and support the concept of multifunctional signal sequences that produce stromal intermediates, these alternate ideas remain to be tested. C. reinhardtii may well be best suited to determine the translocation pathways involved in intact cells.

with mutant signal sequences. Genetics, in press Blobel G and Dobberstein B (1975) Transfer of proteins across membranes. Presence of proteolytically processed and unprocessed nascent immunoglobin light chains on the membrane-bound ribosomes of murine myeloma. J Cell Biol 67: 835–851 Blobel G and Dobberstein B (1977) In vitro synthesis and processing of a putative precursor for the small subunit of ribulose-l,5-bisphosphate carboxylase of Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 74: 1082–1085 Blobel G and Sabatini D (1971) Ribosome-membrane interaction in eukaryotic cells. In: Manson LA (ed) Biomembranes, pp 193–195. Plenum Publishing Corporation, New York Chua N-H and Schmidt G (1978) Post-translational transport into intact chloroplasts of a precursor to the small sub-unit of ribulose-l,5-bisphosphate carboxylase. Proc Natl Acad Sci USA 75: 6110–6111 Cline K and Henry R (1996) Import and routing of nucleusencoded chloroplast proteins. Ann Rev of Cell and Devel Biol 12: 1–26 Cline K, Ettinger W and Theg S (1992) Protein-specific energy requirements for protein transport across or into thylakoid membranes. J Biol Chem 267: 2688–2696 de Vitry C, Olive J, Drapier D, Recouvreur M and Wollman F-A (1989) Posttranslational events leading to the assembly of photosystem II protein complex: A study using photosynthesis mutants from C. reinhardtii. J Cell Biol 109: 991–1006 Emr S, Hanley-Way S and Silhavy T (1981) Suppressor mutations that restore export of a protein with a defective signal sequence. Cell 23: 79–88 Falk H (1969) Rough thylakoids: polysomes attached to chloroplast membranes. J Cell Biol 42: 582–587 Franzen L-G and Falk G (1992) Nucleotide sequence of cDNA clones encoding the beta subunit of mitochondrial ATP synthase from the green alga C. reinhardtii: The precursor protein encoded by the cDNA contains both an N-terminal presequence and a C-terminal extension. Plant Mol Biol 19: 771–780 Franzen L, Rochaix J and von Heijne G (1990) Chloroplast transit peptides from the green alga C. reinhardtii share features with both mitochondrial and higher plant chloroplast presequences. FEBS Lett 260: 165–168 Fujiki M and Verner K (1991) Coupling of protein synthesis and mitochondrial import in a homologous yeast in vitro system. J Biol Chem 266: 6841–6847 Fujiki M and Verner K (1993) Coupling of cytosolic protein synthesis and mitochondrial protein import in yeast. Evidence for cotranslational import in vivo. J Biol Chem 268: 1914– 1920 Funke RP, Kovar JL and Weeks DP (1997) Intracellular carbonic anhydrase is essential to photosynthesis in C. reinhardtii at Demonstration via genomic atmospheric levels of mutant ca-1. Plant complementation of the high Physiol 114: 237–244 Gillham N (1994) Organelle genes and genomes. Oxford University Press New York Gorman D and Levine R (1966) Photosynthetic electron transport of C. reinhardtii VI. Electron transport in mutant strains lacking either cytochrome 553 or plastocyanin. Plant Physiol 41: 1648–1656 Gray JC (1992) Cytochrome f: Structure function and biosynthesis. Photosyn Res 34: 359–374

Acknowledgments We would like to thank Phillip Hartzog, Benoit Baillet and Lib Harris for their help in the work presented, which was supported by the Pew Charitable Trusts, the National Institutes of Health, and the United States Department of Agriculture. We have attempted to represent the field fairly, and apologize to any of those that were excluded, as it was by no means intentional.

References Auchincloss A, Alexander A and Kohorn BD (1992) Requirement for three membrane spanning alpha-helices in the post­ translational insertion ofa thylakoid membrane protein. J Biol Chem 267: 10439–10446 Baillet B and Kohorn BD (1996) Hydrophobic core but not amino-terminal charged residues are required for translocation of an integral thylakoid membrane protein in vivo. J Biol Chem 271: 18375–18378 Bernd KK and Kohorn BD (1998) Tip loci: Six Chlamydomonas nuclear suppressors that permit the translocation of proteins

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Grimm B, Ish-Shalom D, Even D, Glaczinski H, Ottersbach P, Ohad I and Kloppstech K (1989) The nuclear-encoded chloroplast 22kD heat-shock protein of C. reinhardtii. Evidence for translocation into the organelle without a processing step. Eur J Biochem 182: 539–46 Hageman J, Baecke C, Ebskamp M, Pilon R, Smeekens S and Weisbeek P (1990) Protein import into and sorting inside the chloroplast are independent processes. Plant Cell 2: 479–494 Henry R, Kapazoglou A, McCaffery M, and Cline K (1994) Differences between lumen targeting domains of chloroplast transit peptides determine pathway specificity for thylakoid transport. J Biol Chem 269: 10189–10192 Henry R, Carrigan M, McCaffery M, Ma X and Cline K (1997) Targeting determinants and proposed evolutionary basis for the Sec and delta pH protein transport systems in chloroplast thylakoid membranes. J Cell Biol 136: 823–832 Harris EH (1989) The C. reinhardtii Sourcebook: A Compre­ hensive Guide to Biology and Laboratory Use. Academic Press, San Diego High S, Henry R, Mould R, Valent Q, Meacock, S, Cline K, Gray J and Luirink J (1997) Chloroplast SRP54 interacts with a specific subset of thylakoid precursor proteins. J Biol Chem 272: 11622–11628 Hoober J, White R, Marks D and Gabriel J (1994) Biogenesis of thylakoid membranes with emphasis on the process in C. reinhardtii. Photosyn Res 39: 15–31 Horniak L, Pilon M, van’t Hof R and de Kruijff B (1993) The secondary structure of the ferredoxin transit sequence is modulated by its interaction with negatively charged lipids. FEBS Lett 334: 241–246 Howe G and Merchant S (1993) Maturation of thylakoid lumen proteins proceeds post-translationally through an intermediate in vivo. Proc Natl Acad Sci USA 90: 1862–1866 Huang L, Adam Z and Hoffman NE (1992) Deletion mutants of chlorophyll a/b binding proteins are efficiently imported into chloroplasts but do not integrate into thylakoid membranes. Plant Physiol 99: 247–255 Hugosson M, Nurani G, Glaser E and Franzen LG (1995) Peculiar properties of the PsaF Photosystem J protein from the green alga C. reinhardtii: Presequence independent import of the PsaF protein into both chloroplasts and mitochondria. Plant Mol Biol 28: 525–535 Hulford A, Hazell L, Mould R and Robinson C (1994) Two distinct mechanisms forthe translocation ofproteins across the thylakoid membrane, one requiring the presence of a stromal protein factor and nucleotide triphosphates. J Biol Chem 269: 3251–3256 Hurt E, Soltanifar N, Goldschmidt-Clermont M, Rochaix JD and Schatz G (1986) The cleavable pre-sequence of an imported chloroplast protein directs attached polypeptides into yeast mitochondria. EMBO J 5: 1343–1350 Jancro DR and Barrnett R (1981) Cellular and thylakoidmembrane phospholipids of C. reinhardtii 137+. J Lipid Res 22:1126–1130 Kirwin P, Meadows J, Shackleton J, Musgrove J, Elderfield P, Hay N and Robinson C (1989) ATP-dependent import of a lumenal protein by isolated thylakoid vesicles. EMBO J 8: 2251–2255 Kohorn BD, Harel E, Chitnis PR, Thornber JP and Tobin EM (1986) Functional and mutational analysis of the lightharvesting chlorophyll a/b protein of thylakoid membranes.

J Cell Biol 102: 972–981 Kouranov A and Schnell D (1996) Protein translocation at the envelope and thylakoid membranes of chloroplasts. J Biol Chem 271: 31009–31012 Laidler V, Chaddock AM, Knott TG, Walker D and Robinson C (1995) A SecY homolog in Arabidopsis thaliana, J Biol Chem 270: 17664–17667 Li X, Henry R, Yuan J, Cline K and Hoffman N (1995) A chloroplast homologue of the signal recognition particle subunit SRP54 is involved in the posttranslational integration of a protein into thylakoid membranes. Proc Natl Acad Sci USA 92: 3789–3793 Margulies M and Michaels A (1974) Ribosomes bound to chloroplast membranes in C. reinhardtii. J Cell Biol 60: 65–77 Mayfield S, Bennoun P and Rochaix JD (1987a) Expression of the nuclear encoded OEE1 protein is required for oxygen evolution and stability of Photosystem II particles in C. reinhardtii. EMBO J 6: 313–318 Mayfield S, Rahire M, Frank G, Zuber H and Rochaix JD (1987b) Expression of the nuclear gene encoding oxygenevolving enhancer protein 2 is required for high levels of photosynthetic oxygen evolution in C. reinhardtii. Proc Natl Acad Sci USA 84: 749–753 Mayfield S, Schirmer-Rahire M, Frank G, Zuber H and Rochaix J-D (1989) Analysis of the genes of the OEE1 and OEE3 proteins of the Photosystem II complex from C. reinhardtii. Plant Mol Biol 12: 683–693 Merchant S and Bogorad L (1987) The Cu(II)-repressible plastidic cytochrome c: Cloning and sequence of a complementary DNA for the pre-apoprotein. J Biol Chem 262: 9062–9067 Michl D, Robinson C, Schackleton J, Herrmann R and Klosgen R (1994) Targeting of proteins to the thylakoids by bipartite presequences: CFoll is imported by a novel, third pathway. EMBO J 13: 1310–1317 Mishkind M, Wessler S and Schmidt G (1985) Functional determinants in transit sequences: Import and partial maturation by vascular plant chloroplasts ofthe ribulose-1,5-bisphosphate carboxylase small subunit of C. reinhardtii. J Cell Biol 100: 226–234 Mould R and Robinson C (1991) A proton gradient is required for the transport of two lumenal oxygen-evolving proteins across the thylakoid membrane. J Biol Chem 266: 12189– 12193 Nakai M, Tanaka A, Omata T and Endo T (1992) Cloning and characterization of the secY gene from the cyanobacterium Synechococcus PCC7942. Biochem Biophys Acta 1171:113– 116 Nakai M, Goto A, Nohara T, Sugita D and Endo T (1994) Identification of the SecA protein homolog in pea chloroplasts and its possible involvement in thylakoidal protein import. J Biol Chem 269: 31338–31341 Nielsen V, Mant A, Knoetzel J, Moller B and Robinson C (1994) Import of barley photosystem I subunit N into the thylakoid lumen is mediated by a bipartite presequence lacking an intermediate processing site. J Biol Chem 269: 3762–3766 Nohara T, Asai T, Nakai M, Suguira M, and Endo T (1996) Cytochrome f encoded by the chloroplast genome is imported into thylakoids via the SecA-dependent pathway. Biochem Biophys Res Com 224: 474–478 Quinn J, Li H, Singer J, Morimoto B, Mets L, Kindle K and Merchant S (1993) The plastocyanin-deficient phenotype of

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C. reinhardtii ac-208 results from a frame-shift mutation in the nuclear gene encoding plastocyanin. J Biol Chem 268: 7832– 7841 Reed HS (1942) A Short History of the Plant Science. Chronica Botanica Company, Waltham, MA Robinson C and Klosgen R (1994) Targeting of proteins into and across the thylakoid membrane—a multitude of mechanisms. Plant Mol Biol 26: 15–24 Robinson C, Cai D, Hulford A, Brock I, Michl D, Hazell L, Schmidt I, Herrmann R and Klosgen R (1994) The presequence of a chimeric construct dictates which of two mechanisms are utilized for translocation across the thylakoid membrane: evidence for the existence of two distinct translocation systems. EMBO J 13: 279–285 Rufenacht A and Boschetti A (1995) Specificity of processing enzymes in chloroplasts of C. reinhardtii In: P. Mathis (ed) Photosynthesis: From Light to Biosphere, Vol III, pp 767–770. Kluwer Academic Publishers, Dordrecht Schatz G and Dobberstein B (1996) Common principles of protein translocation across membranes. Science 271: 1519– 1526 Schnarrenberger C, Pelzer-Reith B, Yatsuki H, Freund S, Jacobshagen S and Hori K (1994) Expression and sequence of the only detectable aldolase in C. reinhardtii. Arch Biochem Biophys 313: 173–78 Schnell D (1995) Shedding light on the chloroplast protein import machinery. Cell 83: 521–524 Smeekens S, Bauerle C, Hageman J, Keegstra K and Weisbeek P (1986) The role of the transit peptide in the routing of precursors toward different chloroplast compartments. Cell 46: 365–375 Smith T and Kohorn BD (1994) Mutations in a signal sequence for the thylakoid membrane identify multiple protein transport pathways and nuclear suppressors. J Cell Biol 126: 365–374 Su Q and Boschetti A (1993) Partial purification and properties of enzymes involved in the processing of a chloroplast import protein from C. reinhardtii. Eur J Biochem 217: 1039–1047 Su Q and Boschetti A (1994) Substrate- and species- specific processing enzymes for chloroplast precursor proteins. Biochem J 300: 787–792 Theg SM and Geske FJ (1992) Biophysical characterization of a transit peptide directing chloroplast protein import. Bio­ chemistry 31: 5053–5060 Van den Broeck G, Timko MP, Kausch AP, Cashmore AR, Van Montagu M and Herrera-Estrella L (1985) Targeting of a

foreign protein to chloroplasts by fusion to the transit peptide from the small subunit of ribulose 1,5-bisphosphate carboxylase. Nature 313: 358–363 van’t Hof R, van Klompenburg W, Pilon M, Kozubek A, de Korte-Kool G, Demel R, Weisbeek PJ and de Kruijff B (1993) The transit sequence mediates the specific interaction of the precursor of ferredoxin with chloroplast envelope membrane lipids. J Biol Chem 268: 4037–4042 Verner K (1993) Co-translational protein import into mito­ chondria: An alternate view. Trends Biochem Sci 18: 366–371 Voelker R and Barkan A (1995) Two nuclear mutations disrupt distinct pathways for targeting proteins to the chloroplast thylakoid. EMBO J 14: 3905–3914 Voelker R, Mendel-Hartvig J and Barkan A(1997)Transposondisruption ofa maize nuclear gene, tha 1 encoding a chloroplast SecA homologue: In vivo role of cp-SecA in thylakoid protein targeting. Genetics 145: 467–478 von Heijne G, Steppuhn J and Herrmann R (1989) Domain structure of mitochondrial and chloroplast targeting peptides. EurJ Biochem 180: 535–545 von Heijne G, Hirai T, Klosgen RB, Steppuhn J, Bruce B, Keegstra K, and Herrmann R (1991) CHLPEP—A database of chloroplast transit peptides. PMB Reporter 9: 104–126 Wan J, Blakely SD, Dennis DT and Ko K (1995) Import characteristics of a leucoplast pyruvate kinase are influenced by a 19-amino-acid domain within the protein. J Biol Chem 270: 16731–16739 Whelan J and Glaser E (1997) Protein import into plant mitochondria. Plant Mol Biol 33: 771–789 Wood PM (1978) Interchangeable copper and iron proteins in algal photosynthesis. Studies on plastocyanin and cytochrome c-552 in C. reinhardtii. Eur J Biochem 87: 9–19 Yu LM, Merchant S, Theg SM and Selman BR (1988) Isolation of a cDNA for the gamma subunit of the chloroplast ATP synthase of C. reinhardtii import and cleavage of the precursor protein. Proc Natl Acad Sci USA 85: 1369–1373 Yuan J and Cline K (1994) Plastocyanin and the 33-kD a subunit of the oxygen-evolving complex are transported into thylakoids with similar requirements as predicted from path way specificity. J Biol Chem 269: 18463–18467 Yuan J, Henry R, McCaffery M and Cline K (1994) SecA homolog in protein transport within chloroplasts: Evidence for endosymbiont-derived sorting. Science 266: 796–798

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Chapter 14

Supramolecular Organization of the Chloroplast and of the Thylakoid Membranes Jacqueline Olive

Institut Jacques Monod, CNRS/Université Denis Diderot,

2 Place Jussieu, 75251 Paris Cedex, France

Francis-André Wollman

Laboratoire de Photosynthèse, Institut de Biologie Physico-chimique,

13 rue Pierre et Marie Curie, 75005 Paris, France

Summary I. Introduction II. Cell and Chloroplast Morphology A. Cell B. Chloroplast 1. Envelope 2. The Eyespot 3. The Pyrenoid 4. Thylakoid Membranes III. Ultrastructural Organization of Thylakoid Membranes A. Lateral Distribution of Membrane Components 1. PS II Reaction Centers 2. PS I Reaction Center and its Peripheral Antenna 3. Light-Harvesting Complex of PS II 4. Cytochrome Complex 5. ATP Synthase 6. The Static Picture B. Transmembrane Organization IV. Dynamic Aspects of Thylakoid Membrane Organization A. The Stacking-Destacking Process B. State Transition: Phosphorylation-Dependent Changes C. Fusion of Thylakoid Membranes During Sexual Reproduction V. Biogenesis A. Greening: Synthesis of Membrane Complexes B. Assembly of PS II VI. Conclusion Acknowledgment References

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J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 233–254. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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Jacqueline Olive and Francis-André Wollman

Summary From a comparative ultrastructural analysis of the wild-type and mutant strains of Chlamydomonas reinhardtii, it has been established that the complexes involved in photosynthesis are heterogenously distributed between stacked and unstacked membranes: PS II centers and their peripheral antenna are essentially localized in stacked regions of the membranes, whereas PS I centers, their peripheral antenna and ATP synthase are complex is distributed in both membrane domains. exclusively located in unstacked areas; the cytochrome Each of these protein complexes can be visualized by freeze-fracture as individual particles, but they can also be found in association with other proteins: in particular, the peripheral antennae are, in part, associated with their corresponding reaction centers to form large PFu or EFs particles. It has also been suggested that some Cyt complexes may be associated within the same particle with PS I or PS II reaction centers. Another important characteristic of intramembrane complexes is their lateral mobility. Destacking of the membranes by removal of divalent cations induces a random redistribution of the proteins in the membrane. State transitions also cause complexes, along a lateral displacement of the mobile part of the PS II peripheral antenna, and of some Cyt the thylakoid membranes. Fusion of thylakoid membranes from two distinct gametes, upon zygote formation, allows their photosynthetic protein complexes to mix. The identification, as authentic photosynthetic complexes, of a large proportion of the intramembrane thylakoid particles permitted biogenesis studies at the ultrastructural level. Here we discuss the assembly of the Photosystem II complexes.

I. Introduction For two decades, the supramolecular organization of the thylakoid membranes has been the subject of intensive investigations aimed at the identification of substructures corresponding to individual protein complexes and at the description of their spatial and functional integration in the photosynthetic apparatus (for reviews, see Staehelin, 1986; Gantt, 1994; Mustárdy, 1996; Staehelin and van der Staay, 1996). In the chloroplasts of green algae and vascular plants, intramembrane complexes located in specialized membrane discs, the thylakoids, carry out a series of interrelated photochemical and redox reactions by which light energy is trapped and converted into utilizable chemical energy. The complexes involved in the photosynthetic process are the two photosystems, Photosystem I (PS I) and complex, Photosystem II (PS II), a cytochrome operating between PS II and PS I in an electron to nicotinamide-adeninetransfer chain from dinucleotide phosphate and an adenosine triphosphate (ATP) synthase, utilizing the gradient Abbreviations: CPo – chlorophyll-protein complex of Photosystem I antenna; – chlorophyll-protein complex I ; OEE – oxygen evolving enhancer; EFs – stacked external face; EFu – unstacked external face; PS I – Photosystem I reaction center; PS II – Photosystem II reaction center; PFs – stacked protoplasmic face; PFu – unstacked protoplasmic face; RCI – reaction center I; RuBP – ribulose bisphosphate; RuBisCO – ribulose bisphosphate carboxylase/oxygenase

generated by the electron transfer chain to produce ATP. The four protein complexes are supramolecular assemblies of polypeptide chains, pigments and/or electron carriers. In addition, two small electron carriers, plastoquinones and plastocyanin, are located respectively within the hydrophobic core of the membrane and in a soluble form in the thylakoid lumen. The first thin sections of Chlamydomonas spp. were observed by electron microscopy by Sager and Palade (1954). They reported the presence ofa regular laminated organization in the chloroplast, corres­ ponding to thylakoid membranes. Further progress in the knowledge of the ultrastructure of membranes came with the development of the freeze-fracture technique. For this technique, the biological material is frozen at very low temperature (–196 °C) and Torr). fractured under high vacuum conditions ( The membranes of the cells split along an interior plane, exposing face views of the hydrophobic interior of the lipid bilayer. Particles, which are correlated to intramembranous complexes are visualized by shadowing with platinum and carbon layers (Branton, 1966). For Chlamydomonas spp. (Goodenough and Levine, 1969; Goodenough and Staehelin, 1971; Ojakian and Satir, 1974) as well as for vascular plants (Armond and Arntzen, 1977; Armond et al., 1977), distinct membrane domains were described. They were attributed to stacked and unstacked membranes respectively, and were characterized by their content in various classes of particles which

Chapter 14

Supramolecular Organization of Chloroplast and Thylakoid Membranes

differed in size and density. One of the main goals of the freeze-fracture study of thylakoid membranes has been the identification of the different categories of intramembrane particles. For that purpose, research with Chlamydomonas spp. has been of primary importance because of the numerous mutants that were characterized as specifically lacking one type of photosynthetic complex. Thus a correlation could be drawn between the loss of a category of intramembrane particles and the absence of a specific set of polypeptides, corresponding to the different subunits of an oligomeric protein. More recently, the biochemical characterization of the subunits of photosynthetic protein complexes and significant progress in their purification has opened the way to the preparation of specific antibodies. This led to the development of an immunocytochemical approach. For this technique, the cells are embedded at low temperature in a hydrophilic resin (Lowicryl K4M) which preserves the antigenicity of the molecules. Labeling with the antigens is performed on thin sections so that antibodies can access antigens. The bound antibodies are revealed by electron-dense colloidal gold particles, coupled either to protein A or to anti-rabbit IgG antibodies (Carlemalm et al., 1982). This technique provided direct detection ofthe protein complexes in the thylakoid membrane domains. Critical to that technique is the estimation of the proportion of background labeling. The availability of Chlamy­ domonas mutants totally devoid of an antigen has been a most valuable tool to assess the significance of the labeling with the corresponding antibody in the wild type strain.

II. Cell and Chloroplast Morphology

A. Cell There are many species in the Chlamydomonas genus, but mainly four species are used experimentally: C. reinhardtii, C. eugametos, C. moewusii and C. monoica (Harris, 1989). Moreover, the ultra­ structural organization of the thylakoid membranes has been extensively characterized only in C. rein­ hardtii. This is largely due to the availability of a great number of mutant strains of this species. C. reinhardtii was described for the first time by Dangeard (1888). Early descriptions of its architecture were derived from electron micrographs obtained by

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Sager and Palade (1954). It is an ovoid cell, in diameter (Figs. 1A and B). It is delimited by a plasma membrane surrounded by a glycoprotein cell wall. It has a polar structure with two anterior flagella and a single basal Chloroplast that may partially surround the nucleus. The two flagella, each about 10 long, arise from a basal body (Fig. 1C). The

236 chloroplast bounded by a double envelope membrane contains numerous lamellae, the thylakoids, and two typical substructures: the pyrenoid surrounded by starch-containing bodies (Fig. 1B) and the eyespot located just beneath the chloroplast envelope membrane (Fig. 1D). Like other eucaryotic cells, Chlamydomonas spp. cells contain a nucleus, mitochondria, an endoplasmic reticulum and a Golgi apparatus. The cell wall described in a thin section study (Roberts et al., 1972) has also been analyzed by the quick-freeze, deep-etch technique (Goode­ nough and Heuser, 1985). The major wall constituents are hydroxy-proline-rich glycoproteins, with arabinose, mannose, galactose and glucose as the predominant sugars in C. reinhardtii. The intact wall has been shown to consist of a discrete central triplet bisecting a meshwork formed of a distinct set of components. Other species of Chlamydomonas vary in cell shape, thickness of the cell wall or vary in the number of pyrenoids (for a review, see Ettl, 1976).

B. Chloroplast In C. reinhardtii, the chloroplast appears as a highly organized body, with a U-shape profile which occupies about 40% of the volume of the cell (Schötz et al. 1972). It is surrounded by an envelope and contains not only stacks of lamellae which are the site of photosynthesis but also, localized in the stroma, differentiated regions such as the eyespot associated with phototaxis and the pyrenoid associated with starch synthesis and storage, and with accumulation of ribulose-1 -5 bisphosphate carboxylase/oxygenase (Rubisco).

1. Envelope The algal chloroplasts, like those of vascular plants, are delimited by a basic envelope consisting of two parallel membranes, the inner and outer envelope membranes, each with a distinct set of polypeptides. The inner envelope membrane regulates the transport of metabolites into and out of the chloroplast, whereas the outer envelope is highly permeable to many low molecular weight substances. The chloroplast envelope is the site where the organelle interacts with other cellular components. The envelope not only regulates the transport of metabolites between the stroma and the cytosol but also mediates the import of nucleus-encoded chloroplast proteins. Most

Jacqueline Olive and Francis-André Wollman of these are synthesized as precursor proteins with N-terminal extensions, the transit peptides. These transit peptides are necessary and sufficient to direct the import of proteins into the chloroplast. The envelope also plays a major role in the biosynthesis of various lipids including galactolipids, the predominant lipids in chloroplast membranes (Douce and Joyard, 1990; see also Chapter 21, Trémolières). The inner membrane has been shown to have a higher protein to lipid ratio (Douce and Joyard, 1990). Consistent with this result, freezefractured cells show that the inner envelope contains five to ten times more intramembranous particles than the outer envelope membrane (Fig. 2, A and B). As suggested by Staehelin (1986), patches of large particles within the outer envelope membrane make contact with the inner envelope membrane.

2. The Eyespot In C. reinhardtii, the eyespot is found in the chloroplast, midway between the anterior and posterior regions of the cell (Sager and Palade, 1957; Goodenough et al., 1969;Gruber and Rosario, 1974; Melkonian and Robenek, 1980). The eyespot lies directly beneath the chloroplast envelope and consists of plates of large dense granules closely packed together (Figs. 1D and 3A). There is an alternating arrangement of plates and discs. Each plate is composed of a single layer of spherical, uniform bodies, about 100 to 140 nm in diameter (Sager and Palade, 1957). The number of rows of granules is constant for most of the species but can differ from one species to another. The eyespot of C. eugametos possesses only one layer of granules whereas that of C. reinhardtii consists of three or four layers (Lembi and Lang, 1965). The presence of microtubules has been described in the eyespot region but, in the green algae, the eyespot and flagella are separated by a fairly broad expanse cytoplasmic domain (Gruber and Rosario, 1974) and a direct association between the two organelles seems to be unlikely (Walne and Arnott, 1967). Freeze-fracture studies show that the plasmalemma and the outer envelope membranes overlying the eyespot contain a greater number of intramembrane particles than do other areas of both membranes (Nakamura et al., 1973; Bray et al., 1974; Melkonian and Robenek, 1980) (Fig. 3B). It has been shown that the eyespots in all flagellated algae act as a photoreceptor for phototaxis. It has

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been proposed that the eyespot could act as a shading device with the photoreceptor sites lying in the membrane adjacent to the eyespot globules (Arnott and Brown, 1967) or that it could reflect and intensify light of a specific spectral range (Foster and Smyth, 1980).

3. The Pyrenoid The pyrenoid is a differentiated region of the chloroplast stroma of many eucaryotic algae. It is a (Sager spherical body with a diameter of 1.5 to and Palade, 1957; Gibbs, 1962a,b; Griffiths, 1980). It appears in thin sections as uniformly dense (Fig. 1B). Pyrenoids have been isolated from C. reinhardtii and from several species of algae (Holdsworth, 1971; Kerby and Evans, 1978; Sato et al., 1984; Kuchitsu et al., 1988), and their main component was shown to be Rubisco . The functional significance of pyrenoids is controversial. Due to their close spatial relationship with starch granules, a role in starch synthesis has been proposed (Sager and Palade, 1957). Griffiths (1980) and Sato et al. (1984) assumed that the pyrenoid functions as a reservoir of Rubisco protein

whereas Kuchitsu et al. (1988) suggested that RuBP carboxylase is active in the pyrenoid and shows distinct kinetic characteristics. The pyrenoid might function as a specific metabolic compartment providing efficient coupling between carbon fixation and starch metabolism.

4. Thylakoid Membranes The basic structural components of the chloroplast are membranes. They are organized into very long, flat vesicles, called discs (Sager and Palade, 1957; Gibbs, 1962a; Ohad et al., 1967a; Goodenough and Levine, 1969). The discs, in turn, are appressed to one another in such a way as to form an elaborate array of stacked membranes. In thin sections, the membranes inside the wild-type chloroplast are seen either as single discs, or more frequently in stacks of two to ten discs (Fig. 4). A given disc may be

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associated in some places with one stack and in other places with another stack but the length of the intervening unstacked disc region is rarely extensive. In this respect it differs widely from the chloroplast membrane system of vascular plants, in which short, uniform segments of many stacked discs (grana) are interconnected by a network of long, single discs (Staehelin et al., 1977). All components of the photosynthetic electron transport chain are known to be associated with the thylakoid membranes. The freeze-fracture technique, which allows visualization of intramembrane complexes, had been utilized early on to describe the organization of thylakoid membranes. The first pictures of freeze-fractured membranes of C. rein­ hardtii were obtained by Goodenough and Staehelin (1971) and by Ojakian and Satir (1974). In the same period, similar studies were conducted with Euglena gracilis (Miller and Staehelin, 1973) and vascular plant chloroplasts (Sjolung and Smith, 1974; Staehelin, 1975). The two complementary hemi­ membranes, the exoplasmic (EF) and protoplasmic (PF) layers have a different organization in appressed and non-appressed regions, giving rise to four

Jacqueline Olive and Francis-André Wollman

different types of fracture faces: EFs and PFs originate from the stacked regions while EFu and PFu originate from the unstacked membrane domains (Fig. 5). The EF hemi-membrane is facing the exterior of the chloroplast, while the protoplasmic one is facing the lumen. Each fracture face was characterized by classes of particles differing in size and density (Fig. 6 and Table 1). On these grounds a search for the proteins constituting these particles was started, and this search lasted for over two decades. With the help of many Chlamydomonas mutant strains, most of the classes of intramembrane particles were correlated with a defined protein complex. From this supramolecular mapping study has emerged the description of extensive lateral heterogeneity in the distribution of the photosynthetic complexes along the thylakoid membranes. Given this knowledge, it became possible to study how controlled changes in the supramolecular organization of the membranes altered photosynthetic functions.

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III. Ultrastructural Organization of Thylakoid Membranes

A. Lateral Distribution of Membrane Components

1. PS II Reaction Centers The most spectacular correlation between intramembrane particles and protein complexes came with the identification of the PS II-associated particles. In mutant strains of Chlamydomonas devoid of PS II activity, a dramatic loss in EF particles was observed (Olive et al., 1979; Wollman et al., 1980; Olive et al., 1992). The EF particle density decreased in the wild type to 350 from 1450 particles particles in the tbc1-F34 mutant (Fig. 7 A and B) (Wollman et al., 1980). The tbc1-F34 strain lacks the PS II polypeptide P6 (CP43) because the psb C mRNA cannot be translated owing to a mutation in a nuclear regulatory gene (Rochaix et al., 1989) and, as in most mutants defective in PS II primary photoactivity, the whole set of subunits engaged in

the formation of the PS II protein complex is not accumulated in the membranes (Delepelaire, 1984; Bennoun et al., 1986). Moreover, in a partly suppressed strain, F34SU3, which showed restoration of half of the PS II activity, the EF particle density was found to be intermediate between those in the (Wollman et WT and tbc1-F34 mutant al., 1980). In the chloroplast mutant FuD34, which is also devoid of the P6 polypeptide owing to an alteration in the psbC gene (Rochaix et al., 1989), the EF particle density was calculated in both the stacked and unstacked membrane domains (Olive et al., 1992). A decrease in EF particle density was observed in

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Jacqueline Olive and Francis-André Wollman subunits of the PS II reaction center or against the extrinsic oxygen evolving enhancer (OEE) subunits showed that most of the labeling was concentrated in the stacked membrane domains (Vallon et al., 1985). This is illustrated for the labeling of the PS II core antenna subunit P5 (CP47) in Fig. 8A and Table 2. For the various components of the PS II reaction center, 10–20% of the labeling was found in unstacked regions. Similar distribution between stacked and unstacked membranes was also obtained for spinach (a barley (Vallon et al., 1985) whereas for mutant incapable of synthesizing D1), the traces of CP43 (P6) and CP47 (P5) found in this mutant were not concentrated into stacked regions (Simpson et of cytochrome al., 1989). However, the was found at a near normal level and accumulated in stacked regions. The results obtained for C. reinhardtii or spinach were in accordance with numerous fractionation studies, performed with vascular plant thylakoid membranes which indicated a pronounced depletion of PS II in stroma lamellae (Andersson and Anderson, 1980; Andersson and Melis, 1983; Vallon et al., 1987;Bassi et al., 1988).

both membrane domains, which argues for the presence of some PS II reaction centers, not only in the stacked membrane regions where most of the EF particles in the wild-type strain are found but also in the unstacked membrane regions: 533 and 243 p in EFs and EFu respectively versus 1500 and 600 p in wild type. Similar results were observed with another green alga, Chlorella sorokiniana (Lacambra et al., 1984). Taking into account the ratio of unstacked versus stacked membranes, the decreased content in EF particles in the PS II mutants led to an estimate of about 20% of the PS II centers present in the unstacked regions (Olive et al, 1992). From studies in which a double mutant more completely devoid of PS II subunits (Section V.B) was used, it was concluded that PS II centers appear to be responsible for about 95% of the EF particles. Several studies with PS II mutants from vascular plants concluded an even higher heterogeneity in the distribution of PS II, with only minor changes on the EFu faces when there was a considerable drop in the density of the EFs particles (Miller and Cushman, 1979; Simpson et al., 1989). The immunocytochemical approach also supported the heterogeneity in lateral distribution of PS II between the stacked and unstacked domains. Studies using various antibodies against the main integral

2. PS I Reaction Center and its Peripheral Antenna In the freeze-fractured membranes, the PS I reaction center and its peripheral antenna were expected to correspond to a subpopulation of the PF particles, and ATP the other particles corresponding to Cyt synthase. Since, in contrast with PS II mutant strains, neither mutation actually caused the disappearance of only the PS I complex or its peripheral antenna, both complexes were treated together. A comparative freeze-fracture study was conducted with the F14 and Ac40 mutants. The former strain is which contains the PS I primary donor devoid of P700 and the PS I core antenna, and the latter strain which corresponds to the two complexes lacks LHCI-705 and LHCI-680 forming the peripheral antenna of PS I in C. reinhardtii (Wollman and Bennoun, 1982; Ish-Shalom and Ohad, 1983). These two complexes from Chlamydomonas have a very similar polypeptide composition although they are different with respect to Chl a/b and Chl/carotenoid ratio (Bassi et al., 1992). This is in clear contrast to vascular plants where LHCI-730 and LHCI-680 were shown not to have common polypeptides (Bassi et al., 1987).

Chapter 14 Supramolecular Organization of Chloroplast and Thylakoid Membranes

Most of the PF particles above 11 nm in size, which are mostly found on PFu in the wild type, were or complex was lacking. missing when the As a result, PFu and PFs faces could no longer be distinguished on the basis of particle size. The overall particle density on the PF faces of the mutants was similar to that of the pooled PF faces of WT (6800 in the mutant versus 5830 in WT) particles (Olive et al., 1983). It was suggested that the large PFu particles in WT resulted from the association of the two protein complexes. Size measurements on freeze-fracture replicas of isolated PS I complexes

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from pea, containing, in addition to PSI polypeptides, 110 chlorophylls/P700, reconstituted into artificial phospholipid liposomes, led to particles 11 nm in diameter (Mullet et al., 1980). A similar location for PS I reaction centers on the PFu faces in unstacked membrane domains was reported in maize (Miller, 1980) and in barley (Simpson, 1982), based on comparative studies with mutants lacking the PS I complex. In part because of the difficulty in raising antibodies to C. reinhardtii reaction centers, immunocyto­ chemical localization of PS I has proved difficult.

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Using an immunocytochemical approach with an antibody to a spinach reaction center I (RCI) protein which cross-reacted with that of C. reinhardtii, Vallon et al. (1986) observed that most of the label was found over unstacked membranes (Fig. 8B). The weak cross-reaction observed in the wild-type thin sections was proved significant because a control strain, the C3 mutant, totally devoid of the antigen, showed no significant labeling. In contrast, the same workers observed a strong reaction with spinach thylakoid membranes. It confirmed the localization of PS I reaction centers in unstacked membranes. Similar conclusions were derived from phase partition studies with spinach thylakoid membranes (Anders­ son and Hähnel, 1982).

3. Light-Harvesting Complex of PS II As discussed in the section on light-harvesting complexes (Chapter 19, Hoober et al.), the green algae possess Chl a and Chl b, carotenes and xanthophylls linked to a few specific proteins. The light harvesting proteins maintain high concentrations of pigments in a suitable orientation for maximal energy transfer to the reaction centers. Their supramolecular organization is also aimed at protecting the photosynthetic membranes from photodamage. The pigment-protein complexes are embedded in the membrane as components of the supramolecular protein complexes of PS II and PS I. According to the classification derived from studies with vascular plant chloroplasts, C. reinhardtii possesses three types of peripheral antenna complexes, the ‘minor’ antenna complexes, CP24, CP26, CP29, (Bassi and Wollman, 1991), the LHCII

Jacqueline Olive and Francis-André Wollman and LHCI antenna complexes (Delepelaire and Chua, 1981). Antenna complexes from the first and third categories participate respectively to PS II and PS I light harvesting processes. The function of LHCII has been considerably reassessed in the past decade. Originally presented as a major PS II antenna complex, at a time where the existence of the minor antenna proteins was not yet recognized, it is nowadays considered, at least in C. reinhardtii, as a regulating antenna which may associate with PS II as well as PS I depending whether the algae are in state 1 or in state 2 (Delosme et al., 1995; Chapter 30, Ohad). It follows that one may be somewhat confused in reading the original papers on freeze-fractured thylakoid membranes, because of the systematic use of LHC (or LHCII) for some particles which are more likely to correspond to the minor peripheral antenna complexes. We have tried, in the presentation below, to replace the conclusions of these older studies with our more recent knowledge. Greening studies with chloroplasts from vascular plants showed that the variability in EF particle size was, in part, due to the association of PS II core complexes with a variable amount of Chl a/b proteins (Armondetal., 1977). In wild-type Chlamydomonas, the EFs particles have widely different sizes, from 8 to 20 nm, while the EFu particles show a narrower distribution, in the smaller size range, from 6 to 15 nm (Fig. 6). This suggests that PS II centers in nonappressed regions have a smaller antenna, as has been shown in vascular plants. In PS II mutants, the PFs particle density increased upon disappearance of most of the PS II-core containing EFs particles (Wollman et al., 1980), suggesting that antenna complexes can form PFs particles when they are no longer associated with PS II centers. In two other instances, complementary changes were observed between the PFs and EFs faces, with no concomitant changes in the actual content in Chl a/b proteins (Olive et al., 1981). Either with changes in the cationic concentration of the medium in which broken cells of Chlamydomonas were resuspended, or with changes in the light intensity during growth: the larger the EFs particles, the fewer the PFs particles in high salt medium and high light conditions. Thus, the partition coefficient of some proteins—presumably antenna proteins—between the two fracture faces of the stacked membranes varied with the experimental conditions. In agreement with this observation, a C. reinhardtii mutant lacking all classes of Chl a/b containing antennae, the BF4

Chapter 14

Supramolecular Organization of Chloroplast and Thylakoid Membranes

mutant, showed a dramatic loss in PFs particle density, from 6000 particles in WT to 3800 in the mutant, together with a reduction in the size of the EFs particles (Olive et al., 1981). This indicated that Chl a/b proteins were located both as individual particles on PFs and within EFs particles, in association with the PS II reaction centers. Similar effects were reported in barley mutants (Miller et al., 1976; Simpson, 1980), although, in this case, the change in EFs particle size was less marked. This difference reflected the distinct nature of the mutations altering production of the Chl a/b proteins in the various mutants. Whereas the BF4 mutant from C. reinahrdtii has an altered content in all Chl a/b proteins, the chlorina-f2 mutant studied by Simpson (1980) retains normal amounts of the minor antenna complexes. In addition, a maize mutant with normal levels of CP29 and reduced LHCII showed no reduction in the size of EFs particles (Greene et al., 1988). A model thus emerges in which the minor antenna proteins are present together with the PS II reaction centers in the EFs particles, about 12 nm in diameter, while most of the LHCII, although it is functionally associated with the PS II core, is retained on the PFs faces upon freeze-fracture, giving rise to individual LHC particles of 8–10 nm in diameter. In some cases, these LHCII complexes can be retained, upon freeze-fracture, in association with the rest of PS II in the largest EFs particles, 18 nm in diameter. This model is consistent with the findings that CP29 is found more closely (Barbato et al., 1989) and more stably (Dunahay and Staehelin, 1987) associated with the PS II reaction center than LHCII. The presence of the PS II antenna in appressed membranes in State 1 conditions was also demon­ strated by immunocytochemistry. Using antibodies against the polypeptide p 11 (LHCII subunit), 90% of the labeling was found in appressed membranes (Fig. 9 and Table 2) (Vallon et al., 1986).

4. Cytochrome

Complex

The cytochrome complex connects Photosystem II located mainly in the stacked membrane regions to Photosystem I which is restricted to the unstacked membrane regions of the thylakoids membranes. Its presence in both domains has been established in Chlamydomonas by immunogold labeling (Olive et al., 1986, Vallon et al., 1991) and freeze-fracture analysis of mutants lacking the complex (Olive et al., 1986, 1992). A similar conclusion was drawn

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from the freeze-fracture analysis of Chlorella sorokiniana WT and mutant strains (Olive and Wollman, 1987). In two distinct C. reinhardtii mutant strains lacking the complex, a multiplicity of changes was observed both in the stacked and unstacked membrane domains (Olive et al., 1986). There was a decrease in particle densities on PFu, EFs and EFu and a change in particle sizes on PFs, EFs and EFu (Fig. 10). The largest changes were observed on the PFu and EFs faces. These modifications were consistent with the complexes in the stacked and presence of unstacked regions of the thylakoid membranes, some of them in association with the reaction centers PS I and PS II in the largest PFu and EFs particles, respectively (Olive et al., 1986). The presence of 8 nm EF particles corresponding to individual cytochrome complexes present in the stacked membrane regions was further confirmed using a set of double and triple mutants lacking PS II and cytochrome protein complexes (Olive et al., 1992).

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Jacqueline Olive and Francis-André Wollman spinach. Either antibodies unambiguously labeled both the stacked and unstacked regions of the embedded membranes from Chlamydomonas, as illustrated for Cyt f in Fig. 8C, and also from spinach (Olive et al., 1986). Similar conclusions were reached for vascular plant thylakoid membranes by a number of authors using immunocytochemical (Allred and Staehelin, 1985; Goodchild et al., 1985; Shaw and Henwood, 1985) or fractionation (Anderson, 1982; Cox and Anderson, 1982) approaches. In contrast the phase-partitioning technique gave conflicting results, with the cytochrome found only in the stroma lamellae fraction (Henry and Moller, 1981) or in the regions interfacing the stacked and unstacked membranes (Ghirardi and Melis, 1983; Barber, 1984).

5. ATP Synthase The coupling factor complex of the ATP synthase has been shown to be totally excluded from the stacked regions of the thylakoid membranes in C. reinhardtii. This has been demonstrated by an immunocytochemical study, using antibodies directed (Fig. 8D) (Vallon against the and subunits of et al., 1986). These results were confirmed by experiments with spinach membranes where only the stroma and the margins were labeled (Vallon et al., 1986). Previous analysis of vascular plant thylakoid membranes using antibody labeling, enzyme assays and destacking/restacking of thylakoid membranes had also demonstrated that the coupling factor was excluded from grana regions upon membrane stacking (Miller and Staehelin, 1976). The hydrophobic and transmembrane sector of complex has been visualized by the negatively staining them (Staehelin, 1986). When reconstituted into digalactosylipids and freezefractured, they gave rise to 9.5 nm particles falling into the size range of the PFu particles (Mörschel and Staehelin, 1983).

6. The Static Picture

This lateral distribution of cytochrome complexes along the thylakoid membranes was confirmed by immunocytochemistry using antibodies raised against either Cyt f or the Rieske protein from

In summary, the ultrastructural analysis of numerous photosynthesis mutants of Chlamydomonas led to the following conclusions as to the lateral and transverse distribution of the transmembrane protein complexes (Olive and Vallon, 1991) (Table 3). Most of the PS II reaction centers are found in the stacked membrane domains, and are retained on

Chapter 14 Supramolecular Organization of Chloroplast and Thylakoid Membranes

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the exoplasmic faces upon freeze-fracture. The PS I reaction centers, the ATP synthase and the peripheral PS I antenna are located exclusively in the unstacked membrane domain, where they give rise to freeze-fracture particles on the protoplasmic face. The distribution of the PS II peripheral antenna is the same as that of the PS II cores. A part of it partitions to the exoplasmic faces in association with the PS II cores in the same particles while the rest contributes to particle formation on the protoplasmic faces. The cytochrome is present in both stacked and unstacked membranes, mainly on PFu and EFs, in part associated in the same particles with PS II and PS I reaction centers.

B. Transmembrane Organization The vectorial nature of photosynthetic electron transport requires an asymmetric distribution of redox components in the transverse plane of thylakoid membranes. Examples of vectorial processes are proton uptake from the stroma and deposition into the lumen, water-splitting and plastocyanin oxidation which occur only at the lumenal surface whereas reduction and proton-driven ATP synthesis occur only at the stromal surface. It is now recognized that most, and perhaps all, of the intrinsic subunits in thylakoid protein complexes span the membrane and have domains at both the lumenal and stromal surfaces. Intrinsic polypeptides protruding from the membranes have been visualized by the deep-etching technique. The lumenal stacked membrane surface (ESs) contains high density of large protruding particles, each divided into two lobes, whereas the unstacked areas (ESu) show few particles only (Fig. 11). Each particle corresponds to a PS II complex, the protruding elements of which are considered structural equivalents of the OEE subunits making up part of the water splitting enzyme. In the BF25 mutant which lacks most of the OEE subunits, the large ESs particles are absent (O. Vallon, personal communication). Similarly, tetramer-like structures exposed on the lumenal surface were absent in a PS II mutant from tobacco (Miller and Cushman, 1979). In C. reinhardtii, the outer unstacked membrane

surface (PSu) is covered with large particles identified, by analogy with results obtained on higher plants (Miller and Staehelin, 1976; Oleszko and Moudrian­ akis, 1974), as the coupling factor molecules (J. Olive, unpublished). In addition to these large

246 particles, unidentified smaller particles are present which could represent a protruding part of either the LHCI-PS I complex or the cytochrome complex. The two membrane surfaces become accessible to antibodies after cell disruption through a French Press, because of the production of a mixture of inside-out and right-side-out vesicles (Andersson and Akerlund, 1978). When stacked thylakoid membranes are disrupted, they contribute to the formation of inside-out vesicles, which are enriched in PS II, whereas the right-side-out vesicles, originating from the unstacked membranes are enriched in PS I. In C. reinhardtii, cell disruption generates mostly right-side-out vesicles with a minor population of inside-out vesicles. Immunolabeling of these thylakoid membrane vesicles deposited on electron microscope grids can allow the determination of the sidedness of an antibody-binding site. A monoclonal antibody directed against Cyt f, labeled exclusively the insideout vesicles (Fig. 12, O. Vallon, unpublished) indicating that the epitope was accessible on the lumenal surface. This is consistent with the transmembrane organization of the protein, most of which is located in the lumen (Chapter 24, Wollman). The accessibility of an antibody directed against P6 (CP43) has also been observed on one face (undetermined) of the WT membrane vesicles (Vallon, 1986).

IV. Dynamic Aspects of Thylakoid Membrane Organization

A. The Stacking-Destacking Process As mentioned earlier, thylakoid membranes of green algae have well-defined stacked and unstacked domains with distinct protein contents and supramolecular structures, even though their organization is less regular than in vascular plants. Moreover, the ratio of appressed over non-appressed membranes has been shown to vary in mutant strains, depending on the inability of the mutants to synthesize active components of the photosynthetic electron transport chain (Goodenough and Levine, 1969). In particular, in PS I mutants, about 90% of the membranes were appressed versus 65% in WT. By contrast, the ac-115 and ac-141 mutants lacking and Q (the quencher of PS II fluor­ active Cyt escence) contain long, single discs in their chloro-

Jacqueline Olive and Francis-André Wollman

plast (Goodenough and Levine, 1969). The photosynthetic proteins are therefore segre­ gated between these two domains. Barber and coworkers performed an analysis ofthe surface charge density on the thylakoid membranes (Nakatani and Barber, 1980; Barber, 1982) and they reached the conclusion that the unstacked membrane regions carried a higher negative surface charge density than the surfaces in contact in the stacked domains (Barber, 1980; Chow and Barber, 1980; Yamamoto and Ke, 1982). These conclusions were further used in the interpretation of the formation of the two membrane domains, the stacked and unstacked thylakoid membrane regions. Experimentally, it was shown to be a reversible process in vitro: destacking and restacking can be promoted by decreasing then increasing again the concentration of cations. These were proposed to screen the electrostatic surface charges, responsible for repulsive forces, and allow the membranes to stack, due to van der Waals attractive forces. Conversely, removal of cations leads to an increase in repulsive forces due to the net negative charges at the surface of adjacent membranes and to a destacking process. Early studies with Chlamydomonas contributed greatly to the understanding of the relationship between membrane stacking, lateral segregation of complexes and changes in supramolecular organi­ zation. In a pioneering study, Ojakian and Satir (1974) showed by a freeze-fracture analysis of the

Chapter 14 Supramolecular Organization of Chloroplast and Thylakoid Membranes thylakoid membranes from C. reinhardtii that, when destacked by a transfer to zwitterionic buffers at low ionic strength (approximately 50 mM), the lateral segregation of intramembrane particles between grana and stroma membrane regions was lost, due to their intermixing and randomization. Conversely, they showed that segregation of the particles was fully restored when unstacked thylakoids were resuspended in a high salt buffer (more than 3 mM for divalent cations or more than 100 mM for monovalent cations). Similar results were obtained subsequently with vascular plant thylakoid membranes (Staehelin, 1976). The kinetics of light scattering changes which accompany membrane restacking were then studied in C. reinhardtii (Wollman and Diner, 1980). About 80% of the changes in light scatter were completed within 2 s after unstacked membranes were resus­ pended in the cation-containing stacking buffer. In contrast, cation-induced fluorescence changes, which reflect a segregation between PS II and PS I (reviewed in Barber, 1982), showed much slower kinetics, in the time range of several minutes. The segregation of intramembrane particles was not studied in Chlamydomonas. However, similar studies with vascular plant thylakoid membranes showed that complete segregation of the particles requires several minutes and was subsequent to membrane restacking (Staehelin, 1976; Briantais et al., 1983). Thus, thylakoid membrane stacking, which requires screening of surface charges to occur, causes a subsequent lateral segregation of the transmembrane complexes between the two domains, with most of the PS II localized in the stacked domains and PS I in the unstacked domains.

B. State Transition: PhosphorylationDependent Changes As discussed in greater detail by I. Ohad (Chapter 30), plants and algae subjected to light of different spectral quality, undergo state transition, a regulatory mechanism that controls light-energy distribution between the two photosystems (Bonaventura and Myers, 1969; Murata, 1969). It has been suggested that the changes in light energy distribution occur through a lateral displacement of a Chl a/b mobile antenna along the thylakoid membranes (for a review see Allen, 1992). This protein migration would be caused by the reversible phosphorylation of a fraction of the LHCII protein. It was thus proposed that, in

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state 2, phosphorylated antenna proteins move from the Photosystem II-enriched stacked domains to the Photosystem I-enriched unstacked domains. Dephos­ phorylation of the same subunits in state 1 allows the return of these peripheral antenna complexes to the grana regions (Staehelin et al., 1982; Kyle et al., 1983; Barber, 1986). This concept of lateral migration has been studied experimentally in C. reinhardtii, in which particularly extensive state transitions were observed in vivo (Wollman and Delepelaire, 1984). The cells were placed in state 1 or in state 2 and fixed in either state by p-benzoquinone treatment (Bulté and Wollman, 1990). An immunocytochemical study showed extensive changes in the distribution of cytochrome complexes and LHCII between the stacked and unstacked membranes (Vallon et al., 1991). As shown in Table 4, the labeling of LHCII and cytochrome f increased for unstacked membranes in state 2 versus state 1, while the labeling of the core antenna protein P6 (CP43) did not change. Thus, both the lightharvesting antenna and the electron-transport chain undergo a deep reorganization during state transitions. A similar reorganization was observed in maize (Vallon et al., 1991). The driving force for the movement of LHCII is suggested to arise from its change in phosphorylation but there has been no report of reversible phosphory­ lation of the cytochrome complex of C. reinhardtii upon state transitions. However, a cytochrome associated protein of 19.5 kDa (Lemaire et al., 1986) may be part of the process since it has been detected as a phosphoprotein in state 2 (C. de Vitry and F.-A. Wollman, unpublished). In contrast, the minor Chl a/ b antenna protein CP29, which becomes heavily phosphorylated in Chlamydomonas in state 2 (Wollman and Delepelaire, 1984; Bassi and Wollman, 1991) does not undergo lateral displacement upon state transitions.

C. Fusion of Thylakoid Membranes During Sexual Reproduction During the first steps of zygote formation, a number of intracellular rearrangements can be observed including fusion of the nuclei and chloroplasts (Friedman et al., 1968; Cavalier-Smith, 1970, 1975). The question then arises as to how the inner thylakoid membrane system behaves subsequent to chloroplast fusion. Baldan et al. (1991) used two mutant parental strains, one being deficient in the PS I complex and

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Jacqueline Olive and Francis-André Wollman V. Biogenesis

A. Greening: Synthesis of Membrane Complexes

the other in the PS II complex, to follow the possible fusion of their thylakoid membranes upon zygote formation. In order to avoid a genetic comple­ mentation and thus de novo synthesis of the missing chloroplast-encoded polypeptides, the gametes were crossed in the presence of chloramphenicol that inhibits chloroplast translation. Before fusion, the thylakoid membranes originating from the PS II minus-gamete could be distinguished easily from those of the PS I minus-gamete, by their lack in PS II-containing EF particles upon freezefracture as well as by the absence of membrane labeling with antibodies directed against PS II subunits. About 15 h after mating, a complete fusion of the thylakoid membranes from the two parental gametes occurred in the zygote, accompanied by a redistribution of the membrane protein complexes in the fused membranes. The histograms ofthe EFs and EFu particle sizes in the zygote were bimodal with two maxima, each corresponding to that in one type of parental gametes. Also, EF particle densities were intermediate between those recorded in the parental gametes. Immunolabeling of the zygote sections showed that, at an early stage, the thylakoid membranes from the two mutants could still be distinguished on the basis of their differential labeling with antibodies directed against PS II subunits (Fig. 13A). In older zygotes, thylakoid membranes from all regions of the chloroplast were labeled with the antibody, indicating that a redistribution of PS II complexes occurred within the fused membranes (Fig. 13B). Membrane fusion and particle redistribution restored a full functional interaction between PS II and PS I from the two mutant strains as demonstrated by the recovery of linear photosynthetic electron transport in the zygote (Baldan et al., 1991).

The study ofthe biogenesis of membrane components in C. reinhardtii has been facilitated by the use ofthe yellow mutant y-1, unable to synthesize chlorophyll in the dark, which was isolated by Sager and Palade (1954). When grown heterotrophically in the light, the mutant is indistinguishable from the wild type. When grown in the dark, the chloroplast intralamellar system of y-1 is gradually disorganized and drastically reduced in size (Ohad et al., 1967a). Growth in the dark causes loss of thylakoid membranes by dilution but not dedifferentiation ofthe plastid to a proplastid or etioplast(Ohad et al., 1967a; Hoober e tal., 1991). Dark-grown y-1 cells contain less than 5% of the chlorophyll found in light-grown cells. These darkgrown cells offer an appropriate starting point for examining thylakoid membrane formation. Upon return to light at 25 °C,y-1 cells accumulated chlorophyll, after a lag of 1 to 2 h, and developed thylakoid membranes (Hoober and Stegeman, 1973; see also Chapter 19, Hoober et al.). In contrast, the lag disappeared when cells were placed at higher temperature, 37 to 40 °C (Hoober and Stegeman, 1976; Maloney et al., 1989). Chlorophyll concen­ tration, thylakoid membranes and photosynthetic capacity increased in parallel, at linear rates, over a 6–8 h period (Ohad et al., 1967b; Hoober and Stegeman, 1976) when the cells were exposed to light. During this period, no cell division was observed: all the biosynthetic and morphogenetic activities ofthe greening cell are related to chloroplast differentiation. Initially, the system was constituted of single thylakoid membranes. Membrane appres­ sion was a later event in the process and was not required for detection of photosynthetic activity (Ohad et al., 1967b). The rate of increase in PS I activity initially exceeded that of PS II. Since fluorescence remained low (an indication that the PQ pool was oxidized), this indicates that antenna connection to the reaction centers and electron transport chain between the photosystems were fully functional in these membranes (Hoober et al., 1994). Rapid assembly of thylakoid membranes during greening of y-1 cells provided the opportunity to determine directly the site of formation of the membrane. Remnants ofthe chloroplastic disc system are always present which may act as primordia during the greening process (Ohad et al., 1967b). However,

Chapter 14 Supramolecular Organization of Chloroplast and Thylakoid Membranes

electron micrographs of cells exposed to light for only 5 min. revealed extensive arrays of membrane material extending from the inner envelope membrane, suggesting that thylakoid membranes develop by local expansion and infolding ofthe inner membrane (Hoober et al., 1991, 1994). The y-1 mutant, when allowed to green in the presence of chloramphenicol, an inhibitor of protein synthesis on 70s ribosomes, contained Chl a and Chl b but lacked certain membrane polypeptides (Eytan and Ohad, 1970; Bar-Nun and Ohad, 1977) and photosynthetic electron transport. Miller and Ohad (1978) showed that fewer particles were present on the EF fracture face and that they were significantly smaller (11.5 nm) in presence than in the absence of chloramphenicol. A substantial increase in EF particle number and size was observed upon removal of chloramphenicol, correlated with insertion of newly made PS II centers (Cahen et al., 1977). This observation is consistent with the above-described model that most of the EF particles contained PS II centers associated with some peripheral antenna.

B. Assembly of PS II The assembly of the PS II subunits has been studied in some details at the ultrastructural level in C. reinhardtii. This was achieved by comparing various PS II mutants, whose content in specific residual PS II subunits was known with sufficient accuracy that it could be correlated with structural changes. As described in Chapters 15 (Erickson) and 16 (Ruffle and Sayre), the PS II core complex consists

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of the two large reaction center subunits, D1 and D2, encoded by the psbA and psbD chloroplast genes and two core antenna proteins, P5 (CP47) and P6 (CP43), encoded by the psbB and psbC chloroplast genes. Three nucleus-encoded subunits, involved in oxygen evolution, OEE1, OEE2 and OEE3, associate with the PS II core on the lumenal side of the membranes. A number of smaller subunits, which will not be mentioned here, also participate in the assembly of the fully active PS II oligomeric protein. De Vitry et al. (1989) performed an immuno­ cytochemical study of several PS II mutants which were deficient in the synthesis of only one PS II core subunit, but displayed a pleiotropic decrease of all other transmembrane PS II subunits. However, about 10% ofthe WT amount of P5 and some accumulation of D1 and D2 were revealed in the thylakoid membranes of the mutants lacking P6 by application of immunoblotting methods. Similarly 10% of P6 was accumulated in the mutants lacking D1 or D2 (Fig. 14,A and B)(De Vitry et al., 1989). A P5/D1/ D2 subcomplex and the unassembled P6 subunits were inserted independently in the thylakoid membranes. These two parts of the PS II core each had the ability to segregate independently in the stacked membrane regions of the thylakoid mem­ branes. Interestingly, the OEE subunits accumulated in the lumen despite the absence of PS II cores. They were immunodetected as associated with the thylakoid membranes, but in a loose binding state since they were lost during purification of the membranes. The partially assembled P5/D1/D2 subcomplex and the individual P6 subunit each gave rise to

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particles which could be identified upon freezefracture on the EF faces of the thylakoid membranes from PS II mutants. Their densities were 206 and 240 respectively versus 1500 in WT (Olive particles et al., 1992). Comparison of the respective densities of EFs and EFu particles in PS II single mutants and PS II double mutants, totally devoid of the whole set of PS II core subunits, showed that 78% of the EFs particles and 64% of the EFu particles correspond to PS II cores in the WT. Most ofthe EF particles which remain in the PS II double mutant can be attributed to complex, as indicated by the analysis of a the Cyt triple mutant, deficient in all the PS II subunits and in Cyt (Fig. 15). The PS II subcomplexes identified by freezefracturing (Olive et al., 1992) demonstrated the same lateral segregation in stacked membrane domains, as they did when viewed by immunodecoration (de Vitry et al., 1989). Strangely enough, lateral migration of the PS II subcomplexes from the unstacked membrane regions to the stacked domains was not observed in double mutant strains carrying a single PS II mutation and a mutation preventing production complexes. In these strains, the of cytochrome

Jacqueline Olive and Francis-André Wollman

unstacked domains retained most ofthe PS II subunitcontaining EF particles (Olive et al., 1992). This observation suggests that the mechanism of translocation of neosynthesized PS II subunits to the stacked membrane domains requires either a direct complexes or some interaction with cytochrome elements ofa transition to state 2 that cytochrome mutants, which are blocked in state 1 (Wollman and Lemaire, 1988), cannot perform. This latter hypothesis points to a possible requirement for LHCII reversible phosphorylation in the movement of PS II subunits to the stacked membrane domains.

VI. Conclusion Fractionation of thylakoid membranes, performed by breakage or mild detergent solubilization followed by differential centrifugation or phase partitioning, yielded valuable information on the existence of various domains showing membrane protein heterogeneity (for a review, see Albertsson, 1988). However most of our knowledge on the supra­ molecular organization ofthe photosynthetic proteins comes from the two techniques that we have described in some detail here, the immunocytochemical labeling of membranes and their freeze-fracturing. They allowed the characterization in situ of the structural and biochemical heterogeneity between stacked and unstacked membrane regions. A large part of this experimental effort has focused on Chlamydomonas because of its genetics which opened the way to comparative biochemical and ultrastructural analysis

Chapter 14 Supramolecular Organization of Chloroplast and Thylakoid Membranes of mutant and wild-type strains. Mutant strains, in turn, served as controls for immunocytochemistry and freeze-fracturing. At the present time, a drawback of these approaches is that they are difficult to combine efficiently. Although it has been reported that antibody-coupled to colloidal gold can react with freeze-fracture replicas retaining some biological material (Fujimoto, 1995), the technique has not yet been applied successfully to the direct molecular identification of the freeze-fracture particles observed in the thylakoid membranes. Yet, this identification remains a challenging issue for the next years to those who wish to understand the dynamics of the organization of membrane proteins and also to those who are developing methods to solve their threedimensional structures.

Acknowledgment We thank O. Vallon for critical reading of the manuscript.

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Chapter 14 Supramolecular Organization of Chloroplast and Thylakoid Membranes process in Chlamydomonas. Photosynth Res 39: 15–31 Ish-Shalom D and Ohad I (1983) Organization of chlorophyllprotein complexes of Photosystem I in Chlamydomonas reinhardtii. Biochim Biophys Acta 722: 498–507 Kerby NW and Evans LV (1978) Isolation and partial characterization of pyrenoids from the brown alga Pilayella littoralis. Planta 142: 91–95 Kuchitsu K, Tsuzuki M and Miyachi S (1988) Characterization of the pyrenoid isolated from unicellular green alga C. reinhardtii: Paniculate form of rubisco protein. Protoplasma 144: 17–24 Kyle DJ, Staehelin LA and Arntzen CJ (1983) Lateral mobility of the light harvesting complex in chloroplast membranes controls excitation energy distribution in higher plants. Arch Biochem Biophys 222:527–541 Lacambra M, Larsen U, Olive J, Bennoun P and Wollman FA (1984) Characterization of the thylakoid membranes of wild type and mutants of Chorella sorokiniana. Photobiochem Photobiophys 8: 191–205 Lemaire C, Girard-Bascou J, Wollman FA and Bennoun P complex. Charac­ (1986) Studies on the cytochrome terization of the complex subunits in Chlamydomonas reinhardtii. Biochim Biophys Acta 851: 229–238 Lembi CA and Lang NJ (1965) Electron microscopy of Carteria and Chlamydomonas. Amer J Bot 52: 464–467 Maloney MA, Hoober JK and Marks DB (1989) Kinetics of chlorophyll accumulation and formation ofchlorophyll-protein complexes during greening of Chlamydomonas reinhardtii y­ 1 at 30°C. Plant Physiol 91: 1100–1106 Melkonian M and Robenek H (1980) Eyespot of C. reinhardtii: a freeze-fracture study. J Ultrastruct Res 72: 90–102 Miller KR (1980). A chloroplast membrane lacking Photosystem I. Changes in unstacked membrane regions. Biochim Biophys Acta 592: 143–152. Miller KR and Cushman RA (1979) A chloroplast membrane lacking Photosystem II. Thylakoid stacking in the absence of the Photosystem II particles. Biochim Biophys Acta 546: 481– 497 Miller KR and Ohad I (1978) Chloroplast membrane biogenesis in Chlamydomonas: correlation between the formation of membrane components and membrane structure. Cell Biol Reports 2: 537–549 Miller KR and Staehelin LA (1973) Fine structure of the chloroplast membranes of Euglena gracilis as revealed by freeze-cleaving and deep-etching. Protoplasma 77: 55–78 Miller KR and Staehelin LA (1976) Analysis of the thylakoid outer surface: Coupling factor is limited to unstacked membrane regions. J Cell Biol 68: 30–47 Miller KR, Miller GJ and McIntyre KR (1976) The lightharvesting chlorophyll-protein complex of Photosystem II. J Cell Biol 71: 624–638 Mörschel E and Staehelin LA (1983) Reconstitution of Cyt and ATPsynthase complexes into phospholipid and galactolipid liposomes. J Cell Biol 97: 301–310 Mullet JE, Burke JJ and Arntzen CJ (1980) A developmental study of Photosystem I peripheral chlorophyll proteins. Plant Physiol 65: 823–827 M urata N (1969) Control ofexcitation transfer in photosynthesis. I. Light-induced change of chlorophyll a fluorescence in Porphyridium cruentum. Biochim Biophys Acta 172: 242– 251

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Mustárdy L (1996) Development ofthylakoid membrane stacking. In: Ort DR and Yocum CF (eds) Oxygenic Photosynthesis: The Light Reactions, pp 59–68. Kluwer Academic Publishers, Dordrecht Nakamura K, Bray DF, Costerton JW and Wagenaar EB (1973) Infrastructure of Chlamydomonas eugametos as revealed by freeze-etching: cell wall, plasmalemma and chloroplast membrane. Can J Bot 51: 817–819 Nakatami NY and Barber J (1980) Further studies ofthe thylakoid membrane surface charges by particle electrophoresis. Biochim Biophys Acta 591: 82–91 Ohad I, Siekevitz P and Palade GE (1967a) Biogenesis of chloroplast membranes. I. Plastid dedifferentiation in a darkgrown algal mutant (Chlamydomonas reinhardtii). J Cell Biol 35: 521–552 Ohad I, Siekevitz P and Palade GE (1967b) Biogenesis of chloroplast membranes. II Plastid differentiation during greening of a dark-grown algal mutant (C. reinhardtii). J Cell Biol 35: 553–584 Ojakian G and Satir P (1974) Particle movements in chloroplast membranes: quantitative measurements of membrane fluidity by the freeze-fracture technique. Proc Natl Acad Sci USA 21: 2052–2056 Oleszko S and Moudrianakis EN (1974) The visualization of the photosynthetic coupling factor in embedded spinach chloroplasts. J Cell Biol 63: 936–948 Olive J and Vallon O (1991) Structural organization of the thylakoid membranes: freeze-fracture and immunocyto­ chemical analysis. J Electron Microsc Techn 18: 360–374 Olive J and Wollman FA (1987) Localization of the complex by freeze-fracture analysis of the Chlamydomonas reinhardtii and Chlorella sorokiniana mutants lacking in this complex. In: Biggins, J (ed), Progress in Photosynthesis Res Vol II, pp 325– 328, Martinus Nijhoff Publishers, Dordrecht, The Netherlands Olive J, Wollman FA, Bennoun P and Recouvreur M (1979) Ultrastructure-function relationship in Chlamydomonas reinhardtii thylakoids by means of a comparison between the wild type and the F34 mutant which lacks the Photosystem II reaction center. Mol Biol Rep 5: 139–143 Olive J, Wollman FA, Bennoun P and Recouvreur M (1981) infrastructure of thylakoid membranes in C. reinhardtii. Evidence for variations in the partition coefficient of the lightharvesting complex-containing particles upon membrane fracture. Arch Biochem Biophys 208: 456–467 Olive J, Wollman FA, Bennoun P and Recouvreur M (1983). Localization of the core and peripheral antennae of Photosystem I in the thylakoid membranes of Chlamydomonas reinhardtii. Biol Cell 48: 81–84 Olive J, Vallon O, Wollman FA, Recouvreur M and Bennoun P (1986) Studies on the cytochrome complex. II. Localization of the complex in the thylakoid membranes from spinach and Chlamydomonas reinhardtii by immunocytochemistry and freeze-fracture analysis of mutants. Biochim Biophys Acta 851:239–248 Olive J, Recouvreur M, Girard-Bascou J and Wollman FA (1992) Further identification of the exoplasmic face particles on the freeze-fractured thylakoid membranes: A study using double and triple mutants from Chlamydomonas reinhardtii lacking various Photosystem II subunits and the cytochrome complex. European J Cell Biol 59: 176–186 Rochaix JD, Kuchka M, Mayfield S, Girard-Bascou J and Bennoun

254 P (1989) Nuclear and chloroplast mutations affect the synthesis or stability of the chloroplast psbC gene product in Chlamydomonas reinhardtii. EMBO J 8: 1013–1021 Roberts K, Shaw PJ and Hills GJ (1972) Structure, composition and morphogenesis of the cell wall of C. reinhardtii. I. Ultrastructure and preliminary chemical analysis. J Ultrastruct Res 40: 599–613 Sager R and Palade GE (1954) Chloroplast structure in green and yellow strains of Chlamydomonas. Exp Cell Res 7: 584–588 Sager R and Palade GE (1957) Structure and development of the chloroplast in Chlamydomonas. J Biophys Biochem Cytol 3: 463–487 Sato H, Okada M, Nakayama K and Miyaji K (1984) Purification and further characterization ofpyrenoid proteins and ribulose­ 1 -5-biphosphate carboxylase-oxygenase from the green alga Bryopsis maxima. Plant Cell Physiol 25: 1205–1214. Schötz F, Bathelt H, Arnold CG and Schimmer O (1972) Die Architektur und Organisation der Chlamydomonas-Zelle. Ergebnisse der Elektronen Mikroskopie von Serienschnitten und der daraus resultierenden dreidimensionalen Rekon­ struktion Protoplasma75: 229–254 Shaw PJ and Henwood JA (1985) Immuno-gold localization of cytochrome f, light-harvesting complex, ATP synthase and ribulose 1,5-bisphosphate carboxylase/oxygenase. Planta 165: 333–339 Simpson DJ (1980) Freeze-fracture studies on barley plastid membranes. IV. Analysis of freeze-fracture particle size and shape. Carlsberg Res Commun 45: 201–210 Simpson DJ (1982) Freeze-fracture studies on barley plastid membranes. V. a Photosystem I mutant. Carlsberg Res Comm 47: 215–225 Simpson DJ, Vallon O and Von Wettstein D (1989) Freezefracture studies on barley plastid membranes. VIII. In viridis 115 , a mutant completely lacking Photosystem I, oxygen evolution enhancer 1 (OEE1) and the subunit of cytochrome b-559 accumulate in appressed thylakoids. Biochim Biophys Acta 975: 164–174 Sjolung RD and Smith DD (1974) Freeze-fracture studies of photosynthetically deficient supergranal chloroplast in tissue cultures containing virus-like particles. J Cell Biol 60: 285– 292 Staehelin LA (1975) Chloroplast membrane structure. Intramembranous particles of different sizes make contact in stacked membrane regions. Biochim Biophys Acta 408: 1–11 Staehelin LA (1976) Reversible particle movements associated with unstacking and restacking of chloroplast membranes in vitro. J Cell Biol 71: 136–158 Staehelin LA (1986) Chloroplast structure and supramolecular organization of photosynthetic membranes. In: Staehelin LA, Arntzen CJ (ed), Photosynthesis III, pp 1–84, Springer-Verlag Berlin Staehlin LA and van der Staay WM (1996) Structure, composition, functional organization and dynamic properties of thylakoid membranes. In: Ort DR and Yocum CF (eds) Oxygenic Photosynthesis: The Light Reactions, pp 11–30. Kluwer Academic Publishers, Dordrecht

Jacqueline Olive and Francis-André Wollman Staehelin LA, Armond PA and Miller KR (1977) Chloroplast membrane organization at the supramolecular level and its functional implications. Brookhaven Symp Biol 28: 278–315 Staehelin LA, Kyle DJ and Arntzen CJ (1982) Spillover is mediated by reversible migration of LHCP between grana and stroma thylakoids. Plant Physiol 69: 69–81 Vallon O (1986) Organisation supramoléculaire des domaines membranaires engagés dans la communication intercellulaire (cristallin de bovidé) et dans la photosynthèse (membrane des thylacoïdes): Étude immunocytochimique. PhD dissertation, Université Paris VI. Vallon O, Wollman FA and Olive J (1985) Distribution of intrinsic and extrinsic subunits of the PS II protein complex between appressed and non-appressed regions ofthe thylakoid membrane: an immunocytochemical study. FEBS Lett 183: 245–250 Vallon O, Wollman FA and Olive J (1986) Lateral distribution of the main protein complexes of the photosynthetic apparatus in C. reinhardtii and in spinach: An immunocytochemical study using intact thylakoid membranes and a PS II enriched membrane preparation. Photobiochem Photobiophys 12: 203– 220 Vallon O, Hoyer-Hansen G and Simpson DJ (1987) Photo­ system II and cytochrome b-559 in the stroma lamellae of barley chloroplasts. Carlsberg Res Commun 52: 405–421 Vallon O, Bulté L, Dainese P, Olive J, Bassi R and Wollman FA complexes (1991) Lateral redistribution of cytochrome along thylakoid membranes upon state transitions. Proc Natl Acad Sci USA 88: 8262–8266 Walne PL and Arnott HJ (1967) The comparative ultrastaicture and possible function of eyespots: Euglena granulata and Chlamydomonas eugametos. Planta 77: 325–353 Wollman FA and Bennoun P (1982) A new chlorophyll-protein complex related to Photosystem I in C. reinhardtii. Biochim Biophys Acta 680: 352–360 Wollman FA and Delepelaire P (1984) Correlation between changes in light energy distribution and changes in thylakoid membrane polypeptide phosphorylation in C. reinhardtii. J Cell Biol 98: 1–7 Wollman FA and Diner BA (1980) Cation control of fluorescence emission light scatter, and membrane stacking in pigment mutants of Chlamydomonas reinhardtii. Arch Biochem Biophys 201: 646–649 Wollman FA and Lemaire C (1988) Studies on kinase-controlled state transitions in Photosystem II and mutants from Chlamydomonas reinhardtii which lack quinone-binding proteins. Biochim Biophys Acta 933: 85–94 Wollman FA, Olive J, Bennoun P and Recouvreur M (1980) Organization of the Photosystem II centers and their asso­ ciated antennae in the thylakoid membranes: A comparative ultrastructural, biochemical and biophysical study of Chlamydomonas wild-type and mutants lacking in Photo­ system II reaction centers. J Cell Biol 87: 728–735 Yamamoto Y and Ke B (1981) Membrane surface electric properties of Triton-fractionated spinach subchloroplast fragments. Biochim Biophys Acta 636: 175–184

Chapter 15 Assembly of Photosystem II Jeanne Marie Erickson

Departments of Biochemistry and Plant Pathology, 1-87 Agriculture Building,

University of Missouri, Columbia, MO 65211, U.S.A.

Summary 255

I. Introduction 256

II. Developmental Biogenesis of Photosystem II 257

A. Plants 257

B. Algae 259

III. Assembly of Photosystem II Complexes 260

A. Migration of Photosystem II in the Thylakoid Membrane: PS II Damage and Repair 260

B. Photosystem II Assembly in Chlamydomonas 260

1. Three Classes of Mutants Defective in the Assembly or Stability of the PS II Core 260

2. Stepwise Assembly of Photosystem II Intrinsic and Extrinsic Membrane Subunits 262

3. Assembly of Nascent D1 and the OEC Polypeptides with Other PS II Subunits 263

4. Assembly of the PS II Redox Components 264

C. Photosystem II Assembly Intermediates: In Organello and In Vitro 265

1. The Synthesis and Fate of PS II Core Subunits 265

2. D2 as a ‘Docking’-Like Protein Required for D1 Synthesis and/or Stability 266

3. Stepwise Assembly of the Photosystem II Polypeptides in Appressed and

267

Non-appressed Thylakoid Membranes 4. Posttranslational Import of Chimeric D1 into Isolated Chloroplasts and Assembly with PS II 268

D. Processing of the D1 Polypeptide Carboxyl-Terminus is Not a Prerequisite for PS II Assembly 268 270

IV. Assembly of the Extrinsic Membrane Polypeptides of the PS II Oxygen-Evolving Complex A. In Organello Assembly of the OEC Polypeptides with the PS II Core Complex 270

B. In Vivo Requirements for the Extrinsic Membrane Polypeptides of the OEC 272

273

V. Assembly of Manganese: The Catalytic Center of the Oxygen-Evolving Complex A. Photoactivation: Assembly of the Functional Manganese Cluster 273

B. A Role for Bicarbonate in Assembly of the OEC 276

C. Is Oxygen Required for Assembly of the OEC? 276

Acknowledgments 277

277

References

Summary Photosystem II (PS II) is a pigment-protein complex located in the chloroplast thylakoid membrane of plants and algae which acts as a light-driven water-splitting enzyme. At least 16 chloroplast- and 11 nucleus-encoded polypeptide subunits, several types of pigments, and lipid, metal and ion cofactors have been identified as constituents of the chloroplast PS II. Water is split in the oxygen-evolving complex (OEC) of PS II, where a catalytic tetranuclear cluster of manganese ions is required for the photodriven extraction of electrons from water and the release of protons and molecular oxygen. The exciton and electron transfer reactions of PS II are determined by the properties of the redox components themselves, as well as the spatial organization of all

J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 255–285. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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Jeanne Marie Erickson

cofactors within the tertiary structure of the protein environment to which they are bound. Correct assembly and conformation of the photosystem polypeptides, pigments and cofactors is critical for photosystem function and stability. Such assembly depends on the concerted expression of both nuclear and chloroplast genes, with gene expression regulated at the transcriptional, translational and posttranslational levels in response to developmental and environmental cues. Details of PS II assembly have been elucidated through in vivo, in vitro and in organello studies. In developing chloroplasts, PS II polypeptides and pigments are synthesized and assemble during light-induced biogenesis of the thylakoid membrane. In mature chloroplasts, photodamage to PS II subunits, particularly the D1 polypeptide of the PS II reaction center, requires the replacement of damaged subunits and reassembly of PS II complexes. This chapter focuses on the assembly and stability of the PS II polypeptides, with particular emphasis on investigations using the unicellular green alga Chlamydomonas reinhardtii. In addition, the light-dependent activation of the PS II OEC is reviewed. This process, termed photoactivation, involves the assembly of the tetranuclear manganese cluster central to OEC function.

I. Introduction Photosystem II (PS II) is a large, multi-subunit pigment-protein complex located in the chloroplast thylakoid membrane of plants and eukaryotic algae. PS II functions as a light-driven water-splitting enzyme, releasing molecular oxygen and hydrogen ions, and providing electrons for the photosynthetic electron transfer chain (reviewed in Rutherford, 1989). At least 27 different polypeptide subunits, several types of pigments, and lipid, metal and ion cofactors have been identified as constituents of the chloroplast PS II and its antenna complex (Fig. 1, Table 1). The nuclear and chloroplast genes encoding PS II subunits, the subunit organization within PS II, and their functions and associated cofactors and pigments are outlined in Table 1. PS II genes and polypeptides have been reviewed extensively (Erickson and Rochaix, 1992; Ikeuchi, 1992; Bricker and Ghano­ takis, 1996; Hankamer et al., 1997; Chapter 16, Ruffle and Sayre). The subunits of the reaction center (RC), the core complex, and the PS II light harvesting complex (LHCII) are intrinsic membrane poly­ peptides, while the subunits of the oxygen-evolving complex (OEC) are extrinsic membrane polypeptides. The D1 and D2 polypeptides of the PS II RC contain and and the redox-active tyrosine residues Abbreviations: ADP – adenosine monophosphate; ATP – adenosine triphosphate; Cyt – cytochrome; D1 – PS II reaction center polypeptide D1; D2 – PS II reaction center polypeptide D2; EPR – electron paramagnetic resonance; LHCII – lightharvesting complex of Photosystem II; OEC – oxygen-evolving – PS II reaction center chlorophyll; PAGE – complex; polyacrylamide gel electrophoresis; PS I – Photosystem I; PS II – Photosystem I I ; – primary quinone acceptor of PS II; – secondary quinone acceptor of PS II; RC – reaction center; redox-active tyrosine 160 of the D2 polypeptide; – redoxactive tyrosine 161 of the D1 polypeptied

bind the redox-active molecules of pheophytin, a non-heme iron, the and plastoquinones, manganese, and the chlorophyll molecules, including P680, essential for PS II photochemistry (Table 1, Fig. 1). As such, these two polypeptides play a central role in PS II assembly and function. The polypeptide subunits of PS II are generally thought to be present in a 1:1 molar ratio (reviewed in Seidler, 1966). However, there is some evidence that a single (i.e., monomer) PS II core complex may subunits contain two copies each of the Cyt (reviewed in Whitmarsh and Pakrasi, 1996), the 22 kDa S polypeptide, PS II-S (Funk et al., 1995), and the extrinsic OEC subunits (Xu and Bricker, 1992; Bricker and Ghanotakis, 1996). Recent reports by Yocum and coworkers (Betts et al., 1996b, 1997) provide the most compelling genetic and biochemical evidence in support of a model with two OEE1 polypeptides per chloroplast PS II RC. Models for the PS II RC have been made (Trebst, 1986; Santini et al., 1994; Svensson et al., 1996; Xiong et al., 1996, 1998) based on the crystal structures of reaction centers isolated from photosynthetic bacteria. A twodimensional crystal structure of the spinach PS II RC has been resolved at the level of 8 Å (Rhee et al., 1997). For further detail on PS II structure and function, the reader is referred to recent reviews (Diner and Babcock, 1996; Satoh, 1996; Barber et al., 1997; Chapter 16, Ruffle and Sayre). Other catalytic proteins, including catalase and polyphenol oxidase (Sheptovitsky and Brudvig, 1997) appear to be associated at substoichiometric levels with PS II membranes, and may play some role in maintaining a chemical environment favorable to PS II function. Assembly of a multisubunit membrane complex such as PS II, which contains at least 16 subunits translated in the chloroplast, and 11 subunits

Chapter 15 Assembly of Photosystem II

translated in the cytoplasm, imported into the chloroplast and localized to the thylakoid membrane or thylakoid lumen, is clearly a complicated process. Assembly can be affected by levels of transcription and translation, protein processing and modification, import of metal cofactors, synthesis of pigment cofactors, temperature, light quantity, and light quality. The focus of this chapter is the assembly of the PS II core complex and the process of photoactivation, i.e., the light-dependent assembly of the catalytic manganese cluster of the OEC. Emphasis will be given to assembly of PS II polypeptides in the chloroplast thylakoid membrane of plants and of the unicellular green alga, Chlamydomonas reinhardtii, which provides an excellent model system for the study of photo­ synthesis (Rochaix, 1995). Other aspects of the assembly of thylakoid membrane complexes, including LHCII assembly, have been reviewed (Cohen et al., 1995; Webber and Baker, 1996;

257

Chapters 19, Hoober et al.; 25, Strotmann et al.; 17, Webber and Bingham; 24, Wollman).

II. Developmental Biogenesis of Photosystem II

A. Plants In most plants, chlorophyll biosynthesis and plastid development are light-dependent processes that occur during seedling development or during greening of dark-grown, etiolated plants (Mullet, 1988; Barkan et al., 1995). Development of etioplasts into chloroplasts is accompanied by the biogenesis and organization of the thylakoid membrane, including the synthesis and assembly of PS II components. PS II biogenesis is dependent on processes directed by both the nuclear and plastid genomes (Taylor, 1989; Rochaix, 1992, 1996; Mayfield et al., 1995;

258

Jeanne Marie Erickson

Chapter 15 Assembly of Photosystem II Roell and Gruissem, 1996; Goldschmidt-Clermont, 1998; Chapter 12, Hauser et al.; Chapter 10, Stern and Drager). The biogenesis of PS II in greening etioplasts has been studied in many plant systems (Schuster et al, 1985; Sutton et al., 1987; Hird et al., 1991; Meierhoff and Westhoff, 1993; Palomares et al., 1993). Mullet and coworkers analyzed mRNAs and translation products labeled in plastids isolated from etiolated and greening barley seedlings (Klein and Mullet, 1987), or labeled in plastid polysomal fractions separated by sucrose density gradient centrifugation (Klein et al., 1988). These studies showed that psbA transcription and translation are light-regulated; during greening the number of membrane-bound and soluble polysomes increases, and psbA mRNA becomes associated with chloroplast polysomes. Subsequent in vitro translation experi­ ments using lysed barley etioplasts have shown that accumulation of the PS II chlorophyll-binding proteins CP47, CP43 and D1 is dependent on de novo chlorophyll biosynthesis (Eichacker et al., 1990; Franck et al., 1997), Moreover, chlorophyll increases the stability of CP43 and D1 (Mullet et al., 1990), and ribosome ‘pausing’ during Dl translation increases during chloroplast development and may improve the efficiency of chlorophyll binding to nascent D1 (Kim et al., 1994b). Interestingly, the D2 core polypeptide is the only major PS II core subunit that accumulates in dark-grown etioplasts (Gamble et al., 1988). Greening studies with the barley nuclear mutant viridis-115 show that the stability of the CP47, CP43, D1 and D2 polypeptides is reduced after 16 h illumination, although the OEE1 and OEE3 polypeptides accumulate normally, suggesting that a nuclear gene product somehow stabilizes the intrinsic PS II core polypeptides (Gamble and Mullet, 1989). D1 translation initiation, elongation and ribosome pausing are not significantly altered in the vir-115 mutant, but D1 translation intermediates normally detected are lacking. This suggests that the wild-type nuclear viridis-115 gene may encode a factor which stabilizes D1 intermediates, possibly during assembly of D1 with chlorophyll (Kim et al., 1994c). Whether chlorophyll associates with D1 intermediates cotranslationally and affects D1 synthesis is not yet clear. Although chlorophyll did not appear to influence the extent of D1 elongation in barley translational runoff assays, full-length D1 translated on polysomes accumulated only in the presence ofchlorophyll, and

259 chlorophyll was seen to greatly increase the stability of D1 after release from ribosomes (Kim et al., 1994a). These results reaffirm the role of chlorophyll in a posttranslational stabilization of D1.

B. Algae In several organisms, including green algae, chlorophyll synthesis can occur via light-dependent and light-independent pathways, such that wild-type cells are green in the dark (Chapter 20, Timko). Chlamydomonas mutants lacking the light-independent pathway for chlorophyll synthesis have been isolated (Ohad et al., 1967; Bennoun et al., 1995), are ‘yellow in the dark’, and serve as model systems for studying PS II biogenesis during greening (Wettern et al., 1983). The light-dependent assembly of PS II has also been studied in a Scenedesmus obliquus mutant lacking light-independent carotenoid biosynthesis (Humbeck et al., 1990). In a study of wild-type and greening C. reinhardtii y-1 mutants, Malnoë et al. (1988) have shown that immunodetectable levels of the intrinsic PS II core polypeptides D1, D2, CP47 and CP43, as well as the LHCII apoproteins, are similar in light- and darkgrown wild-type cells, but are barely detectable in the dark-grown, chlorophyll-lacking y-1 mutant. In contrast, levels of the OEEC polypeptides are comparable, in light and dark conditions, in both the wild-type and mutant strains, demonstrating that the OEC polypeptides can accumulate independently of the PS II intrinsic core. Synthesis of D1 and D2 is enhanced immediately by light in wild-type cells, and enhanced with a lag in y-1 cells, suggesting that light-induced plastid development must precede lightactivated translation of D1 and D2. A 2 h lag for both accumulation of chlorophyll and accumulation of the D1, D2, CP47, CP43 and LHCII polypeptides during greening suggests that chlorophyll may stabilize these chlorophyll apopolypeptides, as is the case in greening barley (Mullet et al., 1990). Since synthesis of D1 and D2 in wild-type cells is increased in the light, while the abundance of these polypeptides appears constant, the rate of degradation of these polypeptides is assumed to be higher in the light (Malnoë et al., 1988), as seen previously in both Chlamydomonas (Wettern and Ohad, 1984; Section III.A) and in Spirodela oligorrhiza (Fromm et al., 1985). Thus, PS II biosynthesis is affected by both light itself and by the light-dependent stages of plastid development.

260 III. Assembly of Photosystem II Complexes

A. Migration of Photosystem II in the Thylakoid Membrane: PS II Damage and Repair Thylakoid membrane structure and function has been studied extensively, both through electron microscopy freeze-fracture analysis (Staehelin and van der Staay, 1996) and through genetic and biochemical dissection of thylakoid protein components (Cohen et al., 1995; Webber and Baker, 1996; Chapter 11, Olive and Wollman). The nature and function of the thylakoid membrane is determined to a large extent by its glycerolipid, pigment and prenylquinone composition (Douce and Joyard, 1996; Chapter 21, Trémoliéres). In vascular plants and green algae, the thylakoid is characterized by the presence of highly organized helical stacks of membrane, called the appressed membranes or granal lamellae, interspersed by single strands of membrane called non-appressed mem­ branes or stromal lamellae. The terminal membranes of each appressed membrane stack are exposed to the chloroplast stroma and have biochemical properties similar to non-appressed thylakoid membranes. Functional heterogeneity in PS II, between ‘reducing’ and ‘non-reducing’ PS II centers, also called and centers, respectively, has been noted (Guenther et al., 1990; Melis, 1991; Lavergne and Briantais, 1996). Structural heterogeneity is seen also in the segregation of most PS II into the appressed membranes, with only 10–20% of PS II centers found in the non-appressed membranes (Andersson and Anderson, 1980); PS I is found primarily in nonappressed membranes. Such segregation of PS II and PS I may be facilitated by the membrane stacking (Mustárdy, 1996), and may optimize positioning of the two photosystems for efficient energy transfer. Subsequent studies have confirmed this spatial distribution of PS II between the granal and stromal lamellae, and shown that PS II in the non-appressed membranes is depleted of the LHCII-inner antenna complexes, CP26 and CP29 (Vallon et al., 1986, 1987; Bassi et al., 1988; Barbato et al., 1992). A model has emerged in which functional PS II in the appressed membrane is ‘damaged’, dissociates from the LHCII antenna complexes, and migrates to the non-appressed membrane (Mattoo and Edelman, 1987; Adir et al., 1990; Guenther and Melis, 1990; Barbato et al., 1992). There, damaged subunits (primarily D l ) are removed from the inactive PS II

Jeanne Marie Erickson complexes and replaced by newly synthesized subunits. The ‘repaired’ PS II then migrates back to the appressed membrane where it associates with the PS II antenna and is again photoactive (for recent reviews Andersson and Aro, 1997; Barber et al., 1997; Chapter 30, Keren and Ohad). Damage to PS II occurs as a consequence of normal PS II function and is exacerbated during photoinhibitory conditions (Prásil et al., 1992; Aro et al., 1993; Anderson et al., 1997). The D1 reaction center polypeptide appears to be damaged the most (Mattoo et al., 1984; Ohad et al., 1984). D1, and to a lesser extent the D2 polypeptide, are the PS II components with the highest turnover rate (Schuster et al., 1988). Phosphorylation of PS II polypeptides may stabilize PS II in the appressed membranes and protect D1 from degrada­ tion (Ebbert and Godde, 1996; Andersson and Aro, 1997) but perhaps not from damage (Gal et al., 1997). While functional PS II in the granal lamellae may exist as a dimer, the inactive PS II that migrates to the stromal lamellae for repair is most likely a monomer (Santini et al., 1994; Lavergne and Briantais, 1996; Hankamer et al., 1997). In mature chloroplasts, D1 synthesis takes place primarily to replace photodamaged D1 subunits. Hence, most of the PS II ‘assembly’ occurring during steady-state photosynthetic function in mature plant cells may be a specific replacement of the D1 polypeptide rather than the de novo assembly of all newly synthesized PS II subunits, as occurs during greening. The process of photoinhibition is reviewed by Karen and Ohad (Chapter 30).

B. Photosystem II Assembly in Chlamydomonas

1. Three Classes of Mutants Defective in the Assembly or Stability of the PS II Core Proper subunit stoichiometries of the individual polypeptides of soluble, multisubunit protein complexes may in part be obtained by the stabilization of assembled subunits and the proteolysis of unassembled subunits, as shown for C. reinhardtii ribulose-bisphosphate carboxylase by Schmidt and Mishkind (1983). Similar results have been seen with the thylakoid membrane complexes, including the ATP synthase complex (Merchant and Selman, complex (Chapter 24, Wollman), 1984), the Cyt and Photosystem I (PS I) (Takahashi et al., 1991). The PS II intrinsic core polypeptides provide no exception to this rule. However, increasing evidence

Chapter 15

Assembly of Photosystem II

shows that the cell nucleus controls many levels of chloroplast gene expression, and that nuclear control over the temporal and spatial synthesis of polypeptides of multisubunit complexes ultimately controls the coordinate expression and fate of individual subunits (Rochaix, 1996; Rodermeletal., 1996; GoldschmidtClermont, 1998). There appear to be at least three classes of C. reinhardtii PS II assembly/stability mutants that lack one of the intrinsic membrane PS II subunits. In the first group, the absence of one subunit results in the rapid degradation of the other core subunits, the absence of assembled PS II, and obligate heterotrophic growth (reviewed in Rochaix and Erickson, 1988). PS II intrinsic membrane subunits are synthesized but fail to accumulate in the absence of either the D1 polypeptide (Bennoun et al., 1986; de Vitry et al., 1989), the D2 polypeptide (Erickson et al., 1986; Kuchka et al., 1988, 1989; de Vitry et al., 1989), the CP47 apoprotein (P5), (Jensen et al., 1986; Monod et al., 1992), or the CP43 apoprotein (P6), (de Vitry et al., 1989; Rochaix et al., 1989; Zerges et al., 1997). Such a destabilization of PS II is not unique to algae; a similar lack of the other PS II core subunits is seen in a nuclear maize mutant in which the D1 and CP43 polypeptides are rapidly degraded (Leto et al., 1985). The extrinsic membrane OEC subunits and the LHCII subunits appear to accumulate normally in the absence of the PS II core (Greer et al., 1986; Kuchka et al., 1988; de Vitry et al., 1989). The phosphorylated form of D2 (Delepelaire, 1984) is not found in this class of mutants (de Vitry et al., 1989), indicating that D2 phosphorylation may follow core assembly or require PS II function. In addition to the posttranslational destabilization of PS II intrinsic subunits synthesized in the above mutants, there is some evidence for the translational regulation of Dl by D2 (Erickson et al., 1986; de Vitry et al., 1989) and for a tight coupling of D1 and P5 translation (Jensen et al, 1986; de Vitry et al., 1989 and Summer et al., 1997), such that D2 may regulate synthesis of both Dl and P5. Alternatively, interaction with D2 may stabilize a newly synthesized Dl polypeptide (Wu and Kuchka, 1995). A second class of C. reinhardtii PS II mutants includes those obligate heterotrophs missing the psbH or psbK chloroplast gene products. In these mutants, the remaining PS II subunits are degraded more slowly than in the first class of mutants; small amounts of PS II complexes are formed, but are not sufficient to support photoautotrophic growth. Takahashi et al.

261 (1994) showed that disruption of Chlamydomonas psbK did not affect synthesis of the other PS II core subunits, but significantly reduced accumulation of PS II subunits as assessed by pulse chase experiments and immunoblotting. The psbK deletion mutants accumulated <10% of wild-type levels of PS II, and were unable to grow photoautotrophically. Sitedirected C. reinhardtii mutants lacking the H polypeptide (PS II-H) also lack PS II activity (Ruffle et al., 1995; Cheater et al., 1995; Summer et al., 1997) and PS II is unstable even when cells are grown in the dark on acetate (Summer et al., 1997). Deletion of psbH has no influence on the level of accumulation of other PS II mRNAs. Indeed, polypeptides D1, D2 and the CP43 and CP47 apoproteins are synthesized at normal levels during a 10 min pulse, but they are degraded during the chase period in the absence of PS II-H and are not detected in stained gels or immunoblots (Summer et al., 1997). Sucrose density gradient fractionation of polysomal fractions suggests that some assembly of smaller PS II subcomplexes takes place in the absence of PS II-H, and Summer et al. (1997) conclude that either assembly in the mutant is a slow process and the complexes are labile under the sucrose gradient centrifugation conditions, or that assembly is followed by dissociation of PS II in vivo. These authors propose that PS II-H may have a function either in the initial stabilization of the nascent PS II subcomplex, or in the dimerization and subsequent stabilization of PS II. Interestingly, in the absence of PS II-H, the D2 polypeptide is the most unstable of the core subunits. A relationship between PS II-H and the primary on the D2 polypeptide may exist, as has quinone been postulated from studies with cyanobacteria (Mayes et al., 1993). Although several functional roles have been suggested for PS II-H, its contribution to PS II is not clearly understood. PS II-H is one of the phosphorylated subunits of PS II (Ikeuchi et al., 1987; see Bennett 1991 for review), with phos­ phorylation predicted at two different sites. However, lack of PS II-H has no effect on phosphorylation of the Chlamydomonas D2 polypeptide, indicating that if PS II-H is part of a phosphorylation cascade regulating PS II stability/function, it is not essential for phosphorylation of D2 (see discussion in Summer et al., 1997). Site-directed mutagenesis of the presumed N-terminal phosphorylation site in psbH likewise has no obvious effect on PS II stability or function in C. reinhardtii in vivo (Cheater et al., 1995). Both PS II-H and PS II-K (the K polypeptide)

262 may have a more peripheral, yet still essential, role in assembly of a stable PS II particle. It is noted that the Chlamydomonas psbK and psbH deletion mutants have a different phenotype than similar mutants in cyanobacteria, where PS II-K (Ikeuchi et al., 1991; Zhang et al., 1993) or PS II-H (Mayes et al., 1993; Komenda and Barber, 1995) are neither essential for PS II assembly nor for photoautotrophic growth. A third class of Chlamydomonas PS II mutants is able to grow photoautotrophically, under specific conditions, in the absence of either the psbI or psbT (ycf8) chloroplast gene products. A similarphenotype has been reported for a C. reinhardtii psbN deletion mutant by Ruffle and Nugent (Chapter 16, Ruffle and Sayre). Deletion of ycf8 (psbT) led to its identification as a PS II gene, and resulted in mutant cells with an unusual phenotype (Monod et al., 1994). Under conditions of environmental stress, including high light or reduced levels of chloroplast protein synthesis in the presence of antibiotics, PS II is unstable, levels of PS II core subunits are reduced four-fold, and the mutant cells lacking the T subunit (PS II-T) grow poorly. This phenotype is observed only under conditions of stress; under normal growth conditions, photoautotrophic growth rates are the same as for wild-type cells (Monod et al., 1994). Disruption of the C. reinhardtii psbI gene revealed another light-sensitive phenotype. Mutant cells lacking the I subunit (PS II-I) are photoautotrophic in dim light, with both PS II content and oxygenevolution reduced to 10–20% of wild-type levels (Künstner et al., 1995). In high light, mutant cells are photosensitive and unable to grow photosynthetically. The photosensitive phenotype in the absence of PS II-I and PS II-T is in contrast to the total lack of photoautotrophic growth in the absence of Chlamy­ domonas PS II-H and PS II-K. PS II-I may be the only PS II RC component that is not essential for chloroplast PS II assembly or photochemistry. The D1 and D2 polypeptides are absolutely required as described above. And, although C. reinhardtii psbE or psbF deletion mutants have not been reported, the disruption of homologous genes in cyanobacteria leads to loss of PS II and obligate heterotrophy (Pakrasi et al., 1989), may prove to be essential suggesting that Cyt also for chloroplast PS II (Cramer et al., 1996). In summary, PS II-I, PS II-N and PS II-T appear to be involved in the stabilization of the PS II core complex, particularly at higher light intensities and under stress conditions when demands for D1 synthesis and repair

Jeanne Marie Erickson are not met (Sections III.A, III.B.4). These subunits may modulate PS II RC function in ways that further stabilize the functional complex, or may provide stability during or after assembly and play no active role in PS II function per se. In contrast to the lightsensitive phenotype seen in the Chlamydomonas psbI mutant, PS II in the photoautotrophic cyanobacterial psbI deletion mutant is quite stable even at higher light intensities (Ikeuchi et al., 1995).

2. Stepwise Assembly of Photosystem II Intrinsic and Extrinsic Membrane Subunits de Vitry et al. (1989) carried out an extensive study of C. reinhardtii mutants unable to synthesize any one of the PS II core subunits Dl, D2 or CP43, or either of the extrinsic membrane polypeptides OEE1 or OEE2. These mutants were characterized with respect to accumulation of PS II polypeptides, their chlorophyll-binding properties, and their localization by immunocytochemistry. The synthesis and halflife of polypeptides were assessed by in vivo pulsechase experiments, and the in vitro proteasesusceptibility of subunits was measured. Results show that chlorophyll binds to apoP5 (apoCP47) and apoP6 (apoCP43) either during or shortly after synthesis, regardless of the presence of other PS II subunits. P5 and P6 are synthesized and accumulate, independently of each other, in the appressed thylakoid membranes. The stability of the newly synthesized subunits increases stepwise, as assembly occurs initially between D1, D2, P5, and Cyt followed by assembly with P6 and finally with the OEE1 and OEE2 polypeptides. Interestingly, CP43 is more readily detached from the C. reinhardtii PS II core than is CP47 (de Vitry et al., 1984). Spinach PS II assembly-intermediates lacking CP43 have also been identified (van Wijk et al, 1997). In the absence of any CP43, acyanobacterial PS II core is assembled (Rögner et al., 1991). and contains a functional The principle components of the early PS II assembly intermediate—D1, D2 and CP47—appear to be translated in a coordinate manner (Jensen et al., 1986; de Vitry et al., 1989; but see Wu and Kuchka, 1995), while CP43 is translated independently (de Vitry et al., 1989). The OEE polypeptides are processed to the mature form and accumulate in the absence of the PS II core polypeptides (de Vitry et al., 1989), consistent with other studies in C. rein­ hardtii (Greer et al., 1986; Kuchka et al., 1988; Malnoë et al., 1988;Rochaix et al., 1989;Eisenberg-

Chapter 15

Assembly of Photosystem II

Domovich et al., 1995). Interestingly, de Vitry et al. (1989) also observed that, while the OEE polypeptides are found in the lumen loosely bound to the thylakoid membranes regardless of the presence or absence of core subunits, OEE2 is localized to the appressed membranes in the absence of CP43, and to the nonappressed membranes only in strains that synthesized CP43. In contrast, OEE1 is localized to appressed membranes in mutant and in wild-type strains. Thus, a specific interaction between CP43 and OEE2 may be required for the localization of OEE2 to the appressed membranes. The possible assembly of OEE2 with appressed membranes is supported by studies of Hashimoto et al. (1997) using pea chloroplasts (Section IV.A).

3. Assembly of Nascent D1 and the OEC Polypeptides with Other PS II Subunits Ohad and colleagues have studied the synthesis of C. reinhardtii PS II polypeptides and their assembly into PS II complexes, with particular emphasis on the synthesis and assembly of the D1 polypeptide with PS II. D1 has the highest turnover rate of any of the PS II subunits, and D1 damage and replacement is associated with PS II photoinhibition and repair (Ohad et al., 1990a; Chapter 30, Keren and Ohad). Adir et al. (1990) labeled C. reinhardtii chloroplastencoded polypeptides in vivo, fractionated the thylakoid membranes, and identified polypeptides by one- and two-dimensional gel electrophoresis and immunoblotting. They found that light enhanced translocation of the PS II RC from the appressed to the non-appressed membrane fractions, and that the precursor Dl polypeptide (pD1) labeled during a 2 min pulse was integrated into a PS II RC in the nonappressed membranes and subsequently processed. After a 30 min chase, all of the labeled D1 was processed and associated with PS II complexes in the appressed membranes. The failure to detect any free pD1 or D1 polypeptides suggests either that the PS II RC is the receptor for the integration of newly synthesized D1 into the membrane, or that the time between insertion of free pD1 into the membrane and assembly with PS II is extremely short in vivo. Moreover, the PS II RC does not completely disassemble when damaged D1 is removed; the D2, subunits still remain associated CP47, and Cyt (Adir et al, 1990;Eisenberg-Domovich et al, 1995). Three distinct forms of the PS II RC were resolved that appeared to shuttle between the appressed and

263 the non-appressed membranes. These were thought to represent possible changes in the RC organization during light-induced modification of D1, the presence of other PS II polypeptides not identified in their study, or the possible association of PS II RC units into oligomeric complexes (Adir et al., 1990). Using similar techniques, Eisenberg-Domovich et al. (1995) investigated the role of D1 in the reversible association of the extrinsic membrane OEC polypeptides with PS II. They found that the OEE1 polypeptide is released when Dl is degraded, and that the kinetics of D1 degradation correlate with those of OEE1 subunit release. At the same time, CP43 and CP47 are stable, and only a limited amount of D2 is degraded. All three OEC subunits are stable in the lumen for up to 8 h after release from PS II, and can reassociate with PS II following D1 synthesis under low-light conditions, leading to recovery of PS II activity. The OEC polypeptides also reassociate with PS II after D1 is synthesized in the dark. In the presence of a quinone analog that blocks to electron transfer, or in a C. reinhardtii mutant lacking plastocyanin which maintains a high level of reduced plastoquinone in the light, PS II photoinhibition occurs but D1 is not degraded. In this case, the OEC polypeptides are not released. Thus OEE subunit binding is determined by a PS II RC containing D1, and the binding site is not destroyed by photoinhibition (Eisenberg-Domovich et al., 1995). A model for the assembly of newly synthesized D1 into partially-disassembled PS II complexes is given (Fig. 2), based in part on work with Chlamydomonas as described. This model takes into account the association of nascent Dl with other PS II RC polypeptides (de Vitry et al., 1989; Adir et al., 1990), the observed migration of damaged PS II from the appressed membranes to the non-appressed membranes where PS II repair via D1 replacement takes place (Adir et al., 1990; section IIIA), the removal of CP43 prior to degradation of D1 (Barbato et al., 1992) and the observation that OEE 1 release is concomitant with Dl degradation (EisenbergDomovich et al., 1995). While OEE1 may remain localized to appressed thylakoid membranes (de Vitry et al., 1989), evidence from studies on the import of OEE1 precursors into isolated pea chloroplasts suggests that the OEE1 extrinsic membrane subunit can assemble with PS II complexes in the nonappressed thylakoid membranes as well (Hashimoto et al., 1997).

264

4. Assembly of the PS II Redox Components Assembly of a functional PS II requires not only the synthesis and assembly of the polypeptide subunits, but also their functional association with the redoxactive molecules essential for PS II photochemistry, including chlorophyll, pheophytin, plastoquinone, heme and non-heme irons, and manganese (Fig. 1). The identification of putative cofactor binding sites subunits, using siteon the D1, D2 and Cyt directed mutagenesis studies in cyanobacteria and algae, has been reviewed (Diner et al., 1991; Vermaas, 1993; Diner and Babcock, 1996; Chapter 16, Ruffle and Sayre). Chlorophyll-binding to PS II subunits Dl, D2, CP47 and CP43 appears to stabilize these subunits (Section II; Paulsen, 1997; Thomas, 1997).

Jeanne Marie Erickson

In C. reinhardtii mutants blocked at specific steps of carotenoid or chlorophyll synthesis, the PS II core subunits do not accumulate (Herrin et al., 1992). Recent investigations in C. reinhardtii suggest that stabilizes the Dl subunit and is essential for the assembly of functional PS II (Trebst and Depka, 1997). Nuclear maize mutants which have severely reduced levels of the chloroplast PS II plastoquinone (PQ-9) have no detectable PQ-9 in thylakoid membranes, and show no PS II electron transport, but still accumulate normal levels of PS II core subunits (Cook and Miles, 1992). This suggests that and are not the plastoquinone cofactors required for the stable assembly of PS II polypeptides. However, engineering of the cyanobacterial D2

Chapter 15

Assembly of Photosystem II

polypeptide to alter a putative ligand for leads to the destabilization of PS II (Vermaas et al., 1989; reviewed in Vermaas, 1993). Alterations of the binding site of the D1 subunit that permit quinone state, exacerbate lightbinding, but destabilize the induced D1 degradation in site-directed cyanobacterial mutants (Ohad et al., 1990b). Thus, PS II stability may not require the binding of quinones, but may be affected by the redox status of a bound (Gong and quinone and the ratio of PQ to Ohad, 1991). Physical alterations to the quinonebinding sites may result in aberrant PS II photo­ chemistry, elevated subunit damage, and ultimate destabilization of PS II. Many site-directed mutants of cyanobacteria and algae which are unable to assemble the manganese cluster of the OEC still accumulate normal levels of PS II complexes; however, certain amino acid substitutions in D1 or D2 which affect manganese ligation do destabilize the PS II complex of Chlamydomonas (Whitelegge et al., 1995) and of cyanobacteria (reviewed in Diner and Babcock, 1996; Chapter 16, Ruffle and Sayre). Manganese may not be required for the stable assembly of PS II polypeptides, but aberrant configurations of manganese may lead to increased PS II photo­ sensitivity and a reduction in PS II levels. The assembly of the tetranuclear manganese cluster of the PS II OEC is discussed in Section V Using C. reinhardtii, van Wijk et al. (1994) followed the kinetics of reactivation of PS II redox components after light-induced inactivation of PS II in vivo, and correlated this with the kinetics of the synthesis and turnover of the Dl, D2, CP47 and CP43 subunits. Inactivation was carried out at low temperature to allow for accumulation of the damaged D1 subunits, while recovery was carried out at normal growth temperatures. The rate of D1 synthesis during recovery was ~5 times that of D2, and 10–20 times higher than that of CP43 and CP47. No accumulation of any partially-active PS II intermediates was seen, suggesting that assembly/reactivation of the redox components of PS II occurs very quickly after the synthesis and assembly of the PS II core polypeptides. These authors conclude that the rate-limiting step in recovery is not the functional association of the redox components, but rather the degradation of damaged D1 and the synthesis of new subunits, primarily D1 and D2 (van Wijk et al., 1994). This scenario, in which the rapid assembly of the PS II

265 redox components follows a slower reassembly of the PS II core complex, should help prevent the accumulation of partially-activated PS II units that might be damaged by absorption of light-energy in the absence of the full component of essential redox factors.

C. Photosystem II Assembly Intermediates: In Organello and In Vitro 1. The Synthesis and Fate of PS II Core Subunits The synthesis and assembly of chloroplast-encoded PS II polypeptides have been investigated by labeling proteins in isolated chloroplasts (i.e., in organello) or in isolated thylakoid membranes (i.e., in vitro) and following the assembly of these newly synthesized subunits into PS II complexes. Minami et al. (1986) used both in organello and in vitro systems to study the assembly of the PS II core complex of spinach. The in organello synthesis of PS II subunits and their association with larger complexes during greening of barley have been studied (Section II.A). Expanding this type of analysis to spinach grown on a 12 h light/ dark cycle, van Wijk et al. (1995) performed a series of pulse-chase experiments to examine the in organello and in vitro synthesis and subsequent incorporation of nascent PS II polypeptide subunits into different PS II subcomplexes. Membranes were fractionated by sucrose density gradient centri­ fugation, and the fraction components identified (from 15 kDa to 500 kDa) were further subjected to analysis using denaturing polyacrylamide gel electrophoresis, autoradiography, and immunoblotting. Their results suggest that newly synthesized D1 is incorporated primarily into existing PS II subcomplexes, while newly synthesized D2, CP43 and CP47 accumulate in the membrane as unassembled subunits. In contrast to results using isolated chloroplasts, where more of the radiolabeled D1 became associated eventually with PS II core complexes, a high percentage of D1 radiolabeled in isolated thylakoid membranes was unassembled in the membrane or present in smaller PS II subcomplexes but not present in a full PS II core complex, leading these authors to conclude that stromal factors may be required for efficient D1 replacement. The results suggest also that chloroplast synthesis of D1 appears to be regulated in relationship to the ‘need’ for D1 replacement into existing PS II

266 complexes (Adir et al., 1990; Koivuniemi et al., 1995; van Wijk et al., 1995). The effects of light on the synthesis and assembly of PS II polypeptides have been examined in organello (van Wijk and Eichacker, 1996). Chloroplasts isolated from spinach leaves harvested at the end of a twelve hour dark period exhibited an increased accumulation of nascent D1 translation intermediates bound to polysomes when radiolabeling was performed in the dark (in the presence of ATP and reducing agent) as compared to the light. There was also a three-fold reduction in the accumulation of D1 and CP43 radiolabeled in the dark, as compared to that seen when labeling in the light. Moreover, in the dark, most of the radiolabeled D1 that accumulated was present as unassembled polypeptide or in PS II subcomplexes smaller than 100 kDa, while in the light D1 was incorporated primarily into PS II core complexes. Thus, light appears to be required for the efficient translational elongation of Dl and CP43, and for the efficient incorporation of radiolabeled D1 into PS II core complexes. In contrast, the synthesis and accumulation of the D2 polypeptide were not significantly influenced by light (van Wijk and Eichacker, 1996). An interesting aspect of the apparent lightdependent nature of D1 synthesis was revealed in studies by Kuroda et al. (1996). These authors show that D1 can be synthesized in vitro, in total darkness when supplemented with exogenous energy and reducing agents, if the plant leaves are harvested in the light period of a diurnal growth cycle. In contrast, when chloroplasts are isolated from dark-adapted leaves, negligible amounts of D1 synthesis occur in organello in the dark under similar labeling conditions (Kettunen et al., 1997). These results, together with other in vitro (Koivuniemi et al., 1995) or in organello studies (Tanaguchi et al., 1993), as well as in vivo studies on the relationship between D1 synthesis and degradation in Chlamydomonas (Adir et al., 1990; Neale and Melis, 1991) and in plants (Schnettger et al., 1994), suggest that elongation of D1 synthesis may be coupled to and tightly regulated by D1 degradation. Since D1 damage is light-induced (Prásil et al., 1992), degradation and hence D1 synthesis will be affected by light conditions. Given the instability of PS II complexes lacking D1 (Section III.B), the coupling of D1 degradation and synthesis would enhance the stability of a partially-disassembled PS II complex by minimizing the time between removal of damaged D1 and insertion of a newly

Jeanne Marie Erickson synthesized precursor D1 (Kettunen et al., 1997).

2. D2 as a ‘Docking’-Like Protein Required for D1 Synthesis and/or Stability The D2 polypeptide is unique among PS II polypeptides in several respects, and may play a key role in PS II assembly. Evidence suggests that D2 may be a ‘docking’-like protein required for the efficient translation and/or stabilization of Dl. Alternatively or in addition, D2 may play a role in the first stage of D1 assembly into PS II subcomplexes. In contrast to the D1, CP43 and CP47 polypeptides, translational elongation of D2 in pea etioplasts does not appear to require chlorophyll (Gamble et al., 1988). Moreover, D2 is synthesized in spinach chloroplasts, in organello, to similar levels in the light and the dark, while levels of CP43 and D1 synthesis are decreased two-and three-fold, respec­ tively, in the dark (van Wijk and Eichacker, 1996). Dl synthesis has been shown to be controlled primarily at the level of translation by nucleusencoded factors that bind to the 5´ region of psbA mRNA in Chlamydomonas (Danon and Mayfield, 1991; Yohn et al., 1996) and in plants (Staub and Maliga, 1994; Klaff and Gruissem, 1995). The translational regulation of D1 by light may involve redox state and ADP (reviewed in Mayfield et al., 1995). Evidence exists for the translational regulation of D1 by a low redox potential component downstream of PS I in pea (Kuroda et al., 1996). In contrast, the synthesis of D2 may be less tightly controlled by chloroplast factors than that of D1, at least with respect to light conditions, chlorophyll, and photosynthetic redox state. However, D2 gene expression is regulated by nuclear factors (Kuchka et al., 1988; 1989; Wu and Kuchka, 1995; reviewed in Rochaix, 1996; Goldschmidt-Clermont, 1998). In vivo studies show that in a C. reinhardtii psbD frameshift mutant (FUD47) producing an unstable, truncated D2 polypeptide, translation and/or accumulation of Dl is not seen at any significant level (Erickson et al., 1986). Similar effects on D1 synthesis in this same C. reinhardtii mutant were noted also by de Vitry et al. (1989). However, in nuclear mutants with greatly reduced synthesis of D2, attributed to the lack of specific nuclear factors required for D2 translation, Dl is still synthesized at normal rates (Wu and Kuchka, 1995); such results favor a post-translational stabilization of D1 by D2. The relative abundance of the D2 polypeptide in

Chapter 15 Assembly of Photosystem II spinach polysomal-like preparations containing ribosomes, mRNA and other small membrane fragments has been reported (van Wijk and Eichacker, 1996), and suggests that D2 may indeed be required for translation or stabilization of the nascent D1 polypeptide. Further evidence of a special role for D2 comes from the stoichiometry of D1 and D2 in isolated PS II subcomplexes from spinach thylakoids radiolabeled in vitro (van Wijk et al., 1997); a relative surplus of unassembled, stable D2 polypeptides is detected in the membrane leading to the speculation that D2 acts as a “receptor” for the newly synthesized D1 polypeptide. In conclusion, accumulating evidence suggests that unassembled D2 is more stable than unassembled D1, that D2 can accumulate in the membrane in the dark in the absence of chlorophyll, and that stable synthesis and/or accumulation of D1 in vivo requires the presence of a stable D2. A role for D2 as a docking-like protein controlling D1 synthesis and/or stability has been proposed (Erickson et al., 1986; van Wrjk et al., 1997). D2 may facilitate the elongation of D1 through direct interaction with the membranebound polysome, and/or D2 may stabilize Dl either as the nascent D1 polypeptide emerges from the ribosome or posttranslationally. Newly synthesized D1 polypeptides initially found in the thylakoid membrane as unassembled subunits are detected, within minutes, in a subcomplex containing only Dl and D2 (van Wijk et al., 1997). Thus, D2 appears to be the first assembly partner for D1. If the PS IIRC lacking D1 is indeed the membrane receptor for nascent D1 polypeptide (Adir et al., 1990), then the PS II RC-associated D2 polypeptide may be the primary partner for interaction with nascent D1 during the PS II repair cycle (Fig. 2).

3. Stepwise Assembly of the Photosystem II Polypeptides in Appressed and Non-appressed Thylakoid Membranes PS II assembly intermediates were localized to the appressed or non-appressed thylakoid membranes by van Wijk et al. (1996) who used in vitro and in organello pulse-chase protocols to study the incorporation of newly-synthesized D1 polypeptide into PS II subcomplexes. Immediately following a 4 min labeling of spinach chloroplast proteins, radiolabeled D1 was present as a free (unassembled) polypeptide in the thylakoid membrane, in small PS II subcomplexes, and in PS II reaction center particles,

267 but not in full PS II core complexes. Using chase periods from 2.5 to 10 min these authors observed the translational elongation of D1, the rapid carboxylterminal processing of the precursor D1 in chloroplasts, and the kinetics of D1 incorporation into subcomplexes, PS II reaction centers (PS II RC) and eventually PS II core complexes. The PS II core complexes were found almost exclusively in the appressed membranes, while the smaller PS II subcomplexes were found mainly in non-appressed membranes. These authors also examined the fate of radiolabeled D2, CP43, CP47, and three smaller PS II polypeptides of 4, 4.5 and 8 kDa. The labeled D2, CP43 and especially CP47 were rarely incorporated into PS II. In contrast, the three small polypeptides were assembled effectively into full PS II core complexes; they did not accumulate in PS II RC complexes or subcomplexes, and may assemble directly into PS II core complexes. It will be interesting to see whether the turnover rates of these smaller PS II polypeptides are higher than those of D2, CP43 and CP47, or whether the small radiolabeled sub­ units, containing fewer transmembrane helical domains, just exchange more efficiently with PS II core complexes. Given a 5 min delay between the appearance of radiolabeled D1 in PS II reaction center cores, and the subsequent conversion of those subcomplexes into full PS II core complexes, van Wijk et al. (1996) propose that a rate limiting step in PS II assembly is the migration of a PS II reaction center core from the non-appressed to the appressed membranes, where other subunits assemble with it to produce the full core complex. They noted that D1 radiolabeled in isolated thylakoid membrane preparations was found in small PS II subcomplexes, but was not incorporated into full PS II core complexes efficiently, suggesting a requirement for chloroplast stromal factors in the complete assembly of PS II. To investigate the processing and incorporation of radiolabeled precursor D1 into PS II and to identify the polypeptides present in intermediate assembly products as characterized in their previous experi­ ments, van Wijk et al. (1997) further resolved the components of the protein complexes separated by sucrose density gradient centrifugation. The 40 kDa to 360 kDa complexes were characterized by twodimensional gel electrophoresis using non-denaturing and denaturing acrylamide gels, and by isoelectric focusing. Polypeptides were detected by immuno­ blotting with antibodies specific for the PS II subunits and PS II-I. In these D1, D2, CP43, CP47, Cyt

268 studies, mature spinach leaves were harvested in the light period of a diurnal light/dark growth cycle, and intact chloroplasts or isolated thylakoid membranes were radiolabeled under relatively low light intensities of photons ). These conditions favor (50 the repair of D1 in the estimated 1–3% of PS II centers that contain damaged D1, given an 8 h halflife for D1, and 10 min to repair damaged D1. Since D2 Synthesis is greatly reduced in mature leaves, and D1 synthesis is associated primarily with PS II repair, the use of mature leaves gives a greater resolution of labeled D1 and precursor D1 on denaturing gels. The migration of pulse-labeled polypeptides into different PS II subcomplexes was observed during chase periods of up to 60 min, and the identities of the polypeptide subunits in each subcomplex were examined. A model for PS II assembly emerges (Fig. 3) in which the nascent D1 polypeptide (pD1) assembles in the non-appressed membranes, sequentially, with polypeptides D2, Cyt and PS II-I to form the PS IIRC. This PS IIRC then assembles with the subunits CP47 and, finally, CP43 to complete the PS II RC and inner antenna complex. Subsequent assembly steps involve the migration of the PS II subcomplex to the appressed membranes, the assembly of the PS II core complex and the OEC, and the association of LHCII as described in the legend of Fig. 3.

4. Posttranslational Import of Chimeric D1 into Isolated Chloroplasts and Assembly with PS II Elegant experiments were conducted by Wu and Watanabe (1997), who studied the import of a chimeric precursor protein—containing the rbcS transit peptide and a small portion of the N-terminus of the Rubisco small subunit (SSU) fused to the D1 polypeptide—into isolated pea chloroplasts. The transit peptide and the carboxyl-terminus of D1 were properly processed in the stroma and lumen, respectively, and the chimeric SSU-D1 was assembled in the thylakoid membrane into PS II complexes which had a configuration similar to those containing native D1. These results clearly show that the D1 polypeptide sequence provides information for targeting D1 to the thylakoid, and that proper insertion of D1 into the thylakoid membrane can occur independently of D1 translation on thylakoid ribosomes. Moreover, as chlorophyll has been shown to stabilize D1 (Kim et al., 1994a), it would appear that chlorophyll can assemble with the imported, chimeric D1 posttranslationally. The results of Wu

Jeanne Marie Erickson andWatanabe (1997) donot imply that cotranslational assembly of chlorophyll with D1 and cotranslational D1 membrane insertion do not occur in vivo, but simply that these D1 assembly processes can take place in organello independently of the translation process. Indeed, the in organello and in vitro studies with spinach led van Wijk et al. (1996) to propose both a cotranslational and a posttranslational pathway for the incorporation of D1 into PS II complexes.

D. Processing of the D1 Polypeptide CarboxylTerminus is Not a Prerequisite for PS II Assembly In all cyanobacteria and eukaryotes whose psbA. sequence has been examined (Svensson et al., 1991), except Euglena gracilis, D1 is synthesized as a precursor polypeptide which undergoes proteolytic processing at the carboxyl-terminus (Marder et al., 1984) to form the mature truncated polypeptide ending with alanine residue 344 (Takahashi et al., 1990; Trost et al., 1997). Processing can occur normally in Chlamydomonas when serine residue 345 is replaced with glycine, cysteine, valine or phenylalanine, suggesting that protease cleavage is indifferent to the substitutions at this position, or that the in vivo processing rate is not limited by enzymatic cleavage (Takahashi et al., 1996). The enzyme responsible for carboxyl-terminal processing of D1 (Ctp-protease) is located in the thylakoid lumen (Fujita et al., 1995). This protease was isolated from spinach, characterized and partially sequenced (Fujita et al., 1995); the complete protein sequence of the Scenedesmus obliquus Ctp-protease has been determined (Trost et al., 1997). CtpA cDNAs encoding the Ctp-protease have been characterized from barley (Oelmüller et al., 1996), spinach (Oelmüller et al., 1996; Inagaki et al., 1996) and S. obliquus (Trost et al., 1997), and a homologue from cyanobacteria has been cloned and characterized (Anbudurai et al., 1994). Several lines of in vivo evidence suggest that carboxyl terminal processing of D1 is not a prerequisite for incorporation of D1 into PS II. The LF1 mutant of S. obliquus is unable to evolve oxygen and does not process the carboxyl terminus of D1, but does incorporate precursor D1 into PS II complexes that lack a functional OEC (Metz et al., 1980; Diner et al, 1988; Seibert et al., 1989; Trost et al., 1997 and references therein). In other studies, it was shown that mutations in psbA which eliminate the processing of precursor D1 in Synechocystis sp. 6803 (Nixon et al., 1992), or mutations in ctpA itself

Chapter 15 Assembly of Photosystem II

269

270 (Shegtakov et al., 1994), can produce a mutant phenotype similar to that of the LF1 mutant, i.e., precursor D1 is assembled into nonfunctional PS II complexes. In C. reinhardtii, the D1 precursor can be inserted into PS II RC complexes prior to processing (Adir et al., 1990). Analysis of a CtpA cDNA from the S. obliquus LF1 mutant has shown that a single base pair deletion results in a frame-shift mutation leading to the production of a truncated, nonfunctional D1 protease (Trost et al., 1997). These authors have also isolated and characterized the CtpA cDNA from a photo­ autotrophic suppressor strain derived from LF1, and show that a single base pair insertion close to the initial mutation site restores the reading frame and restores Ctp-protease activity. Their results are the first definitive experiments showing that the nuclear CtpA gene product encodes the protease actually responsible for C-terminal processing of the D1 polypeptide (Trost et al., 1997). It is likely that the reduction in D1 carboxyl-terminal processing seen in isolated thylakoid membranes compared to isolated chloroplasts (van Wijk et al., 1996; 1997) may be related to a physical loss or loss of activity of the lumenally located, D1-specific C-terminal peptidase encoded by CtpA. Site-directed mutagenesis in Synechocystis sp. 6803 (Nixon et al., 1992) and Chlamydomonas (Lers et al., 1992; Schrader and Johanningmeier, 1992), aimed at deletion of the carboxyl-terminal residues of D1 which are normally removed by processing, produced mutants containing engineered ‘pre­ processed’ D1 polypeptides which are incorporated efficiently into PS II. Although no change in PS II function or photosensitivity was seen in the algal mutants, the cyanobacterial mutants containing genetically ‘pre-processed’ D1 are more photo­ sensitive than wild-type strains at high light intensities, i.e., when D1 turnover is rapid (Trost et al., 1997). Thus, proteolytic processing of the carboxyl-terminus of the D1 precursor may be coordinated with PS II repair (Fig. 2) to favor PS II assembly/stability under conditions of light stress. The effect of thylakoid membrane lipid unsatur­ ation levels on D1 processing was studied in an engineered cyanobacterial mutant lacking the genes for fatty acid desaturation (Kanervo et al., 1997). At low temperatures, this mutant was unable to process the D1 precursor and no photoactivation of the PS II OEC occurred. A C. reinhardtii mutant, likewise impaired in chloroplast fatty acid desaturation,

Jeanne Marie Erickson exhibited a high temperature tolerance for PS II function and photosynthesis (Sato et al., 1996). The effect of low temperature on D1 processing in the algal mutant was not reported; it may be that, just as in the cyanobacterial mutant, low temperature inhibits chloroplast D1 processing when thylakoid mem­ branes contain low levels of unsaturated fatty acids. The high temperature tolerance, conversely, may be due in part to the ability to process D1 and assemble PS II at temperatures prohibitive for PS II assembly in wild-type strains. Clearly, the lipid composition of the membrane affects D1 processing and PS II function. In summary, all lines of evidence suggest that the removal of carboxyl-terminal residues from precursor D1 and the assembly of D1 with other PS II polypeptides are two distinct, and possibly unrelated, processes. However, under certain environmental conditions, the coordination of these two events may enhance PS II assembly or stability (Trost et al., 1997).

IV. Assembly of the Extrinsic Membrane Polypeptides of the PS II Oxygen-Evolving Complex

A. In Organello Assembly of the OEC Polypeptides with the PS II Core Complex The three extrinsic membrane polypeptides of the chloroplast OEC are encoded in the nucleus, translated in the cytoplasm, and imported into the chloroplast thylakoid where they bind to the PS II intrinsic membrane core polypeptides at the lumenal surface of the thylakoid membrane. In Chlamy­ domonas, these subunits are referred to as the ‘oxygen-evolving enhancer’ polypeptides OEE1 (33 kDa), OEE2 (23 kDa) and OEE3 (16 kDa). The structure, function and PS II-binding of the OEC polypeptides has been extensively reviewed (Vermaas et al., 1993; Bricker and Ghanotakis, 1996; Seidler, 1996; Bricker and Frankel, 1998; Chapter 16, Ruffle and Sayre). The OEE1 polypeptide is tightly bound to the intrinsic subunits of PS II; evidence suggests that OEE1 binds to CP47, and possibly to CP43, D1, and PS II-I. OEE2 binds to PS II in the D2, Cyt absence of OEE1, but with reduced affinity, suggesting that OEE1 provides part of the OEE2 binding site or causes a conformational change that creates a high-affinity PS II binding site for OEE2.

Chapter 15 Assembly of Photosystem II The binding site for OEE3 appears to be located on or determined by the presence of OEE2 (Miyao and Murata, 1989). Although OEE1 can bind to PS II regardless of whether or not any manganese is assembled with PS II, as shown by in vivo studies in Chlamydomonas (Yamamoto, 1988) and in vitro studies in spinach (Leuschner and Bricker, 1996), OEE1 binding to PS II is reduced in the absence of the manganese cluster (Miyao and Murata, 1989). Unlike most of the PS II core polypeptides, which are unstable if not assembled with each other, the OEE polypeptides appear to be quite stable as individual, unassembled subunits in Chlamydomonas mutants lacking PS II cores (Greer et al., 1986; Kuchka et al., 1988; Mayfield et al., 1987a; Rochaix et al., 1989; de Vitry et al., 1989). Moreover, in pea chloroplasts, 20 to 50% of the OEE subunits of PS II are present as a soluble pool in the thylakoid lumen (Ettinger and Theg, 1991). Hashimoto et al. (1996) studied the import of labeled OEE1 and OEE2 precursor polypeptides into isolated pea chloroplasts, and followed the processing, localization and stability of these subunits under light and dark conditions. Although newly imported OEEs were more stable in the dark, the precursors were efficiently processed to the mature form and localized to the thylakoids in both light and dark, where they partitioned between the thylakoid membrane and the soluble thylakoid fractions. Their work demonstrates that a pool of relatively stable, unassembled OEE subunits exists in the thylakoid lumen, and that the individual unassembled subunits are able to assemble subse­ quently with PS II. The observed decreased stability of newly imported OEE1 in the light vs. dark may reflect the increased turnover of D1 in the light (Prásil et al., 1992), and a close association of D1 with the OEE1 extrinsic polypeptide. Import studies with a series of truncated OEE2 precursor poly­ peptides lacking portions of the OEE2 carboxyl terminus revealed that, while truncation of OEE2 impairs import to some degree, the truncated forms of OEE2 could be properly processed and imported into the lumen where they were rapidly degraded (Roffey and Theg, 1996). Thus, there appears to be a specific and rapid proteolytic degradation of truncated, unassembled OEE2, even though wildtype OEE2 is fairly stable. It may be that the conformation of the wild-type OEE polypeptides protects them from such proteolysis while in a soluble, lumenal pool of unassembled extrinsic subunits. To further explore the temporal and spatial

271 regulation of OEE subunit assembly with PS II, Hashimoto et al. (1997) conducted a series of experiments in which labeled precursor OEE subunits from pea, spinach or maize were imported into isolated pea, spinach or maize chloroplasts. After 40 min of import in the light, chloroplasts were subfractionated to determine the distribution of the mature OEE subunits between the stroma, the thylakoid lumen, and the thylakoid membrane fractions. In species-homologous polypeptide-import systems, the OEE1, OEE2 and OEE3 precursors were properly imported, processed, and localized to the thylakoid lumen and the thylakoid membrane fractions with a partitioning similar to that seen for the indigenous, pre-existing OEE subunits. Studies on the release of pre-existing and newly imported OEE1 and OEE2 subunits from thylakoid membranes, under conditions shown to remove these subunits from the OEC, showed no difference in binding properties of the indigenous vs. newly imported subunits. This suggests that the newly imported subunits associated with the thylakoid membrane are most likely incorporated into the OEC of PS II. To determine whether the newly imported OEE subunits assemble with pre-existing or nascent PS II cores, similar import studies were carried out in the presence of inhibitors of chloroplast translation (Hashimoto et al., 1997). No difference was seen in partitioning of imported OEE1 and OEE2 subunits in the absence of chloroplast translation, suggesting that these extrinsic subunits can assemble with pre­ existing PS II complexes. Interestingly, labeled OEE2, newly imported and bound to the thylakoid membrane in isolated chloroplasts, could be chased from the thylakoid membrane fraction to the soluble fraction by importing excess, non-labeled OEE2. These authors propose that newly imported OEE subunits actively displace existing OEE subunits bound to the PS II core, rather than assembling only with PS II cores lacking the OEE subunits (Hashimoto et al., 1997). The fate and oflabeled OEE1 and OEE2 precursors was assayed during the first hour after initiation of import into pea chloroplasts in the light (Hashimoto et al., 1997). While protease-protected, thylakoidmembrane-localized mature subunits of both OEE1 and OEE2 eventually partitioned ~80% to the appressed and ~20% to the non-appressed mem­ branes, there was a great difference in the time course of subunit migration from non-appressed to appressed membranes. Ninety percent of imported

272 OEE2 which localized to the thylakoid membrane was found in the appressed membranes at the earliest time points; this slowly decreased to 80% and within 10 min no further change was seen. In contrast, 60% of the newly imported, protected OEE1 was found primarily in the non-appressed membranes at the earliest time points, and this slowly decreased over the course of an hour to 20%, with a concomitant rise in the amount of OEE1 associated with appressed membranes. This suggests that OEE1 is imported into the non-appressed thylakoid membranes, and is translocated subsequently to the appressed mem­ branes. As OEE2 appears very quickly in the appressed membranes, it is not clear whether that is the initial site of import, or whether migration from the stromal to the granal lamellae occurs much more quickly for the OEE2 extrinsic subunit. In Chlamy­ domonas, de Vitry et al. (1989) found that OEE2 was localized by immunocytochemistry to the appressed membranes in wild-type cells, but was found in nonappressed membranes in a mutant lacking CP43. Thus, the location ofOEE2 may depend on its binding to PS II in appressed membranes at a site determined to some extent by CP43. It is not yet clear whether OEE2 import occurs in the non-appressed mem­ branes, as hypothesized for OEE1 and other lumenally localized proteins, followed by the very rapid migration to the appressed membranes where it is stabilized in PS II containing CP43, or whether OEE2 import occurs initially in the appressed membranes followed by migration to the nonappressed membranes if CP43 is absent. A model for the assembly ofthe OEC polypeptides with PS II is incorporated into Fig. 3, based on the above work, on in vivo studies in Chlamydomonas (de Vitry et al., 1989; Eisenberg-Domovich et al., 1995) and on the studies of van Wijk et al. (1997) as discussed (Section III). For further discussion of chloroplast protein import and targeting, see Perret et al. (Chapter 13).

B. In Vivo Requirements for the Extrinsic Membrane Polypeptides of the OEC In the absence of any or all three of the extrinsic membrane polypeptides ofthe OEC, purified spinach PS II membranes are able to evolve oxygen in the presence of elevated levels of calcium and chloride ions (reviewed in Bricker, 1992; Debus, 1992). Thus, the OEC extrinsic membrane subunits are not essential for PS II catalysis of water oxidation in

Jeanne Marie Erickson vitro. The in vivo requirements for the OEC subunits, with respect to photoautotrophic growth of Chlamy­ domonas and cyanobacteria, are discussed below. C. reinhardtii nuclear mutants lacking the extrinsic membrane subunits OEE1 (FUD44 mutant – Mayfield et al., 1987a; de Vitry et al., 1989) or OEE2 (BF25 mutant—Mayfield et al., 1987b; de Vitry et al., 1989: FUD39 mutant—de Vitry et al., 1989; Rova et al., 1994; Rova et al., 1996) have been characterized. Results show that in the absence of the OEE1 polypeptide, mutant cells are completely deficient in photosynthetic oxygen-evolution and are unable to grow photoautotrophically. In contrast, mutants lacking the OEE2 polypeptide can evolve oxygen, albeit at rates lower than wild-type cells, and are able to grow photoautotrophically. In the absence ofeither OEE1 or OEE2, the other OEE subunits appear to accumulate normally in the chloroplast. However, thylakoid membranes isolated from the FUD39 mutant (OEE2-minus) contain slightly lower levels ofOEE3 than those of wild-type cells (de Vitry et al., 1989). This might be expected if the OEE3 bindingsite on PS II is determined by the presence of OEE2, as in vitro results with spinach indicate (Miyao and Murata, 1989). Interestingly, an excess of OEE1 subunits appears to have no effect on PS II function. In engineered transformants of FUD44 that contain multiple copies of psbO and accumulate OEE1 at levels ~3 times that of wild-type cells, excess OEE1 is localized to the thylakoid lumen but does not bind in excess to the thylakoid membrane, accumulation of the other PS II polypeptides is normal, and photosynthetic oxygen-evolution is equivalent to that of wild-type cells (Mayfield, 1991). The absence of either one of the OEE1 or OEE2 subunits does affect the accumulation of the core PS II polypeptides. A substantial reduction in the accumulation ofthe PS II core subunits was reported to occur via their rapid turnover in the absence of the extrinsic OEE1 subunit (Mayfield et al., 1987a; de Vitry et al., 1989). A less dramatic reduction in the accumulation of the core polypeptides was seen by de Vitry et al. (1989) in both the FUD39 and BF25 mutants lacking OEE2, while Mayfield et al. (1987b) reported nearly wild-type levels of the PS II core in BF25. Recently, Rova et al. (1994, 1996) have reexamined FUD39 and shown that this OEE2-minus mutant has an increased chloride requirement for photoactivation and is particularly sensitive to photoinhibition under normal growth conditions, resulting in reduced accumulation of PS II subunits.

Chapter 15

Assembly of Photosystem II

However, these authors find that FUD39 contains wild-type levels of PS II under low-light growth conditions. Moreover, elevated levels of chloride decrease the frequency of irreversible PS II photoinhibition in the absence of OEE2 (Rova et al., 1994). Photoactivation in FUD39 was also found to ), be optimal under low light incubation ( and the photoactivation process in this mutant has a very narrow light intensity range, suggesting that photoactivation competes with photoinhibition (Rova et al., 1996). Based on their studies with FUD39, these authors proposed a model for OEC assembly in which photoligation of the second manganese in the absence of chloride can lead to the irreversible photoinhibition of PS II. In the absence of an active oxygen-evolving complex, PS II is more sensitive to photoinhibition (Blubaugh and Cheniae, 1990; Klimov et al., 1990; Minagawa et al., 1996), and suffers increased damage to the D1 reaction center polypeptide (Jegerschöld et al., 1990; Chapter 30, Keren and Ohad). Thus, it is not surprising that lack of the OEE1 arid OEE2 polypeptides in vivo, which results in the absence of or reduction in PS II photoactivation, respectively, leads to the loss of PS II polypeptides due to increased photoinhibition. The chloride-dependent phenotype of the OEE2­ minus mutant reaffirms the role of OEE2 in promoting the effect of chloride on photoactivation and oxygenevolution (reviewed in Debus, 1992). While the C. reinhardtii OEE1 -minus mutant is an obligate heterotroph (Mayfield et al., 1987a), cyanobacterial mutants lacking the homologue of the OEE1 polypeptide are photoautotrophic, though with a reduced number of PS II reaction centers (Burnap et al., 1992), an increased sensitivity to photoinhibition (Mayes et al., 1991; Philbrick et al., 1991), an increased OEC instability in the dark (Engels et al., 1994; Burnap et al., 1996) and a in the growth medium (Philbrick requirementfor et al., 1991; Engels et al., 1994). A chloride requirement for growth of the OEE 1-minus cyanobacterial strains is controversial. OEE1 is the only OEC polypeptide which is common to eukaryotes and cyanobacteria. Cyanobacteria lack homologues for OEE2 and OEE3, but do contain two other smaller subunits—the psbU (12 kDa poly­ peptide) and psbN (Cyt c-550) gene products (Shen and Inoue, 1993; reviewed in Seidler, 1996)—which are not found in chloroplasts. These differences may account in part for the fact that OEE1 is required for the photoautotrophic growth of Chlamydomonas,

273 but not of cyanobacteria. An extensive comparative analysis of the extrinsic subunits of the OEC and the phenotype of OEE 1-minus mutants in Chlamy­ domonas and cyanobacteria has been made (Seidler, 1996; Trost et al., 1997; Chapter 16, Ruffle and Sayre).

V. Assembly of Manganese: The Catalytic Center of the Oxygen-Evolving Complex

A. Photoactivation: Assembly of the Functional Manganese Cluster The functional oxygen-evolving complex (OEC) is the catalytic center of PS II water oxidation. In conjunction with photooxidation of P680, the assembled OEC cycles between five oxidation states, identified as sequential transitions from the least oxidized state ( ) to the most oxidized state ( ), i.e., with light being required for each of these four transitions (Joliot and Kok, 1975; reviewed in Cheniae, 1993; Britt, 1966; Chapter 16, Ruffle and Sayre). The state is highly unstable and the transition occurs with concomitant release of molecular oxygen. is the dark-stable state of the OEC. The S-states most likely represent increasingly oxidized states of a tetranuclear manganese cluster (reviewed in Britt, 1996). Current models for the chemistry of PS II manganese-mediated water oxidation include the radical catalytic involvement of the D1 tyrosine in the extraction of hydrogen atoms or protons from water, and suggest that the OEC is a metalloradical enzyme (Gilchrist et al., 1995; Hoganson et al., 1995; Hoganson and Babcock, 1997; Tommos and Babcock, 1998). Studies with site-directed mutants of cyanobacteria (Chu et al., 1995) and Chlamy­ domonas (Roffey et al., 1994a; 1994b) implicate histidine residue 190 of the D1 polypeptide as an acceptor of protons from (reviewed in Diner and Babcock, 1996; Chapter 16, Ruffle and Sayre). The structural environment ofthe chloroplast OEC is provided by polypeptides of the intrinsic PS II core complex and by the three OEE extrinsic membrane polypeptides associated with PS II at the lumenal surface of the thylakoid membrane. Water, amino acids, and possibly calcium and chloride ions are predicted to provide the estimated 24 ligands necessary for octahedral coordination of each Mn in the cluster, including possibly 14 bridging ligands

274 between Mn ions (Yachandra et al., 1993). Ligands may be provided by inorganic oxygen, as well as amino acid residues (Britt, 1996; Diner and Babcock, 1996). Extensive site-directed mutagenesis studies aimed at genetic engineering of the D1 polypeptide in Chlamydomonas (Whitelegge et al., 1995) and in cyanobacteria strongly suggest that at least six amino acid residues of D1 (D170, H332, E333, H337, D342, and A344) provide ligands to manganese (reviewed in Diner et al., 1991; Britt, 1996; Diner and Babcock, 1996; Chapter 16, Ruffle and Sayre). D2 may also provide ligands to manganese (Vermaas, 1993). Assembly of functional manganese complexes in PS II membranes or thylakoid membrane preparations depleted of Mn and lacking one or more of the OEE1, OEE2 and OEE3 polypeptides has allowed for dissection of the assembly process (Miyao and Murata, 1984; 1989; Tamura and Cheniae, 1987; Miller and Brudvig, 1989; 1990; Chen et al., 1995 and discussion therein). As light is required for Mn assembly, this process has been called photoactivation (Radmer and Cheniae, 1971). A two-step model for the assembly of the tetranuclear manganese complex was proposed by Tamura and Cheniae (1987). Presumably, a similar process takes place in vivo during PS II assembly. In the first step (Fig. 4A), is bound to the PS II reaction center core, presumably to amino acid residues of the D1 polypeptide, and photooxidized to . A second binds in the same vicinity to form an unstable intermediate. Photooxidation of the second bound manganese results in the formation of a semistable dimer. Alternatively, a dimer may be formed as an intermediate prior to photooxidation (Ananyev and Dismukes, 1997). The quantum efficiency ofphotooxidation of the first two Mn ions appears to be very low. In the second step, two additional ions bind (Fig. 4B). This binding of manganese does not require light and is thought to occur ‘spontaneously’ in the dark, leading to the formation of a tetranuclear Mn complex (Tamura and Cheniae, 1987). S states more reduced than (i.e., to observed in vitro and outlined in Fig. 4(Riggs-Gelascoetal., 1996), may exist in vivo during assembly of the OEC. The most reduced (semi)stable state of the tetranuclear manganese and two cluster may contain two state. In the dark, when Mn equivalent to an photooxidation is not competing with loss of from the cluster, the tetranuclear cluster is destabilized and Mn is released from the membrane.

Jeanne Marie Erickson The dark-stable state of the manganese cluster is proposed to consist of one dimer and one initially binds to dimer (Fig. 4). Assuming that PS II, six successive oxidations of manganese are required for the production of the state.Although the last two ions bind stably in the dark, it is generally accepted that all Mn oxidations are in fact photooxidations mediated through the lightand that the last dependent oxidation of by two photooxidations of are quite efficient (Diner and Babcock, 1996; Riggs-Gelasco et al., 1996; see however, Section V.C). Tamura et al. (1997) conducted a detailed study of the effects of reducing reagents and redox potential on photoactivation in PS II membranes and isolated chloroplasts, and speculate that the lower quantum efficiency of photoactivation observed in reducing environments results from the in partially and reduction and solubilization of fully assembled Mn clusters. The manganese cluster can assemble in the absence of the extrinsic OEC polypeptides, but two of the manganese ions are unstable in the absence of OEE1. Betts et al. (1996a) propose that light-dependent interactions occur between OEE1 and the PS II core during redox cycling of the manganese cluster, based on in vitro studies of PS II thylakoid membrane preparations reconstituted with mutant OEE1 protein. Cyanobacterial mutants lacking OEE1 but containing specific amino acid substitutions in the D1 polypeptide exhibit increased binding or photo­ oxidation of manganese in the absence of OEE 1 in vivo(Chu et al., 1994). OEE1 has significant buffering capacity, and may be involved in a proton pathway between the site of water-splitting and the thylakoid lumen (Shutova et al., 1997). Wydrzynski et al. (1996) propose that the OEE1 extrinsic subunit provides a hydrophobic protein pocket that protects the catalytic site of the OEC and allows for the controlled binding of water substrate molecules. Calcium and chloride ions are essential for OEC function (reviewed in Debus, 1992). They may provide direct ligands to the catalytic Mn cluster, or alter the surrounding protein structural environment of the OEC to facilitate assembly and/or function of the and manganese cluster. The effects of levels on photoactivation have been studied by Ananyev and Dismukes (1996a), who developed a photoactivation assay utilizing an integrated Clarktype oxygen electrode and illumination cell that is highly sensitive, has a fast response time, essentially eliminates photoinhibitory effects, and allows for moles) the detection of minute amounts (~5 x

Chapter 15 Assembly of Photosystem II

275

276 of oxygen from a 5 ml sample of PS II membranes. The membranes are depleted of manganese, calcium and the three extrinsic OEE polypeptides using a lipophilic chelator, and photoactivation is assayed in the presence or absence of specific levels of manganese and calcium ions. Titration ofthe steadystate yield of upon photoactivation indicates that 4 Mn are incorporated per PS II to effect 95% restoration of activity, and that ~1 Mn is involved in the rate-limiting photolytic step (Ananyev and Dismukes, 1996b). Calcium may be bound to PS II prior to each photolytic step in Mn-cluster assembly, as shown in Fig. 4A (Zaltsman et al., 1997). Studies ofphotoactivation in PS II membranes isolated from light-grown, dark-grown, and photoactivated pine cotyledons show that integration of calcium into the OEC precedes integration of Mn during photomay organize activation (Shinohara et al., 1992). the binding site for Mn ions in the apo-OEC complex (Chen et al., 1995), and may interact directly with Mn or through protein- or solvent-derived bridges (Ananyev and Dismukes, 1996b). Cheniae and coworkers propose that calcium may modulate the photoassembly of the Mn cluster by preventing ligation of nonfunctional, high-valency states of manganese (Chen et al., 1995 and references therein). The role of chloride in the photoactivation process is not entirely clear. In vitro studies of binding to PS II membranes depleted of chloride suggest that PS II has one chloride binding site with two distinct states (Lindberg and Andréasson, 1996). Once chloride has been removed from a functional, highrebinds with low-affinity affinity binding site, and exchanges rapidly until, over time, the binding site is converted to the high-affinity state. In vivo studies in a C. reinhardtii mutant lacking the OEE2 polypeptide suggest that chloride is required for (Rova productive photoligation of the second et al., 1996), as shown in Fig. 4B. In the absence of chloride at this step, a non-productive intermediate is formed which leads to irreversible photoinacti­ vation of the OEC. Wincencjusz et al. (1997) have shown that chloride is required for the S-state redox transitions from through but not from through It is noteworthy that there is a correlation between photoactivation of the donor side of P680 and PS II acceptor-side function. Johnson et al. (1995) have shown that PS II photoactivation is associated with a lowering in the midpoint potential ofthe first quinone acceptor, suggesting that donor-side function

Jeanne Marie Erickson may regulate acceptor-side function. Alternatively, modification of the PS II reducing-side may lead to and thus favoring an increased stability of photoactivation (Tamura et al., 1997). The fine regulation between donor- and acceptor-side function may be part of a protective mechanism during PS II assembly, to avoid photoinhibition in the absence of a fully functional OEC (Section IV.B). Clearly, these two functional domains of PS II are intimately linked.

B. A Role for Bicarbonate in Assembly of the OEC

Bicarbonate has long been known to stimulate photosynthetic electron transfer and has been shown to reverse the inhibitory effects of anions and nitric oxide on PS II activity. Evidence exists for donorside (El-Shintinawy and Govindjee, 1990) and acceptor-side binding of bicarbonate (Diner and Petrouleas, 1990; Govindjee et al., 1997). Klimov and coworkers (Klimov et al., 1995; Allakhverdiev et al., 1997) now have experimental evidence suggest­ ing that bicarbonate is a constituent of the wateroxidizing complex of Photosystem II, required for the assembly and function of the OEC. They find that restoration ofoxygen-evolution in Mn-depleted PS II membranes is increased in the presence of bicarbonate and that the effect is independent of pH (pH 5–8). Bicarbonate may modify the binding site for Mn, or may provide a carboxyl group as one of the bridging ligands to Mn. In the case of structural changes on the acceptor-side of PS II that alter donor-side function (Hutchison et al., 1996), it is difficult to unravel the nature of the joint donoracceptor side effects (Johnson et al., 1995). Likewise, in the case of bicarbonate binding, it is possible that bicarbonate binds at both sides of PS II; it is also possible that bicarbonate binding on one side causes structural and functional changes on the other side of the membrane. X-ray crystal studies show that relatively large changes in physical structure are effected by the light-induced redox state of the bacterial reaction center quinones (Stowell et al., 1997).

C. Is Oxygen Required for Assembly of the OEC? Oxygen consumption by thylakoids has long been noted. The ‘Mehler’ reaction (Mehler, 1951) is characterized by electron transfer from PSI to oxygen,

Chapter 15

Assembly of Photosystem II

is coupled to ATP production, and has been observed in many systems. Park et al. (1996) suggest that photoinactivation of PS II in pea leaves is mitigated by electron transfer to oxygen through the Mehler reaction and photorespiration. Provocative work by Ebina and Yamashita (1996) suggests that recovery of oxygen-evolving activity, i.e., assembly of a functional OEC, involves the consumption of oxygen. These authors examined photoactivation of Tris­ inactivated, manganese-depleted spinach thylakoid membranes, using PS II electron donors that did not reduce oxygen directly. The reconstitution of a functional manganese cluster in the presence of micromolar levels of Mn was always accompanied by low levels of oxygen consumption. Ebina and Yamashita (1996) note that electron transfer through PS II is required for the observed oxygen consump­ tion, and invoke a Mehler-type electron transfer from PS II to oxygen, via PS I, as playing an important role in OEC photoreactivation. However, no direct evidence for the involvement of PS I in the observed reduction of oxygen was reported. Indeed, in vivo consumption of oxygen in Chlamydomonas has been observed in the mutant F15 lacking PS I (Peltier and Thibault, 1988) and in site-directed mutants deleted for the PS I reaction center genes psaA or psaB (Cournac et al., 1997), as reviewed (Chapter 18, Redding and Peltier; Chapter 35, Bennoun). Evidence for the consumption of oxygen by PS II comes from studies of Beck et al. (1985) using lowtemperature EPR spectroscopy and measurements of oxygen. These authors correlated changes in PS II oxygen-consumption with changes in the Mn site of the OEC during dark-adaptation of isolated PS II membranes or thylakoid membranes. Following illumination, membrane preparations placed in the dark consumed oxygen when a functional OEC and consumption excess reductant were present; decreased during dark adaptation, with the same time constant as an EPR-detected change in the Mn site of the OEC. Beck et al. (1985) attribute the slow consumption of in the dark to the catalytic reduction of by the OEC, and suggest that during dark-adaptation, a structural change occurs in the Mn site of the OEC that alters PS II donor-side electron transfer causing a conversion of the OEC from the active, -consuming state to a resting state incapable of consumption. Regardless of the mechanism, it is clear that light may activate an -consuming process dependent on electron flow through PS II. Further experiments are

277 needed to identify the site and mechanism of oxygen consumption during photoactivation. The question of whether it takes oxygen to make oxygen in PS II is not a trivial one to answer. It will be interesting to see whether specific chemical reactions utilizing oxygen are intrinsic to PS II itself, and what their relationship, if any, is to the assembly or maintenance of a functional oxygen-evolving manganese cluster.

Acknowledgments I thank Bruce Diner, Gary Brudvig, Charles Yocum, Gerald Babcock, and Terry Bricker for helpful discussions and clarifications, Paul Mahoney for excellent assistance with the figures and table, and the editors for critical reading of this chapter.

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Chapter 15

Assembly of Photosystem II

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283 16255–16263 Shestakov SV, Anbudurai PR, Stanbekova GE, Gadzhiev A, Lind LK and Pakrasi HB (1994) Molecular cloning and characterization ofthe ctpA gene encoding a carboxyl-terminal processing protease: Analysis ofa spontaneous Photosystem IIdeficient mutant strain of the cyanobacterium Synechocystis sp. PCC 6803. J Biol Chem 269: 19354–19359 Shinohara K, Ono TA and Inoue Y (1992) Photoactivation of oxygen-evolving enzyme in dark-grown pine cotyledons: Relationship between assembly of Photosystem II proteins and integration of manganese and calcium. Plant and Cell Physiol 33: 281–289 Shutova T, Irrgang K-D, Shubin V, Klimov VV and Renger G (1997) Analysis of pH-induced structural changes of the isolated extrinsic 33 kilodalton protein of Photosystem II. Biochemistry 36: 6350–6358 Staehelin LA and van der Staay GWM (1996) Structure, composition, functional organization and dynamic properties of thylakoid membranes. In: Yocum CF and Ort DR (eds) Oxygenic Photosynthesis: The Light Reactions, pp 11–30. Kluwer Academic Publishers, Dordrecht. Staub J and Maliga P (1994) Translation of psbA. mRNA is regulated by light via the 5´-untranslated region in tobacco plastids. Plant J 6: 547–553 Stowell MHB, McPhillips TM, Rees DC, Soltis SM, Abresch E and Feher G (1997) Light-induced structural changes in photosynthetic reaction center: Implications for mechanism of electron-proton transfer. Science 276: 812–816 Summer EJ, Schmid VHR, Bruns BU and Schmidt GW (1997) Requirement for the H phosphoprotein in Photosystem II of Chlamydomonas reinhardtii. Plant Physiol 113: 1359–1368 Sutton A, Sieburth LE and Bennett J (1987) Light-dependent accumulation and localization of Photosystem II proteins in maize. Eur J Biochem 164: 571–578 Svensson B, Vass I and Styring S (1991) Sequence analysis ofthe D1 and D2 reaction center proteins of Photosystem II. Z Naturforsch 46c: 765–776 Svensson B, Etchebest C, Tuffery P, van Kan P, Smith J and Styring S (1996) A model for the Photosystem II reaction center core including the structure of the primary donor Biochemistry 35: 14486–14502 Takahashi Y, Nakane H, Kojima H and Satoh K (1990) Chromatographic purification and determination of the carboxy­ terminal sequences of Photosystem II reaction center proteins, D1 and D2. Plant Cell Physiol 31: 273–280 Takahashi Y, Goldschmidt-Clermont M, Soen SY, Franzen LG and Rochaix J-D (1991) Directed chloroplast transformation in Chlamydomonas reinhardtii: Insertional inactivation of the psaC gene encoding the iron sulfur protein destabilizes Photosystem I. EMBO J 10: 2033–2040 Takahashi Y, Matsumoto H, Goldschmidt-Clermont M and Rochaix J-D (1994) Directed disruption ofthe Chlamydomonas chloroplast psbK gene destabilizes the Photosystem II reaction center complex. Plant Mol Biol 24: 779–788 Takahashi Y, Utsumi K, Yamamoto Y, Hatano A and Satoh K (1996) Genetic engineering of the processing site of D1 precursor protein of Photosystem II reaction center in Chlamydomonas reinhardtii. Plant Cell Physiol 37: 161–168 Tamura N and Cheniae G (1987) Photoactivation of the wateroxidizing complex in Photosystem II membranes depleted of Mn and extrinsic proteins. I. Biochemical and kinetic

284 characterization. Biochim Biophys Acta 890: 179–194 Tamura N, Kuwahara M, Sasaki Y, Wakamatsu K, and Oku T (1997) Redox dependence for photoligation of manganese to the apo-water-oxidizing complex in chloroplasts and Photosystem II membranes. Biochemistry 36: 6171–6177 Tanaguchi M, Kuroda H and Satoh K (1993) ATP-dependent protein synthesis in isolated pea chloroplasts. Evidence for accumulation of a translation intermediate of the D1 protein. FEBS Lett 317: 57–61 Taylor WC (1989) Regulatory interactions between nuclear and plastid genomes. Annu Rev Plant Physiol Plant Mol Biol 40: 211–233 Thomas H (1997) Chlorophyll: a symptom and a regulator of plastid development. Tansley Review No. 92, pp 163–181 Tommos C and Babcock GT (1998) Oxygen production in nature: a light-driven metalloradical enzyme process. Accts Chem Res, in press Trebst A (1986) The topology of the plastoquinone and herbicide binding peptides of Photosystem II in the thylakoid membrane. Z Naturforsch 41c: 240–245 Trebst A and Depka B (1997) Role of carotene in the rapid turnover and assembly of Photosystem II in Chlamydomonas reinhardtii. FEBS Lett 400: 359–362 Trost JT, Chisholm DA, Jordan DB and Diner BA (1997) The D1 C-terminal processing protease of Photosystem II from Scenedesmus obliquus: Protein purification and gene characterization in wild type and processing mutants. J Biol Chem 272: 20348–20356 Vallon O, Wollman F-A and Olive J (1986) Lateral distribution of the main protein complexes of the photosynthetic apparatus in Chlamydomonas reinhardtii and in spinach: An immuno­ histochemical study using intact thylakoid membranes and a PS II enriched membrane preparation. Photobiochem Photobiophys 12: 203–220 Vallon O, Hoyer-Hansen G and Simpson DJ (1987) Photo­ system II and cytochrome b-559 in the stroma lamellae of barley chloroplasts. Carlsberg Res Commun 52: 405–421 van Wijk KJ and Eichacker L (1996) Light is required for efficient translation elongation and subsequent integration of the D1-protein into Photosystem II. FEBS Lett 388: 89–93 van Wijk KJ, Nilsson LO and Styring S (1994) Synthesis of reaction center proteins and reactivation of redox components during repair ofPhotosystem II after light-induced inactivation. J Biol Chem 269: 28382–28392 van Wijk KJ, Bingsmark S, Aro E-M and Andersson B (1995) In vitro synthesis and assembly of Photosystem II core proteins. J Biol Chem 270: 25685–25695 van Wijk KJ, Andersson B and Aro E-M (1996) Kinetic resolution of the incorporation of the D1 protein into Photosystem II and localization ofassembly intermediates in thylakoid membranes of spinach chloroplasts. J Biol Chem 271: 9627–9636 van Wijk KJ, Roobol-Boza M, Kettunen R, Andersson B and Aro E-M (1997) Synthesis and assembly of the D1 protein into Photosystem II: Processing of the C-terminus and identification of the initial assembly partners and complexes during Photosystem II repair. Biochemistry 36: 6178–6186 Vermaas W (1993) Molecular-biological approaches to analyze Photosystem II structure and function. Annu Rev Plant Physiol Plant Mol Biol 44: 457–481 Vermaas W, Charité J and Shen G (1989) binding to D2 contributes to the functional and structural integrity of

Jeanne Marie Erickson Photosystem II. Z Naturforsch 45c: 359–365 Vermaas WFJ, Styring S, Schröder WP and Andersson B (1993) Photosynthetic water oxidation: The protein framework. Photosynth Res 38: 249–263 Webber AN and Baker NR (1996) Control of thylakoid membrane development and assembly. In: Yocum CF and Ort DR (eds) Oxygenic Photosynthesis: The Light Reactions, pp 41–58. Kluwer Academic Publishers, Dordrecht Wettern M and Ohad I (1984) Light induced turnover of thylakoid polypeptides in Chlamydomonas reinhardtii. Isr J Bot 33: 253–263 Wettern M, Owens JC and Ohad I (1983) Role of thylakoid polypeptide phosphorylation and turnover in the assembly and function of Photosystem II. Methods in Enzymology 97: 554– 567 Whitelegge JP, Koo D, Diner BA, Domian I and Erickson JM (1995) Assembly of the Photosystem II oxygen-evolving complex is inhibited in psbA. site-directed mutants of Chlamydomonas reinhardtii: Aspartate 170 of the D1 polypeptide. J Biol Chem 270: 225–235 Whitmarsh J and Pakrasi HB (1996) Form and function of cytochrome b-559. In: Yocum CF and Ort DR (eds) Oxygenic Photosynthesis: The Light Reactions, pp 249–264. Kluwer Academic Publishers, Dordrecht Wincencjusz H, van Gorkom HJ and Yocum CF (1997) The photosynthetic oxygen evolving complex requires chloride for its redox state transitions but not for transitions. Biochemistry 36: 3663–3670 Wu G-J and Watanabe A (1997) Import of modified D1 protein and its assembly into Photosystem II by isolated chloroplasts. Plant Cell Physiol 38: 243–247 Wu HY and Kuchka MR (1995) A nuclear suppressor overcomes defects in the synthesis of the chloroplast psbD gene product caused by mutations in two distinct nuclear genes of Chlamydomonas. Curr Genet 27: 263–269 Wydrzynski T, Hillier W and Messinger J (1996) On the functional significance of substrate accessibility in the photosynthetic water oxidation mechanism. Physiologia Plantarum 96: 342– 350 Xiong J, Subramaniam S and Govindjee (1996) Modeling of the D1/D2 proteins and cofactors of the Photosystem II reaction center: Implications for herbicide and bicarbonate binding. Protein Science 5: 2054–2073 Xiong J, Subramaniam S and Govindjee (1998) A knowledgebased three-dimensional model of the Photosystem II reaction center of Chlamydomonas reinhardtii. Photosynth Res, in press Xu Q and Bricker TM (1992) Structural organization of proteins on the oxidizing side of Photosystem II: Two molecules of the 33 kDa manganese-stabilizing protein per reaction center. J Biol Chem 267: 25816–25821 Yachandra VK, DeRose VJ, Latimer MJ, Mukerji I, Sauer K and Klein MP (1993) Where plants make oxygen: A structural model for the photosynthetic oxygen-evolving manganese cluster. Science 260: 675–679 Yamamoto Y (1988) Analysis of the relationship between the extrinsic 30-kDa protein, manganese and oxygen evolution in the thylakoid of Chlamydomonas reinhardtii grown under manganese-deficient conditions. Biochim Biophys Acta 933: 165–171 Yamamoto HE and Bassi R (1996) Carotenoids: localization and

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Assembly of Photosystem II

function. In: Yocum CF and Ort DR (eds) Oxygenic Photosynthesis: The Light Reactions, pp 539–563. Kluwer Academic Publishers, Dordrecht Yohn CB, Cohen A, Danon A and Mayfield SP (1996) Altered mRNA binding activity and decreased translation initiation in a nuclear mutant lacking translation of the chloroplast psbA mRNA. Mol Cell Biol 16: 3560–3566 Zaltsman L, Ananyev GM, Bruntrager E and Dismukes GC (1997) Quantitative kinetic model for photoassembly of the photosynthetic water oxidase from its inorganic constituents: Requirements for manganese and calcium in the kinetically

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Chapter 16 Functional Analysis of Photosystem II Stuart V. Ruffle and Richard T. Sayre

Departments of Biochemistry and Plant Biology, 2021 Coffey Road,

Ohio State University, Columbus, OH 43210, U.S.A.

Summary I. Introduction II. The Photosystem II Complex A. Photosystem II Particles B. D1 and D2 Polypeptides C. P680, the Primary Donor D. Chlorophyll Monomers and

an Extra Pair of Chlorophylls in Photosystem II E. Pheophytin Redox Active Components of the D1 and D2 Proteins F. and G. Water Oxidation and the Tetra-Manganese Complex H.

the Stable Electron Acceptor of Photosystem II and the Herbicide Binding Site I. Cytochrome J. The PS ll-l Protein K. CP43 and CP47, the Antenna Chlorophyll-Protein Complex of Photosystem II III. The Chloroplast DNA Encoded Small Polypeptides of Photosystem II

A. The PS II-H Protein B. The PS II-J Protein C. The PS II-K Protein D. The PS II-L Protein E. The PS II-M Protein F. The PS II-N Protein G. The PS II-T Protein IV. The Nucleus Encoded Polypeptides of the Photosystem II Complex A. OEE1, the 33 kD Oxygen Evolving Complex Protein B. OEE2, the 23 kD Oxygen Evolving Complex Protein C. OEE3, the 17 kD Oxygen Evolving Complex Protein D. Additional Nucleus-Encoded Small Photosystem II Subunits V. Perspectives Acknowledgments References

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Summary The structural and functional analysis of the Photosystem II complex has been facilitated by the use of model organisms which are amenable to experimental manipulation. The unicellular alga, Chlamydomonas reinhardtii, was initially chosen as a model organism based on the relative ease with which Photosystem II mutants could be generated and characterized. Furthermore, cells grown in the dark (heterotrophically) were demonstrated to synthesize chlorophyll, and assemble Photosystem II complexes which were competent for charge separation J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 287–322. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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and photoactivation of the water splitting complex. These traits proved to be particularly useful for the phenotypic characterization of mutants susceptible to photoinhibition. With the recent development of chloroplast and nuclear DNA transformation technologies, however, it has been possible to engineer genes encoding polypeptides of the Chlamydomonas Photosystem II complex. As a result ofthese developments Chlamydomonas has become the dominant model system for genetic manipulation of the eukaryotic Photosystem II complex. In this chapter we describe the structure and primary processes of the Photosystem II complex with emphasis on results obtained from Chlamydomonas. Models for the organization of the Chlamydomonas reaction center, core, and thylakoid membrane Photosystem II complexes are presented. These models are discussed in the context of the partial reactions which each Photosystem II complex type is capable of supporting, both in wild type and mutant cells. Finally, comparisons are drawn between Chlamydomonas (eukaryotic) and cyanobacterial (prokaryotic) Photosystem II mutants of similar genotypes. It is evident from these studies that the two Photosystem II reaction center types are similar but not identical in structure and function.

I. Introduction The primary event in photosynthesis is the transfer of an electron from a light generated excited state chlorophyll(s) (Chl) to a stable electron acceptor. This primary reaction takes place in membrane embedded protein-chromophore complexes known as photosystems. The photosystems or photosynthetic reaction center complexes bind the chromophores and primary and intermediate electron donors and acceptors involved in charge separation. The complexes stabilize the charge separated state and allow for forward electron transfer with high quantum efficiency. Not all photosynthetic reaction center complexes are identical, however. Results from biochemical and biophysical surveys of photosynthetic complexes from a variety ofbacterial and eukaryotic organisms has lead to the realization that there are two different reaction center types (reviewed by Blankenship, 1992; Golbeck, 1993). Members of each reaction center Abbreviations: Chl – chlorophyll; – chlorophyll special – pair; CP – chlorophyll binding polypeptide; cytochrome DCBQ – 2,6-dimethylbenzoquinone; DCMU – 3-(3,4 dichlorophenyl)-l,1-dimethylurea; DPIP – 2,6-dichloroindophenol; – midpoint potential; ENDOR – electron nuclear double resonance; EPR – electron paramagnetic resonance; EXAFS – extended X-ray absorption and fine structure spec­ troscopy; Fe-S – iron-sulfur; FTIR – Fourier transform infrared spectroscopy; kD – kilodalton; Km– Michaelis constant; nicotinamide-adenine dinucleotide phosphate (oxidized form); OEE1 – oxygen evolution enhancer 1; OEE2 – oxygen evolution enhancer 2; OEE3 – oxygen evolution enhancer 3; PS I – Photosystem I; PS II – Photosystem II; SDS-PAGE – sodium dodecyl maltoside polyacrylamide gel eletrophoresis; XANES – X-ray absorbance near-edge spectroscopy

type share some degree of protein structural similarity and have common redox active components. The two reaction center types can be readily distinguished by the identity of their first stable electron acceptor and are known as the quinone and Fe-S reducing photosynthetic reaction center types (reviewed by Blankenship, 1992). The primary charge separation event in both reaction center types is the transfer of an electron from a chlorophyll excited state singlet to a nearby electron acceptor. Beyond this primary event, however, the quinone and Fe-S reaction center types have little in common. Each reaction center type has unique protein subunits, redox active intermediates, and sources of electrons for the reduction of the oxidized primary donor. With the exception of the cyanobacteria and Prochlorophytes, all prokaryotes have either a quinone or Fe-S reaction center complex type but not both. The eukaryotic photosynthetic organisms, however, have both reaction center types (Photosystem II and I) and share the distinction with the cyanobacteria that they can oxidize water. The Photosystem II complex (quinone type) of oxygenic photosynthetic organisms carries out the light dependent oxidation of water and reduction of plastoquinone. Note, however, that only cyanobacterial and eukaryotic quinone type reaction centers are capable of water oxidation. All other bacterial quinone type reaction centers do not oxidize water. The Photosystem I complex (Fe-S type) carries out the light dependent oxidation of a soluble copper containing (plastocyanain) or heme protein and the reduction of an Fe-S protein The two (ferredoxin) which in turn reduces photosystems (PS I and PS II) function in series

Chapter 16

Photosystem II

transferring electrons from water to via the complex, while releasing or cytochrome translocating protons across a membrane to drive ATP synthesis. Historically, our knowledge of the biochemistry and biophysics of PS II has relied on a limited number of plant and algae as sources of abundant and stable PS II complexes. Spinach, pea, corn and tobacco have been some of the more common sources of PS II material. Vascular plant sources of PS II require long growing periods and have not been particularly amenable to genetic manipulation. Since the early 1980s cyanobacteria have moved to the forefront as the organisms of choice for PS II studies. The shift in emphasis from higher plant to cyanobacterial sources of PS II complexes was based primarily on the ease with which cyanobacteria could be genetically modified. In addition, it has been demonstrated that cyanobacteria can selectively incorporate isotopically labeled amino acids into their reaction center polypeptides. In the last five years, however, the green alga C. reinhardtii, has re-emerged as a model eukaryotic PS II organism, based primarily on its ability to be genetically engineered (Boynton and Gillham, 1993; Hutchison et al., 1996a). Historically, Chlamy­ domonas has been a rich source of photosynthetic material and randomly generated PS II mutants. With the growing realization that prokaryotic and eukaryotic PS II complexes share many features but also have many differences, the future of Chlamy­ domonas as a tool for PS II research appears bright. In the following sections we describe energy and electron transfer reactions which are carried out within the framework ofsimple and more complicated PS II particles with a focus on Chlamydomonas. For recent descriptions of PS II the reader is directed to the excellent reviews on PS II presented in recent volumes in this series (Barry et al, 1994; Bricker and Ghanotakis, 1996; Britt, 1996; Diner and Babcock, 1996; Satoh, 1996) and by Debus (1992). We begin with a general description ofthe structure and function of the PS II complex.

II.The Photosystem II Complex The oxygen evolving PS II complex is a multisubunit protein-chromophore complex containing at least 20 polypeptides and five different redox active components (chlorophyll, pheophytin, plastoquinone,

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tyrosine and manganese) (Fig. 1A). The primary electron donors and acceptors of the PS II complex are bound by two highly similar 32 kD polypeptides known as the D1 and D2 proteins (Fig. 2). Associated with the reaction center complex are the antenna chlorophyll binding polypeptides, CP43 and CP47, which transfer energy to the primary electron donor. Many small polypeptides, whose function is being determined largely by deletion mutagenesis studies and comparative analysis of PS II particles, are also associated with the complex (Table 1). The primary donor of PS II is known as P680 which may be one, to several, chlorins. P680, when excited by visible light, transfers an electron from to a pheophytin on the the excited state, picosecond time scale (Fig. 3). The pheophytin reduces the first stable electron acceptor, a bound plastoquinone molecule, which typically accepts only subsequently transa single electron. Reduced fers an electron to a stable two electron acceptor site. The plastoquinone molecule bound to the

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site plastoquinone moves out of the site when it is doubly reduced and protonated. The oxidized primary donor, , is reduced by a tyrosine residue

(Y161) located on the reaction center D1 polypeptide.

is in turn reduced by extraction of an electron

from the tetra-Mn, water oxidizing complex. Following the sequential extraction of four electrons (S-state transitions) from the water oxidizing complex, molecular oxygen is released (Kok et al., 1970; Britt, 1996). Four protons are also released during water oxidation in a step-wise process associated with the extraction of each electron.

A. Photosystem II Particles The determination of the structural composition of the PS II complex has benefited from comparative analyses of different PS II particle preparations which are capable of carrying out water oxidation and

Stuart V. Ruffle and Richard T. Sayre

reduction of or some subset of the PS II electron transfer reactions. The earliest oxygen evolving PS II particles isolated from chloroplasts were obtained by detergent (Triton X-100) solubilization of spinach thylakoid membranes in the presence of5 mM (Berthold et al., 1981). These so called ‘BBY’ type PS II particles were capable of high rates (approx. of oxygen evolution and presumably contained all ofthe proteins and cofactors required for maximum rates of oxygen evolution. In 1990, Shim et al. adapted the BBY particle preparation protocol from higher plants for the isolation of oxygen evolving particles from Chlamydomonas thylakoid membranes. The Chlamy­ domonas BBY-type particles contained approx­ imately 250 chlorophylls per reaction center but had a more complex polypeptide profile than BBY particles isolated from spinach.

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Photosystem II

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Smaller PS II core particles having 40–50 chlorophylls per reaction center (40–50 chlorophylls/ ) were isolated from Chlamydomonas membranes following solubilization with the detergents digitonin and Triton X-100, sucrose density gradient fraction­ ation, and ion exchange chromatography in the presence of lauryl maltoside (de Vitry et al., 1991). The particles contained the D1, D2, CP43, CP47, subunits plus several small cytochrome polypeptides. The particles did not evolve oxygen to but were capable of electron transfer from On size exclusion chromatography columns the particles behaved as monomers with a molecular weight of 440–510 kD. The first oxygen evolving PS II core particles isolated from Chlamydomonas were prepared by solubilization with dodecyl maltoside in the presence followed by density gradient centrifugation of and anion exchange chromatography. These particles had 40 Chl per 2 pheophytins and in the presence of and DCBQ (2,6-dimethylbenzoquinone) were capable of oxygen evolution rates of site herbicide, DCMU (3-(3,4 The dichlorophenyl)-1,1-dimethylurea), reduced the rate

Stuart V. Ruffle and Richard T. Sayre by 40% indicating some damage to the site. The particles contained the following polypeptides: CP47, CP43, D1 , D2, OEE1 (oxygen evolution enhancer 1), and four small polypeptides including one Cyt per two pheophytins (Bumann and Oesterhelt, 1996) (Fig. 1B). This method for isolating oxygen evolving PS II particles is particularly attractive for potential crystallization attempts since only one detergent is used. The least complex and smallest particle capable of charge separation is the reaction center preparation. PS II reaction center particles capable of oxidizing P680 and reducing pheophytin were first isolated by Triton X-100 solubilization of spinach thylakoid membranes and anion exchange chromatography. These particles contained the D1, D2, Cyt subunits and the psbI protein (Fig. 1C) (Nanba and Satoh, 1987). Alizadeh et al. (1995) reported the isolation of a PS II reaction center particle from a Chlamydomonas mutant (F54-14). This mutant lacked the PS I and ATP synthase complexes. Polypeptides from both complexes were frequently found as contaminants in earlier PS II reaction center preparations. A higher Triton X-100 to chlorophyll ratio (50:1) was required for solubilization of Chlamydomonas thylakoid membranes than for the corresponding treatment in spinach. The solubilized fractions were then bound to an anion exchange column and washed with buffer containing dodecyl maltoside to remove the antennae chlorophyll protein complexes. The reaction center particles were eluted using a salt gradient. The particles contained: 6 chlorophyll a/2 pheophytin/1 Cyt /l-2carotenoids and were capable of the light dependent oxidation of P680 and reduction ofpheophytin. Such particles are particularly useful for spectroscopic investigations of primary charge separation events.

B. D1 and D2 Polypeptides The initial identification of many of the PS II reaction center polypeptide components was accomplished using C. reinhardtii PS II mutants. The identification of the 32 kD D1 protein (diffuse band-1) as a PS II component was achieved by comparative analyses of the polypeptide profiles of Chlamydomonas PS II (minus) mutants and wild-type thylakoids (Chua and Bennoun, 1975). The D1 protein was further recognized as the most rapidly synthesized and turning over protein in thylakoids and as the protein which was radiolabeled by

Chapter 16 Photosystem II analogues of atrazine, a site herbicide (Grebanier et al., 1978; Pfister et al., 1981) Subsequent isolation of the D1 protein and determination of its partial protein sequence eventually lead to the recognition of its chloroplast gene (psbA) and determination of its complete protein sequence from the gene sequence (Fig.4)(Zurawski et al., 1982; Erickson et al., 1984). The identification of C. reinhardtii site-specific mutations in the psbA gene which were linked with resistance to site herbicides further demonstrated that the D1 protein was a component of the PS II complex. The analysis of PS II mutants of Chlamydomonas also was instrumental in identification of the D2

293 protein. A second diffuseband, the 33 kD D2 thylakoid protein rapidly turned over in Chlamydomonas PS II mutants which did not synthesize the D1 polypeptide. The gene encoding the D2 polypeptide was identified subsequently in C. reinhardtii by J.-D. Rochaix’s group. Using the psbA gene as a probe on low stringency hybridization Southern blots a chloroplast gene (psbD) encoding the D2 protein was identified, cloned and sequenced (Rochaix et al, 1984). It was apparent from the predicted protein sequence of the psbA and psbD genes that the D1 and D2 proteins had 25% identity and shared similar protein sequence domains (Figs. 4 and 5). The high degree of protein sequence similarity

294

between the PS II D1 and D2 proteins and the Rhodobacter sp. bacterial photosynthetic reaction center L and M subunits was first recognized by Williams et al. (1984, 1986). The significance of the sequence similarity was not fully appreciated, however, until the resolution of the crystal structure of the Rhodobacter (Rhodopseudomonas) viridis quinone-type photosynthetic reaction center. In 1985, the crystal structure of the Rp. viridis bacterial reaction center complex was resolved at 3 Å, and then refined

Stuart V. Ruffle and Richard T. Sayre

to 1.6 Å resolution by X-ray diffraction (Deisenhofer et al., 1985; Michel and Deisenhofer, 1988). The bacterial photosynthetic reaction center complex is composed of two membrane embedded polypeptides, the L and M subunits, a largely extrinsic H subunit, and cytochrome c. The L and M subunits coordinate and/or bind the chromophores (four bacterio­ chlorophylls, two bacteriopheophytins, and a and involved carotenoid) and two quinones in primary charge transfer. The cytochrome c contains

Chapter 16 Photosystem II four hemes, functionally linked in series, which reduce the oxidized primary donor or bacteriochlorophyll special pair. Charge separation leads to the transient reduction of bacteriopheophytin, possibly mediated by a bacteriochlorophyll monomer (Arlt et al., 1993). Similar to the chlorophyll special pair, the chlorophyll monomers and pheophytins display C2 symmetry; however, the electron moves on only one side of the complex from the primary donor to bacterio­ pheophytin. Bacteriopheophytin subsequently reduces the first stable electron acceptor, a quinone quinone is a located in the binding site. The single electron acceptor which reduces the two electron acceptor quinone in the binding site. Williams et al. (1983, 1984) recognized that the D1 and D2 polypeptides had substantial (25%) sequence identity with the L and M subunits, respectively. Furthermore, much of the conserved sequence was clustered in domains common in all four (L, M, D1, and D2) polypeptides. These domains included histidine residues which coordinated the chlorophylls of the special pair and the non-heme iron in the bacterial photosynthetic reaction center. Furthermore, it was apparent from hydropathy plot analyses that the L and M and D1 and D2 polypeptides probably shared similar secondary structures, i.e. each protein contained several (5) hydrophobic potentially membrane spanning domains. Based on the conserved structural and functional similarities site (L and D1 mutants conferring resistance to herbicides), Trebst (1986a,b) predicted the folding topology of the D1 and D2 polypeptides. Similar to the L and M subunits, the D1 and D2 proteins were predicted to have five transmembrane spanning alpha helices. The folding topology of the D1 and D2 proteins was later confirmed by mapping proteolytic peptide fragments of the D1 and D2 proteins which were identified by antibodies generated against synthetic peptides (corresponding to predicted extrinsic domains) (Sayre et al., 1986; Trebst, 1986a). Demonstration that the D1 and D2 polypeptides bound the non-protein cofactors (P680 and pheo­ phytin), involved in primary charge separation, was confirmed with the isolation of a PS II reaction center complex (containing the D1, D2, Cyt subunits, and the PS II-I protein) which was capable of charge separation (Nanba and Satoh, 1987). This minimal complex necessary for charge separation was capable of carrying out the light-dependent oxidation of P680 and the reduction of pheophytin. an sites were damaged, however, in the The

295 reaction center preparation. Based on the bacterial photosynthetic reaction center crystal structure, models of the tertiary and quaternary structure ofthe D1 and D2 reaction center polypeptides have been proposed (Svensson et al., 1990, Brookhaven Protein Databank-1DOP; Ruffle et al., 1992, Xiong et al., 1996). In agreement with experimental observations, these PS II reaction center models predict that D1 and D2 polypeptides have five transmembrane spans, and coordinate or bind the primary and secondary electron donors and acceptors (chlorophylls, pheophytins and quinones) between the heterodimeric complex (Fig. 2). It is apparent, however, that these models do not adequately address the functional differences between the two reaction center types. Unlike the bacterial reaction center, the PS II reaction center oxidizes water, catalyzed by a tetra-Mn complex. Therefore, protein ligands which coordinate the tetra-Mn water splitting complex are not likely to be conserved in the bacterial reaction center complex. In addition, the structure and redox properties ofthe PS II primary donor, P680, and intermediate electron donors and differ from the bacterial reaction acceptors (e.g. center (Barry and Babcock, 1987; Durrant et al., 1993; Schelvis et al., 1994). Other significant differences between the two reaction center types include: the apparent lack of conservation of the L and M subunit accessory bacteriochlorophyll ligands in the D1 and D2 proteins, and the lack of sequence similarity between the amino-terminal transmem­ brane spanning domains (span A) of the bacterial L and M and PS II D1 and D2 polypeptides (Svensson et al., 1990).

C. P680, the Primary Donor The primary donor of PS II, P680, is unique among primary donor chlorophylls in that its mid-point potential is shifted approximately 370 mV from the midpoint potential of oxidized free chlorophyll in solution. The high (1.1 V) midpoint potential of provides the driving force for water oxidation (average midpoint potential for water oxidation = 0.86 V). With the exception of PS II, the midpoint potential of the primary donor chlorophylls is negatively shifted in other reaction center types. It is apparent from comparative spectroscopic analyses of the Rp. viridis primary donor and P680 that the primary donors are dissimilar. In contrast to the Rp. viridis chlorophyll special pair, the chlorophyll(s) of

296 P680, are not strongly coupled (van der Vos et al., 1992; Durrant et al., 1993; Noguchi et al., 1993; Rigby et al., 1994; Schelvis et al., 1994). In fact, P680, has properties of both a chlorophyll monomer and a chlorophyll dimer (reviewed in van Gorkom and Schelvis, 1993). Properties which suggest P680 is a chlorophyll monomer include: 1) optical and EPR (electron paramagnetic resonance) properties of the cation and triplet states which suggest that these states are localized on a monomer (tilted 30° relative to the membrane plane similar to the bacterial accessory chlorophyll monomers), 2) the absence of absorbance an appreciable red shift in the P680 band as expected for a chlorophyll dimer, and 3) Stark effect and hole burning measurements which do not show features predicted for a chlorophyll dimer (reviewed in van Gorkom and Schelvis, 1993). There are, however, features which suggest that P680 is a dimer including: 1) conservation of the histidine ligands on the D1 and D2 proteins which coordinate the (chlorophyll special pair) on the L and M subunits, and 2) the observation that the initial bleaching of P680, upon selective excitation, has twice the amplitude expected for a monomer (van Gorkom and Schelvis, 1993; Schelvis et al., 1994). To account for these discrepancies, Shelvis et al. (1994) proposed that P680 is a chlorophyll dimer with monomeric properties attributed to an anti­ transition parallel orientation of the chlorophyll moments. They proposed that this dimeric structure could account for the observations that the excited state transition is localized on one molecule. Recently, Noguchi et al. (1993) have proposed, on the basis of FTIR (Fourier transform infrared spectroscopy) studies of the P680 triplet, that the hydrogen bonding to the chlorophylls of the special pair is asymmetric and that most of the triplet population is localized on one chlorophyll of the dimer. They propose that the P680 chlorophylls are coordinated by the conserved histidines (D1-H198 and D2-H198) and that the chlorophyll monomer located near the D1 protein is tilted 30° relative to the membrane plane as reported by van Mieghem et al., (1991). In contrast, the chlorophyll located near the D2 protein is oriented perpendicular to the membrane plane. These orientations are, in part, determined by asymmetric hydrogen bonding interactions between the chloro­ phylls and proteins. It was not apparent from this model, however, what the orientation of the macrocycle rings is (see Shelvis et al., 1993). The comparative effects of mutagenesis of the

Stuart V. Ruffle and Richard T. Sayre ligands in bacterial and PS II reaction centers has lead to insights into the organization of P680. ligand, D2-H197 Mutagenesis of the PS II (corresponding to D2-H197 in C. reinhardtii) to a tyrosine residue in the cyanobacterium, Synechocystis sp. PCC 6803, resulted in a loss of the ability to evolve oxygen, altered chlorophyll fluorescence emission kinetics and an inability to bind the herbicides atrazine and diuron (indicating a loss of the PS II complex) (Vermaas et al., 1987). Muta­ genesis of the corresponding residue (M-H200) in the bacterial photosynthetic reaction center to a leucine did not destabilize the complex but resulted in the replacement of a bacteriochlorophyll in the special pair with a bacteriopheophytin. Several additional mutations were also introduced at the D2 ligand site. The D2-H197Q mutant grew photosynthetically but had oxygen evolution rates which were 40% of wild-type rates. The D2-H197L mutant was unable to evolve oxygen, similar to the D1-H198L mutant. A number of substitutions in the ligands (D1-H198 and D2-H197) D1 and D2 have been made in Synechocystis sp. PCC 6803 (Coleman et al., 1995). The substitution of D1-H198 with Gln, Lys, Cys and Ala had little affect on oxygen evolution rates, whereas the D1-H198 Glu, Ser, Thr and Val mutants had oxygen evolution rates less than 20% of wild type and were unable to grow photoautotrophically. Only the D1-H198L mutant, among those substitutions tested, did not evolve oxygen. Conservative replacements (D1-H198Q and D1-H198N) of the D1 chlorophyll ligands had little effect on the optical properties of P680 (a small shift was observed in the Soret absorbance peak), however, the D1-H198A mutation had a 50-fold faster than wild type and a recombination rate for It was seven fold faster recombination rate for not apparent from pigment analyses whether there was a change in the pigment composition of the as observed in bacterial reaction center mutants. These results suggest that some other molecule, such as water, may substitute as a ligand and coordinate chlorophylls of the special pair in the D1-H198A mutant. One of the first site-directed mutants introduced into the Chlamydomonas chloroplast genome (D1­ H195 Tyr, Asp, and Asn) was targeted to address the issue of the high midpoint potential of (Roffey et al., 1991, 1994a, Kramer et al, 1994). A conserved D1 histidine residue (H195) present in all psbA sequences determined to date and absent from

Chapter 16 Photosystem II the corresponding Rp. viridis L-subunit was targeted for mutagenesis based on the hypothesis that a ligand(D1positively charged residue near the of P680. Each H198) could affect the midpoint of the D1-H195 mutants evolved oxygen and the reduction in hydroxylamine kinetics of extracted thylakoids were similar to wild type for the D1-H195N and D1-H195Y mutants. The reduction kinetics ofthe D1-H198D mutant, however, were 100-fold slower than wild type. The introduction and of a negatively charged residue between of either P680 could presumably shift the or resulting in slower charge transfer kinetics (Roffey et al., 1994a,b).

D. Chlorophyll Monomers and , an Extra Pair of Chlorophylls in Photosystem II There has been some controversy regarding the number of chlorophylls per two pheophytins in the PS II reaction center. Values between four and six chlorophylls per two pheophytins have been reported. A consensus is building, however, which supports the presence of an extra pair of chlorophylls in PS II which are not present in the Rp. viridis photosynthetic reaction center. Eijckelhoff and Dekker (1995) isolated reaction center preparations that contain six chlorophylls per two pheophytins using several of the established methods in the literature. Some of these preparations were previously reported as yielding four chlorophylls per two pheophytins. These workers attribute the variation between the results of the different groups to errors in the extinction coefficients for chlorophyll and pheophytin used for calculating the chlorophyll/pheophytin ratio. It would appear that all of the purification protocols yield similar results when prepared and assayed in one laboratory. This still does not exclude the possibility that there is contamination of the reaction center preparations with other chlorophyll binding proteins or that preparations vary due to the removal of less tightly bound pigments. Molecular modeling of the PS II reaction center, based upon the three dimensional crystal structure from photosynthetic bacteria (Svennson et al., 1990; Ruffle et al., 1992) placed two chlorophyll molecules as the special pair and a pair of chlorophylls as the accessory pigments at a site equivalent to that occupied by the chlorophyll monomers in the bacterial photosynthetic reaction center. The placement of these accessory chlorophyll molecules was deter­

297 mined primarily by the restrictions placed upon the modeling process since the coordinating histidine residues ofthe bacterial accessory chlorophylls were missing in the D1 and D2 polypeptides. In addition to these four chlorophylls it is possible to place two additional chlorophyll molecules perpendicular to the membrane associated with the B helix of the Dl and D2 polypeptides. These chlorophylls have been proposed to be ligated by histidine residues that are conserved in all D1 and D2 sequences (Deisenhofer et al., 1985) (see Fig. 2). There is increasing evidence that the arrangements of the accessory pigments in PS II is different from that of the bacterial reaction center. The presence of an accessory chlorophyll close to the primary donor, as in the bacterial reaction center model, could lead to its oxidation by P680, however. This could only be avoided if the midpoint potential of the accessory pigment was raised by some 600 mV, possibly by hydrogen bonding interactions (Mulkidjanian et al., 1996). Recent evidence indicates, however, that a single hydrogen bond to a bacteriochlorophyll shifts its by only 100 mV (Lin et al., 1994). It is evident from models of the D1 and D2 structure that there are insufficient hydrogen bonding residues in the vicinity of the putative accessory chlorophyll binding sites which are equivalent to that occupied by the accessory bacteriochlorophylls to account for a hydrogen bond of the PS II accessory induced shift in the chlorophylls. Alternative sites (ligated by the D1-H118 and D2­ H117residues of Chlamydomonas) for the accessory chlorophylls of PS II have also been proposed (Michel and Deisenhofer, 1988; Ruffle et al., 1992; Schelvis et al., 1994; Hutchison and Sayre, 1995; Mulkidjanian et al., 1996) (Fig. 2). Placement of the accessory chlorophylls perpendicular to the membrane plane at the periphery of the D1 and D2 proteins would satisfy the predicted arrangement of a chlorophyll or whose spectral features are shifted by (Mulkidjanian et al., 1996). Recently, Koulougliotis et al., (1994) proposed that D1-H118 or D2-H117 residues may coordinate a chlorophyll whose EPR is presumably generated detectable radical following oxidation by P680. Previously, had been tentatively assigned to a position occupied by one of the accessory bacteriochlorophylls close to has been proposed to oxidize the primary donor. and donate electrons to the primary donor (Thompson and Brudvig, 1988) (Fig. 3).Using power saturation and saturation-recovery EPR spectroscopy

298 measurements in manganese depleted samples Koulougliotis and coworkers (1994) determined that the distance between the non-heme iron and was approximately 39Å. The enhancement of the relaxation of the radical by dysprosium ions in preparations where the extrinsic polypeptides had been removed indicates that the pigment anion is equally spaced between the stromal and lumenal membrane surfaces. This geometry places in the lipid bilayer and in a region where coordination by the D1-H118 or D2-H117 residue(s) would be possible. Any model for the organization of P680 must also account for the means by which energy is transferred between the antennae and reaction center chloro­ phylls. As previously indicated, the antennae chlorophylls need to be far enough away from so as to avoid oxidation but close enough to permit energy transfer (Schelvis et al., 1994). Under optimal conditions ofspectral overlap and orientation Schelvis et al., (1994) argue that the center to center distance between the antennae and P680 must be <20 Å to Å to avoid oxidation of the avoid exciton loss but accessory chlorophylls. As previously indicated, the ligands which coordinate the accessory chlorophylls in the Rp. viridis reaction center, at a center to center distance of <20 Å, are not conserved in the D1 and D2 proteins. H. van Gorkom’s group has proposed that the symmetry related histidine residues (D1­ H118, D2-H117) in the B transmembrane spans of the D1 and D2 proteins coordinate the accessory chlorophylls at a center to center distance of (Schelvis et al., approximately 30 Å from the 1994). Significantly, the distance between the chlorophylls presumably coordinated by the B span symmetry related histidine residues (D1-H118 and coordination site D2-H117) and the putative meet the criteria of distance to be sufficiently far removed from P680 to avoid oxidation but close enough to permit excitonic coupling. Mutagenesis of the D2-H118 residue in Synecho­ cystis sp. PCC 6803 resulted in an inability to evolve oxygen (Pakrasi and Vermaas, 1992). Mutagenesis of the symmetry related residue (H118) in the D1 protein of Chlamydomonas to a leucine (non­ conservative) or asparagine (conservative) also resulted in a phenotype which was unable to evolve oxygen (Hutchison and Sayre, 1995). Based on the absence of a variable chlorophyll fluorescence decay component following a flash, the absence of the thermoluminescence band, the failure of the light

Stuart V. Ruffle and Richard T. Sayre dependent reduction of DPIP (2,6-dichloroindophenol) using the PS II donor hydroxylamine, and the failure to accumulate the radical, it was proposed that the D1 -H118 mutants were unable to form a stable charge separated state. However, there was one notable difference between the two mutants, the D1-H118L mutant, when grown in the light, failed to accumulate the D1 protein in thylakoids or in PS II particles. In contrast, light grown D1-H118R mutant cells had near wild-type levels of the D1 protein in thylakoid and PS II particle fractions. These results suggest a differential sensitivity to photoinhibition in the D1-H118 mutants. However, the issues remains open as to the location and number (two or four) of accessory chlorophylls in PS II.

E. Pheophytin Pheophytin is the first detected electron acceptor in PS II. The forward rate of electron transfer between P680 and pheophytin is in the picosecond time range, however, there has been considerable debate about the precise time constant for this reaction. A forward rate of electron transfer of 3 ps has been proposed by several groups (Fig.3). Wasielewski and coworkers (1989) proposed this time constant based upon the appearance of the oxidized state of P680 (increase in absorbance at 820 nm) following excitation at 610 nm. This was supported by the observation that the time constant for the decay of the lowest excited state singlet of P680 was similar. Alternatively, Durrant et al. (1993) have proposed a 21 ps time constant for this reaction. This is based upon the rate of bleaching of the pheophytin ground state (545 nm), the appearance of the pheophytin anion (460 nm), and the loss of stimulated emission from chlorophyll , all proposed to be caused by the formation of of which have time constants of about 21 ps. Biochemical analyses indicate that there are at least two pheophytins per P680 in PS II reaction center preparations (Nanba and Satoh, 1987). Only one of the two pheophytins participates in electron transfer, however. Recent evidence indicates that the two pheophytins can be distinguished on the basis of their optical absorbance properties, one absorbs near 676 nm and the other near 680 nm (Breton, 1990; Tang et al., 1990; van Kan et al., 1990; Otte et al., 1992; van der Voos et al., 1992; van Gorkom and Shelvis, 1993). The pheophytin active in electron transfer appears to be the 676 nm form (van Kan et al., 1990).

Chapter 16 Photosystem II The location of the pheophytin involved with the forward electron transfer in PS II has been modeled to be homologous to that in the bacterial reaction center (Svensson et al., 1990 and Ruffle et al., 1992). The quinone-type reaction center structure places the pheophytin between the primary donor and the primary acceptor, at an angle of 60° relative to the membrane plane. It has been proposed that the pheophytins are hydrogen bonded to the D1 and D2 polypeptides and occupy a non-polar pocket associated with the transmembrane helices B and C. The active branch pigment forms a hydrogen bond from the keto-carbonyl group of ring V and the carboxylic acid side chain of glutamate 104 in Rp. viridis. This residue is totally conserved in higher plant and algal D1 sequences at position 130. Resonance Raman spectroscopy has also inferred the interaction of a hydrogen bond between the 9­ keto carbonyl group of the pheophytin and a glutamate carboxyl side chain in PS II (Möenne-Loccoz, 1990) It is interesting to note that the residue at position D1-130 is a glutamine in Synechocystis sp. PCC 6803. Site-directed mutagenesis ofthis residue in the cyanobacterium has revealed a difference between the cyanobacterial and higher plant PS II. Giorgi and coworkers (1996) compared the absorbance of the Qx transition of the pheophytin, the quantum efficiency of charge separation, and the rate of forward electron transfer in wild type and site-directed substitutions (D1-Q130E and D1-Q130L) in Synechocystis sp. PCC 6803. Conversion of the D1­ Q130 residue to a glutamate (identical to the residue present at residue 130 in the eukaryotic D1 protein) increased the quantum yield for charge separation to near 1, as opposed to 0.6 in wild-type cyanobacteria. The D1-Q130E mutation also caused a shift in the pheophytin Qx band to 544 nm, as opposed to 541.5 nm. None ofthe mutations affected the time constant pheophytinelectron transfer. The leucine for substitutions resembled the cyanobacterial wild type with respect to the Qx absorption maximum and quantum efficiency. These data support the location of the pheophytin around D1-130, that is, in a homologous position to that seen in the bacterial reaction center.

F. and Redox Active Components of the D1 and D2 Proteins Kinetic studies on the reduction of indicate that it is reduced by an intermediate electron donor

299 referred to as Z which in turn is reduced by the tetra-Mn water oxidizing complex (Babcock et al., 1989; reviewed in Debus, 1992). Following removal of the water-splitting Mn complex the transiently formed radical (Z) is easily detected by EPR in illuminated samples but rapidly decays in the dark at room temperature. Its spectrum is known as signal (Babcock, et al., 1989). A spectrally similar radical can be generated in non-oxygen evolving PS II particles but gives rise to a dark stable radical (Babcock et al., 1989). This denoted EPR signal alternate donor, D, does not participate in steady state electron transfer from the water oxidizing and has a markedly lower redox complex to potential than does Z. In 1987, Barry and Babcock provided strong evidence that the species giving rise to the D radical was a tyrosine based on the demonstration that deuteration of tyrosines in cyanobacteria had a substantial effect on the line shape of signal On the basis of PS II structural models, two symmetrically related and conserved tyrosine residues, located on the lumenal end of the third transmembrane segment of the D1 and D2 proteins, were proposed to be respectively. Due to the ease with which and cyanobacteria could be genetically transformed, sitedirected mutagenesis of these residues was carried out in Synechocystis sp. PCC 6803 to test these predictions. Substitution of residue D2-Y160 with and phenylalanine resulted in the loss of signal led to the identification of Y160 as (Debus et al., 1988a; Vermaas et al., 1988; Metz et al., 1989). Subsequent mutagenesis of the D1 protein confirmed (Debus 1988b, the identification ofD1-Y161 as Metz et al., 1989). Recently, Minagawa et al. (1996) have mutated D1-Y161 in C. reinhardtii to a phenyl­ alanine. Flash dependent chlorophyll fluorescence rise and decay kinetics indicated that the rapid donor to was absent in the mutant. In a series of flashes, however, a high fluorescent state could be generated resulting from the oxidation of a slow donor. A slow, oxygen dependent bleaching of carotenoid (438 nm) and a chlorophyll (435 nm) species was observed in this mutant. The bleaching of the carotenoid could be blocked by DCMU, hydroquinone, anaerobiosis, superoxide dismutase was required for or catalase, indicating that the bleaching as well as oxygen radicals. and Dipolar coupling experiments between the non-heme Fe, and the tetra-Mn complex have established distance relationships

300 between each of these cofactors. Hirsch and Brudvig (1993) estimated by EPR pulsed relaxation and power saturation experiments that the distance between and the non-heme Fe was 37 ± 5 Å. Using D2­ mutants Koulougliotis et al. (1995) Y160F obtained similar distance relationships for the low and the non-heme Fe. A temperature trapped has been broadening of the EPR signal by observed consistent with a distance between and P680 of 10–15 Å. Using the models of PS II based on the coordinates of the bacterial photo­ synthetic reaction center it is apparent that is closer to P680 than in part accounting for the preferential oxidation of In addition, the midpoint potentials of the two redox active tyrosines differ. Vass and Styring (1991) and Boussac and Etienne (1987) have estimated the midpoint potential to be 0.72–0.76V whereas the midpoint of is 1.03 V potential of Low temperature EPR, transient proton ENDOR (electron nuclear double resonance) spectroscopy and molecular modeling studies of the environments near (hydrophilic environment) and (hydrophobic environment) indicate that they reside in different environments (Ruffle et al., 1992; Tommos et al., 1995; Tang et al., 1996). apparently has an exchangeable proton hydrogen bonded to a nearby residue. The apparent hydrogen bonding partner is D2-H189. Replacement of D2-H189 with Asp, Gln, Leu (Tang et al., 1993) and Leu and Tyr (Tommos et EPR al., 1993) results in a narrowing of the signal. In contrast, mutagenesis of the symmetry related histidine residue in the D1 protein of C. reinhardtii and Synechocystis sp. PCC 6803 does not affect the lineshape of the radical (Roffey et al, 1994a, Tang et al., 1996). The rate of oxidation of in the D1-H190F mutant (Chlamydomonas) was 100-fold slower than in hydroxylamine extracted wild-type thylakoids. In addition, the D1-H190F mutant did not contain a tetra-Mn complex, lacked thermoluminescence band attributed to the the radical charge recombination between and the (in the presence of DCMU), but was capable of oxidizing hydroxylamine at 45% of the wild-type rate. The origin of the thermoluminescence band remains controversial. It has been argued that the oxidized donor involved in band charge recom­ bination is either or D1-H190. Debus (1992) has suggested that conditions which shift the equilibrium between and may reduce the quantum yield of produced at the low temperatures

Stuart V. Ruffle and Richard T. Sayre (–20 °C). Thus the D1-H190F mutant may be unable to generate at the low temperatures used for thermoluminescence measurements. A loss ofthe thermoluminescence band has also been observed, however, in diethylpyrocarbonate treated (modifies histidine residues) spinach membranes suggesting that the band may arise from an oxidized histidine residue (Ono and Inoue, 1990). Further evidence in comes from support of a donor other then analyses of mutations (D1-H195D in Chlamy­ domonas) which decreases the equilibrium constant and (by 50-fold). Mutagenesis of between the D1-H195 residue to an aspartate did not shift the peak temperature of the thermoluminescence band suggestingthat does not give rise to this feature (Kramer et al., 1994). It has been argued, however, recombination that the activation energy for may be substantially greater than the precluding a shift in the peak temperature of the band in the D1-H195D mutant (Diner and Babcock, 1996).

G. Water Oxidation and the Tetra-Manganese Complex The catalyst for water oxidation is the charge accumulating S-state or tetra-Mn complex bound to the D1 and possibly D2 proteins and stabilized by chloride, calcium and the extrinsic 33 kD OEE1 protein (reviewed in Debus, 1992; Britt, 1996). The driving force for water oxidation is the oxidation of P680 which oxidizes the tetra-Mn complex via The oxidation of two molecules of water requires four quantum events which drive the S-state complex through five redox transitions beginning with the dark stable state (Kok et al., 1970). The kinetics of the S-state transitions are slower with each S-state transition (Fig. 3). Oxygen is released after the final redox transition (the third flash). A proton is released at each S-state transition, however, proton release to the lumen may be buffered by amino acid side chains and influenced by the (Haumann and Junge, electrostatic effects of 1992). The structure of the tetra-Mn complex and its protein interactions remain one ofthe more enigmatic aspects of the water oxidizing complex. The participation of a mixed valence state Mn cluster in water oxidation was first demonstrated by Dismukes and Siderer (1980). They detected a flash or S-state dependent g=2.0 multiline EPR spectrum at cryogenic

Chapter 16 Photosystem II temperatures arising from the state. A broad (1,000 gauss) signal at g=4.1 is also formed at cryogenic state (Zimmerman and temperatures from the Rutherford, 1986; Kim et al., 1992). Structural information on the organization of the tetra-Mn complex has been provided by EXAFS (extended Xray absorption and fine structure spectroscopy) studies. Yachandra et al. (1993) have proposed that bridged the Mn complex is organized as two Mn dimers linked by a bridge. XANES (X-ray absorbance near-edge spectroscopy) has been used to assign the oxidation states ofthe Mn atoms in the complex. In the dark stable state there appears to be 2 Mn (III) and 2 Mn (IV) ions. Further but may not occur oxidation of Mn occurs at at Instead an organic radical may be generated transition (Andrews et al., 1995). on the Boussac et al (1990) have favored the formation of a transition, however, histidine radical for the this assignment has been questioned (Hallahan et al., 1992; Gilchrist et al., 1995). In addition to Mn, the inorganic cofactors, chloride and calcium, are required for water oxidation. PS II and high particles contain both low affinity calcium binding sites. Depletion of calcium from PS II particles blocks oxygen evolution and results in the loss ofthe normal g=2 multiline EPR signal and its replacement by a unique dark stable multiline spectrum. Calcium has also been shown to be required for the photoactivation or light dependent assembly of the tetra-Mn complex (Ananyev and Dismukes, 1996). EXAFS and FTIR experiments have provided evidence that calcium is in close proximity to manganese although these results remain controversial (Latimer et al., 1995; Noguchi et al., 1995). Interestingly, calcium depletion also dramatically slows electron transfer suggesting a structural role for the cation (Adelrath et al., 1995). Chloride depletion also inhibits oxygen evolution. Unlike calcium, however, chloride may be replaced by other anions though with less effectiveness (Kelley and Izawa, 1979). A modified state can be formed in chloride depleted PS II complexes but it does not advance to without re-addition of chloride (Sandusky and Yocum, 1986; Homann, 1988). Whether chloride depleted membranes are blocked or the transition remains at the controversial. The tetra-Mn complex requires at least 24 ligands if it is octahedrally coordinated. The identification of

301 ligands contributions by protein side chains has been an area of intense interest. Mutagenesis and isotopic labeling experiments coupled with pulsed EPR experiments have led to the identification ofpossible Mn ligands. The mutagenesis experiments have been performed in both Synechocystis sp. PCC 6803 and C. reinhardtii. Any interpretation of these results must be constrained, however, by the possible secondary affects of mutations on protein structure. At least twelve residues on the lumenal side of the D1 and D2 proteins have been implicated as possible Mn ligands. On the D1 protein these residues include; D59, D61, E65, D170, E189, H190, H331, E332, H336, D342 and the C-terminal residue, A344 (Chlamydomonas numbering) (Diner et al., 1991; Nixon et al., 1992; Debus, 1992). On the D2 protein, only E69 has been implicated as a Mn ligand (Vermass et al., 1990). There is also direct biophysical evidence for the coordination of Mn by both imidazole and carboxylate residues (Tang et al., 1994, Noguchi et al., 1995). While most of the potential lumenal side D1 and D2 Mn ligands have been identified through site-directed mutagenesis studies in Synechocystis sp. PCC 6803 several potential Mn ligand mutations have also been generated in Chlamydomonas. As mentioned previously, a D1-H190F mutant was unable to evolve oxygen or bind Mn of the water oxidizing complex. The mutant was, however, capable of oxidizing the artificial donor hydroxylamine (45% of wild-type rates) and exogenous manganese chloride. As in the wild-type strain, the H190F mutant had both high (H190, H190F, and low affinity (H190, Mn oxidation sites (Roffey et al., 1994a,b; Kullander et al., 1995). A D1-H190Y mutant, however, only had a low affinity Mn oxidation site. A Chlamydomonas (D1-Y161F) mutant had the lowest affinity for Mn and was also unable to evolve oxygen. A D1-P186L mutant had only a low affinity Mn oxidation site but was still capable of oxygen evolution at reduced rates. It was evident from the analyses of these D1 protein mutants that there was no relationship between the presence and absence of the high and low affinity Mn oxidation sites and the ability to evolve oxygen. It is now apparent that assembly of the functional water oxidizing complex involves multiple peptide residues. These residues may be involved in Mn oxidation, Mn ligation, calcium ligation and proton exchange. Mutations which perturb these processes or alter structural

302 relationships between residues may preclude assembly or destabilize the tetra-Mn water oxidizing complex. Recently, it has been suggested that the D1-H190 residue may participate indirectly in water oxidation following its oxidation as a proton acceptor from by D1-H190 would act as a weak base to shuttle protons from the water oxidizing complex to the thylakoid lumen (Britt, 1996). According to this radical would act as a strong model, the neutral base and extract a proton from water. This model requires that the tetra-Mn complex be in close Recent studies from D. Britt’s proximity to group suggest that this is the case (Gilchrist et al., 1995). Thus, the D1-H190 residue may have multiple functions in donor side processes of PS II. Another potential manganese ligand, D1-D170, has also been targeted for mutagenesis in Chlamydomonas (Whitelegge et al., 1995). Similar to Synechocystis sp. PCC 6803 mutants, the Chlamydomonas D1-D170H and D170N mutants had wildtype levels of PS II centers and 45% and 5–15% of the oxygen evolution rate of wild type, respectively (Chu et al., 1994). The D1-D170T Chlamydomonas mutant phenotype was also similar to the Synecho­ cystis sp. PCC 6803 mutant and was unable to evolve oxygen but had near wild-type levels of PS II. These results suggest that identical mutants in both organisms behaved similarly. Finally, mutations which introduce a premature stop codon at the C-terminal proteolytic processing site of the Chlamydomonas D1 protein or mutations which introduce either a Gly, Cys or Val at the S345 processing site do not affect oxygen evolution or the stability of the PS II complex (Schrader and Johanningmeier, 1992; Lers et al., 1992; Takahashi et al., 1996). These results suggest that the C-terminal processing is not required for assembly of the PS II complex and that the protease which cleaves the D1 protein (between residues A344 and S345) does not require a serine residue at position 345.

H. and the Stable Electron Acceptor of Photosystem II and the Herbicide Binding Site A molecule of plastoquinone-9 occupies the site on the D2 protein and stabilizes the charge separated The at pH7) of the state couple is –150 mV, approximately 430 mV less reducing than P680* (Fig. 3). Typically is a single

Stuart V. Ruffle and Richard T. Sayre electron acceptor, however, under high light conditions can be doubly reduced initiating photoinhibition or the turnover of the D1 protein (Keren et al., 1995). Significantly, is not protonated even at a stromal pH of 4.0 (Pulles et al., 1976). Initial electron transfer from occurs in but is slower from presumably due to electrostatic effects (Fig. 3). The structure of the site has been modeled on the basis of the Rp. viridis photosynthetic reaction site is most closely center crystal structure. The associated with the D2 protein (Trebst, 1986b). Several residues are spatially conserved between the predicted PS II and Rp. viridis crystal structure. These residues include: D2-T217, F252, W253 and H214 which correspond to the M subunit residues: T220, F249, W250 and H217. EPR measurements of the coupled signal (g=1.82) indicate that is in close proximity (7 Å) to the non-heme iron (Nugent et al., 1981). Addition of formate, which displaces a bound bicarbonate (see below), results in signal and slower an enhanced (100-fold) electron transfer.

The structure of the , plastoquinone-9, binding site has also been modeled after the structure of the Rp. viridis photosynthetic reaction center crystal structure. Residues which are conserved between the PS II and Rp. viridis structures include D1-H215 and S264. These residues correspond to L-H190 and Lstructure. Residues in the S223 in the Rp. viridis Rp. viridis site structure which may be involved in protonation of the semiquinone and quinol are not apparently conserved in the structure (Trebst, 1986). site is also a target for herbicide binding, The e.g. atrazine, diuron (DCMU), bromacil, and metribuzin. Many site-specific Chlamydomonas herbicide resistant mutants have been characterized from spontaneous and chemically induced mutants as well as genetically engineered cells (reviewed in Erickson and Rochaix, 1992). These mutations are clustered in the -binding region of the D1 protein which includes residues 211–275. (Diner et al., 1991). The phenotypes ofthe mutants were not all equivalent, however. The D1-S264A mutation was shown to increase tolerance to the herbicides metribuzin (5,000 × wild type) and atrazine (100 × wild type), and slowed both and electron transfer (Erickson et al., 1989; Govindjee et al., 1992; Crofts et al., 1993). The D1-G256D mutant conferred

Chapter 16 Photosystem II resistance to atrazine (15 × wild type) and slowed by but not reduction by reduction of These results suggested that the D1-S264A and D1G256D mutants had a differential effect on protonation (Crofts et al., 1993). A D1-A251V mutant, which conferred resistance to atrazine (1,000 electron transfer × wild type), had slower kinetics but electron transfer kinetics were unchanged relative to wild type. (Johanningmeier et al., 1987, Govindjee et al., 1992). In contrast, the D1V219I, F255Y, and L275F herbicide resistance or mutants had little effect on electron transfer (Erickson et al., 1989, Govindjee et al., 1991, 1992, Strasser et al., 1992). One of the first sets of site-directed mutants generated in Chlamydomonas by chloroplast DNA transformation were multiple psbA mutants (D1N266T/I259S and D1-N266T/S264A/I259S) which conferred herbicide resistance (Przibilla et al., 1991). Transformants were selected on metribuzin plates after introduction of a psbA fragment containing the engineered mutations. The triple mutant had reduced metribuzin resistance (1,580 × wild type) compared to the D1-S264A mutant (5,000 × wild type) but increased phemediphan resistance (1,000 × wild type) compared to the D1-S264A mutant (200 × wild type). The double mutant behaved similarly to the triple mutant. Recently, twelve different amino acid substitutions were introduced into the D1-A251 residue by chloroplast transformation (Lardans et al., 1997). As discussed above, the D1-A251V mutant confers atrazine resistance. The five charged or polar amino acid substitutions (Arg, Asp, Gln, Glu and His) at position D1-A251 resulted in non-photosynthetic phenotypes, whereas the non-polar substitutions were capable of photosynthetic growth. With the exception ofthe D1-A251R mutant, all the non-photosynthetic mutants accumulated either a 33–34 kD form of the D1 protein or an immunologically related form 24– 25 kD N-terminal form ofthe protein. In each mutant which synthesized the 24–25 kD form of the D1 protein it rapidly turned over. In contrast, the D1A251R mutant synthesized an apparently mature form of the D1 protein, however, it had a short half life in the membrane. These results were in contrast to similar mutations introduced into this domain (between transmembrane spans IV and V) in cyanobacteria where all mutants introduced result in photosynthetic phenotypes. Thus, there are

303 intrinsic differences between the structure and or stability of the cyanobacterial and chloroplastic PS II reaction center types. and sites is the nonLocated between the heme Fe. The non-heme Fe of PS II is most likely coordinated by residues D1-H215, D1-H272, D2H214 and D2-H268 which are conserved from the bacterial reaction center crystal structure (Figs. 2, 4, 5) (Deisenhofer and Michel, 1988). Bicarbonate and possibly other small anions are thought to provide a fifth ligand to the non-heme Fe(II) which in the Rp. viridis crystal structure is provided by residue ME232 (see review by Blubaugh and Govindjee, 1988). Bicarbonate is required for normal electron transfer Its displacement by formate rates between and reduces the kinetics of reduction by 100-fold. In addition, NO can form an adduct with the non-heme Fe(II) giving rise to a unique EPR signal (g=4.0) (Petrouleas and Diner, 1990). The possible functions of the non-heme Fe(II) and associated small anions remains controversial. Bicarbonate has been proposed to be involved in protonation of since the kinetics of electron transfer are less affected by the absence of bicarbonate than the kinetics of protonation (Xu et al., 1991). Additional roles for the and the non-heme Fe(II) include: stabilization of PS II structure (Dutton et al., 1978). The binding site for bicarbonate has been investigated extensively by site-directed mutagenesis. Obvious candidates for bicarbonate binding include positively charged residues near the non-heme Fe(II). Several arginine residues are located near this site (Diner et al., 1991). To determine whether a possible positively charged residue (D1-R269) located near the non-heme Fe(II) bound bicarbonate, a nonconservative (D1-R269G) mutant was introduced into C. reinhardtii by chloroplast DNA transformation (Hutchison et al., 1996b). Interestingly, the D1R269G mutant was unable to grow photosynthetically, did not evolve oxygen with DCBQ as an electron acceptor, and lacked manganese. The D1-R269G mutant had a decreased level of the D1 protein when grown in light, but normal D1 protein levels when grown in dark, implicating a greater sensitivity to photoinhibition. In dark grown cells the amplitudes and of the EPR detectable signals were only 30% that of wild type indicating that there were fewer functional PS II complexes in the mutant. Chlorophyll a fluorescence decay kinetics also indicated a blockage in the electron transfer

304 from to similar to that induced by the loss of bicarbonate. The presence of a formate enhanced signal indicated, however, that the D1-R269G mutant was able to bind bicarbonate. Furthermore, the magnitude of the formate affect on the EPR signal in the D1-R269G mutant was consistent with the predicted levels of functional PS II complexes.

l. Cytochrome Associated with the D1 and D2 proteins in the PS II reaction center particle preparations is Cyt The cytochrome is composed of two polypeptides ( and ) that are proposed to coordinate a heme via histidine axial ligands. The and subunits are coded for by the chloroplast genes psbE and psbF. It is unclear, at this time, whether the structure of the cytochrome is heterodimeric or a combination of homodimers. There is evidence to suggest that the amino terminal ends of the and polypeptides are located on the stromal side of the thylakoid membrane (Tae et al., 1988; Tae and Cramer, 1994); these observations would place the heme associated with the polypeptides on the stromal side of the membrane. If the conclusions drawn by these authors are correct it is important to note that the positive inside rule (von Heijne, 1986) for the insertion of transmembrane spans across membranes is not followed. Most of the sequences described for the subunits of follow this rule, however, the Chlamydomonas subunit, or the heterodimer taken as a whole, have an excess of positive charges in the C-terminal portions of their sequences suggesting the opposite orientation of insertion. Presumably the topology of the PS II reaction center in Chlamydomonas is the same as other higher plants and cyanobacteria, and some other mechanism, possibly the increased number of negatively charged residues around the positively charged ones, may explain this enigma. The psbE gene encodes the of Cyt which ranges in size between 82 to 86 amino acids. The Chlamydomonas chloroplast psbE gene was described independently by Alizadeh et al. (1994) and Mor et al. (1995). Both gene sequences reveal a protein of 82 amino acids, one residue less than the higher plant sequences, which would yield a mature protein with a molecular mass of approximately 9.2 kD. The protein sequence is 71% identical to that of spinach and 65% identical to that of Synechocystis sp. PCC 6803. The predicted transmembrane

Stuart V. Ruffle and Richard T. Sayre spanning segment is highly conserved with only two amino acid substitutions to similar residues (Herrmann et al., 1984; Tae et al., 1988). All of the sequences available to date have a histidine residue (H22 in C. reinhardtii) in the fifth position of the putative transmembrane spanning region. A conserved histidine residue located in the transmembrane span has been predicted to be one of two histidines that coordinate the heme cofactor associated with Cyt (Babcock et al., 1985). The second heme ligand is provided either from a second PS II-E polypeptide or from a conserved histidine residue located in transmembrane spanning domain of the psbF gene product. A more detailed discussion of the relationship between the and subunits of Cyt is presented by Whitmarsh and Pakrasi (1996). Disruption of the psbE gene has been reported for the cyanobacterium Synechocystis sp. PCC 6803 (Shukla et al., 1992). The introduction of stop codon in the coding sequence leads to a non-photoautotrophic phenotype with a lack of detectable D2 protein in the photosynthetic membrane. Similarly, site-directed mutagenesis of the axial heme ligand (histidine) to a leucine, in the psbE sequence results in a loss of PS II activity in the prokaryote (Pakrasi et al., 1991). The subunit of Cyt is encoded by the psbF gene. In higher plants and cyanobacteria it is cotranscribed with psbE but, as discussed below, the genes for the two cytochrome subunits are separated in the Chlamydomonas genome. First described by Fong and Surzycki (1992, Genbank-CRPSBFLG), the psbF gene encodes a 44 amino acid polypeptide which was reported to be about 60% identical to the protein sequences from a variety of higher plants, liverwort, Euglena and Synechocystis sp. PCC 6803. The polypeptide has a calculated molecular mass of 5.2 kD. The higher plant genes encode a protein of only 38 amino acids. Notably, the Chlamydomonas psbF sequence presented by Fong and Surzycki did not have any histidine residues. The residue at an equivalent position to the conserved histidine in all other sequences was a threonine. This region of the genome has now been analyzed by several other laboratories and a different sequence had been proposed. Alizadeh et al. (1994) and Mor et al. (1995) independently sequenced the psbF gene and found it to encode a 44 amino acid polypeptide that had a single histidine residue in the predicted transmembrane spanning domain. The Chlamydomonas psbF gene sequences have between 64 and

Chapter 16 Photosystem II 79% identity with the corresponding sequences from pea, liverwort, Euglena and Synechocystis sp. PCC 6803. The psbF gene product subunit) has been deleted in Synechocystis sp. PCC 6803 (Pakrasi et al., 1990) and the resulting phenotype is non-photosynthetic, with barely detectable levels of the reaction center core proteins (D1 and D2) suggesting the psbF gene product is essential for the stable assembly of PS II. The removal of the subunit histidine ligand by sitedirected mutagenesis also led to a loss of PS II activity with no detectable D2 protein despite the presence of D1 (Pakrasi et al., 1991). In higher plants and in Synechocystis, psbE is cotranscribed with psbF, psbL and psbJ (Herrmann et al., 1984; Pakrasi et al., 1988; reviewed in Erickson and Rochaix, 1992) as the sequences are in tandem with a small intergenic spaces. However, in Chlamydomonas the arrangement of the genes encoding the subunits of the cytochrome are quite different. The coding regions for psbE and psbF are approximately 7.3 kb apart and on opposite strands of the chloroplast genome (Alizadeh et al., 1994). Mor et al. (1995) demonstrated, by northern analysis, that psbE is transcribed alone on a mRNA of approximately 0.3 kb, whereas psbF is found on a 0.9 kb molecule. A similar rearrangement of the operon is also seen in the related species C. moewusii (Boudreau et al., 1994) and C. eugamatos (Turmel and Otis, 1994). The different transcripts for the psbE and psbF in green algae poses an interesting subunits difference in the regulation of the Cyt when compared to land plants. The stoichiometry of to subunits is 1:1 in reaction center particles (reviewed in Whitmarsh and Pakrasi, 1996). With co-transcription it is assumed that this 1:1 ratio of to subunits can be maintained simply, however, in Chlamydomonas with different transcripts for each polypeptide, some other control mechanism is presumed to exist to ensure that there are sufficient and subunits to maintain this ratio.

p er PS II The stoichiometry of cytochrome reaction center is also in some doubt. There are as many reports of one cytochrome per PS II as there are for two per PS II. The majority of the confusion surrounding this issue seems to come from the different investigators using different PS II preparations, with different detergent treatments, to determine the stoichiometry. This is further compounded by researchers using different extinction coefficients for the hemes and different deter-

305 minations of the number of PS II centers in their preparations (reviewed in Whitmarsh and Pakrasi, 1996). Buser and coworkers (1992a) determined that there per PS II. Using differis only one cytochrome ence spectroscopy to assay the chlorophyll and cytochrome, and EPR spectroscopy to quantify the and cytochrome in detergent treated preparations and thylakoid membranes, they per PS II in determined that there was 1.2 Cyt detergent preparations, and one Cyt per PS II in thylakoid membranes. This apparent enrichment upon detergent treatment is significant as it argues against other observations that propose that the 1:1 ratio is due to the loss of cytochromes caused by over solubilization of PS II particles (van Leeuwen et al., 1991). As well as a possible role as a nucleating site for have been PS II formation, other functions ofCyt proposed. The hemes can exist in two distinct redox potential forms; high potential (approximately 350– 400 mV) and low potential (20–80 mV) (Cramer and Whitmarsh, 1977). The rates of oxidation and reduction ofthe cytochrome under normal conditions are slow and this probably precludes it from being actively involved in forward transfer of electrons through PS II or the splitting of water (Whitmarsh and Cramer, 1977). Many workers have proposed that the cytochrome is involved in a protective mechanism including a cyclic electron flow around PS II. This flow would dissipate excess energy and decrease the likelihood ofthe formation ofdamaging singlet and triplet species capable of destructive oxidation reactions. PS II is known to be damaged by high light stress conditions. This photoinhibition occurs in several stages, some of which are reversible. The first step is the photoinactivation of the electron transfer process which is followed by the degradation of the D1 protein. The D2 and other polypeptides of the PS II complex may then also be damaged. This damage may be caused by radicals formed on either the donor or acceptor side of PS II (reviewed in Aro et al., 1993; Chapter 30, Keren and Ohad). The oxidation state of Cyt may be a signal for the degradation of the D1 protein during photoinhibition. Nedbal and coworkers (1992) have shown that an unidentified redox component can decrease photoinhibitory damage approximately ten-fold after its oxidation. The most likely candidate for this component is the low potential form of Cyt These authors propose that it is only after has

306 been doubly reduced and pheophytin has accepted a further electron, that the low potential form of the cytochrome is reduced and that is the signal for D1 degradation. Barber and De Las Rivas (1993) have can be fully reduced by demonstrated that Cyt pheophytin under continuous illumination. These observations also obviate the need for a fast turnover of the oxidation states of the cytochrome, indeed it is likely that a slow turnover of the redox states is necessary as the accumulation of the stable, reduced form of pheophytin only occurs with low quantum efficiency (Klimov et al., 1977). has been proposed to act as Alternatively, Cyt an electron donor to protect from damage caused by highly oxidizing components ofthe PS II donor side. The high potential form of the cytochrome can be oxidized when the forward rate of electron transfer between the water oxidizing complex to the primary donor has been impaired (Buser et al., 1990). This oxidation has also been demonstrated at cryogenic temperatures, which also inhibits forward electron transport from water. Under these conditions the yield of oxidized cytochrome increases, and the size EPR multiline signal associated with the of the water oxidizing complex decreases (Thompson and Brudvig, 1988). Further illumination of the PS II reaction center after the cytochrome is oxidized leads to the photooxidation of a chlorophyll molecule, (Visser et al., 1977). Cyt has been shown to be oxidized by the primary donor via a photooxidized (Thompson et al., 1988) which itself could be a damaging radical. These observations led Buser and operates coworkers (1992b) to propose that in cyclic electron flow around PS II to protect it is against photodamage. The involvement of also significant as its oxidation state may affect excitation of P680 (see Section II.C). Other proposed roles for the cytochrome include: the photoactivation of the water oxidation complex (following the oxidation of a manganese ion) (Cramer et al., 1986); involvement as a redox active proton pump across the thylakoid membrane (Arnon and Tang, 1988); and possible interactions with superoxide dismutase (Ananyev et al., 1994).

J. The PS II-I Protein The PS II-I protein is one of several small polypeptides present in PS II reaction center particles (Nanba and Satoh, 1987). Based on its gene sequence (psbI) the PS II-I protein is predicted to have 37 amino acids

Stuart V. Ruffle and Richard T. Sayre (4.8 kD) and a single transmembrane spanning helical domain. The predicted Chlamydomonas protein sequence shares considerable identity with psbI gene products from cyanobacteria (65%) and higher plants (76%) (reviewed by Erickson and Rochaix, 1992), but is one amino acid longer than the higher plant sequences. An additional leucine residue is present in the Chlamydomonas sequence near the carboxyl-terminus. From the analysis of the sequence and the composition of various PS II preparations there is no evidence that PS II-I binds any cofactor or pigment. The deletion of the psbI gene has been reported in C. reinhardtii (Künstner et al., 1995). This mutant has its psbI gene sequence replaced with a modified spectinomycin/streptomycin antibiotic resistance cassette (Goldschmidt-Clermont, 1991). PS II-I minus mutants were able to evolve oxygen, however, the amount of D1 protein was reduced to 10–20% of wild-type levels. The psbI deficient lines were also capable of photoautotrophic growth in dim light but cell growth was significantly reduced and the cells died at a light by growth at These data suggest that the intensity of PS II-I polypeptide stabilizes the PS II complex and may protect the complex from photoinhibition. Similar deletion mutants have been reported in the cyanobacterium Synechocystis sp. PCC 6803 (Ikeuchi et al., 1995). The cyanobacterial PS II-I deficient cells also grew photoautotrophically but at rates only slightly slower than the control cells. The mutants had 75% ofthe wild-type level ofsteady state oxygen evolution activity and had wild-type levels of other PS II polypeptides (D2, CP47, PS II-H and PS II-K) indicating that the psbI deletion has less of an effect on destabilization of the prokaryotic PS II complex than the eukaryotic PS II complex.

K. CP43 and CP47, the Antenna ChlorophyllProtein Complex of Photosystem II In addition to the reaction center polypeptides, oxygen evolving PS II core particles contain two structurally related 43 and 47 kD chlorophyll a binding proteins known as CP47 (Fig. 6) and CP43 (Fig. 7). These proteins transfer energy from the light harvesting chlorophyll proteins to P680 but are also required for oxygen evolution (Bricker and Ghanotakis, 1996). Each polypeptide is highly conserved (80–85% identity) between different organisms and binds either 12–14 or 20–25 chlorophylls (Tang and Satoh, 1984;

Chapter 16

Photosystem II

de Vitry et al., 1984). As shown by immunological and protein labeling studies (D-biotin-N-hydroxysuccinimide) CP43 and CP47 each have six transmembrane spanning domains with a large lumenal extrinsic domain (134 residues in CP43 and 180 residues in CP47) located between the fifth (E) and sixth (F) transmembrane spanning domains (Fig. 6) (Vermaas et al., 1987, Bricker, 1990, Sayre and Wrobel-Boerner, 1994). A notable feature of the predicted CP43 and CP47 secondary structures is

307

the location of conserved and possibly symmetry related histidine residues at the margins of each transmembrane spanning domain. These residues have been proposed to bind chlorophylls (Vermaas et al. 1987; Bricker, 1990; Sayre and Wrobel-Boerner, 1994). Both CP43 and CP47 are present in oxygen evolving PS II core particles but are lacking in reaction center particles (Figs. 1B and 1C). Deletion or inactivation mutants of the psbC gene encoding

308 CP43 are unable to evolve oxygen in either cyanobacteria or Chlamydomonas. CP43 deficient mutants (cyanobacteria and Chlamydomonas) contain PS II complexes capable of charge separation, and retain CP47 protein, but do not associate with the nucleus encoded light harvesting chlorophyll a/b binding protein (de Vitry et al., 1984; Vermaas et al., 1988; Rögner et al., 1991). In contrast to the CP43 mutants, cyanobacterial and Chlamydomonas mutants which fail to accumulate CP47 do not accumulate a reaction center complex and are incapable of charge separation (Monod et al., 1992). The results of these studies as well as studies on the 77 K chlorophyll fluorescence emission spectra of ) and CP47 ( CP43 (fluorescence ) suggest that CP47 is most closely associated with the D1 and D2 polypeptides and that energy transfer is from CP43 to CP47 (Bricker and Ghanotakis, 1996). Numerous lines of evidence indicate that CP43 and CP47 are also required for oxygen evolution. Monoclonal antibodies directed against the large lumenal extrinsic domain of CP47 fail to bind (in situ) when the 33 kD (OEE1) protein is bound to the core complex or when some of the manganese of the water oxidizing complex is present (in the absence of the 33 kD protein) (Frankel and Bricker, 1989). Furthermore, CP47 can be cross-linked to the 33 kD protein (Enami et al., 1987; Bricker et al., 1988; Odom and Bricker, 1992). Site-directed mutagenesis of residues located on the large and small lumenallyexposed domains of CP47, in Synechocystis sp. PCC6803, also lead to losses in oxygen evolution (Putnam-Evans and Bricker, 1994; Eaton-Rye and Vermaas, 1991; Haag et al., 1993; Gleiter et al., 1994). These results suggest that the large extrinsic domain may be involved in binding the 33 kD protein of the oxygen evolving complex. III. The Chloroplast DNA Encoded Small Polypeptides of Photosystem II

A. The PS II-H Protein The psbH gene (Johnson and Schmidt, 1993; H. O’Connor, J. Nugent and S. Purton, 1993, GenbankCHCRPSBHA) is located on the chloroplast genome and encodes an 88 amino acid polypeptide (PS II-H) with a predicted molecular mass of 9.3 kD. In higher plants the psbH gene encodes a smaller, 73 amino

Stuart V. Ruffle and Richard T. Sayre acid protein. The Chlamydomonas PS II-H protein has eight additional residues at the amino-terminus and six extra amino acids at the C-terminus which are not present in the vascular plant sequences. The PS II-H protein contains a single transmembrane spanning helix based on hydropathy plot analyses, which is presumably phosphorylated at the second threonine (Dedner et al., 1988). In order to determine the function of the PS II-H phospho-protein in Chlamydomonas the gene was disrupted using the spectinomycin resistance cassette (Ruffle et al., 1995). The resulting mutant algae were unable to grow photoautotrophically concomitant with a loss of a phosphorylated 9 kD thylakoid protein. Cells lacking the PS II-H protein do not EPR signal associated evolve oxygen and have no with a functional PS II complex (Ruffle et al., 1995). Recently, it has been demonstrated that PS II-H deletion mutants fail to accumulate the PS II core complex proteins (D1, D2, CP47 and CP43) in dark grown cells, suggesting that lack of PS II activity in these mutants is unrelated to photoinhibition (Summer et al., 1997). These observations imply that PS II-H may have a structural role in the assembly or the stability of the PS II complex. The role of the phosphorylated threonine residue at the second position (PS II-H-T2) in the polypeptide has also been investigated by site-directed mutagenesis in Chlamydomonas (Cheater et al., 1995). SDS-PAGE analysis of thylakoid membranes treated with radiolabeled phosphate showed that the substitution of the threonine with alanine (PS II-HT2A) prevented phosphorylation of the protein. This implies that a second threonine at position 4 cannot be phosphorylated or that the phosphorylation of PS II-H is sequential and dependent on phosphorylation of PS II-H-T2. Significantly, analysis of the total phosphorylation profile of the thylakoid membranes in the PS II-H-T2A mutant indicated that there was almost no phosphorylation of any of the thylakoid proteins. Other measures of PS II activity showed that the site-directed mutant was indistinguishable from wild type with respect to oxygen evolution, variable chlorophyll fluorescence and EPR. Overall it is apparent that PS II-H has a structural role in PS II and that phosphorylation of other thylakoid membrane proteins is not required for maintenance of steady state photosynthesis. In contrast to Chlamydomonas, psbH deletion mutants in the cyanobacterium Synechocystis sp.

Chapter 16 Photosystem II PCC 6803 (Mayes et al., 1993) were able to grow photoautotrophically. The mutants were, however, to electron transport, but it is impaired in apparent that PS II-H protein was not required for functional PS II complexes in cyanobacteria.

B. The PS II-J Protein The psbJ gene has been identified in higher plants and cyanobacteria (Ohyama et al., 1986; Shinozaki et al., 1986; Hiratsuka et al., 1989 and Kaneko et al., 1996). The gene sequence has yet to be reported in Chlamydomonas but is believed to be on the chloroplast DNA fragment EcoRI-17 or PstI-3, close to the atpI gene (Chlamydomonas database at the National Agriculture Library, United States Department of Agriculture). A 5 kD protein assigned to be PS II-J has been detected in the photosynthetic membranes of cyanobacteria (Lind et al., 1993). Early termination in the gene does not prevent photoautotrophic growth in Synechocystis sp. PCC 6803. Herbicide binding studies show that, in the mutant, the PS II per chlorophyll ratio is only half of that of wild-type cells and that PS II mediated electron transport is only 46% of that of wild type. PS I electron transfer was unaffected by the mutation. The protein has yet to be detected in any thylakoid membrane preparations of the oxygenic eukaryotes.

C. The PS ll-K Protein In the Chlamydomonas chloroplast genome the gene (psbK) encoding the PS II-K protein lies adjacent to the elongation factor gene tufA. The sequence for the psbK gene was first described by Silk et al. (1990) and it encodes a 46 amino acid polypeptide with over 80% identity to the higher plant PS II-K proteins after post-translational processing. The gene appears on a mono-cistronic mRNA (Takahashi et al., 1994) unlike the situation in higher plants where psbK is co-transcribed with psbI (Neuhaus and Link, 1990). The predicted PS II-K protein sequence has a single hydrophobic transmembrane spanning domain. In vascular plants the PS II-K protein is posttranslationally processed such that the initial residue in the mature form, purified from PS II complexes, is a lysine (Murata et al., 1988). In the Chlamydomonas sequence the leader peptide up to the first lysine residue is only half of the length of the higher plant type. The N-terminus of the mature protein from

309 Chlamydomonas PS II particles was sequenced and was found to begin with a lysine residue (de Vitry et al., 1991). Disruption of the coding sequence of psbK with the spectinomycin resistance cassette in Chlamydomonas caused a loss of PS II activity with respect to oxygen evolution and chlorophyll fluorescence measurements (Takahashi et al., 1994). Mutated cells did not grow photoautotrophically. Immunological studies showed that only 10% of the wild-type levels ofthe D1 protein were accumulated, however, pulsechase radiolabeling ofcells, lacking PS II-K, showed that the rates of D1 and D2 protein synthesis were normal. These results suggest that PS II-K was necessary for the stable assembly or the stability of the PS II complex. The disruption mutation in Chlamydomonas is quite different from that reported for the prokaryote Synechocystis sp. PCC 6803 (Ikeuchi et al., 1991). The mutant cyanobacterium retains the ability to grow photoautotrophically although at a reduced rate compared to wild type, despite the fact that they appear to have normal levels of PS II as measured on a per chlorophyll basis.

D. The PS II-L Protein The coding sequence for the PS II-L polypeptide (Fong and Surzycki, 1992) is located adjacent to, and downstream of, the psbF gene. The gene encodes a 44-amino acid polypeptide which is predicted to have a single transmembrane span. Compared to the sequences from vascular plants and cyanobacteria, there are an additional six amino acids located near the N-terminus in Chlamydomonas. Correspondingly, the predicted molecular mass for the PS II-L protein is approximately 0.7 kD larger (5.2 kD for Chlamydomonas and 4.5 kD for the plant species and Synechocystis sp. PCC 6803) (Ohyama et al., 1986; Shinozaki et al., 1986; Hiratsuka et al., 1989; Anbudurai and Pakrasi, 1991). A sequence alignment of PS II-L polypeptides indicates a high degree of homology in all photosynthetic genera (74–80% identity). The protein was not found in non-oxygen evolving PS II core preparations of Chlamydomonas but has been identified as a 4.8 to 5 kD band in higher plant and cyanobacterial core complexes (de Vitry et al., 1991; Ikeuchi et al., 1989a, b; Webber et al., 1989; Nagatsuka et al., 1991). There have been no reports of deletion or insertional inactivation mutants of

310 psbL in C. reinhardtii to date. The psbL gene, initially named psbI by Cantrell and Bryant (1988), has been deleted in the cyanobacterium Synechocystis sp. PCC 6803 (Anbudurai and Pakrasi, 1993). Loss of the PS II-L polypeptide correlated with a loss of oxygen evolving capacity and the ability to grow under photoautotrophic conditions. The cells had reduced (<75% of wild type) levels of the D1 and D2 proteins and could not bind radioactive diuron. This indicates that the psbL gene product is essential for the stable assembly of the PS II reaction center complex. Reconstitution experiments with depleted spinach reaction center particles indicate that the addition of PS II-L with another unidentified 4.1 kD protein reduction (Nagatsuka et al., 1991). restored Kitamura and coworkers (1994) demonstrated that with highly purified PS II-L, plastoquinone-9 could be reconstituted into isolated reaction center core molecule in the presence complexes as a functional of thylakoid lipids. The lipids and plastoquinone These data themselves could not reconstitute show that the unidentified 4.1 kD protein used by T. Nagatsuka and coworkers was probably not involved in the reconstitution.

E. The PS II-M Protein The sequence for the psbM gene has recently been reported from Chlamydomonas (Higgs et al., 1996). This sequence encodes a 31 amino acid polypeptide that shares 74% identity, at the protein level, to the published sequence of the tobacco PS II-M protein (Shinozaki et al., 1986). The N-terminal sequencing of a 4.7 kD polypeptide from PS II particles (de Vitry et al., 1991) showed some similarity to the predicted PS II-M polypeptide sequences from vascular plants and cyanobacteria, with four of the eight sequenced residues showing identity. The partial sequence shows little homology to the predicted polypeptide sequence derived from the putative Chlamydomonas psbM gene (Higgs et al., 1996), so it is unlikely that this 4.7 kD polypeptide is a product of the psbM gene. There have been no reports of deletion mutants for the PS II-M protein. The gene and protein have also been identified in cyanobacteria (Ikeuchi et al., 1989c).

F. The PS II-N Protein The psbN gene is located in the psbB gene cluster along with psbH and psbT (previously named ycf8) (Johnson and Schmidt, 1993). The organization of

Stuart V. Ruffle and Richard T. Sayre the gene cluster is similar to that of vascular plants, however, the intergenic distances are greater. The coding sequence is predicted to yield a polypeptide of 43 amino acids with a putative single transmembrane spanning helix. This protein was not detected by protein sequencing of small polypeptides in Chlamydomonas PS II particles presumably because the N-terminus is blocked (de Vitry et al., 1991). The protein has been detected in the cyanobacterium Synechocystis sp. PCC 6803 with an apparent molecular mass of4.7 kD (Ikeuchi et al., 1989c). The psbN gene has been disrupted by cloning the spectinomycin resistance cassette into the coding sequence (S. V. Ruffle and J. H. A. Nugent, unpublished). Initial characterization of this mutant indicates that it is not photoautotophic but does show some PS II activity when grown heterotrophically.

G. The PS II-T Protein The gene (psbT) encoding the PS II-T protein has previously been called ORF31 and ycf8 and has been sequenced in Chlamydomonas (Monod et al., 1992; Johnson and Schmidt, 1993). This gene is located in the psbB/H/N gene cluster, is co-transcribed with psbB, and is expressed constitutively (Monod et al., 1994). It encodes for a 31 amino acid polypeptide with a predicted molecular mass of approximately 3.3 kD. A deletion mutant has been reported where the chloroplast gene had been replaced with an antibiotic resistance cassette (Monod et al., 1994). This mutant could grow photoautotrophically but was sensitive to certain high stress conditions. Under high light or high concentrations of intensity spectinomycin (which reduced overall chloroplast protein synthesis) cell growth rates were depressed. It appears that this protein is dispensable under normal growth conditions bu t is required for maintaining optimal growth under adverse growth conditions. A similar polypeptide is predicted from the psbT sequence found in the Synechocystis sp. PCC 6803 genome. It encodes a 31 amino acid protein which shares 40% identity with the algal sequence; there have been no reports of a deletion mutant. Another PsbT gene from the nuclear genome of cotton has been reported (Kapazoglou et al., 1995). These researchers report that a mature protein of approximately 28 residues is targeted to the thylakoid lumen following its processing from a 7.5 kD

Chapter 16

Photosystem II

intermediate. This species is processed, after chloroplast import, from an 11 kD precursor. This PS II-T polypeptide has a high degree of homology to a protein designated PS II-T from spinach PS II. This PsbT gene product has not been detected in Chlamydomonas. It is clear that the PS II-T polypeptides from cotton and spinach have no homology with the chloroplast encoded or Synecho­ cystis sp. PCC 6803 psbT genes and there needs to be a renaming of one or other of the genes and their gene products.

IV. The Nucleus Encoded Polypeptides of the Photosystem II Complex

A. OEE1, the 33 kD Oxygen Evolving Complex Protein The product of the nuclear PsbO gene (Mayfield et al., 1989) is targeted to the thylakoid lumen where it has been shown to be essential for water oxidation in oxygenic eukaryotes (Mayfield et al., 1987a). This protein is also known as OEE1 (oxygen evolution enhancer 1) and as shown in reconstitution experiments, binds to the lumenal or donor side of PS II (Andersson et al., 1984). It appears that the protein is more crucial for oxygen evolution in Chlamydomonas than in Synechocystis sp. PCC 6803. In the cyanobacterium, the deletion of the PsbO gene, which encodes a 33 kD mature protein, reduces the rate of oxygen evolution to about 70% of the wild-type level (Burnap and Sherman, 1991). The deletion mutants are also more sensitive to photoinhibitory treatments than wild type. The mutants could grow autotrophically at reduced rates, however, when the cells are starved of calcium ions photosynthetic growth is lost (Philbrick et al., 1991). In the absence of calcium the manganese water oxidizing complex is less stable in the mutant. In contrast to Synechocystis sp. PCC 6803, the Chlamydomonas PsbO transposon insertion strain, FUD44, cannot grow photoautotrophically and does not evolve oxygen (Mayfield et al., 1987a). The mutant specifically lacks the transcript and gene product for PsbO but still has the other two extrinsic polypeptides associated with the donor side of PS II, the 23 and 17 kD proteins. However, it is not apparent whether they are bound to the membrane. The PsbO gene encodes a 291 amino acid precursor protein that has both a chloroplast and a lumenal targeting peptide at the amino terminal end of the sequence. The

311 mature protein has 239 amino acids and shows 67% identity to the spinach sequence (Mayfield et al., 1987b). The loss of the OEE1 protein does not affect the expression of any of the other nucleus encoded subunits of PS II but does cause a decrease in the level of the PS II core proteins presumably due to an increased rate of protein turnover (Mayfield et al., 1987a). Site-directed mutations within the PsbO gene of Synechocystis sp. PCC 6803 have been used to investigate the functional and structural roles of specific residues (Burnap et al., 1994). The mutation of a highly conserved aspartate residue (D9K) near the amino-terminus of the protein had no significant effect on the accumulation of the protein or the rates of oxygen evolution. It had been previously demonstrated that protease treatment of the amino terminus inhibited the binding of OEE1 to PS II (Eaton-Rye and Murata, 1989). The mutation of a cysteine residue to a serine (C20S) had a profound effect on OEE1 stability. This mutant did not accumulate any detectable levels of OEE1. The transcript of the PsbO gene was present at levels comparable to wild type so the lesion appears to be at the protein structural level presumably associated with the destruction of a disulfide bridge. The mutants have impaired rates of oxygen evolution and exhibit characteristics comparable to the PsbO deletion strain. Mutation of the conserved aspartate residue at position 159 to asparagine appears to have no adverse effect on the structure of the protein with normal levels of OEE1 accumulated. However, the rate of water oxidation activity is reduced, although not as significantly as in the deletion strain (Burnap et al., 1994). These observations reveal that it is both the overall structure of OEE1, as well as key amino acids, that presumably bind calcium and/or chloride ions, that are important for the role of the protein as an enhancer of water oxidation. The OEE1 protein has also been shown to be closely associated with CP47, the gene product of the psbB gene. Bricker (1990) proposed that the OEE1 interacts with a long hydrophilic loop between predicted transmembrane helices V and VI. Other workers have isolated crosslinked fragments of PS II that included the OEE1. The subunits found to be associated with 33 kD polypeptide included; D1, D2 (Mei et al., 1989), CP47 (Enami et al., 1987), Cyt and PS II-I (Enami et al., 1992). Immunoprecipitation reactions have also determined that the 33 kD has neighboring interactions with a 24 (23) kD protein (Ljungberg et

312 al., 1984). A structural model ofthe OEE1 has been generated (Xu et al., 1994) based upon the method ofconstrained Chou-Fasman sequence analysis. This model proposes that the structure contains a large proportion of sheet (39%) and only a small amount of helix (8.5%). It is also in good agreement with circular dichroism measurements of the solution structure of purified spinach OEE1 (48% sheet; 9% helix) (Xu et al., 1994). Both the solution structure and the model structure give little insight as to likely interactions between the OEE1 and the PS II reaction center or ion binding sites. The organization of the extrinsic polypeptides on the donor side of PS II has been investigated in Synechocystis sp. PCC 6803 with respect to the stoichiometry of the OEE1. There has been controversy in the literature over the number of OEE1 subunits per PS II reaction center. Using immuno-quantification of western blots of both oxygen evolving PS II membranes and enriched reaction center complexes, Xu and Bricker (1992) and Leuschner and Bricker (1996) have determined that there are two cooperatively binding OEE1s per four manganese. These workers report that previous reports of a 1:1 ratio (Murata et al., 1984; Andersson et al., 1984) of OEE1 to PS II is due to an underestimation of OEE1 content. This underestimation is believed to be due the masking effect of the D2 protein, which co-migrates with OEE1, during SDS-PAGE (Xu and Bricker, 1992). Western blot analysis of calcium chloride washed PS II preparations shows that the D2 protein does indeed mask OEE1 and decreases its apparent abundance when assayed using immuno-quantification. The OEE1 appears to stabilize the inorganic cofactors of the water oxidizing complex. It has been shown to be non-essential for oxygen evolution in cyanobacterial deletion strains. Biochemical depletion of the OEE1 from isolated PS II preparations by calcium chloride or sodium chloride/urea washing has shown that up to 99% of OEE1 can be removed and yet 20% ofthe untreated rate ofoxygen evolution remains in the presence of exogenous calcium and chloride ions (Ono and Inoue, 1984; Bricker, 1992). Other reported effects of OEE1 depletion include the loss of two of the four manganese atoms of the water oxidizing complex, however, this loss could be prevented by exogenous calcium ions (Miyao and Murata, 1984). It has to be noted that treatments which remove the OEE1 are also known to deplete PS II of the 23 and 17 kD polypeptides and some or

Stuart V. Ruffle and Richard T. Sayre all ofthe effects seen with washed PS II preparations could be due to these pleiotropic effects. Reconstitution experiments with cyanobacterial and higher plant OEE1s and core complexes have demonstrated that they are interchangeable although they do not restore as much oxygen evolving activity as the native subunit. Koike and Inoue (1985) used the OEE1 from the thermophile Synechococcus vulcanus to reconstitute 28% (compared to reconstitution with the S. vulcanus protein) of oxygen evolution activity to OEE1 lacking spinach PS II particles. In the reverse experiment the plant OEE1 could reconstitute 60% activity to the cyanobacterial PS II. This latter reconstitution was not possible at the higher growth temperatures of the cyanobacterium. These observations suggest that some of the interactions between the OEE1 and PS II reaction center are shared between higher plants and cyanobacteria. It is possible that the OEE1 coordinates some of the manganese of the water oxidizing complex. However, biophysical measurements of the water oxidizing complex indicate that the structure is essentially unchanged upon the removal ofthe 33 kD protein. These observations include a multiline signal state of the water that is associated with the oxidizing complex (Miller et al., 1987) as well as an X-ray absorption K-edge spectrum from the state (Cole et al., 1987). The OEE1 may bind calcium ions which have been shown to be essential for maximizing the rates of oxygen evolution (Tamura and Chenaie, 1985). Wales et al., (1989) analyzed the OEE1 sequence and proposed a calcium binding site based upon a similarity with a calcium binding E-F hand motif. Biochemical evidence for the ability of the OEE1 to bind calcium is uncertain. Webber and Gray (1989) proposed that a 32 kD protein had affinity for calcium, on nitrocellulose after SDS-PAGE, but did not positively identify this polypeptide as the PsbO gene product.

B. OEE2, the 23 kD Oxygen Evolving Complex Protein The PsbP gene encodes the 23 kD OEE2 (oxygen evolution enhancer 2) protein in Chlamydomonas and other photosynthetic eukaryotes (Mayfield et al., 1987b). The PsbP gene is not present in cyanobacteria. The OEE2 pre-polypeptide contains 245 amino acids and has a bipartite thylakoid lumen targeting sequence at its N-terminal portion. The mature polypeptide is

Chapter 16

Photosystem II

found in the thylakoid lumen and is 188 amino acids in length, it shares about 60% identity with vascular plant sequences. Mayfield et al. (1987b) reported that Chlamydomonas mutants (BF25) lacking the 23 kD protein had lower rates of oxygen evolution (5% of wild type) and a decreased abundance of PS II compared to wild type. They could, however, grow photoautotrophically. Miyao and Murata (1989) demonstrated by reconstitution experiments in isolated spinach PS II particles that the 23 kD was required for the binding of the 17 kD subunit. It was only when the 33, 23 and 17 kD subunits were present that high rates of oxygen evolution could be observed (discussed in Ghanotakis and Yocum, 1990; Debus, 1992). Efficient rates of oxygen evolution were obtained in the mutant lacking the 23 kD protein but only in the presence of exogenous chloride ions (Rova et al., 1994). The lack ofthe transcript for PsbP had no effect on the level of PS II accumulated when compared to wild type. The PS II centers in the mutant, however, had a decreased affinity for chloride ions. It had been previously demonstrated that the removal ofthe extrinsic 23 kD protein decreased the chloride affinity of PS II by several orders of magnitude (Andersson et al., 1984; Miyao and Murata, 1985) Recently, Rova and coworkers (1996) have studied the process ofphotoactivation in a Chlamydomonas mutant strain that lacks the PsbP gene product (FUD39). Photoactivation or the light dependent assembly of the tetra-Mn complex is a two-quantum event process with a dark stable step in between (Tamura and Chenaie, 1985). Both calcium and chloride ions are required for the correct assembly of the tetra-Mn water oxidizing complex. The deletion mutants were shown to be more sensitive to photodamage during photoactivation. Wild-type cells typically photoactivate at light intensities from 0.2 to with near 100% efficiency. The optimal light intensity for photoactivation of the mutant, measured by electron transfer activity after a light and the yield of the treatment, was only process was only 60% when compared to wild-type cells under the same conditions (Rova et al., 1996). The light intensity used for photoactivation was critical in this mutant, 0.8 and only yielded 50 and 40%, respectively, and 60 yielded less than 10% ofthe electron transfer activity of wild type. Continued illumination ofthe mutant at the higher light intensities led to a photoinhibitory loss of electron transfer activity. From these

313 observations, the authors describe a model for photoactivation, suggesting that the 23 kD protein is required on the donor side of PS II to sequester chloride ions during PS II assembly. The loss of the 23 kD protein presumably leads to the inability to assemble an oxygen evolving tetra-Mn complex resulting in increased sensitivity to donor side photoinhibition and PS II turnover.

C. OEE3, the 17 kD Oxygen Evolving Complex Protein The PsbQ gene encodes a 199-amino acid precursor for the 17 kD lumenal protein OEE3 (oxygen evolution enhancer 3) or PS II-Q (Mayfield et al., 1989). The pre-polypeptide has a bipartite transit sequence at its N-terminus that targets the mature form of the protein to the thylakoid lumen. The mature protein of 149 amino acids shares only about 28% identity with vascular plant sequences. This low level of homology probably reflects the fact that it is the least closely associated of the extrinsic polypeptides to the PS II core, and consequently being less constrained, it has diverged to a greater degree during phylogeny. There have been no reports of OEE3 mutants in Chlamydomonas. The role of the PsbQ gene product is unknown. Removal of the protein from isolated PS II particles has been achieved using calcium chloride and sodium chloride washes. Reconstitution experiments have shown that efficient rates of oxygen evolution can only be achieved when the 33, 23 and 17 kD subunits are all bound to PS II(Åkerlund et al., 1982; discussed in Ghanotakis and Yocum, 1990) unless exogenous calcium and chloride ions are present. As described above (Section IV. B.) the 17 kD polypeptide can only associate with the PS II core after the 23 kD protein is bound (Miyao and Murata, 1989). Presumably the 23 kD polypeptide has a cooperative role in binding the PsbQ gene product to PS II or the 17 kD extrinsic polypeptide is only bound to the PsbP gene product.

D. Additional Nucleus-Encoded Small Photosystem II Subunits There have been several reports of genes encoding PS II subunits of cyanobacteria and photosynthetic eukaryotes that have yet to be reported for C. reinhardtii. These include PsbS, a nuclear gene encoding the 22 kD polypeptide of PS II (Kim et al., 1992b; Wedel et al., 1992). This protein has been

314 identified in vascular plants and cyanobacteria and has some sequence similarities to chlorophyll binding proteins LHCI/II (light harvesting complex), CP27 and 29 (Wedel et al., 1992). The gene encodes a 274 residue precursor which is processed during import into the organelle to a 200 amino acid mature form. Based on hydropathy plot analyses, the PS II-S protein is predicted to have four membrane spanning helices. The protein appears to have considerable internal homology in two domains suggesting that it may have been the result of an internal gene duplication of the original PsbS gene. In spinach, the PS II-S protein appears to bind four or five chlorophyll molecules as well as carotenoids but is stable in the absence of the pigments. There appear to be two PS II-S proteins per PS II reaction center. In etiolated seedlings the amount of PS II-S protein increases with time and also increases following exposure to light (Funk et al., 1995). The PS II-S protein appears to be regulated at several levels during plant development. The transcript of PsbS increases during an eight day etiolation period in spinach cotyledons. During the first 48 hours after germination the steady state PsbS transcript level appears to be determined by increased transcriptional activity. After an extended period in the dark, control of its expression is exerted at the post-transcriptional level. Beside this lightindependent regulation of PS II-S production, the expression of the gene is positively up-regulated by phytochrome after etiolated seedlings are exposed to light. As may be expected, red light down-regulates the number of PsbS transcripts, a classic characteristic of a phytochrome mediated response (Adamska et al., 1996). The chlorophylls bound to PS II-S show very weak excitonic coupling indicating that it is unlikely to be involved in energy or electron transfer (Adamska et al., 1996). The PsbS transcript is present only in photosynthetic tissues. These observations, along with its developmental regulation, suggest that it has a role as a pigment chaperone, acting as a buffer between pigment biogenesis and assembly of active photosynthetic units, thus decreasing the likelihood of the toxic effects of non-bound chlorophyll molecules. The nuclear gene PsbW encodes a 137 amino acid precursor protein that contains an 83 amino acid bipartite transit peptide. Interestingly, the mature form of the protein is directed to the thylakoid membrane in Arabidopsis thalliana instead of the

Stuart V. Ruffle and Richard T. Sayre lumen as might be expected (Lorkovic et al., 1995). The 6.1 kD protein is predicted to have a single transmembrane span and is found in reaction center core particles. The transcript ofthe PsbW gene appears to be light regulated. It is believed to be the only nucleus encoded component of the reaction center core (Fig. 1C), however, the role of this subunit is unknown. The nucleotide sequence for PsbW has also been described in Odontella sinensis (a diatom) (Kowallik et al., 1995), the red alga Porphyra purpurea (Reith and Munholland, 1995), and spinach. The protein sequence derived from the gene shares a high degree of identity to the partial protein sequence reported by de Vitry and coworkers (1991) and designated as the 6.1 kD nucleus encoded subunit. There are sequences published of a gene and product designated PsbX. In tomato, this nuclear gene encodes a 257 residue precursor polypeptide which is processed to a mature form of 185 amino acids with an apparent molecular mass of 23 kD (Betts and Pichersky, 1992). The protein sequence has a high degree of homology to the OEE2 protein and should probably be considered the PsbP gene in tomato. Another gene designated psbX has been reported from the chloroplast genomes of O. sinensis, P. purpurea, Cyanophora paradoxa and the genome of Synechocystis sp. PCC 6803 (Kowallik et al., 1995; Reith and Munholland, 1995; Stirewalt et al., 1995). The gene products have considerable identity to each other and all are predicted to yield a 4.1 kD polypeptide which is almost entirely hydrophobic except the final 8 amino acids. The role of this protein is unknown. These sequences display similarity to the partial protein sequences of a 4.1 kD PS II-X polypeptide from wheat and spinach. Recently, a cDNA has been identified from Arabidopsis thaliana (Kim et al., 1996) which encodes a nuclear gene, that after processing, yields a 42residue polypeptide. It is predicted to have a single transmembrane span, with a high degree of sequence homology to the PS II-X protein predicted from the psbX gene from Odontella as well as the partial protein sequences from higher plants. This gene and its protein are of particular interest as the gene appears to have ‘moved’ recently in the course of evolutionary history, as it is a plastid gene in the unicellular eukaryote and a nuclear gene (with presequences for lumenal targeting) in a vascular plant. The protein seems to insert into the thylakoid membrane independently ofthe delta pH or Sec-type

Chapter 16 Photosystem II translocases, as inhibitors to each of these import systems do not block its insertion. It is therefore proposed that this third type of presequence has arisen recently in evolutionary terms when compared with the Sec-dependent signals (Kim et al., 1996).

V. Perspectives It is evident that the tools required to engineer and characterize PS II mutants in Chlamydomonas are maturing. Routine chloroplast transformation is now possible (Boynton and Gillham, 1993; Hutchison et al., 1996b; Chapter 8, Goldschmidt-Clermont) and nuclear gene knock-out transformation systems are available. In addition, PS II particles of various complexities and functional properties are available. It is also evident that the phenotypes of PS II deletion and site-directed mutations in Synechocystis sp. PCC 6803 and Chlamydomonas, representing the prokaryotic and eukaryotic PS II types, are not always equivalent. This may reflect differences in the subunit architecture and the apparent greater plasticity in copy number (e.g. psbA) and expression of cyanobacterial PS II genes. Various works in progress suggest that these differences may be greater than currently appreciated. Finally, in contrast to higher plant chloroplastic PS II complexes, the Chlamydomonas PS II can be greened or assembled in the dark. This feature is particularly useful for the characterization of mutants which display secondary effects when exposed to light (photoinhibition). Overall, it is apparent that Chlamydomonas is the paradigm for analysis of chloroplastic PS II complexes.

Acknowledgments We acknowledge Dr. Jonathan Nugent for assistance with the structural model of the PS II complex. We thank Drs David Kramer, Ron Hutchison,Tony Crofts, and Robin Roffey for many useful discussions. The writing of this chapter was supported in part by the U.S. Dept. of Energy (RTS).

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320 Nedbal L, Samson G and Whitmarsh J (1992) Redox state of a one-electron component controls the rate of photoinhibition of Photosystem II. Proc Natl Acad Sci USA 89: 7929–7933 Neuhaus H and Link G (1990) The chloroplast psbk operon from mustard (Sinapis alba L.): Multiple transcripts during seedling development and evidence for divergent overlapping transcription. Curr Genet 18: 377–383 Nixon PJ and Diner BA (1992) Aspartate 170 of Photosystem II reaction center polypeptide D1 is involved in the assembly of the oxygen evolving manganese cluster. Biochemistry 31: 942–948 Nixon PJ, Trost JT and Diner BA (1992) Role of the carboxy terminus of polypeptide D1 in the assembly of a functional water-oxidizing manganese cluster in Photosystem II of the cyanobacterium Synechocystis sp. PCC 6803: Assembly requires a free carboxyl group at C-terminal position 344. Biochemistry 31: 10859–10871 Noguchi T, Inoue Y and Satoh K (1993) FTIR studies on the triplet state of P680 in the Photosystem II reaction center: Triplet equilibrium within a chlorophyll dimer. Biochemistry 32: 7186–7195 Noguchi T, Ono TA and Inoue Y (1995) Direct detection of a carboxylate bridge between Mn and in the photosynthetic oxygen-evolving center by means of Fourier transform infrared spectroscopy. Biochim Biophys Acta 1228: 189–200. Nugent JHA, Diner BA and Evans MCW (1981) Direct detection of the electron acceptor of Photosystem II. Evidence that Q is an iron-quinone complex. FEBS Lett 124: 241–244 Odom WR and Bricker TM (1992) Interaction of CPa-1 with the manganese-stabilizing protein of Photosystem II: Identification of domains cross-linked by l-ethyl-3-[3-(dimethylamino) propyl]carbodiimide. Biochemistry 31: 5616–5620 Ohyama K, Fukuzawa H, Kohchi T, Shirai H, Sano T, Sano S, Umesono K, Shiki Y, Takeuchi M, Chang Z, Aota S, Inokuchi H and Ozeki H (1986) Chloroplast gene organization deduced from complete sequence of liverwort Marchantia polymorpha chloroplast DNA. Nature 322: 572–574 Ono T and Inoue Y (1991) Biochemical evidence for histidine oxidation in Photosystem II depleted of the Mn-cluster for evolution. FEBS Lett 278: 183–186 Otte SCM, van der Vos R and van Gorkom HJ (1992) Steady state spectroscopy at 6K of the isolated Photosystem II reaction centre: Analysis of the red absorption band. J Photochem Photobiol B: Biol 13: 5–14 Pakrasi HB, Williams JGK and Arntzen CJ (1988) Targeted mutagenesis of the psbE and psbF genes blocks photosynthetic electron transport: Evidence for a functional role ofcytochrome b-559 in Photosystem II. EMBO J 7: 325–332 Pakrasi HB, Nyhus KJ and Granok H (1990) Targeted deletion mutagenesis of the of cytochrome 6-559 protein destabilizes the reaction center of Photosystem II. Z Naturforsch 45c: 423–429 Pakrasi HB, de Ciechi P and Whitmarsh J (1991) Site-directed mutagenesis of the heme axial ligands of Cytochrome b-559 affects the stability of the Photosystem II complex. EMBO J 10: 1619–1627 Petrouleas V and Diner BA (1990) Formation by NO of nitrosyl adducts of redox components of the Photosystem II reaction center. 1. NO binds to the acceptor-side non-heme iron. Biochim Biophys Acta 1015: 131–140 Pfister K, Steinback KE, Gradner G and Arntzen CF (1981)

Stuart V. Ruffle and Richard T. Sayre Photoaffinitylabeling of a herbicide receptor protein in chloroplast membranes. Proc Natl Acad Sci USA 78: 981–985 Philbrick JB, Diner BA and Zilinskas BA (1991) Construction and characterization of cyanobacterial mutants lacking the manganese-stabilizing polypeptide of Photosystem II. J Biol Chem, 266: 13370–13376 Przibilla E, Heiss S, Johanningmeier U and Trebst A (1991) Sitespecific mutagenesis of the D1 subunit of Photosystem II in wild-type Chlamydomonas. Plant Cell 3: 169–174 Pulles MPJ, van Gorkom HJ and Verschoor GAM (1976) Primary reactions of Photosystem II at low pH. 2. Light induced changes of absorbance and electron spin resonance in spinach chloroplasts. Biochim Biophys Acta 440: 98–106 Putnam-Evans C and Bricker TM (1994) Site-directed mutagenesis of the CP47 protein of Photosystem II: Alteration of the basic residue 448R to 448G prevents the assembly of functional Photosystem II centers under chloride-limiting conditions. Biochemistry 33: 10770–10776 Reith ME and Munholland J (1995) Complete nucleotide sequence of the Porphyra purpurea chloroplast genome. Plant Mol Biol Rep 13: 333–335 Rigby SE, Nugent JHA and O’Malley PJ (1994) ENDOR and special triple resonance studies of chlorophyll cation radicals in photosystem 2. Biochemistry 33: 10043–10050 Rochaix J-D, Dron M, Rahire M and Malone PM (1984) Sequence homology between the 32 kDa and D2 chloroplast membrane polypeptides of Chlamydomonas reinhardtii. Plant Mol Biol 3: 363–370 Roffey RA, Golbeck JH, Hille CR and Sayre RT (1991) Photosynthetic electron transport in genetically altered Photosystem II reaction centers of chloroplasts. Proc Natl Acad Sci USA 88: 9122–9126 Roffey RA, Kramer DM, Govindjee and Sayre RT (1994a) Lumenal side histidine mutations in the D1 protein of Photosystem II affect donor side electron transfer in Chlamydomonas reinhardtii. Biochim Biophys Acta 1185: 257–270 Roffey R, van Wijk K, Sayre R and Styring S (1994b) Spectroscopic characterization of Tyrosine-Z in Histidine 190 mutants of the D1 protein in Photosystem II (PS II) in Chlamydomonas reinhardtii. J Biol Chem 269: 5115–5121 Rögner M, Chisholm DA and Diner BA (1991) Site-directed mutagenesis of the psbC gene of Photosystem II: Isolation and functional characterization of CP43-less Photosystem II core complexes. Biochemistry 30: 5387–5395 Rova EM, Franzen L-G, Fredriksson P-O and Styring S (1994) Photosystem II in a mutant of Chlamydomonas reinhardtii lacking the 23 kDa PsbP protein shows increased sensitivity to photoinhibition in the absence of chloride. Photosynth Res 39: 75–83 Rova EM, McEwen B, Fredriksson P-O and Styring S (1996) Photoactivation and photoinhibition are competing in a mutant of Chlamydomonas reinhardtii lacking the 23-kDa extrinsic subunit of Photosystem I I . J Biol Chem 271: 28918–28924 Ruffle SV, Donnelly D, Blundell TL and Nugent JHA (1992) A three-dimensional model of the Photosystem II reaction centre of Pisum sativum. Photosynth Res 34: 287–300 Ruffle SV, O’Connor H, Cheater AJ, Purton S and Nugent JHA (1995) The construction and analysis of a disruption mutant of psbH in Chlamydomonas reinhardtii. In: Mathis P (ed) Photosynthesis: From Light to Biosphere, Vol III, pp 663–666.

Chapter 16 Photosystem II Kluwer Academic Publishers, Dordrecht Sandusky PO and Yocum CF (1986) The chloride requirement for photosynthetic oxygen evolution: Factors affecting the nucleophilic displacement of chloride from the oxygen evolving complex. Biochim Biophys Acta 849: 85–93 Satoh K (1996) Introduction of the Photosystem II reaction center—isolation and biochemical and biophysical characterization. In: Ort DR and Yocum CF (eds) Oxygenic Photosynthesis: The Light Reactions, pp 193–211. Kluwer Academic Publishers, Dordrecht Sayre RT and Wrobel-Boerner EA (1994) Molecular topology of the Photosystem II chlorophyll a binding protein, CP43: Topology of a thylakoid membrane protein. Photosynth Res 40: 11–19 Sayre RT, Andersson B and Bogorad L (1986) The topology of a membrane protein: The orientation of the 32 kd Qb-binding chloroplast thylakoid membrane protein. Cell 47: 601–608 Schelvis J, van Noort P, Aartsma T and van Gorkom H (1994) Energy transfer, charge separation and pigment arrangement in the reaction center of Photosystem II. Biochim Biophys Acta 1184: 242–250 Schrader S and Johanningmeier U (1992) The carboxy-terminal extension of the D1-precursor protein is dispensable for a functional Photosystem II complex in Chlamydomonas reinhardtii. Plant Mol Biol 19: 251–256 Shen JR, Vermaas W and Inoue Y (1995) The role of cytochrome c-550 as studied through reverse genetics and mutant characterization in Synechocystis sp. PCC 6803. J Biol Chem 270: 6901–6907 Shim H, Cao J, Govindjee and Debrunner P (1990) Purification of highly active oxygen-evolving Photosystem II from Chlamydomonas-reinhardtii. Photosynth Res 26: 223–228 Shinozaki K, Ohme M, Tanaka M, Wakasugi T, Hayashida N, Matsubayashi T, Zaita N, Chunwongse J, Obokata J, Yamaguchi-Shinozaki K, Ohto C, Torazawa K, Meng BY, Sugita M, Deno H, Kamogashira T, Yamada K, Kusuda J, Takaiwa F, Kato A, Tohdoh N, Shimada H and Sugiura M (1986) The complete nucleotide sequence of tobacco chloroplast genome: Its gene organization and expression. EMBO J 5: 2043–2049 Shukla VK, Stanbekova GE, Shestakov SV and Pakrasi HB (1992) The D1 protein of the Photosystem II reaction-centre complex accumulates in the absence of D2: Analysis of a mutant of the cyanobacterium Synechocystis sp. PCC 6803 Mol Micro Biol 6: 947–956 lacking cytochrome Silk G, dela Cruz F and Wu M (1990) Nucleotide sequence of the chloroplast gene for the 4kD K polypeptide of Photosystem II (psbK) and the psbK-tufA intergenic region ofChlamydomonas reinhardtii. Nuc Acids Res 18: 4930 Stirewalt VL, Michalowski CB, Luffelhardt W, Bohnert HJ and Bryant DA (1995)Nucleotide sequence of the cyanelle genome from Cyanophora paradoxa. Plant Mol Biol Rep 13: 327–332 Strasser RJ, Eggenberg P, Pfister K and Govindjee (1992) An equilibrium model for electron transfer in the Photosystem II acceptor complex: An application to Chlamydomonas reinhardtii cells of D1 mutants and those treated with formate. Archs Sci Genève 42: 207–224 Summer EJ, Schmid VHR, Bruns BU and Schmidt GW (1997) Requirement for the PS II-H Phosphoprotein in Photosystem II of Chlamydomonas reinhardtii. Plant Phys 113: 1359–1368 Svensson B, Vass I, Cedergren E and Styring S (1990) Structure

321 of donor side components in Photosystem II predicted by computer modelling. EMBO J 9: 2051–2059 Tae G-S and Cramer WA (1994) Topography of the heme prosthetic group of cytochrome b-559 in the Photosystem II reaction center. Biochemistry 33: 10060–10068 Tae G-S, Black MT, Cramer WA, Vallon O and Bogorad L (1988) Thylakoid membrane protein topology: Transmembrane orientation of the chloroplast cytochrome b-559 in the Photosystem II reaction center. Biochemistry 27: 9075–9080 Takahashi Y, Matsumoto H, Goldschmidt-Clermont M and Rochaix J-D (1994) Directed disruption of the Chlamydomonas chloroplast psbK gene destabilizes the Photosystem II reaction center complex. Plant Mol Biol 24: 779–788 Takahashi Y, Utsumi K, Yamamoto Y, Hatano A and Satoh K (1996) Genetic engineering of the processing site of D1 precursor protein of Photosystem II reaction center in Chlamydomonas reinhardtii. Plant Cell Physiol 37: 161–168 Tamura N and Chenaie G (1985) Effects of Photosystem II extrinsic proteins on microstructure of the oxygen evolving complex and its reactivity to water analogs. Biochim Biophys Acta 809: 245–259 Tang D, Jankowiak R, Seibert M, Yocum CF and Small GJ (1990) Excited-state structure and energy-transfer dynamics of two different preparations of the reaction center of Photosystem II: A hole-burning study. J Phys Chem 94: 6519– 6522 Tang X-S and Satoh K (1984) Characterization of a 47 kD chlorophyll-binding polypeptide isolated from a Photosystem II core complex. Plant Cell Physiol 25: 935–945. Tang X-S, Chisolm DA, Dismukes GC, Brudvig GW and Diner BA (1993) Spectroscopic evidence from site-directed mutants of Synechocystis PCC6803 in favor of a close interaction between Histidine 189 and redox-active Tyrosine 160, both of polypeptide D2 of the Photosystem II reaction center. Biochemistry 32: 13742–13748 Tang X-S, Diner BA, Larsen BS, Gilchrist ML Jr, Lorigan GA and Britt RD (1994) Identification of histidine at the catalytic site of the photosynthetic oxygen-evolving complex. Proc Natl Acad Sci USA 91: 704–708 Tang X-S, Zheng M, Chisholm DA, Dismukes GC and Diner BA (1996) Investigation of the differences in the local protein environments surrounding tyrosine radicals, and in Photosystem II using wild type and the D2-Tyr 160Phe mutant of Synechocystis 6803. Biochemistry 35: 1475–1484 Thompson LK and Brudvig GW (1988) Cytochrome b-559 may function to protect Photosystem II from photoinhibition. Biochemistry 27: 6653–6658 Thompson LK, Miller A-F, de Paula JC and Brudvig GW (1988) Electron Transport in Photosystem II. Is J Chem 28: 121–128 Tommos C, Davidsson L, Svensson B, Madsen C, Vermaas W and Styring S (1993) Modified EPR spectra of the radical in Photosystem II in site-directed mutants of Synechocystis sp. PCC 6803: Identification of side chains in on the D2 protein. the immediate vicinity of Biochemistry 32: 5436–5441 Tommos C, Tang X-S, Warncke K, Hoganson CW, Styring S, McCracken J, Diner BA and Babcock GT (1995) Spin density distribution, conformation and hydrogen bonding of the redoxactive tyrosine, in Photosystem II from multiple electron magnetic-resonance spectroscopies: Implications for photosynthetic oxygen evolution. J Am Chem Soc 117: 10325–

322 10335 Trebst A (1986a) The topology of plastoquinone and herbicide binding polypeptides of Photosystem II in the thylakoid membrane. Z Naturforsch 42c: 240–245 Trebst A (1986b) The three dimensional structure of the herbicide binding niche on the reaction center polypeptides of Photosystem II. Z Naturforsch 42c: 742–750 Turmel M and Otis C (1994) The chloroplast gene cluster containing psbF, psbL, petG and rps3 is conserved in Chlamydomonas. Curr Genet 27: 54–61 van der Vos R, van Leeuwen PJ, Braun P and Hoff AJ (1992) Analysis of the optical absorbance spectra of D1-D2cytochrome b-559 complexes by absorbance-detected magnetic resonance. Structural properties of P680. Biochim Biophys Acta 1140: 184–198 van Gorkom HJ and Schelvis J (1993) Kok’s oxygen clock: What makes it tick? The structure of P680 and consequences of its oxidizing power. Photosynth Res 38: 297–301 van Kan PJM, Otte SCM, Kleinherenbrink FAM, Nieveen MC, Aartsma TJ and van Gorkom HJ (1990) Time-resolved spectroscopy at 10K of the Photosystem II reaction center; deconvolution of the red absorption band. Biochim Biophys Acta 1020: 146–152 van Leeuwan PJ, Nieveen MC, van de Meent EJ, Dekker JP and van Gorkom HJ (1991) Rapid and simple isolation of pure Photosystem II core and reaction center particles from spinach. Photosynth Res 28: 149–153 van Mieghem FJE, Satoh K and Rutherford AW (1991) A chlorophyll tilted 30° relative to the membrane in the Photosystem II reaction centre. Biochim Biophys Acta 1058: 379–385 Vass I and Styring S (1991) pH-dependent charge equilibria between tyrosine-D and the S states in Photosystem II. Estimation of relative midpoint redox potentials. Biochemistry 30: 830–839 Vermaas WFJ, Williams JGK and Arntzen CJ (1987) Sequencing and modification ofpsbB, the gene encoding the CP-47 protein of Photosystem II in the cyanobacterium Synechocystis 6803. Plant Mol Biol 8: 317–326 Vermaas WFJ, Rutherford AW and Hanson O (1988) Sitedirected mutagenesis in Photosystem II of the cyanobacterium Synechocystis sp. PCC 6803: Donor D is a tyrosine residue in the D2 protein. Proc Natl Acad Sci USA 85: 8477–8481 Vermaas W, Charite J and Shen GZ (1990) Glu-69 of the D2 protein in Photosystem II is a potential ligand to Mn involved in photosynthetic oxygen evolution. Biochemistry 29: 5325– 5332 Visser JWM, Rijgersberg CP and Gast P (1977) Photooxidation of chlorophyll in spinach chloroplasts between 10 and 180 K. Biochim Biophys Acta 460: 36–46 von Heijne G, (1986) The distribution of positively charged residues in bacterial inner membrane proteins correlates with the trans-membrane topology. EMBO J 5: 3021–3027 Wales R, Newman BJ, Pappin D and Gray JC (1989) The extrinsic 33 kDa polypeptide of the oxygen-evolving complex of Photosystem II is a putative calcium-binding protein and is encoded buy a multi-gene family in pea. Plant Mol Biol 12: 439–451 Wasielewski MR, Johnson DG, Seibert M and Govindjee (1989) Determination of the primary charge separation rate in isolated Photosystem II reaction centers with 500-fs time resolution.

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Chapter 17 Structure and Function of Photosystem I Andrew N. Webber and Scott E. Bingham

Department of Plant Biology and Center for the Study of Early Events in Photosynthesis,

Arizona State University, Box 85287-1601, Tempe, AZ 85287-1601, U.S.A.

Summary I. Introduction A. Photosystem I Function B. Photosystem I Proteins C. Evolutionary Aspects of Photosystem I II. Structure of Photosystem I A. PsaA and PsaB Subunits B. Electron Transfer Cofactors C. Antenna Chlorophylls III. Nature and Function of Electron Transfer Cofactors A. B. A and C. D.

E. and IV. Antenna Structure and Function V. Function of Photosystem I Subunits A. Peripheral Subunits Not Binding Electron Transfer Cofactors 1. Acceptor Side 2. Donor Side 3. Intrinsic Subunits B. Subunits Binding Electron Transfer Cofactors 1. PsaA and PsaB Binding Domain a. Binding Domain b. Binding Domain c. 2. PsaC VI. Biogenesis of Photosystem I Acknowledgments References

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Andrew N. Webber and Scott E. Bingham

Summary Photosystem I, a membrane protein complex found in all oxygenic photosynthetic organisms, uses light energy to transfer electrons from plastocyanin to ferredoxin. Light energy captured by antenna chlorophylls is The excited primary donor then initiates electron transfer transferred rapidly to the primary electron donor, and A three-dimensional structural model of the through a series of secondary acceptors Photosystem I complex at 4 Å resolution is used to describe the spatial organization of cofactors. The chemical identity of each of the electron transfer cofactors based on current spectroscopic information is discussed in relation to the available structural information. The use of specific mutagenesis to probe the structure and function of individual subunits and to investigate the interaction of cofactors with the protein environment is then described. The combined results from structural, spectroscopic and molecular analysis are generating a unified model of the structure, function and evolution of Photosystem I and related reaction centers.

I. Introduction

A. Photosystem I Function Photosystem I is a multimeric membrane protein complex that uses light energy to initiate electron transfer from plastocyanin (PC), located in the thylakoid lumen, to ferredoxin (Fd) located on the stromal side of the thylakoid membrane (reviewed by Golbeck, 1992; Sétif, 1992; Nechushtai et al., 1996). Reduced ferredoxin is used for reduction of or other alternate electron acceptors (Knaff, 1996). In Chlamydomonas and other green algae cytochrome can replace plastocyanin under copper deficient growth conditions (Chapter 31, Merchant). Light energy captured by antenna pigments in PS I is rapidly transferred to the primary electron donor, where charge separation is initiated. The excited transfers an electron primary donor molecule, to the first spectroscopically detectable electron in 1.5 ps (Fig. 1). The initial charge acceptor, separated state, is then rapidly stabilized by electron transfer through a series of secondary and Either electron acceptors termed is then able to reduce Fd. Spectroscopic or analysis has been used to assign a chemical composition to each cofactor (Golbeck, 1992; Sétif, and 1992; Malkin, 1996). The primary donor, first detectable acceptor, are Chl a molecules. Abbreviations: A – Chl a molecule positioned between and – The first electron acceptor in PS I; – The second electron acceptor in PS I; BChl – bacteriochlorophyll; Chl – chlorophyll; ENDOR – electron nuclear double resonance; EPR – electron paramagnetic resonance; – iron-sulfur center A; – iron-sulfur center B; Fd – Ferredoxin; – primary electron donor in PS I; PC – Plastocyanin; PS I – Photosystem I; PS II – Photosystem II; RC – reaction center

is a phylloquinone (vitamin ). and are each [4Fe-4S] iron-sulfur clusters. An additional and Chl a molecule, termed A, located between was also identified in a 4 Å structural model of PS I (Krauss et al., 1996; Section II). The 4 Å structure also shows that the cofactors between and form two symmetrical branches, either one or both of which may be involved in electron transfer (Krauss et al., 1996).

B. Photosystem I Proteins The PS I complex (Fig. 2) is composed of 10–14 individual polypeptides, depending on the organism studied (Nechushtai et al., 1996). These subunits have been named PsaA to PsaN. The PsaG, PsaH and PsaN subunits are found only in PS I complexes from eukaryotes while PsaN has so far only been found in complexes from prokaryotes. The PS I complex in Chlamydomonas is encoded by genes located in both the nuclear and chloroplast genome (Fig. 2). Polypeptides encoded by genes in the nucleus are translated in the cytosol and post-translationally imported into the chloroplast. The two largest subunits, PsaA and PsaB, form a heterodimeric reaction center complex (Fig. 2) and coordinate the through (Golbeck, electron transfer cofactors 1992). In addition, the PsaA and PsaB polypeptides coordinate approximately 100 Chl a molecules and 10–16 carotenoids (Nechushtai et al., 1996). The electron acceptors and are coordinated by a 9 kDa subunit, PsaC, associated with the stromal side of PS I (Dunn and Gray, 1988). The remaining polypeptides are important for normal assembly and accumulation of PS I and several may be involved in correct binding of soluble electron donors and acceptors such as PC and Fd

Chapter 17 Photosystem I

(Section V). The PsaD and PsaE subunits are located on the stromal side of the complex where they have been shown to provide a docking site for Fd (Fig. 2). PsaE may also be required for cyclic electron transport. On the lumenal side of PS I the PsaF and PsaN subunits may be involved in docking PC. The PS I complex also contains two hydrophobic low molecular weight polypeptides, PsaI and PsaJ, that are important for normal levels of PS I accumulation (Section V.A.3). PS I from plants and green algae is also associated with additional pigment-protein complexes referred to as light harvesting complex I (LHCI). The LHC complex is composed of several distinct polypeptides which together coordinate an additional 100 Chls (Mullet et al., 1980). The LHC I complex contains both Chla and Chlb at a ratio of approximately 3.5:1.

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(Nitschke and Rutherford, 1991). Early evidence that reaction center proteins from Chlorobium limicola contained iron-sulfur centers similar to and was obtained by EPR spectroscopy (Nitschke et al., 1990). Later, the reaction center genes from Chlorobium limicola (Büttner et al., 1992) and Heliobacillus mobilis (Liebl et al., 1993) were cloned and sequenced and shown to encode proteins with some sequence identity to PsaA and PsaB. A cysteine rich region known to coordinate the iron-sulfur center was the most highly conserved region (Büttner et al., 1992; Liebl et al., 1993). Only a single gene encoding the reaction center protein has been identified in Chlorobium limicola and Heliobacillus mobilis supporting the current notion that the bacterial reaction center is a homodimer (Büttner et al., 1992; Liebl et al., 1993). During the course of evolution the gene encoding the bacterial RC has presumably duplicated and diversified to give rise to thepsaA and psaB genes. Although there is very little amino acid sequence identity between the different classes of RC, it is becoming increasingly clear that the cofactor arrangement and protein domains involved in cofactor coordination are similar between proteins in type I and type II (quinone as terminal electron acceptor) RCs. This indicates that both type I and type II reaction centers arose from a common ancestor (Vermaas, 1994).

C. Evolutionary Aspects of Photosystem I PS I is a member ofthe iron-sulfur center containing (type I) reaction centers that also include the reaction centers of green-sulfur bacteria and Heliobacteria

II. Structure of Photosystem I The PS I complex isolated from the thermophylic

326 cyanobacterium Synechococcus elongatus has been crystallized and a model generated from X-ray diffraction data at approximately 4 Å resolution (Krauss et al., 1996). The PS I complex from S. elongatus contained 11 polypeptides (PsaA-F and PsaI-M) and was isolated and crystallized in a trimeric form. Although the cyanobacterial complex lacks several subunits found in PS I from eukaryotic sources, it is still expected that the basic structure of the complex, especially of the core reaction center subunits, is very similar to that found in plants and green algae. Two different views of the PS I monomer are shown in Fig. 3A. The figure also includes a portion of the 2nd and 3rd monomers of the trimer to show the interface region. Each monomer contains 31 transmembrane and 12 non-membrane spanning Eleven transmembrane and four surface are each assigned to PsaA and PsaB were assigned to the (Fig. 3A). Other remaining subunits of the PS I complex (Krauss et al., 1996). At the interface of each monomer (the center of the trimer) are three helices, p, q and r, that form the connecting domain (Fig. 3A). Two of these helices have been assigned to the PsaL subunit (Krauss et al., 1996) based on the finding that cyanobacterial mutants lacking PsaL cannot form trimers (Chitnis and Chitnis, 1993). Because PsaL was shown to be closely associated with PsaI in PS I of Synechocystis sp. 6803 (Xu et al, 1995), the third helix was assigned to PsaI (Krauss et al., 1996). Hydrophobicity analysis of the derived PsaF amino acid sequence indicates that it contains at least one hydrophobic membrane spanning domain and PsaF was found to be associated with psaJ (Xu et al., 1994c). This has led to the assignment of helices u, v, w, and x to PsaF and PsaJ (Krauss et al., 1996). Fig. 3B shows that 3 form a large ridge (rising almost 35 Å from the membrane surface) on the stromal surface. These helices were assigned to the PsaC and PsaD subunits (Krauss et al., 1996).

A. PsaA and PsaB Subunits The 4 Å structure shows a total of 30 helices that lie on either side of a pseudo-C2 symmetry axis (Krauss et al., l996). Fifteen ofthese helices were assigned to PsaA and 15 to PsaB. The helices were labeled a-o and to reflect this pseudo symmetry. Each

Andrew N. Webber and Scott E. Bingham subunit contains eleven transmembrane and four surface This agrees with previous predictions based on hydropathy analysis ofthe PsaA and PsaB amino acid sequences indicating that PsaA and PsaB each have 11 membrane spanning regions (Fish et al., 1985). The helices a-o and were labeled in sequence from the N-terminus to the Cterminus of the protein (Krauss et al., 1996). The ten inner-most membrane spanning helices i, j, k, m and o (and and ) surround the electron through (Krauss et al., transfer cofactors 1996). Since thehelices i, j, k, m and o are sequentially connected, the inter-helical loops connecting helices j, k and are inferred to contain the cysteine residues coordinating (Krauss et al., 1996).

B. Electron Transfer Cofactors The electron transfer cofactors are arranged on either side of a pseudo-C2 symmetry axis that runs between on the stromal side, and on the lumenal side are (Fig. 4). The two Chl a molecules that form parallel but offset from each other so that the ring overlap is very small. The center to center distance is Å and the interplane distance Å (Krauss et al., 1996). Four additional Chl monomers have been identified in the structure and these form two potential branches for electron transfer. One or both ofthe two were assigned as Chl monomers furthest from (Krauss et al., 1996). The Chls are 16 Å from and separated from each other by 27 Å. Between and are two Chls named A and A´. These have been termed accessory Chls by analogy to a similar pair of BChl a molecules found in reaction centers from purple bacteria. The iron-sulfur center assigned to and two ironsulfur centers, and assigned to either or lies on the are shown in Fig. 3B and Fig. 4. pseudo-C2 axis of symmetry with the axis between and at an angle of 54°. This places 15 Å from and 22 Å from and are 12 Å apart (Krauss et al., 1996). Assignment of or to or is still unclear. The two short observed close to and are thought to be associated with PsaC (Krauss et al., 1996). The exact position of the two has not phylloquinone molecules, one ofwhich is been identified. The head group ofthe phylloquinone is smaller than Chl and can not be distinguished at the current resolution (Krauss et al., 1996).

Chapter 17 Photosystem I

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Andrew N. Webber and Scott E. Bingham present structural model, we anticipate that additional Chls will be identified. As the resolution improves the model will, naturally, be re-evaluated.

III. Nature and Function of Electron Transfer Cofactors The spectroscopic evidence for the chemical nature of PS I cofactors has been well reviewed (Golbeck, 1992, 1994; Sétif, 1992; Malkin, 1996). This section will briefly address each of the PS I cofactors with emphasis on results obtained during the past few years.

C. Antenna Chlorophylls The organization of the Chls associated with the antenna of PS I is shown in Fig. 5. A total of 65 antenna Chl a molecules and 6 electron transfer Chl a were identified. Viewed from the stromal side the 65 antenna Chl a molecules form an ellipsoid ring around the central core of electron transfer cofactors (Krauss et al., 1996). The closest distance between and any antenna Chl a is > 20 Å. Two Chl a molecules called cC and cC´ are 14 Å from and and may serve to connect Chls between the antenna and electron transfer Chls. It was speculated that the pathway of excitation energy transfer from may be via the Chls cC, and A the antenna to (Krauss et al., 1996). Since all the Chls associated with PsaA and PsaB (up to 100 Chl based on spectroscopic studies) are not accounted for in the

The assignment of as a monomer or a dimer of Chl a molecules has been long debated in the literature (Sétif, 1992). While much evidence now supports that is a Chl a dimer there are still some issues that require clarification. The crystal structure shows are that the two Chl a molecules assigned to within 4 Å, but with only minimal overlap of the ring systems (Krauss et al., 1996). The two Chls are further apart than the BChl dimer in the purple bacterial reaction centers indicating that a weaker interaction between the two Chls would be expected. At least part of the confusion lies in the fact that the interaction between the two Chls may be different depending on whether the ground state, triplet state or oxidized state is studied. Recent work has focused on the electronic structure of the cation radical, using EPR and ENDOR (electron nuclear double resonance) spectroscopy. ENDOR spectroscopy provides quantitative information on the interactions between the spin of the unpaired electron and spins of various magnetic Such interactions are referred to as nuclei spectra of PS I hyperfine couplings. The from Chlamydomonas reinhardtii is shown in Fig. 6. The largest hyperfine couplings observed by ENDOR of monomeric Chl in vitro, or in vivo, were assigned (Käß et al., 1995) to the methyl protons at positions 12 of Chl a (Fig. 7). Progressively weaker hyperfine couplings were assigned to methyl protons at position 7 and 2, respectively (Käß et al. 1995). The average reduction in spin distribution in compared to Chl in vitro was 15% (Käß et al., 1995). This could be interpreted to indicate that the spin density is distributed between the two Chl a

Chapter 17 Photosystem I

molecules of a dimer, with one Chl carrying at least 85% of the spin distribution. However, the hyperfine couplings for the Chl carrying the smaller (15%) portion of charge could not be detected in these experiments. Previous measurements as monomeric Chl a perturbed have interpreted by the protein environment (O’Malley and Babcock, 1984), and this interpretation cannot be ruled out at the present time. The reduced average hyperfine compared to monomeric Chl a, coupling in observed in the most recent studies, could in fact be due to perturbation of the molecular spin geometry by the protein.

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B. A and The first component that has been identified clearly is The chemical as the electron acceptor from identity of has been confused by the lack of a clear spectroscopic signal that could be attributed to without significant interference from other radicals (reviewed by Malkin, 1996). The current consensus is a monomeric Chl a, which is also is that consistent with the 4 Å structural model (Krauss et al., 1996). Time resolved kinetic studies have examined the under physiological optical characteristics of

Andrew N. Webber and Scott E. Bingham

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conditions using PS I preparations from cyanobacteria (Holzwarth et al., 1993; Hastings et al., 1994 a,b), Chlamydomonas (Hastings et al., 1995; Melkozernov et al., 1997 and spinach (Hastings et al., 1994 a,b; White et al., 1996). Under low intensity flashes no absorption changes could be observed because the is faster than reduction of the reoxidation of initial acceptor ( formation) This limitation was overcome by using high intensity flashes which to measure allowed sufficient accumulation of in PS I absorption changes. The spectrum of preparations isolated from Chlamydomonas (Hastings et al., 1995, Melkozernov et al., 1997) showed a maximum bleaching at approximately 686 nm (Fig. 8). This spectrum was similar to previously reported spectra for obtained from ethyl ether extracted PS I preparations that lack (Mathis et al., 1988). The spectrum of showed a shoulder at approximately 670 nm, and this was taken as an indication that more than one Chl a contributes to the absorption spectrum (Hastings et al., 1995; Melkozernov et al., 1997). Hastings et al. (1995) speculated that this * The first two letters of the mutant name indicate the amino acid change (e.g., HN; His to Asn). This is followed in parenthesis by the subunit and amino acid number (e.g., B656; PsaB residue 656).

second Chl a may represent the acceptor A observed in the crystal structure. However, no kinetic evidence obtained so far supports the notion that an additional Chlfunctions as a real intermediate in electron transfer (Hastings et al., 1995; Malkin, between 1996; Melkozernov et al., 1997).

C. It is now generally acceptedthat is a phylloquinone molecule although the current evidence (vitamin is not definitive (Malkin, 1996). If is a quinone, the protein environment must significantly perturb the molecule to lower the midpoint potential of phylloquinone from ca. –100 mV, typical of the midpoint potential observed in vitro, to ca. –800 mV observed in isolated PS I. The generally accepted stoichiometry is two moles of phylloquinone per Only one quinone is thought to particimole of pate as an electron transfer intermediate between and The presence of phylloquinone in PS I and the effects of extraction and/or reconstitution on PS I activity has been discussed in detail in several recent reviews (Golbeck, 1992; 1994, Malkin, 1996). The most convincing evidence that is a quinone comes from EPR spectroscopy ofphotoaccumulated photoaccumulated at 205 K in the presence of

Chapter 17 Photosystem I dithionite shows an asymmetric EPR spectrum (g=2.0048, peak to peak linewidth 0.95 mT) which is compatible with reduced phylloquinone (Mansfield and Evans, 1988). Following treatment that leads to a double reduction of (Heathcote et al., 1993), the EPR spectrum cannot be produced, further suggesting that is a quinone. Another EPR technique, called electron spin polarized (ESP) EPR, was used to detect a transient light induced signal that arises from the radical pair state Double reduction of also led to a loss of the ESP-EPR signal (Snyder et al., 1991). Replacement of with deuterated phylloquinone led to a narrowing of the spectrum providing strong evidence that the phylloquinone contributes to the spin polarized signal (Rustandi et al., 1990). Very recently, pulsed EPR spectroscopy has been used to compare the out of phase electron spin echo of PS I with similar signal arising from the signals from the state of Zn substituted reaction centers from Rhodobacter sphaeroides (Bittle and Zech, 1997). The magnetic dipolar interaction has a very strong distance dependence that allowed a determination of the distance between the radicals. The crystal structure of the purple bacterial reaction center is well resolved allowing the accuracy of the interpretation to be assessed. The spin coupling in PS I was found to be significantly stronger than in the bacterial reaction centers indicating a shorter distance and than between and (Bittle between and Zech, 1997). Calculation determined the distance between the primary donor of PS I and the quinone acceptor at 25.4 A ± 0.3 A, approximately 3 Å and (Bittle and Zech, shorter than between 1997).

D. The historically controversial assignment of as either a [2Fe-2S] or [4Fe-4S] iron-sulfur center, has been resolved in the past few years. All evidence indicates that is a [4Fe-4S] (reviewed by Golbeck, 1992, 1994). The cluster is ligated by two cysteines from PsaA and two cysteines from PsaB forming an inter-polypeptide iron-sulfur cluster. The role of the cysteines in ligating has been confirmed by sitedirected mutagenesis (Section V.B.1.a). A role for as a real intermediate in forward electron transfer between and has only recently been resolved. This has been addressed by studying and have the kinetics of reoxidation when

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been removed by chemical treatments which deplete the PS I complex of PsaC. In the absence of transient EPR (Moenne-Locoz, 1994), electron spin echo (van der Est et al., 1994) and transient optical spectroscopy (Lünberg et al., 1994) showed that reoxidation kinetics were not changed. Mutants with an altered due to replacement of one of the cysteine ligands with serine showed impaired forward electron transfer (Vassilev et al., 1996). These results support a sequence of electron transfer from via to which is also consistent with the spatial organization of revealed by the X-ray and structure.

E.

and

The iron-sulfur centers and are both [4Fe-4S] centers coordinated by the PsaC protein. The role of each iron-sulfur cluster in forward electron flow is still unresolved. Based on the midpoint potentials it is expected that electronflowwill occurfrom to to The crystal structure showed three iron-sulfur clusters, but only could be assigned unambiguously. The identity of and could not be assigned, and therefore the clusters were termed The distance between and is 15 Å and and between and is 22 Å. Based on this structure and knowledge of the relative midpoint potentials, it would be expected that the most distal cluster is However, the assignment of the electron transfer pathway is confused by potentially conflicting experimental data (reviewed by Golbeck, 1992;

332 Malkin, 1996). For example, selective denaturation does not impair room temperature or low of (Golbeck and temperature photoreduction of Warden, 1982; Fujii et al., 1990; Jung et al., 1995). Using site-directed mutagenesis to replace Cys 14, a with Asp, the center could be ligand to inactivated in Synechocystis (Zhao et al., 1992) and Anabena PsaC (Mannan et al., 1996). Analysis of these mutants showed that the presumed inactivation (Zhao et al., of did not impair photoreduction of 1992; He et al., 1995; Mannan et al., 1996). However, replacement of Cys 14 with Asp has been shown to lead to assembly of an altered iron-sulfur cluster that may still serve as an electron sink (Yu et al., 1995), complicating interpretation of results from such site-directed mutants. In Chlamydomonas, specific mutation of K52 and R53 of PsaC to serine and alanine, respectively, removes two positive In the K52S/K53A PsaC charges close to center mutant is preferentially reduced at 77K while at room temperature electron transfer from PS I to Fd proceeds at normal rates (Fischer et al., 1997). Taking a slightly different approach, Rodday et al. (1996) have examined the role of surface exposed acidic residues of PsaC in mediating interaction with the PsaA and PsaB core subunits. Specific mutation of D9 of the PsaC subunit to alanine impaired reconstitution of the mutant PsaC protein, produced by overexpression in E. coli, to a PsaC depleted PS I core preparation (Rodday et al., 1996). Mutation of other surface exposed residues did not impair reconstitution. Rodday et al. (1996) proposed a model With a large amount of where is proximal to conflicting information, it is likely that assignment of and will await a higher resolution structure. IV. Antenna Structure and Function PS I is associated with an accessory antenna complex that contains Chl a and b, referred to as the LHCI complex (Mullet, 1980). The Chl a/b ratio is approximately 3.5:1. In plants, the LHCI complex is composed of 4 individual polypeptides referred to as Lhcal-4 with molecular weights ranging from 17– 26 kDa. The LHCI complex from Chlamydomonas has been isolated and characterized (Bassi et al. 1992). The LHCI complex can be resolved into two Chl a/b binding fractions with fluorescence emission maxima at 680 nm and 705 nm (Bassi et al., 1992). The precise number and stoichiometry of poly-

Andrew N. Webber and Scott E. Bingham peptides associated with each Chl complex is somewhat unclear, although Bassi et al (1992) propose as many as ten polypeptides in the molecular weight range 21–26 kDa. A cDNA clone for a 20 kDa LHCI component (sometimes termed p22) of Chlamydomonas has been characterized by Hwang and Herrin (1992). The PS I reaction center complex, often referred to as the PS I core complex, contains approximately 100 Chl a molecules (Golbeck, 1992). All of these Chls are coordinated by the PsaA and PsaB reaction center subunits. Sixty-five antenna Chl a molecules have been identified in the 4 Å model of PS I that form a funnel-like network to transfer excitation energy to the electron transfer cofactors (Fig. 5). PS I also contains several Chls that absorb at wavelengths longer than that absorbed by the primary donor (these are often termed red Chls). The function of the long wavelength-absorbing Chls is unclear, although it has been suggested that they may focus energy transfer to the reaction center (Van Grondelle and Sundström, 1988) or increase the absorption cross section (Trissl, 1993). The antenna Chls function to capture light energy and effectively transfer it to the reaction center Chls. Excitation energy transfer in the antenna of PS I includes several phases of different life times. Excitation of individual Chl a molecules leads to a rapid transfer ofexcitation energy between individual Chl a molecules that occurs within 0.2 ps (Du et al., 1993). There is also excitation energy transfer between small pools of pigments that occurs within 2–8 ps (Klug et al, 1989; Du et al., 1993; Holzwarth et al., 1993; Hastings et al., 1994a,b, 1995). As a result of these processes the excitation energy is transferred in 20–30 ps to the primary electron donor, (Owens et al, 1988, 1989; Holzwarth et al 1993; Hastings 1994a,b, 1995). Due to considerable spectral overlap between heterogeneous antenna Chl a it has been difficult to develop molecules and appropriate models that describe the energy transfer process in PS I. Two models have been developed to describe excitation energy transfer from the antenna One model, called the diffusion limited model, to assumes that there is a bottle-neck in the antenna that limits the rate of overall excitation energy transfer to (Van Grondelle and Sundström, 1988). The alternate trap limited model assumes that the intrinsic rate of primary charge separation is the limiting factor and so the excitation can visit the trap several times before being used for photochemistry (Pearlstein, 1982; Trissl, 1993). A variation of the

Chapter 17 Photosystem I latter model, termed the special trap limited model, assumes that energy transfer from the longwavelength absorbing Chls is the limiting factor (Van Grondelle and Sundström, 1988; Otte et al., 1993). In order to test the above models, the decay of excitation energy in mutants of Chlamydomonas that have an increased midpoint potential of the primary donor molecule was studied (Krabben et al., 1995; Webber et al., 1996). These Chlamydomonas mutants have a site-directed change ofresidue 656 of PsaB that substitutes His for Asn (described in more detail in Section V.B.1.b). The increased midpoint potential (40 mV more positive) was expected to decrease the driving force for electron transfer and slow the rate of primary charge separation (Melkozernov et al., 1997). This would be manifest in the decay of excitation energy in the antenna if a trap limited process operates in PS I. Analysis of the transient absorption spectra from wild-type PS I (prepared from a PS II minus strain, FuD7) indicated that the excitation energy decayed with a half time of 30 ps. In the HN(B656) mutant PS I the half time of excitation energy decay was increased to 65 ps (Melkozernov et al., 1997). The results thus supported the notion that excitation energy transfer in PS I is best described by a trap limited model. However, a caveat is that it was not possible to directly measure the rate of primary charge separation in either wildtype PS I or mutants. Therefore, the possibility that a special trap limited model best describes PS I excitation energy trapping could not be excluded. It is possible that the red pigments are coupled to and that a change in the midpoint potential of may change the lifetime of the trapping process (Melkozernov et al., 1997). In this regard it is interesting to note that the two pigments, cC and cC´ (Figs. 4 and 5) are close to the electron transfer chain. More work is needed to fully understand the excitation energy dynamics in PS I. A complete understanding will require a much higher resolution structure combined with further studies of specific mutants that effect the antenna excitation energy dynamics and electron transfer steps.

V. Function of Photosystem I Subunits In recent years molecular genetics has played an important role in defining the function of individual PS I subunits. Much of the in vivo work in this area

333 has involved site-directed mutagenesis or deletion mutagenesis of PS I genes in cyanobacteria, and of chloroplast and nuclear genes in Chlamydomonas reinhardtii. Reconstitution ofdepleted PS I complexes with mutant proteins expressed in E. coli has also been a valuable approach to understanding the function of acceptor side subunits.

A. Peripheral Subunits Not Binding Electron Transfer Cofactors The topography of the accessory subunits in PS I has been studied in detail using a range of biochemical techniques. Since the gene sequence encoding each known PS I protein has been obtained, in many cases from a number of different organisms, predictions as to whether the protein is integral or extrinsic to the membrane, or lies on the stromal or lumenal side of the membrane, have been made and are summarized in Table 1. A large body of information has also been obtained on the function ofthe accessory subunits by analysis ofdeletion mutants and site-directed mutants. Much of this has been authoritatively reviewed in recent years and readers are referred to these reviews for more detail (Golbeck 1994; Chitnis et al, 1995; Chitnis 1996).

1. Acceptor Side Subunits on the acceptor side of PS I are expected to be involved in mediating interaction between the core proteins and PsaC (electron transfer between and also between PsaC and ferredoxin or and flavodoxin. A combination of studies involving protease susceptibility, chemical crosslinking and protein modifications have demonstrated that PsaD and PsaE, along with PsaC, are peripheral proteins associated with the stromal side of cyanobacterial and plant PS I complexes (Fig. 2; Golbeck 1994; Chitnis et al, 1995; Chitnis 1996). These subunits are suggested to form the 35 Å high ridge on the stromal side of PS I observed in the crystal structure (Krauss, 1996). PS I from plants and green algae contain an additional polypeptide, PsaH, also located on the stromal side of the complex. Studies with zero length cross linkers have shown that Fd (Zanetti and Merati, 1987; Zilber and Malkin, 1988) and flavodoxin (Mühlenhoff et al., 1996) can be cross linked with PsaD. PsaE has also been shown to cross-link with PsaD. A PsaD-less mutant Synechocystis sp. 6803

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(Chitnis et al., 1989a) grew at normal rates under photoheterotrophic growth conditions but only poorly photoautotrophically. Thylakoid membranes isolated from the psaD mutant were unable to reduce via Fd (Xu et al., 1994e) but were still able to reduce flavodoxin at a reduced rate. These results demonstrated that PsaD is important for optimal PS I activity consistent with its proposed role in docking Fd and binding PsaC. Deletion of psaE from Synechocystis sp. 6803 resulted in a mutant that showed normal rates of photoautotrophic growth (Xu et al., 1994e). Ferredoxin-mediated and flavodoxin-mediated photoreduction in the mutant were 5% and 35% of wild-type, respectively. A PsaE deletion mutant of Synechococcus sp. PCC 7002 grow photoautotrophically at high light intensity (Yu et al., 1993) but not photoheterotrophically in the presence

Andrew N. Webber and Scott E. Bingham

of glucose and DCMU, or photoautotrophically under low light and air concentration of (Yu et al., 1993). Similar results were reported for the PsaEless Synechocystis mutant (Yu et al., 1993). It was suggested that cyclic electron flow around PS I was impaired in the PsaE-less mutants. The ndhF gene, encoding a component of the respiratory complex, was deleted from the PsaE-less Synechococcus sp. PCC 7002 mutant to generate a double mutant. Cyclic electron transport was then measured reduction in the presence by comparing rates of and absence of methyl viologen. In the mutant the rate of reduction of was the same in the presence and absence of methyl viologen indicating that cyclic electron flow was absent (Yu et al., 1993).

Chapter 17 Photosystem I

2. Donor Side In plants and green algae, the PsaF and PsaN polypeptides (He and Malkin, 1992; Knoetzel and Simpson, 1993) are located on the lumenal side of PS I. The PsaN polypeptide is not found in cyanobacteria. PsaF has typically been considered an extrinsic polypeptide located on the lumenal side of PS I. That the bulk of PsaF is located on the lumenal side is supported by biochemical evidence that PsaF could be cross-linked to PC in spinach PS I (Wynn and Malkin, 1988) and to cytochrome in Synechococcus sp. PCC 7002 and 6301 PS I (Wynn et al., 1989). The C-terminal region of PsaF, however, was susceptible to proteolysis in the absence of PsaD and PsaE, and could be cross-linked to PsaE (Armbrust et al, 1996) indicating it is located on the stromal side of PS I (Xu and Chitnis, 1996). The thylakoid targeting sequence on the N-terminus of the PsaF pre-protein is presumably required to transfer the bulk of the polypeptide across the thylakoid membrane and the hydrophobic ammo acids toward the C-terminus are assumed to function as a stoptransfer signal, as proposed also for Cyt f. Based on results from chemical cross linking it was suggested that PsaF provides a docking site for orienting the protein for PC and cytochrome (Wynn and Malkin, efficient electron transfer to 1988). The necessity of PsaF for docking plastocyanin has been questioned since a PsaF-less mutant of Synechocystis sp. PCC 6803 was found to grow at rates equivalent to wild-type (Chitnis et al., 1991). However, more recent analysis of a PsaF deletion mutant ofChlamydomonas indicated altered electron (Farah et al., 1996). transfer from reduced PC to is In Chlamydomonas, electron donation to biphasic with a fast phase of approx. 3 and a slower phase with a half-time of 100–200 (Farah et al., 1996; Hippler et al., 1997). The fast phase is attributed to a first order electron transfer between The slower phase is representative bound PC and of electron transfer from unbound PC to PS I. In cyanobacteria, the fast phase of electron transfer is absent. The PsaF deletion mutant of Chlamydomonas lacked the fast phase of electron transfer, probably because a PC-PS I complex that allowed rapid electron transfer did not form (Farah et al., 1996; Hippler et al. 1997). The N-terminus of PsaF from plants and algae contains a conserved domain with several positively charged amino acids (Hippler et al., 1996). This domain would be located on the lumenal side of

335 PS I and could interact with negatively charged residues on PC. Interestingly, this N-terminal domain is missing from cyanobacterial PsaF, leading Hippler et al. (1996) to suggest that the N-terminal domain is required for complex formation between PC and PS I.

3. Intrinsic Subunits Deletion of the psaL gene has no impact on photoautotrophic growth of Synechocystis sp. PCC 6803 although the mutant grows faster than wildtype under photoheterotrophic conditions in the DCMU (Chitnis et presence of glucose and al., 1993). An interesting observation was that the PsaL-less PS I complexes, isolated by solubilization failed to form a trimeric with PS I complex (Chitnis and Chitnis, 1993). This has led to the suggestion that PsaL may form the small ‘connecting’ domain, observed in the crystal structure of trimeric PS I complexes (Fig. 3 A), that links the PS I monomers to form a trimer (Krauss et al., 1996). Based on this the PsaL subunit was assigned to two of the three helices, p q and r in the connecting domain (Krauss et al., 1996). Several low molecular weight subunits PsaI, PsaJ and PsaK are also associated with plant and algal complexes. A PsaI-less strain of Synechocystis sp. PCC 6803 grew at normal rates and isolated membranes showed only a small decrease in photoreduction (Xu et al., 1995). Membranes from the PsaI-less mutant contained only 20% of the normal level of PsaL. The level of psaL mRNA was reduced by 95% in the PsaI-less mutant. Since the psaI gene is located only 90 bp downstream from psaL, insertional inactivation of psaI may have affected psaL transcription, or stability of the mRNA, leading to a reduced level of synthesis of PsaL. PsaL in the PsaI-less membranes showed an increased sensitivity to proteolysis, leading to the conclusion that PsaI is important for the structural organization of PsaL. When psaJ was deleted in Synechocystis sp. PCC 6803 there was again no impairment of photoautotrophic growth. However, there was loss of PsaF during detergent purification of PS I and it was suggested that PsaJ is important for stable association of PsaF with the complex (Xu et al., 1994c,d). The function ofthe remaining PS I polypeptides is mostly unknown. The psaK gene product is predicted to span the thylakoid membrane twice and is synthesized with a presequence that may target the

Andrew N. Webber and Scott E. Bingham

336 N-terminus towards the lumenal side of PS I. PsaK is closely associated with the PsaA and PsaB polypeptides and can be removed by treatment with reducing agents without affecting primary charge separation (Wynn and Malkin, 1990). PsaK and PsaG have been shown to cross link to Lhca3 and Lhca2 in PS I of spinach and barley (Jansson et al., 1996). The PsaG, PsaH and PsaN polypeptides are found only in PS I complexes from eukaryotic algae and plants. There are no known function for these subunits, although it is possible that they are involved in optimizing structural and functional interaction between the reaction center and LHCI complexes (Jansson et al., 1995). The ability to make nuclear knock-out mutants in Chlamydomonas, such as the PsaF deletion mutant described earlier, may enable the production of mutants that lack the PsaG, PsaH and PsaN proteins.

B. Subunits Binding Electron Transfer Cofactors

1. PsaA and PsaB The PsaA and PsaB polypeptides coordinate the and early electron primary electron donor, acceptors and The psaA and psaB genes have been sequenced from a number of plant and cyanobacteria species. The derived amino acid sequences of PsaA and PsaB show an overall identity of 45%. Hydrophobicity profiles for both PsaA and PsaB are very similar and predict that each polypeptide contains eleven membrane spanning domains, consistent with the current structural model. The reaction center protein of both Chlorobium limicola (Büttner et al., 1992) and Heliobacillus mobilis (Liebl et al., 1993) is predicted to be highly hydrophobic and to contain up to eleven membrane spanning regions. The overall amino acid sequence identity between the PsaA and PsaB proteins with the C. limicola reaction center is approximately 14% (Büttner et al., 1992) and with the H. mobilis reaction center approximately 16% (Liebl et al., 1993). This is close to the 17.4% identity between the reaction center proteins of H. mobilis and C. limicola (Liebl et al., 1993). The much higher identity between PsaA and PsaB than to the H. mobilis or C. limicola polypeptides suggests that divergence between the bacterial reaction center genes and PS I reaction center genes occurred before a gene duplication that gave rise to psaA and psaB (Liebl et al., 1993; Vermaas, 1994).

a.

Binding Domain

The most highly conserved region in PsaA and PsaB is in the membrane spanning helix j and the connecting sequences between helix j and k (Golbeck, 1992). Cysteine residues located in the connecting region have been predicted to coordinate the iron-sulfur is a [4Fe-4S] iron-sulfur center, center Since four cysteines are required for coordination (Golbeck, 1992). PsaA and PsaB each have two cysteines in this domain necessitating that is an interpolypeptide [4Fe-4S] iron-sulfur center. The binding region, PCDGPGRGGTC, is identical in all PsaA and PsaB sequences and is approximately 75% identical to a similar region in the reaction center protein of H. mobilis and C. limicola (Fig. 9). The role of the cysteine containing domain (Fig. was tested by site-directed 9) in coordinating mutagenesis of psaB in Chlamydomonas and Synechocystis sp. PCC 6803. When the first cysteine of this domain, C561 of Chlamydomonas PsaB, was replaced by a histidine, a PS I complex did not accumulate in the mutants (Webber et al., 1992, 1993). The PsaB protein was translated indicating that the mutation affects a post-translational stage in assembly or stability of PS I. Substitution of the adjacent proline with alanine or leucine had no effect on accumulation of the PS I complex (Webber et al., 1992, 1993). This result showed that the cysteine played a crucial role in PS I assembly and provided some of the first experimental evidence that this A similar proregion is involved incoordinating cys motif has been found in the [Fe-S] binding region of nearly all ferredoxins and it was surprising that substitution of the proline did not affect PS I. Similar results were reported with the ferredoxin iron-sulfur protein from Clostridium pasteurianum (Gillard et al, 1993; Quinkal et al., 1994). Mutagenesis of the two prolines adjacent to cysteine ligands in Clostridium pasteurianum ferredoxin was not found to alter the EPR characteristics of the reduced ironsulfur centers (Gillard et al, 1993; Quintal et al., 1994). Site-directed mutants have also been generated in the cyanobacterium Synechocystis sp. PCC 6803. Conversion ofcysteine 565 (this is the second cysteine in PsaB) to serine allowed assembly of PS I, although to a significantly reduced extent. Analysis ofthe cluster by low temperature EPR revealed both a mixed ligand [4Fe-4S] iron-sulfur center, capable of and a [3Fe-4S] cluster electron transfer to

Chapter 17 Photosystem I

(Smart et al., 1993). This provided evidence that C565 is a ligand to A CS(B556) PsaB mutant was also constructed in Synechocystis sp. PCC 6803 and found to accumulate a mixed ligand [4Fe-4S] cluster. In both the CS(B565) and CS(B556) mutants of Synechocystis sp. PCC 6803, forward electron transfer to centers and was decreased following a single

337

saturating flash (Vassiliev et al., 1996). Together, these results show that both cysteines in PsaB are involved in coordinating the [4Fe-4S] iron-sulfur center, and that is required for electron transfer from to

region Molecular modeling of the

predicted that the intercysteine amino acids of PsaA

Andrew N. Webber and Scott E. Bingham

338 and PsaB form two large loop structures that provide a docking site for PsaC (Rodday et al., 1993). When folded according to this model, positively charged arginine residues are located at the apex of both the PsaA and PsaB loops. Chemical modification of surface exposed arginine residues on spinach PS I with phenylglyoxal prevented the reconstitution of the PsaC and PsaD proteins with the modified PS I core complex in support of this model (Rodday et al., 1993). In Synechcystis sp. PCC 6803 R561 in the loop of PsaB was mutated to Glu. The RE(B561) mutant accumulated PS I at a significantly reduced level showing that the complex was destabilized (Rodday et al., 1994). The flash-induced absorption decayed with normal transient associated with kinetics. However, the mutant PS I complex was more susceptible to dissociation by urea suggesting the RE(B561) mutation impaired interaction between the PS I core and PsaC (Rodday et al., 1993). Low concentrations of divalent cations facilitated reconstitution of the mutant PS I core with subsaturating concentrations of PsaC (Rodday et al., 1993). A Chlamydomonas mutant, RE(B566), in which the equivalent arginine in PsaB was mutated, did not assemble PS I (Rodday et al., 1995). A proline to leucine mutation within the PsaB region, PL(B564), also increased the susceptibility of the PS I complex to dissociation by urea (Rodday et al., 1995). Together, these results indicate that the loop region is important for forward electron transfer between and the centers. Results from the Chlamydomonas PsaA mutant CD(A576), in which interaction between the PS I core and PsaC is impaired, suggests a similar role for the analogous loop region of PsaA (Hallahan et al., 1995). The stability of the PS I complex from the various proline mutants of PsaB in Chlamydomonas was investigated by measuring the extent and rate of dissociation of the PS I reaction center following treatment with urea and subsequent reconstitution of the urea-treated complex with PsaC expressed in E. coli (Rodday et al., 1995). The extent of dissociation and reconstitution was followed spectrophotometrically by measuring the recomand following a bination kinetics between brief activating flash. In wild-type, the for decay of the flash induced absorption change was approximately 30 ms, which is diagnostic for charge and The two PsaB recombination between proline mutants, PA(B560) and PL(B560), both disfor recombination of 30 ms indicating played a

complete electron transfer to the acceptors on PsaC (Rodday et al., 1995). The flash transient from PL(B560) showed a minor contribution of a faster ms phase indicating some disruption in electron transfer between and (Rodday et al., 1995). Following dissociation of the reaction center with urea the PA(B560) and PL(B560) mutants reconstituted poorly with PsaC indicating that the interaction with PsaC was impaired (Rodday et al., 1995). Similar results were observed following mutagenesis of proline residues in ferredoxin of Clostridium pasterianium—the redox properties of the iron-sulfur centers were not affected but the centers were significantly destabilized at increased temperatures (Gillard et al, 1993; Quinkal et al., 1994). NMR studies of the mutants showed that destabilization was due to changes in hydrogen bonding within the cluster (Gillard et al, 1993; Quinkal et al., 1994). It was suggested that the proline mutations in PsaB of PS I also led to altered domain causing hydrogen bonding in the decreased stability (Rodday et al., 1995). How the mutant PS I complexes assemble in vivo remains an interesting question. It is very likely that additional factors are present in the cell that aid in PS I assembly.

b.

Binding Domain

The reaction center electron transfer cofactors are surrounded by a central core of 10 membrane and spanning helices, contributed by PsaA and PsaB. The helices closest to the Chl dimer constituting are and Helices and correspond to what was previously named helix X of PsaA and PsaB predicted on the basis of hydrophobicity profile analysis (Fig. 9). Analysis ofthe derived amino acid sequence of PsaA or PsaB from Chlamydomonas shows that several residues within the stretch of amino acids that constitute helix m or m´ are highly conserved, even in reaction center proteins from H. mobilis and C. limicola. Histidine 656 of PsaB and H676 of PsaA are analogous conserved residues that are most The likely to provide the axial ligands to combined evidence from X-ray structure analysis and the study of site-directed mutants in Chlamydomonas, discussed below, together with recent spectroscopic results that indicate the axial ligand to is an unidentified His residue (Mac et al., 1996), all support such an assignment. Furthermore, mutation of several other conserved histidines in

Chapter 17 Photosystem I helix previously suggested to coordinate did not lead to any altered properties of (Cui et al., 1995). The function of His (A676) and His (B656) as has been tested by sitepossible ligands to directed mutagenesis in Chlamydomonas. The mutants were constructed in several genetic backgrounds of Chlamydomonas lacking either PS (strain II (the FuD7 strain) or PS II and Chl CC2696), to assist with biochemical and biophysical characterization of the mutant PS I complexes. In the PS II lacking strain FuD7, H(B656) has been mutated to many amino acids with a wide range of physical properties (Webber et al., 1996; H. Lee and A. Webber, unpublished). In general, substitution of H(B656) with amino acids of a similar or smaller physical size, that can still potentially ligate the central Mg of Chl such as Asn, Ser, Cys, and Tyr, still allows accumulation of significant although reduced amounts of PS I. Substitution with residues that cannot ligate the central Mg, such as Phe and Leu, results in a much reduced level of PS I (< 10 %). Substitution of H(B656) with Gly, not expected to provide a fifth ligand to Mg, resulted in significant accumulation of PS I. In the reaction center of Rhodobacter sphaeroides, substitution of either of the axial ligands to the primary donor, H(L173) or H(M202), with Gly allowed accumulation ofa BChl dimer rather than the expected BPhe/BChl heterodimer (Goldsmith et al, 1996). It was suggested that water is able to slip into a cavity generated when His is replaced by a small amino acid such as Gly (Goldsmith et al, 1996), and that this water molecule provides the fifth ligand to the Mg. A similar situation may also occur in the Gly substitute mutants in PS I. Mutation of H(A676) to the same set of amino acids described above leads to a similar altered pattern of PS I accumulation (H. Lee, unpublished). The biophysical properties of the HN(B656) and HS(B656) mutants of Chlamydomonas have been studied in some detail. Titration of the recombination kinetics with ferri- and ferro-cyanide showed a 40 mV increase in the redox midpoint (Webber et al., 1996). Analysis potential of of the electron spin distribution in by spectroscopy showed that the mutation causes a very specific increase in the hyperfine coupling associated with the methyl protons of the C12 methyl group (Fig. 6). Together with the increased midpoint potential of this data supports the hypothesis that His 656 of the PsaB subunit from Chlamy-

339 domonas closely interacts with the PS I primary donor. An interesting observation is that the mutation of H656 resulted in a very localized change in electron and that spin distribution in only one Chl a of this is the Chl carrying the majority of the charge. This could suggest that the Chl carrying the majority of the charge is located on the PsaB side of the dimer. Recently we have found that identical mutations in the PsaA subunit do not change the spectra (H. Lee, L. Krabben, W. Lubitz and A. Webber, unpublished). This observation is expected if the majority of the electron spin density is located on the B-side. In purple bacteria, the majority of electron transfer occurs along the L-branch. Interestingly, the is also asymmetrically spin distribution in distributed towards the L-side. The question of whether electron transfer occurs preferentially down the A- or B-side branch of cofactors, or equally down both branches, is a difficult question to address in PS I because of the overlapping optical signals from the donor and acceptor pigments. The absorption difference spectra from PS I preparations of HN(B656), HS(B656) and HQ(B656) showed a new bleaching band at approx. 670 nm both at femtosecond and millisecond time resolution (Webber et al., 1996; Melkozernov et al., 1997; Redding et al., 1998). The origin of this new is unclear. absorption decrease on oxidation of One possibility is that the mutation alters the relative orientation of one of the Chl molecules of changing the exciton interaction between the dimer pair. It cannot be ruled out at this stage that the with a neighboring mutations alter interaction of Chl molecule that is not part of the dimer, possibly A or A´. The decay of excitation energy in PS I can be monitored by picosecond fluorescence and absorbance spectroscopy techniques. The rate of excitation energy trapping in PS I was shown to decrease from 30 ps in wild type to 60 ps in the HN(B656) (Melkozernov et al., 1997). In wild-type, this rate of excitation energy decay has been used to estimate an and initial rate of charge separation between of 1.5 ps. In the HN(B656) the excitation energy decay was doubled and may reflect the decreased rate of forward electron transport. The increased redox would be expected to lower the potential of driving force for electron transfer from to resulting in an estimated charge separation rate of 3.5 ps. This in turn would result in an increase in the rate of excitation energy dissipation from the antenna.

340

These results support a trap-limited model for excitation energy trapping discussed previously in Section IV. Using picosecond fluorescence spectroscopy it was possible to study excitation energy trapping in thylakoid membranes from mutants that accumulate very low levels of PS I. In the HF(B656) mutant the rapid decay of fluorescence associated

Andrew N. Webber and Scott E. Bingham

with trapping by the reaction center is slowed to 70 ps. This would indicate that the mid-point potential in this mutant is raised even further than for HN(B656) which is consistent with the conversion of one of the Chl a to a pheophytin (Melkozernov et al., 1998). A model for the folding of helices j-o of PsaB is presented in Fig. 10 based on data discussed above,

Chapter 17 Photosystem I together with the information from the crystal structure. The axial ligand to has been identified as a His residue based on electron spin echo envelope modulation (ESEEM) spectroscopy of His labeled PS I from Synechocystis sp. PCC6803 (Mac et al., 1996). The mutant studies demonstrate that the most Chls likely histidines to act as axial ligands to the are His 656 of PsaB and His 676 of PsaA. The folding of the polypeptide is very similar to that of the L and M subunits of the purple bacterial reaction center (Fig. 10). As well as a similar folding, the same protein domains in both RC complexes coordinate the special pair. Whether this similarity holds for is discussed additional PS I cofactors such as below but remains to be tested experimentally.

c.

Binding Domain

At the current 4 Å resolution of the PS I crystal structure the location of the molecules cannot be determined. With the assignment of His (B656) as a it became clear that the ligand to one Chl a of folding of the reaction center binding region of PsaB (and PsaA) resembles quite strongly the bacterial reaction center from purple bacteria (Fig. 10). In the purple bacterial reaction center, the quinone binding domain is located between helices D and E, analogous to membrane spanning regions m and o of PsaA/B. Based on this analogy, it might be expected that the ammo acid sequences located between the m and o membrane spans may form the phylloquinone binding pocket (Figs. 9 and 10). It has been suggested that the unusual electronic structure and midpoint potential of is induced by the protein environment, possibly stacking interthrough hydrogen-bond and/or actions (van der Est et al., 1995, 1997; Rigby et al., 1996). A comparison of the amino acids in this region shows that several residues are highly conserved between PsaA/B from evolutionary diverse organisms and perhaps the bacterial reaction center protein (Fig. 9). Trp (B682) and Trp (A702) may be interactions with the napthoquinone involved in head group, and His (B684) and His (A704) may Hbond with the carbonyls. These residues predicted to provide the quinone binding region are highly conserved in the H. mobilis reaction center protein but not in C. limicola.

2. PsaC PsaC is a 9 kDa polypeptide that coordinates the

341 [4Fe-4S] iron-sulfur centers and one of which serves as the immediate acceptor of electrons from (Dunn and Gray, 1988; Golbeck and Bryant, 1991). The protein contains two cysteine rich motifs with the sequence CxxCxxCxxxCP which, in bacterial Fds, is involved in coordinating the two [4Fe-4S] iron-sulfur centers. The psaC gene has been inactivated by insertional mutagenesis in Anabaena variabilis (Mannan et al., 1991) and Chlamydomonas (Takahashi et al., 1991). In PsaC-less Chlamydomonas the PS I complex does not assemble to yield a stable complex and the subunits are rapidly degraded. In PsaC-less cyanobacteria the PS I complex accumulates, although to a reduced level. Thus, in both prokaryotes and eukaryotes PsaC is important for the normal accumulation and stability of the PS I complex. core complexes with Reconstitution of PsaC and PsaD, generated by expression of the cyanobacterial psaC and psaD genes in E. coli (Zhao et al., 1990), has provided a valuable tool for studies ofmutant PsaC polypeptides. Site-directed mutations were made in the psaC gene that changed either cysteine 14 or cysteine 51 to aspartate (Zhao et al., 1992). The mutated psaC genes were expressed in E. coli and the resulting protein used to reconstitute the core complex. In the C14D mutant, EPR analysis of the reconstituted complex revealed the presence of a [3Fe-4S] cluster and a [4Fe-4S] cluster that had a characteristic of (Zhao et al., 1990). In the C51D mutant the unmodified cluster characteristic of (Zhao et al., 1990). had a By analogy to the coordination of the iron-sulfur centers in bacterial ferredoxins it was suggested that is ligated by Cys 21, 48, 51 and 54 and that is ligated by Cys 11, 14, 17 and 58 (Zhao etal., 1990).

VI. Biogenesis of Photosystem I PS I is a multimeric subunit complex encoded by genes located in the chloroplast and the nucleus (Table 2). The chloroplast genome contains thepsaA, psaB, psaC, psaI and psaJ genes. In Chlamydomonas the psaA and psaB genes are spatially separated and are not part of a transcriptional unit as commonly found in plant chloroplasts and cyanobacteria (Kück et al., 1987). The psaA gene is further split into three spatially separated exons (Kuck et al., 1987). Following independent transcription of each exon, the three mRNAs are spliced in trans to form the

342

mature psaA mRNA (Choquet et al 1988). At least one chloroplast locus, tscA, has been shown to be involved in this trans-splicing event (GoldschmidtClermont et al., 1991; Chapter 8, GoldschmidtClermont). The remaining genes encoding PS I are located in the nucleus. The nucleotide and deduced amino acid sequences for PsaD, E, F, G and H have been derived from analysis of cDNA clones (Franzen et al., 1989a,b). A genomic clone for PsaF from Chlamydomonas has been characterized (Farah et al., 1996). Biogenesis of PS I in Chlamydomonas therefore requires the coordinated expression of genes located in different cellular compartments (Rochaix, 1992). In addition a large number of cofactors, including chlorophylls, carotenoids, quinones and iron-sulfur clusters need to be incorporated into PS I subunits for assembly of a functional complex. A number of genetic and biochemical approaches are beginning to provide a limited understanding of some steps of PS I biogenesis. However, the information in the literature is quite scant and much more work needs to be done in this important area. Nucleus-encoded proteins are first synthesized in the cytosol with an N-terminal transit sequence. The transit sequence serves to target the polypeptide into

Andrew N. Webber and Scott E. Bingham

the chloroplast where it is cleaved by a processing protease to produce the mature protein. The mature protein then assembles into the PS I complex. The exact sequence of the later two events is unclear in most cases and may vary for different polypeptides. In the case of lumenal polypeptides such as PsaF and PsaN (Knoetzel and Simpson, 1993), the polypeptide is encoded with a bipartite transit sequence. Following removal of the chloroplast targeting sequence, the thylakoid transfer sequence functions to target the protein to the thylakoid lumen. In the case of PsaN it is unclear if the chloroplast transit sequence is removed before transfer across the thylakoid membrane because an intermediate processed form has not been detected (Nielsen et al., 1994). PsaD was shown to integrate into the thylakoid membrane as a precursor form that could only be processed by the stromal processing protease after assembly into the PS I complex (Cohen et al., 1992; Cohen and Nechushthai, 1992; Cohen et al., 1993). Light has been shown to play an important role in signaling the initiation of transcription and/or translation for nuclear and chloroplast proteins, especially during development of chloroplasts from etioplasts. The involvement of phytochrome, blue

Chapter 17 Photosystem I light and blue-UVA light receptors in expression of PS I genes was demonstrated in wild-type tomato and in phytochrome-deficient mutants (Oelmuller et al., 1989; Oelmuller and Kendrick, 1991). Exposure of etiolated seedlings with red light was shown to induce mRNA accumulation from psaD but no protein accumulation was detected until exposure to several hours of white light (Lotan et al., 1993). In etiolated leaves, chloroplast mRNAs encoding many thylakoid proteins were present (Klein and Mullet, 1986; 1987; Kreuz et al, 1986). However, the polypeptides, particularly those involved in Chl binding, such as PsaA and PsaB in PS I, did not accumulate until the leaves were exposed to light (Klein and Mullet, 1986; 1987; Kreuz et al, 1986). In the y–l mutant of Chlamydomonas, that is unable to synthesize Chl without light, Chl-binding proteins are also absent in the dark (Malnoe et al., 1988; Herrin et al., 1992). The importance of Chl synthesis in PsaA and PsaB apoprotein accumulation was directly tested in etioplasts where it was demonstrated that addition of Chl precursors, which induce Chl synthesis in the dark, resulted in accumulation of PsaA and PsaB (Eichacker, et al., 1990). It was suggested that binding of Chl a to the apoprotein facilitates translation and/ or release of the protein from the ribosome complex. However, these assays could be limited by the inability to detect very rapid degradation of newly translated intermediates. In an apparently contradictory study, Kim et al. (1994) showed that the abundance of translation initiation complexes, and the extent of translational run-off in the presence of lincomycin, was unaffected by Chl availability suggesting that Chl binding to newly synthesized apoproteins leads rapidly to formation of a stable protein complex. There is some limited evidence for coordinated translation of PsaA and PsaB polypeptides in Chlamydomonas. The FUD26 mutant with a frameshift in psaB did not synthesize a full length PsaB polypeptide, nor did it synthesize PsaA (GirardBascou et al., 1987). However, mutants unable to synthesize PsaA were found to synthesize PsaB (Girard-Bascoue et al., 1987; Takahashi et al., 1991), although the PsaB polypeptide was subsequently rapidly degraded. Many directed mutations in PS I reaction center proteins in both cyanobacteria and Chlamydomonas lead to reduced accumulation of the complex to some extent. This could be due to impaired expression of genes or decreased stability of the reaction center complex. In the case of one such mutant of the PsaB

343 subunit, HL(B523), rapid pulse-chase-labeling of proteins in Chlamydomonas has been used to address the question of whether the mutant PsaB protein was synthesized to the same extent as wild-type, and if the synthesized proteins assembled into a reaction center complex. In the HL(B523) mutant it was shown that normal levels of radiolabeled PsaA and PsaB polypeptides could be detected following SDSPAGE, indicating that translation ofthe proteins was unaffected by the mutation (Cui et al., 1995). However, when the same thylakoid membranes were analyzed on LDS-PAGE ‘green’ gels, it was found that a reduced level of labeled proteins accumulated in the green band corresponding to the PS I reaction center complex (Cui et al., 1995). This showed that the HL(B523) mutation impaired the ability of the fulllength polypeptide to assemble into the reaction center complex. Two chloroplast open reading frames, ycf3 and ycf4, are required for stable accumulation of the PS I complex in Chlamydomonas (Boudreau et al., 1997). Both the ycf3 and ycf4 gene products are located in the thylakoid membrane although neither co-isolate with PS I. Deletion of either ycf3 or ycf4 leads to a complete loss of PS I. The exact roles of the ycf3 and ycf4 products are unknown, but they are presumably involved in the synthesis, assembly or stability of PS I. Nuclear genes are required for the synthesis and assembly of PS I and expression of chloroplast genes of PS I subunits. Girard et al. (1980) isolated 25 nuclear mutants that belonged to 13 different complementation groups. Each mutant lacked the PsaA and PsaB polypeptides as well as six low molecular weight polypeptides which we now know to be identical to the PS I associated polypeptides discussed previously (Section V.A.). It is likely that some mutants isolated were impaired in the expression of these nuclear encoded PS I polypeptides. However, other loci may represent genes involved in stability of mRNAs for chloroplast encoded PS I polypeptides or in translation of these mRNAs. Nuclear mutants have been identified that lack mRNA for psaA, psaB and psaC genes (K. Breuschweiler, S. Bingham and A. Webber, unpublished) and trans-splicing of psaA mRNAs (Choquet et al., 1988). Stampacchia et al. (1997) recently characterized a PS I deficient strain of C. reinhardtii, F15, that contained a mutation, called tab1-F 15, that impairs initiation of translation of psaB mRNA. Translation of both psaB and psaA mRNAs are impaired in this mutant similar to earlier

344 observations of Girard-Bascou et al. (1987). A suppressor mutation localized to the 5´ UTR of psaB mRNA restored translation of both polypeptides suggesting that PsaA translation is dependent on translation of PsaB. Mutations in five chloroplast loci have also led to a PS I deficient phenotype. One chloroplast locus has been identified as the psaB gene based upon DNA sequence analysis of a mutated gene from two independent mutants (Girard-Bascou, 1987; Bingham, et al., 1991). Another locus has been identified as the site of transcription of a small RNA, tscA, that is involved in trans-splicing of psaA mRNA (Goldschmidt-Clermont et al., 1991). The other loci have not been identified but presumably represent the sites of other chloroplast PS I genes.

Acknowledgments Work from the authors’ laboratories was supported by grants from National Research Initiatives Competitive Grants Program of the USDA.

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Andrew N. Webber and Scott E. Bingham polypeptides: Identification of PsaB domains that may interact with PsaD. Plant Physiol 108: 1067–1072 Xu Q, Guikema JA, and Chitnis PR (1994a) Identification of surface-exposed domains on the reducing side of Photosystem I Plant Physiol 106: 617–624 Xu Q, Trent S, Guikema JA and Chitnis PR (1994b) Organization of Photosystem I polypeptides: A structural interaction between the PsaD and PsaL subunits. Plant Physiol 106: 1057–1063 Xu Q, Yu L, Chitnis VP and Chitnis PR (1994c) Function and organization of Photosystem I in a cyanobacterial mutant that lacks PsaF and PsaJ subunits. J Biol Chem 269: 3205–3211 Xu Q, Odom WR, Guikema JA, Chitnis VP and Chitnis PR (1994d) Targeted deletion of psaJ from the cyanobacterium Synechocystis sp. PCC 6803 indicates structural interactions between PsaJ and PsaF subunits of Photosystem I. Plant Mol Biol 26: 291–302 Xu Q, Jung Y-S, Chitnis VP, Guikema JA, Golbeck JH and Chitnis, PR (1994e) Mutational analysis of Photosystem I polypeptides in Synechocystis sp. PCC 6803: Subunit requirements for reduction of mediated by ferredoxin and flavodoxin. J Biol Chem 269: 21512–21518 Xu Q, Hoppe D, Odom WR, Guikema JA, Chitnis VP and Chitnis PR (1995) Mutational analysis of the Photosystem I polypeptides in the cyanobacterium Synechocystis sp. PCC 6803. Targeted inactivation of psal reveals the function of Psal in the structural organization of PsaL. J Biol Chem 270: 1–8 Xu R, Bingham SE and Webber AN (1993) Increased mRNA accumulation in a psaB frame- shift mutant of Chlamydomonas reinhardtii suggests a role for translation in psaB mRNA stability. Plant Mol Biol 22: 465–474 Yu L, Zhao J, Mühlenhoff U, Bryant DA and Golbeck JH (1993) PsaE is required for in vivo cyclic electron flow around Photosystem I in the cyanobacterium Synechococcus sp. PCC 7002. Plant Physiol 103, 171–180 Yu J, Smart LB, Jung Y-S, Golbeck J and McIntosh L (1995) Absence of PsaC subunit allows assembly of Photosystem I core but prevents the binding of PsaD and PsaE in Synechocystis sp. PCC6803. Plant Mol Biol 29: 331–342 Zanetti G and Merati G (1987) Interaction between Photosystem I and ferredoxin. Identification by chemical crosslinking of the polypeptide which binds ferredoxin. Eur J Biochem 169: 143–146 Zhao J, Warren PV, Li N, Bryant DA and Golbeck JH (1990) Reconstitution of electron transport in Photosystem I with PsaC and PsaD proteins expressed in Escherichia coli. FEBS Lett 267: 175–180 Zhao J, Li N, Warren PV, Golbeck JH and Bryant DA (1992) Site-directed conversion of a cysteine to aspartate leads to the assembly of a [3Fe-4S] cluster in PsaC of Photosystem I. The photoreduction of FA is independent of FB. Biochemistry 31: 5093–5099 Zhao J, Snyder WB,Mühlenhoff U, Rhiel E, Warren PV, Golbeck JH and Bryant DA (1993) Cloning and characterization of the psaE gene of the cyanobacterium Synechoccocus sp. PCC 7002: Characterization of a psaE mutant and overproduction of the protein in Escherichia coli. Mol Microbioi 9: 183–194 Zilber AL and Malkin R (1988) Ferredoxin crosslinks to a 22 kDa subunit of Photosystem I. Plant Physiol 88: 810–814

Chapter 18 Reexamining the Validity of the Z-Scheme: Is Photosystem I Required for Oxygenic Photosynthesis in Chlamydomonas? Kevin Redding

University of Geneva, Departments of Molecular Biology and Plant Biology,

30, quai Ernest-Ansermet, CH1211 Geneva 4, Switzerland

Gilles Peltier

CEA/Cadarache - DSV- DEVM - Laboratoire d’Ecophysiologie de la Photosynthèse,

Bâtiment 161, 13108 Saint Paul-lez-Durance, France

Summary I. The Z-Scheme of Oxygenic Photosynthesis and Alternative Schemes II. Electron Transport in the Absence of PS II III. Photosynthesis in the Absence of PS I IV. Putative Electron Transport Pathways Outside of the Z-Scheme V. Thermodynamic Considerations VI. Evolutionary Considerations VII. Conclusions Acknowledgments References

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Summary The ‘Z-scheme’ is the prevailing model for photosynthetic electron transport, in which electrons from Photosystem II (PS II) are transferred to Photosystem I (PS I) with the participation of the plastoquinone pool, complex, and plastocyanin. PS II can accomplish water oxidation due to the low potential the cytochrome of its oxidized primary donor, and PS I reduces ferredoxin by virtue of its low potential iron-sulfur centers. There have been several claims over the last two decades that PS II can carry out the reduction of ferredoxin, lately using Chlamydomonas reinhardtii mutants defective in synthesis of PS I (Greenbaum et al., 1995; Lee et al., 1996). However, recent studies performed on mutants harboring deletions of genes encoding PS I subunits demonstrated that photoautotrophic growth and fixation require the presence of PS I (Cournac et al., 1997). When observed in ‘PS I-deficient’ mutants, photoautotrophic growth and fixation are most likely attributable to small amounts of PS I in those mutants. We discuss thermodynamic and evolutionary implications of the Z-scheme and possible exceptions to it.

J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 349–362. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

350 I.The Z-Scheme of Oxygenic Photosynthesis and Alternative Schemes The discovery of two different light reactions in oxygenic photosynthesis was initially based on studies of the enhancement effect in algae by Emerson and co-workers, observed by combining two beams of different wavelengths (Emerson, 1958; Myers, 1971). After the discovery of cytochromes b (Cyt b) and f (Cyt f) as components of the photosynthetic machinery, Hill and Bendall (1960) proposed that the two light reactions were operating in series to achieve linear electron transfer reactions from with Cyts b and f serving as the to intermediates between the photosystems. The essence of this model, referred to as the ‘Z-scheme’, is that Photosystem II (PS II) accomplishes the oxidation of water and the reduction of plastoquinone (PQ) and cytochromes, but is unable to reduce ferredoxin (Fd) . According to this scheme, Photosystem I or and the (PS I) is required for the reduction of oxidation of Cyt f through plastocyanin (PC). Although other models could be proposed to explain the enhancement effect and the cooperativity between photosystems (Arnon, 1995), the Z-scheme of photosynthesis was supported by experimental data, such as the light-induced redox changes of Cyt f (Duysens et al., 1961). The Z-scheme is now considered as the central dogma of oxygenic photosynthesis (see Fig. 1A; Trebst, 1974; Stryer, 1995). Since the formulation of the Z-scheme, various facts have come to light that do not fit easily into this model. It was found that PS II is mainly localized in appressed (granal) membranes, while PS I is mainly located in nonappressed (stromal) membranes (for a review see Staehelin and De Witt, 1984). This surprising finding questioned the validity of the Zscheme, as it became clear that the physical arrangement ofelectron carriers within the membrane is different from that supposed by the Z-scheme. Abbreviations: CCCP – carbonyl cyanide-chlorophenylhydrazone; Cyt–cytochrome; DCMU–3-(3,4-dichlorophenyl)1,1-dimethylurea; DBMIB – 2,5-dibromo-3-methyl-6-isopropylp-benzoquinone; FCCP – carbonyl cyanide-p-trifluoromethoxyphenylhydrazone; Fd – ferredoxin; FNR–ferredoxin: reductase; OEC – oxygen evolving complex; PC – plastocyanin; PQ – plastoquinone; PS I – Photosystem I; PS II – Photosystem II;

RC – reaction center; RC1 – reaction center of type 1 (quinone); RCII – reaction center of type 2 (Fe-S); SQR – sulflde:quinone oxidoreductase

Kevin Redding and Gilles Peltier According to the supramolecular organization of photosynthetic complexes, the operation of the Zscheme would necessarily involve the existence of at least one mobile electron carrier to carry electrons from grana membranes to stroma lamellae. The nature of this carrier (PQ or PC) has been a subject of controversy (Haehnel, 1984; Lavergne and Joliot, 1991). Initially, PS II was considered as a strong oxidant capable of oxidizing water and only a poor reductant, The discovery of unable to reduce Fd and pheophytin as the first electron acceptor of PS II (Klimov et al., 1977; Klimov and Krasnovsky, 1982) provoked the question of whether PS II was able to carry out reduction of and/or and several authors suggested that such a reaction was possible (see Fig. 1B; Albertsson et al., 1983; Klimov et al., 1986; Arnon and Barber, 1990). However, although PS I-dependent reactions can be studied easily in the absence of PS II contribution due to the existence of potent and specific inhibitors of PS II such as DCMU, and also because ofthe possibility of independently exciting PS I using far-red illumination, the study of PS II-dependent reactions in the absence of PS I contribution is a much more difficult task. Indeed, no specific inhibitor of PS I is known and it is not possible to specifically excite PS II using particular wavelengths. By studying Fd reduction in unfractionated thylakoid membrane preparations, Arnon et al. (1980a,b) proposed the existence of two different pathways of Fd reduction, one of these pathways being able to reduce Fd without concomitant reduction of the iron-sulfur centers of PS I. This pathway was proposed to involve PS II without significant participation of PS I (see Fig. 1B for one possible scheme). Later, Albertsson et al. (1983) used inside-out vesicles, originating from appressed thylakoids greatly enriched in PS II, to investigate the ability of PS II to reduce These authors observed that such preparations were able to reduce as electron donor. Surprisingly, using this reaction required PC and was inhibited by an (DBMIB), strongly suggesting inhibitor of Cyt the involvement of residual PS I present in the preparation. Arnon (1995) proposed that PC was not only a donor to PS I but also a donor to PS II. Finally, Arnon and Barber (1990) presented evidence that isolated PS II reaction centers (RCs), stripped of their oxygen evolving complexes (OECs), quinones,

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and antenna chlorophyll proteins, can reduce in a reaction that required 2,5-diphenylcarbazide and PC (presumably as donors), Fd (presumably as reductase (FNR). acceptor), and ferredoxin: The work of Arnon and co-workers, which has not been confirmed by other laboratories, has been criticized for several reasons. The results related to the reduction of Fd without concomitant reduction centers (Arnon et al, 1980a,b) is most of the probably explained by the fact that electrons reside only temporarily on and are rapidly ( in the fastest cases; Sétif and Bottin, 1994; Sétif and Bottin, 1995; Fischer et al., 1997) transferred to Fd, which can accumulate in the reduced state. On the other hand, one can imagine that the data of Arnon and Barber (1990) describe a direct transfer of electrons to Fd from reduced pheophytin in PS II particles from which the quinone terminal electron acceptors have been removed. However, on structural grounds, it is difficult to imagine how Fd docks to PS II in such a way as to accept electrons from pheophytin, given the position of the analogous bacteriopheophytin of the purple bacterial RC in the middle of the membrane bilayer (Deisenhofer and Michel, 1989). In any case, the physiological relevance of such a reaction is questionable. The most important criticism is the possibility ofresidual amounts of PS I in the preparations used and the requirement for PC, a well-known PS I electron donor. A demonstration of Fd reduction by PS II can only be convincing using a well-characterized experimental system completely devoid of PS I activity. In this regard, the availability ofgenetic mutants is a great advantage for microalgal systems. Since C. reinhardtii mutants deficient in photosynthesis can grow heterotrophically using acetate as a carbon source and still develop green chloroplasts containing thylakoid membranes (Harris, 1989), they have been widely used in the past to study photosynthesis (Levine, 1968; Togasaki and Whitmarsh, 1986). Several mutants of C. reinhardtii that specifically lack PS I, PS II, Cyt -ATPase, Rubisco, etc. have been described (Harris, 1989). Active PS I or PS II can be assembled in mutants that fail to synthesize the other photosystem (Givan and Levine, 1967; Bennoun and Levine, 1967), thus making possible the analysis of electron transport occurring in the absence of one of the photosystems.

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II. Electron Transport in the Absence of PS II It is worth beginning with a discussion of the photosynthetic activities possible in the absence of PS II. Probably the best-known example is cyclic electron transport. The term ‘cyclic’ denotes the fact that electrons received by Fd are used to re-reduce As PS II is not used, there is no net source of electrons from water-splitting. However, as the product of PS I (reduced Fd) has a lower redox potential than that of PS II (plastoquinol), the first can be used to generate the second. The protons pumped across the thylakoid membrane by action of complex, as it reduces PC using electrons the Cyt from plastoquinol, are used to make ATP (Boyer, 1993). Thus, it has been postulated that the partitioning of electrons between cyclic and linear pathways can be used by the cell to fine-tune the NADPH/ATP ratio (see Bendall and Manasse, 1995 and references therein). An in vivo analysis in C. reinhardtii using photoacoustics confirmed that energy storage by cyclic electron flow was not inhibited by DCMU (a specific inhibitor of PS II), that it could be driven by far-red light (>715 nm), and that it took place in a PS II-less mutant (Ravenel et al., 1994). Results with other inhibitors were consistent with earlier in vitro results that demonin strated an absolute requirement for Fd and Cyt cyclic electron transport (Tagawa et al., 1963; Hosier and Yocum, 1985; Bendall and Manasse, 1995). Interestingly, Chlamydomonas appears to have two ways to transfer electrons from Fd to the PQ pool (see Fig. 1B), either one of which is able to fully accommodate cyclic electron flow in the absence of the other. Based on inhibitor studies, Ravenel et al. (1994) suggested that one involves an antimycin Asensitive Fd:PQ oxidoreductase activity (cycle 1), while the other appears to involve FNR-catalyzed and then to the PQ transfer of electrons to pool (cycle 2). This latter step might utilize the elusive NAD(P)H dehydrogenase of the thylakoid membrane (discussed below). Under normal conditions, cycle 2 would be responsible for the majority of the electron flow. However, cycle 1 might be called upon when the capacity to recycle electrons via NADPH becomes saturated and electrons accumulate at the level ofFd. Such conditions might occur during photosynthetic induction conditions, where the ratio becomes very high due to the initially inactive state of the Calvin cycle enzymes (Slovacek et al., 1980); cycle 1 is highly stimulated in isolated spinach

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354 thylakoids by a high ratio (Hosler and Yocum, 1987). without C. reinhardtii is capable of fixing PS II under certain conditions. Like many other green algae, upon a shift to anaerobic conditions Chlamydomonas induces the enzyme hydrogenase, which catalyzes the reaction:

The electron acceptor/donor is Fd, which has a redox (Happe and Naber, potential close to that of 1993). The use of as an electron donor in algae was first discovered by Gaffron (1939), who observed with the concomitant uptake of the fixation of in anaerobic conditions and termed the process ‘photoreduction.’ Maione and Gibbs (1986) found that DCMU does not inhibit photoreduction in isolated C. reinhardtii chloroplasts, consistent with hydrogenase supplanting PS II as the source of electrons. However, photoreduction is inhibited by DBMIB, indicating a role for Cyt (Maione and Gibbs, 1986), and the uncoupler FCCP, which can be at least partially reversed by the addition of ATP (to chloroplasts). Taken together, these results suggest that the role of PS I is to produce ATP by cyclic transport, while NADPH is generated by hydrogenase/ Fd/FNR, and fixation occurs by the usual reductive pentose phosphate cycle (Russel and Gibbs, 1968). When Chlamydomonas cells are subjected to conditions where electrons from the light reactions fixation, reduced Fd can are not consumed by serve as a donor to hydrogenase with the concomitant (Gaffron and Rubin, 1942; Stuart generation of and Gaffron, 1972a,b). Using a continuous gas flow in system, it was found that the ratio of evolved conditions was roughly 2:1, as anaerobic one would expect, although deviations from this were observed at high light intensities or using farred light which excites PS I preferentially (Greenbaum, 1984). Although electrons originating from PS II is PS II can contribute to the production of not required for this process (Stuart and Gaffron, 1972a,b; Gfeller and Gibbs, 1984). That electrons consumed by evolution can originate from organic reductants, such as starch, is consistent with the observation that fermentation in the dark can drive (Healey, 1970; Gfeller and the production of Gibbs, 1984; Graves et al., 1989).

Kevin Redding and Gilles Peltier III. Photosynthesis in the Absence of PS I A large collection of C. reinhardtii mutants deficient in PS I has been described in the literature (Girard et al., 1980; Goldschmidt-Clermont et al., 1990). The psaA gene is split into three exons that are widely separated and located on different strands of the C. reinhardtii chloroplast genome (Kück et al., 1987). These exons are separately transcribed and then joined together in a process known as ‘trans-splicing’ (Choquet et al., 1988; also see Chapter 11, Herrin et al.), a process which requires many of the genes identified as necessary for PS I biosynthesis (Goldschmidt-Clermont et al., 1990). Some of the nuclear mutants have been shown to contain residual amounts of PS I. For instance, Girard-Bascou et al. (1980) found among their collection of 25 mutants, three leaky mutants that contained from 5 to 20% of wild-type PS I activity. Although PS I-deficient mutantshave been reported to be unable to grow photosynthetically and to require acetate for growth (Harris, 1989), it is possible that this property was due to biases imposed by the genetic screens that were employed. In other words, if one screens for acetate-requiring mutants and isolates PS I-deficient mutants, then one assumes that the loss of phototrophy is due to the loss of PS I. A reverse genetics approach could remove this bias, as one could attempt to eliminate PS I without any expectation as to the phenotypic consequences of the mutation. The psaA, psaB and psaC genes have been deleted in C. reinhardtii (Takahashi et al., 1994; Fischer et al., 1996), and the deletion mutants were nonphotosynthetic. In addition, a few PS I-deficient mutants that are impaired in chloroplast genes have been described as nonphotosynthetic (Girard-Bascou, 1987; Girard-Bascou et al., 1987). PS I subunit genes have also been deleted in two cyanobacterial species, with concomitant loss of photoautotrophic growth (Smart et al., 1991; Toelge et al., 1991). Besides the lack of photoautotrophic growth, it has been observed that PS I-deficient mutants of algae or cyanobacteria are exquisitely photosensitive, even on media containing a source of reduced carbon (Spreitzer and Mets, 1981; Vermaas et al., 1994). and Measurements of light-induced exchange by mass spectrometry in the F15 mutant was deficient in PS I led to the conclusion that produced at low rates by PS II during steady state illumination, but that fixation did not occur (Peltier and Thibault, 1988). In contrast, photo-

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Validity of the Z-Scheme

synthetic fixation and photoautotrophic growth were reported in two other PS I-deficient mutants (Greenbaum et al., 1995; Lee et al., 1996), both of which are nuclear mutants deficient in the transsplicing of the psaA mRNA (Girard et al., 1980; Goldschmidt-Clermont et al., 1990; Greenbaum et al., 1995). With the use of a gas flow apparatus, Greenbaum et al. (1995) were able to measure an coupled to fixation or to evolution of evolution (Fig. 2A). The amount of fixed by these mutants strains (B4 and F8) was roughly equivalent to that measured in wild-type cells. Initially, fixation in the PS I-deficient strains appeared to be limited to anaerobic conditions (Greenbaum et al., 1995), but photoautotrophic growth was later reported in the presence of (Lee et al., 1996). Several controls were carried out by the authors to eliminate the possibility of trace amounts of PS I. Irradiation with far red light which excites PS I but not PS II, produced a small amount of evolution in WT cells but not in the B4 mutant (Greenbaum et al., 1995). Photobleaching experiments also failed to detect P700 in thylakoid membranes from the mutants (Lee et al., 1996). More recently, C. reinhardtii strains deleted in the chloroplast genes psaA or psaB encoding the two main subunits of the PS I reaction center were found fixation or photoautotrophic to be incapable of growth (Fig. 2B,C), although limited steady state production by PS II could be measured by mass spectrometry (Cournac et al., 1997). The discrepancy between the ability of different PS I-deficient mutants to grow photoautotrophically was troubling. Two hypotheses could be envisaged to explain these results: either the B4 and F8 strains had small amounts of PS I (undetected by the authors) that allowed them and grow to synthesize enough NADPH to fix photoautotrophically, or the genetic background of the strains in which the gene deletions had been made was deficient in some way and did not allow the operation of a PS I-independent pathway. These alternate possibilities could be distinguished by a simple genetic test. If a gene for a PS I major subunit were disrupted in the B4 or F8 background, the first hypothesis predicts that photoautotrophic growth would cease, while the second predicts an unabated photoautotrophic growth. After deletion of fixation and photopsaA in the F8 mutant, autotrophic growth both ceased (Cournac et al., 1997; see Fig. 2B,C). The same phenotype was observed after deleting the psaC gene (K. Redding, L. Cournac,

355 and G. Peltier, unpublished), which encodes a small protein binding PS I electron acceptors and (Golbeck and Bryant, 1991). Unrelated genetic damage caused by transformation could not explain these results, as the reintroduction of psaA could restore slow photoautotrophic growth to Moreover, immunological analysis indicated that F8 expressed some PsaA polypeptide (ca. 10–15% the WT level), which disappeared upon disruption ofthe psaA gene (Cournac et al., 1997), and spectroscopic analysis confirmed that F8 harbored active PS I (I. Vassiliev, K. Redding, and J. Golbeck, unpublished). In the case of B4, photoautotrophic growth could not be repeated (Cournac et al., 1997). Instead, phenotypic revertants that expressed PsaA were obtained from B4 upon selection for photoautotrophic growth; such photoautotrophic variants could not be selected from strain (Cournac et al., 1997). Taken the together, this indicates that fixation in the B4 strain required the presence of PS I, at least in small amounts. Greenbaum and coworkers (Greenbaum et al., 1995; Lee et al., 1996) initially reported that evolution occurred in both F8 and B4 strains. However, as in the case of fixation,

photoevolution was undetectable in the

and

mutants as well as in F8 and B4 strains deleted in the psaA gene (L. Cournac, K. Redding, fixation and G. Peltier, unpublished). While requires both NADPH and ATP, production only requires reduced Fd and is thus a more sensitive test for the reduction of Fd in vivo. The absence of evolution in PS I-deficient strains is therefore a good indication that Fd is not reduced in the absence of PS I. As PS I has an inherently faster electron transfer from the primary donor to the terminal acceptor, a small amount of PS I is theoretically able to handle the electron flow from a relatively larger amount of PS II (Boichenko, 1996). However, under conditions where illumination saturates both photosystems, overall electron flow would be limited by the amount of PS I present, if it were much less abundant than PS II. In recent experiments, Greenbaum and evolution in a mutant coworkers have examined containing low levels of PS I and observed quantities of larger than would be predicted by Boichenko’s hypothesis (Greenbaum et al., 1997). Thus, although it is clear that the presence of PS I is required for photoautotrophic growth, fixation, and evolution, more work needs to be done in the case of mutants with low amounts of PS I to definitively

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decide if PS I-independent and PS I-dependent electron flows exist side-by-side in such mutants. If this turns out to be the case, then one must explain why such a PS I-independent flow does not occur in mutants without any PS I.

IV. Putative Electron Transport Pathways Outside of the Z-Scheme Although fixation is not observed in C. rein­ hardtii mutants deficient in PS I, it is true that some electron transport activity dependent solely upon PS II has been observed. By using a C. reinhardtii mutant deficient in PS I (ACC-1), Klimov et al. (1986) reported that addition to intact cells of benzyl viologen or methyl viologen (typical acceptors of PS I) induced an acceleration of the dark oxidation of reduced pheophytin. These authors concluded that pheophytin was able to directly reduce could enter although it is questionable that intact cells. Peltier and Thibault (1988) reported the existence of a continuous evolution of in the PS Ideficient F15 mutant. This production was completely blocked by DCMU, thus demonstrating the involvement of the PQ pool and arguing against a phenomenon involving direct reduction of by pheophytin. These results were recently confirmed in C. reinhardtii strains deleted in the psaA or in the

357 psaB gene, where it was found that the amount of evolved was exactly matched by an increase in uptake (Cournac et al., 1997). Based on the effect of inhibitors of the mitochondrial electron transport chain, these authors suggested that a transportable form of reducing power generated by PS II was transported towards mitochondria and subsequently oxidized by the respiratory electron transport chain (Chapter 35, Bennoun). Because oxygen evolution was inhibited by uncouplers such as CCCP (Peltier and Thibault, 1988), FCCP or crown (L. Cournac and G. Peltier, unpublished), it was suggested that NAD(P)H was synthesized by a proton gradientdependent reverse electron flow operating through a NAD(P)H dehydrogenase complex (Fig. 1C). Measurements of light-induced variations in bluedeletion green fluorescence in or mutants are also consistent with a PS II-dependent reduction of the pool (L. Cournac and G. Peltier, unpublished). However, we must mention that a convincing demonstration of a thylakoid NAD(P)H dehydrogenase has been problematic (discussed in Chapter 35, Bennoun). An alternate possibility is the use of fumarate which could be reduced to succinate by PQ using a chloroplastic succinate dehydrogenase (Willeford et al., 1989). If NAD(P)H is synthesized by a NAD(P)H dehydrogenase complex operating in a reverse mode

358 (Fig. 1C), the absence of fixation could be explained by two hypotheses. In the first one, the electrochemical gradient generated by water splitting would be used to generate NAD(P)H by reverse functioning of a NAD(P)H dehydrogenase complex and thus could not be used for ATP synthesis (see next section). In the second, fixation would be impossible even in the presence of NADPH and ATP, if the enzymes of the Calvin cycle were in an inactive state. Several of the key enzymes are activated by thioredoxin, which receives electrons from Fd (Buchanan, 1980; Hoober, 1984). Thus, the lack of significant amounts of reduced Fd in PS I-deficient mutants might explain their inability to fix

V.Thermodynamic Considerations The energy of a red light photon absorbed by chlorophyll is approximately 1800 meV However, if one considers a photosystem as a Carnot machine, the maximal power that it can deliver is (ca. 1240 meV; Lavergne and Joliot, about 0.69 1996). This would be in the range sufficient to just bridge the gap between oxidation of and reduction of on However, since the potential must be the donor side of PS II in order to oxidize water, the potential on the acceptor side has to be almost all of Therefore, in order to reduce the free energy deliverable by PS II would have to be consumed, leaving almost nothing for building up the electrochemical gradient used to synthesize ATP. Therefore, even if PS II appears theoretically capable of directly reducing ATP synthesis would not be possible, perhaps explaining why fixation is not observed in the absence of PS I (Cournac et al., 1997). In any event, experimental data obtained in PS I-deficient C. reinhardtii mutants argue against the existence of a direct reduction of by PS II, since electron transfer reactions in the absence of PS I are completely inhibited by DCMU (Peltier and Thibault, 1988), which blocks photosynthetic and (i.e. electron transfer reactions between beyond pheophytin).

VI. Evolutionary Considerations PS II and PS I are evolutionarily related to RCs occurring in extant photosynthetic bacteria, where a

Kevin Redding and Gilles Peltier single type of RC is used and no oxygen is evolved. The quinone-reducing type 2 reaction centers (RCII) found in Proteobacteria (purple bacteria) and Chloroflexus are structurally related to PS II, while the Fe/S center-containing type 1 RCs (RCI) of Heliobacteria and Chlorobium resemble PS I (Golbeck, 1993; Rutherford and Nitschke, 1996; Nitschke et al., 1996). In both types of bacteria, cyclic electron transport is used to create a proton gradient for ATP production (see Fig. 1D, E). Electrons exit the cycle in the form of NAD(P)H. Although in a straightforward RCI bacteria can reduce manner, RCII bacteria must make use of a NAD:quinone oxidoreductase with the help of the proton gradient to drive electrons from the relatively (see Fig. 1D). If high potential quinol to experimental data confirm that NAD(P)H synthesis linked to evolution can be driven in plants by PS II alone using a NADH dehydrogenase operating in the reverse mode (Fig. 1C), one might consider this reaction as a ‘living fossil’ from more primitive forms ofphotosynthesis. The existence oflimitations in inter-system electron transfer reactions (due either to the diffusion of mobile electron carriers and/or to electron transfer reactions through the Cyt complex) could explain the persistence ofa bacterialtype photosynthesis in which PS II achieves reduction by dissipating the electrochemical gradient through the NAD(P)H dehydrogenase complex. Thus, the ‘branchpoint’ between cyclic and linear electron transport is at the level of Fd and/or NADPH for RCI bacteria and at the level of the proton gradient for RCII bacteria (see Fig. 1D, E). Of course, for every electron that exits the cycle, one must enter it. Photosynthetic bacteria have found several ways to extract electrons from their environment. Sources include sulfide, sulfur, thiosulfate, sulfite (Brune, 1995), and ( Ehrenreich and Widdel, 1994). Although electrons could potentially enter the cycle at any point, in all known cases they do so at the level of one of the mobile electron carriers (i.e. quinone or cytochrome). We have invoked sulfide:quinone oxidoreductase (SQR) in the schemes of Fig. 1E and 1D, as this enzyme has been identified in both RCI (Shahak et al., 1992) and RCII (Schütz et al., 1997) bacteria, as well as a cyanobacteria (Arieli et al., 1994). In addition, some bacteria probably use flavocytochrome c to catalyze electron transfer from sulfide to a variety of small c-type cytochromes (Brune, 1995). If one were to join the electron transport chains of

Chapter 18

Validity of the Z-Scheme

RCI and RCII bacteria, one would not have the Zscheme, but rather two cycles joined at the Cyt bc complex. The other key step in the evolution of the Zscheme was the invention of water oxidation. It is impossible at this time to determine which occurred first. One can imagine an RCII bacteria that developed a new type of RCII with a more strongly-oxidizing primary donor and which associated with manganese ions in a way that allowed the oxidation of water. A duplication of the genes encoding the core subunits would have allowed the differentiation of one of the RCIIs into this hypothetical ‘proto-PS II,’ since such a bacteria would have been obliged to retain its original RCII in order to perform cyclic electron transport and generate a proton gradient. Gene transfer would then allow the introduction into the mix of an RCI, which may or may not have already evolved into a PS I-like complex. Such a gene transfer could have also occurred before the development of oxygen evolution. Alternatively, the ancestor of all photosynthetic bacteria may have already contained both, but the lack of any identified anoxygenic bacteria with both an RCI and an RCII argue against this. In any event, the common ancestor of cyanobacteria and chloroplasts lost its ‘cycling’ RCII without a firmly attached OEC. This last point explains why mutant algae that even possess only PS I or PS II cannot fix though photosynthetic bacteria possessing only one type of RC are able to perform this feat. They do so by using an exogenous source of electrons that is not intimately linked to photochemistry, such as sulfide, in concert with cyclic electron transport. Cyanobacteria, algae, and plants use water, whose oxidation is intimately linked to PS II photochemistry. The OEC, which is required for stable water oxidation, also shields P680 from Cyt c or PC, which would otherwise be thermodynamically capable of reThus, organisms using PS II are reducing dedicated water-oxidizers. There are exceptions to as an this generalization, such as the use of electron source in algae (discussed above), and a cyanobacterium (Oscillatoria limnetica) that can facultatatively use sulfide as a source of electrons to without the use ofPS II (Cohen et al., 1975b; fix Cohen et al., 1975a). The fact that 11 of 21 cyanobacterial strains tested by Garlick et al. (1977) were capable of facultative sulfide-dependent, anoxygenic photosynthesis indicates that this ability likely reflects an ancestral trait rather than one transferred recently, as was suggested by Blankenship

359 (1992). PS I should be incapable of oxidizing water is of on both thermodynamic (the +447 mV; Webber et al., 1996) and structural (it has no OEC) grounds. However, cyanobacteria and algae can revert to a ‘RCI-bacterial’ mode if given another source ofelectrons, because PS I is capable of cycling. PS II is not, and therefore is less flexible, as the amount of protons pumped (by water splitting and PQ reduction) per electron transferred to PQ is a constant (1:1). If PS II could directly reduce Fd or one would expect to see fixation in the absence of PS I. The fact that this does not occur in vivo implies that PS II is incapable of performing this feat to any significant degree. The absence of observable photoevolution, which does not require ATP, in PS I-deficient Chlamydomonas mutants (L. Cournac, G. Peltier, and K. Redding, unpublished) adds greater weight to this conclusion.

VII. Conclusions Oxygenic fixation cannot be driven by either PS I or PS II in isolation. It is clear that the simplest view of the Z-scheme with a single linear pathway (Fig. 1 A) does not explain all from water to of the data. Electrons can enter and exit the pathway at several points. Cyclic electron transport around PS I (as shown in Fig. 1B) has been experimentally verified and is analogous to similar bioenergetic pathways in anoxygenic photosynthetic bacteria. In or starch can be used as the absence of PS II, electron donors. Studies with PS I-deficient mutants have also demonstrated that electrons can exit the PQ pool. The nature of the electron carriers involved in these alternative electron transfer pathways and the contribution of these pathways to photosynthesis will require further work for their elucidation. fixation does not occur in the absence Oxygenic of PS I in vivo in Chlamydomonas, but the fact that surprisingly small amounts of PS I appear to be required for fixation and evolution will be an interesting subject for further study.

Acknowledgments The authors wish to thank N. Roggli for preparation of figures, M. Goldschmidt-Clermont, M. Hippler, S. Merchant, and J.-D. Rochaix for critical reading of the manuscript. Our thinking has been stimulated by

360 interesting discussions with A.W. Rutherford, L. Cournac, S. Merchant, E. Greenbaum, J.W. Lee, T.G. Owens, and L.J. Mets. K. Redding also wishes to thank the National Science Foundation (USA) for financial support via a Plant Biology Fellowship and J.-D. Rochaix for his support during a post-doctoral stay in his laboratory. References Albertsson P-Å, Hsu BD, Tang GMS and Arnon DI (1983) driven Photosynthetic electron transport from water to by Photosystem II in inside-out chloroplast vesicles. Proc Natl Acad Sci USA 80:3971–3975 Amesz J (1995) The antenna-reaction center complex from heliobacteria. In: Blankenship RE, Madigan MT and Bauer CE (eds) Anoxygenic Photosynthetic Bacteria, pp 687–697. Kluwer Academic Publishers, Dordrecht Arieli B, Shahak Y, Taglicht D, Hauska G and Padan E (1994) Purification and characterization of sulfide-quinone reductase, a novel enzyme driving anoxygenic photosynthesis in Oscillatoria limnetica. J Biol Chem 269:5705–5711 Arnon DI (1995) Divergent pathways of photosynthetic electron transfer: The autonomous oxygenic and anoxygenic photosystems. Photosynth Res 46:47–71 Arnon DI and Barber J (1990) Photoreduction of by isolated reaction centers of Photosystem II: Requirement for plastocyanin. Proc Natl Acad Sci USA 87:5930–5934 Arnon DI, Tsujimoto HY and Tang GM (1980a) Contrasts between oxygenic and anoxygenic photoreduction of ferredoxin: Incompatibilities with prevailing concepts of photosynthetic electron transport. Proc Natl Acad Sci USA 77:2676–2680 Arnon DI, Tsujimoto HY and Tang GM (1980b) Photoreduction of ferredoxin by chloroplasts with or without an accompanying photoreduction of the bound iron-sulfur centers. FEBS Lett 120:119–124 Bendall DS and Manasse RS (1995) Cyclic photophosphorylation and electron transport. Biochim Biophys Acta 1229:23–38 Bennoun P and Levine RP (1967) Detecting mutants that have impaired photosynthesis by their increased level of fluorescence. Plant Physiol 42:1284–1287 Blankenship RE (1992) Origin and early evolution of photosynthesis. Photosynth Res 33:91–111 Boichenko VA (1996) Can Photosystem II be a photogenerator of low potential reductant for fixation and evolution? Photosynth Res 47:291–292 Boyer PD (1993) The binding change mechanism for ATP synthase—some probabilities and possibilities. Biochim Biophys Acta 1140:215–250 Brune DC (1995) Sulfur compounds as photosynthetic electron donors. In: Blankenship RE, Madigan MT and Bauer CE (eds) Anoxygenic Photosynthetic Bacteria, pp 847–870. Kluwer Academic Publishers, Dordrecht Buchanan B B (1980) Role of light in the regulation of chloroplast enzymes. Ann Rev Plant Physiol 31:341–374 Choquet Y, Goldschmidt-Clermont M, Girard-Bascou J, Kück U, Bennoun P and Rochaix JD (1988) Mutant phenotypes

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361 Kück U, Choquet Y, Schneider M, Dron M and Bennoun P (1987) Structural and transcriptional analysis of two homologous genes for the P700 chlorophyll a-apoproteins in Chlamydomonas reinhardtii: Evidence for in vivo transsplicing. EMBO J. 6:2185–2192 Lavergne J and Joliot P (1991) Restricted diffusion in photosynthetic membranes. Trends Biochem Sci 16:129–134 Lavergne J and Joliot P (1996) Dissipation in bioenergetic electron transfer chains. Photosynth Res 48:127–138 Lee JW, Tevault CV, Owens TG and Greenbaum E (1996) Oxygenic photoautotrophic growth without Photosystem I. Science 273:364–367 Levine RP (1968) Genetic dissection of photosynthesis. Science 162:768–771 Maione TE and Gibbs M (1986) Hydrogenase-mediated activities in isolated chloroplasts of Chlamydomonas reinhardii. Plant Physiol 80:360–363 Myers J (1971) Enhancement studies in photosynthesis. Ann Rev Plant Physiol 22:289–312 Nitschke W, Mattioli T and Rutherford AW (1996) The FeS-type photosystems and the evolution of photosynthetic reaction centers. In: Baltscheffsky H (ed) Origin and Evolution of Biological Energy Conversion, pp 177–203. VCH Publishers, New York Peltier G and Thibault P (1988) Oxygen-exchange studies in Chlamydomonas mutants deficient in photosynthetic electron transport: evidence for a Photosystem II-dependent oxygen uptake in vivo. Biochim Biophys Acta 936:319–324 Ravenel J, Peltier G and Havaux M (1994) The cyclic electron pathways around Photosystem I in Chlamydomonas reinhardtii as determined in vivo by photoacoustic measurements of energy storage. Planta 193:251–259 Russel GK and Gibbs M (1968) Evidence for the participation of the reductive pentose phosphate cycle in photoreduction and the oxyhydrogen reaction. Plant Physiol 43:649–652 Rutherford AW and Nitschke W (1996) Photosystem II and the quinone-iron-containing reaction centers: comparisons and evolutionary perspectives. In: Baltscheffsky H (ed) Origin and Evolution of Biological Energy Conversion, pp 143–175. VCH Publishers, New York Schütz M, Shahak Y, Padan E and Hauska G (1997) Sulfidequinone reductase from Rhodobacter capsulatus. Purification, cloning, and expression. J Biol Chem 272:9890–9894 Sétif PQ and Bottin H (1994) Laser flash absorption spectroscopy study offerredoxin reduction by Photosystem I in Synechocystis sp. PCC 6803: Evidence for submicrosecond and microsecond kinetics. Biochemistry 33:8495–504 Sétif PQ and Bottin H (1995) Laser flash absorption spectroscopy study of ferredoxin reduction by Photosystem I: Spectral and kinetic evidence for the existence of several Photosystem Iferredoxin complexes. Biochemistry 34:9059–9070 Shahak Y, Arieli B, Binder B and Padan E (1987) Sulfidedependent photosynthetic electron flow coupled to proton translocation in thylakoids of the cyanobacterium Oscillatoria limnetica. Arch Biochem Biophys 259:605–615 Shahak Y, Arieli B, Padan E and Hauska G (1992) Sulfide quinone reductase (SQR) activity in Chlorobium. FEBS Lett 299:127–130 Slovacek RE, Crowther D and Hind G (1980) Relative activities of linear and cyclic electron flows during chloroplast 2fixation, Biochim Biophys Acta 592:495–505

362 Smart LB, Anderson SL and McIntosh L (1991) Targeted genetic inactivation of the Photosystem I reaction center in the cyanobacterium Synechocystis sp. PCC 6803. EMBO J. 10:3289–3296 Spreitzer RJ and Mets L (1981) Photosynthesis-deficient mutants of Chlamydomonas reinhardtii with associated light-sensitive phenotypes. Plant Physiol 67:565–569 Staehelin LA, De Witt M (1984) Correlation of structure and function of chloroplast membranes at the supramolecular level. J Cell Biochcm 24:261–269 Stryer L (1995) Biochemistry, Fourth Edition. W. H. Freeman and Company, New York Stuart TS and Gaffron H (I972a) The mechanism of hydrogen production by several algae. I. The effect of inhibitors of photophosphorylation. Planta 106:91–100 Stuart TS and Gaffron H (1972b) The mechanism of hydrogen production by several algae. II. The contribution of Photosystem I I . Planta 106:101–112 Tagawa K, Tsujimoto HY and Arnon DI (1963) Role of chloroplast ferredoxin in the energy conversion process of photosynthesis. Proc Natl Acad Sci USA 49:567–572 Takahashi Y, Matsumoto H, Goldschmidt-Clermont M and Rochaix JD (1994) Directed disruption of the Chlamydomonas chloroplast psbK gene destabilizes the Photosystem II reaction center complex. Plant Mol Biol 24:779–788

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Chapter 19 Assembly of Light-Harvesting Systems J. Kenneth Hoober, Hyoungshin Park, Gregory R. Wolfe*, Yutaka Komine and Laura L. Eggink Center for the Study of Early Events in Photosynthesis, and Department of Plant Biology, Arizona State University, Tempe, AZ 85287-1601, U.S.A.

Summary I. Thylakoid Biogenesis in Chlamydomonas A. Advantages of the System B. Kinetics of Greening II. Analysis of LHCII Assembly A. Structural Model of LHCII B. Synthesis of LHCII Apoproteins and Assembly of the Complex III. Site of Assembly of LHCII During Initial Greening A. Ultrastructure of the Developing Chloroplast B. Immunolocalization of LHCII Apoproteins 1. Strain y1 2. Strain cw15 arg7A 3. Strain cbn1-113 arg2 y 4. Strain MC9 C. Fate of Cytoplasmic LHCII Apoproteins IV. Conclusions Acknowledgments References

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Summary Plastid development in ‘yellow-in-the-dark’ strains of Chlamydomonas reinhardtii provides a unique system to investigate assembly of light-harvesting complexes (LHC) in situ. The ultrastructure of the plastid in degreened cells exposed to only a few minutes of light at 38°C indicated that thylakoid membrane formation was initiated by expansion of the envelope. The kinetics of accumulation of LHCII were consistent with assembly of the complex concomitant with import of the apoproteins into the chloroplast. The initial fluorescence of newly formed LHCII was quenched within the first minute of greening as the energy transducing system also assembled within initial membranes. Most of the chlorophyll (Chl) in early greening cells was distributed in foci at the periphery of the chloroplast. In such cells, LHCII apoproteins were detected adjacent to the inner surface of the envelope by immunoelectron microscopy. Interestingly, LHCII apoproteins were also detected outside of the chloroplast in vacuoles, rather than in the chloroplast stroma, when synthesized in the absence of synthesis of Chl. Wall-deficient cw15 cells, grown in the light or dark at 28°C, contain large granules within these vacuoles that were immunoreactive to antibodies against LHCII apoproteins. In a Chl b-less strain, cbn1­ 113 arg2 y, which is an algal analogue of chlorina mutants of plants when grown in medium lacking acetate, *Current Address: USDA-ARS Western Cotton Research Laboratory, 4135 E. Broadway Road, Phoenix, AZ 85040–8830, U.S.A. J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 363–376. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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LHCII apoproteins accumulated also in cytoplasmic vacuoles. The apoproteins were quantitatively recovered as the mature-sized species in each case, which suggested that even those outside of the chloroplast were processed. A possible interpretation of these results is that assembly of LHCII in the chloroplast envelope, by association of apoproteins with Chl and xanthophylls, is required to prevent retrograde movement of the proteins into the cytosol. I.Thylakoid Biogenesis in Chlamydomonas

A. Advantages of the System Biochemical and ultrastructural studies of initial thylakoid formation during conversion of etioplasts to chloroplasts in several-day-old seedlings of barley (Hordeum vulgare) showed that chlorophyll (Chl) synthesis from the protochlorophyllide (Pchlide) pool within the prolamellar bodies was followed by dispersal of these structures into prothylakoid membranes (Henningsen and Boynton, 1969, 1970, 1974; Whatley et al., 1982). Light-harvesting Chl a/bprotein complexes (LHCII) associated primarily with Photosystem II (PS II) in thylakoid membranes of greening barley seedlings accumulated in these membranes initially as monomers, which over a period of 6 to 12 h formed the trimers found in mature chloroplasts (Dreyfuss andThornber, 1994a). The site of assembly of these monomers in the chloroplast, with apoproteins synthesized on cytoplasmic ribosomes (Hoober and Stegeman, 1976), has been the subject of intensive study. The different compositions of the two halves of the thylakoid membrane bilayer suggests that lipids are transferred from their site ofsynthesis in the envelope (Douce and Joyard, 1990; Ohlrogge and Browse, 1995) to thylakoid membranes through transient fusions of these membranes (Rawyler et al., 1995). Whether complexes of proteins and pigments would also be transferred through such fusions, after assembly in the envelope, or are assembled within the developing thylakoids, is uncertain. The slow progression of chloroplast development in plant systems has precluded direct analysis of the mechanism of LHCII assembly and thylakoid formation. Algal cells, however, are more easily manipulated for experimental work and provide amenable systems for such studies (Hoober et al., 1994). Use of Chlamydomonas reinhardtii as a Abbreviations: CHL – chlorophyll; Chlide – chlorophyllide; LHC – light-harvesting complexes; LHCII – light-harvesting Chl a/b-protein complexes; Pchlide – protochlorophyllide; PS I – Photosystem I; PS II – Photosystem II

favorite organism for studies on the biophysical, biochemical and physiological processes in chloroplast development was initiated by the discovery of a mutation that causes a ‘yellow-in-the-dark’ (y1) phenotype inherited in a Mendelian pattern (Sager, 1955). Although the algal chloroplast genome contains chlB, chlL and chlN genes that encode a light-independent NADPH:Pchlide oxidoreductase, mutations in the nuclear genome apparently affect expression of one or more of these genes. Consequently, Chl synthesis in such y mutants requires the light-dependent NADPH:Pchlide oxidoreductase encoded by the nuclear CRlpcr-1 gene (Li and Timko, 1996). Cells of y strains, unable to synthesize Chl, make little or no thylakoid membranes in the chloroplast when grown in the dark (Ohad et al., 1965a). After growth for 3 to 4 d in the dark (four to six doubling times), the amount of Chl and residual membranes are reduced to only a few percent of the light-grown, green cell level. This condition provides a unique starting point for studying thylakoid formation (Ohad et al., 1965b). The major LHCII contains the predominant protein components, one-half to two-thirds of the total Chl, and nearly all of the Chl b of thylakoid membranes. Assembly of this complex is an excellent system to approach the study of membrane biogenesis. Two models were proposed for the pathway from synthesis of LHCII apoprotein precursors to integration of the mature forms into thylakoid membranes. Pathway 1 (Fig. 1),the ‘membrane-vesicle intermediate’model, proposed assembly of LHCII within the inner membrane ofthe envelope concomitant with transport of apoproteins into the plastid and subsequent transport to thylakoid membranes as membrane vesicles (Hoober, 1987). Pathway 2, the ‘soluble intermediate’ model (Cline et al., 1989; Reed et al., 1990), proposed that precursors of LHCII apoproteins are transported across the envelope into the stroma and subsequently integrated into thylakoid membranes. In the first case, processing of the precursors would occur while the proteins reside within the envelope membranes, whereas in the latter model processing may occur before or after integration into

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on a Chl b-less mutant of C. reinhardtii, which has a chlorina-type phenotype, have suggested an alternative route of elimination by secretion of apoproteins from the cell (Komine et al., 1996; Park and Hoober, 1997).

B. Kinetics of Greening

thylakoid membranes (Chitnis et al., 1986). The ‘membrane-vesicle intermediate’ model predicts that initiation of biogenesis of thylakoid membranes occurs when LHCII apoproteins interact with newly synthesized Chl (or chlorophyllide [Chlide]) and xanthophylls within the envelope during import. Synthesis of additional pigments and lipids would lead to expansion of the inner membrane and discharge of thylakoid domains as vesicles. Because Pathway 1 can be studied only in systems that generate bulk quantities of membrane, i.e., synthesize sufficient amounts of the proteins, pigments and lipids needed for measurable de novo assembly of thylakoid membranes, in vivo systems are required. These systems can also be used to test Pathway 2 by investigating whether soluble intermediates occur within the chloroplast during development. The fate of LHCII apoproteins in mutants lacking Chl b has also been a provocative issue. In higher plants, Chl b is required for accumulation of the major apoproteins, Lhcb 1 and Lhcb2 (Jansson, 1994; Priess and Thornber, 1995). Chl b-less strains result in a Chl-deficient (chlorina) phenotype. Some results suggested that these major apoproteins are rapidly degraded after import into the chloroplast in the absence of Chl b (Bellemare et al., 1982; Terao and Satoh, 1989), although identification of the responsible protease has not been achieved. Studies

At 25°C, Chl synthesis resumed after a lag of several hours when dark-grown, yellow cells were returned to light (Ohad et al., 1965b; Hoober, 1972). Accumulation of mRNA for LHCII apoproteins and expression of other light-inducible genes correlated with the gradual increase in Chl (Hoober et al., 1982; Malnoë et al., 1988). The lag period, however, made biochemical studies of chloroplast development— in particular, the initial events in the biogenesis of thylakoid membranes—difficult and offered few advantages over plant systems other than the ease of manipulation ofthe cells and the generation ofmutant strains. An adventitious finding was that messenger RNA for the major LHCII apoproteins accumulated in the dark when yellow cells were incubated at higher temperatures (Hoober et al., 1982). The mechanism of induction by an increase in temperature has not been determined, but the recent discovery that Lhcb genes in plants are repressed in the dark (Wei and Deng, 1996; Chory et al., 1996) suggests that at higher temperatures a factor required for repression is inactivated. Although 3 8 °C is above the temperature at which C. reinhardtii cells normally grow, cultures show no obvious symptoms of heat shock or metabolic stress, and protein synthesis proceeds at rates expected for the higher temperature. Sato et al. (1996) found that the optimal temperature for photosynthesis in C. reinhardtii is 35 °C to 38 °C. A feature unique to these higher temperatures was the immediate greening of cells exposed to monochromatic light of 645 to 655 nm, the wavelengths maximally absorbed by the ternary complex of NADPH, Pchlide, and Pchlide oxidoreductase (Hoober and Stegeman, 1976). This observation was particularly significant because it demonstrated that only synthesis of Chl was required to induce assembly of thylakoid membranes under these conditions. Thus, a study of events in the assembly of LHCII and formation of thylakoid membranes could be undertaken within the first few minutes after the process was triggered by light. The starting material for biochemical experiments were cells degreened by growth in the dark for 3 d,

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incubated at 38 °C in the dark for 1.5 to 2 h, and then exposed to light (Maloney et al., 1989). Studies of ultrastructural or immunological changes were done with cells grown at 28 °C or 38 °C.

II. Analysis of LHCII Assembly

A. Structural Model of LHCll The structural model that guides investigations of LHCII assembly was derived from electron crystallographic data by Kühlbrandt et al. (1994). Monomeric units ofthe major peripheral LHCII from pea contain a single apoprotein, which in the structural studies was a polypeptide chain 232 amino acids in length, with three helical, membrane-spanning regions. The first and third helical segments, nearest the N- and Ctermini, respectively, are positioned next to each other in the folded structure and interact with two lutein molecules that also span the membrane. The middle membrane-spanning segment forms an outer boundary on one side of the complex and may be involved in the association of monomers to form trimers, which seems to be the native arrangement and the form used for structural analysis (Kühlbrandt and Wang, 1991; Green and Durnford, 1996). Approximately seven Chl a and six Chl b molecules are bound to each apoprotein. Kühlbrandt et al. (1994) suggested from fluorescent energy transfer data that Chl a molecules are near the lutein molecules at the center of the complex. However, Connelly et al. (1997) showed that singlet energy transfer from lutein to Chl b was more rapid than that to Chl a, which indicated that some of the Chl b reside near the center ofthe complex. Only two histidine residues in the protein are in position to serve as ligands for Chl. Functional groups ofother amino residues, such as amide groups of asparagine and glutamine, also serve this purpose. Additional ligands may be provided by three glutamate residues in ion-pair arrangement with arginine residues (Kühlbrandt et al., 1994). Reconstitution of the complex from isolated components has shown that xanthophylls and Chl are required for assembly. Studies by Paulsen et al. (1993) indicated that the pigments are required for initiation of folding of the polypeptide chain. Complexes reconstituted with lutein and Chl b were more stable than those containing lutein and Chl a,

although those with lutein and both Chls were most stable (Cammarata et al., 1997). Loroxanthin, an algal xanthophyll, as well as violaxanthin or neoxanthin will replace lutein in vitro (Plumley et al., 1987; Cammarata et al., 1997). Reconstituted LHCII, made with a major apoprotein, Lhcbl, trimerizes in vitro in the presence of the thylakoid lipid phosphatidyl glycerol (Nussberger et al., 1993; Hobe et al., 1994). Nonpolar residues were identified in the N- and C-terminal domains of the apoprotein that are essential for trimer formation (Hobe et al., 1995; Kuttkat et al., 1996). Arabidopsis mutants that contain zeaxanthin as the only xanthophyll green slowly but show otherwise no phenotypic characteristics, which suggests that zeaxanthin can replace lutein in LHCII (Pogson et al., 1996). In mutant strains ofthe alga Scenedesmus, monomeric LHCII were assembled with other xanthophylls when lutein was absent (Bishop, 1996). These results suggest that lutein is preferred in normal assembly of LHCII but that other xanthophylls will substitute in its absence. The xanthophyll requirement in Scenedesmus was clarified by recent studies that showed that lutein is required for assembly of the major LHCII in vivo but can be replaced by violaxanthin, zeaxanthin or antheraxanthin during assembly of the minor complexes (Heinze et al., 1997). In support of this conclusion, the luteindeficient strain does not make trimeric forms of LHCII (Bishop, 1996; Heinze et al., 1997), which are generally composed of complexes containing only the major species, Lhcb1 and Lhcb2 (Bassi and Wollman, 1991; Peter and Thornber, 1991; Jansson, 1994). In plants, six families of Lhcb genes have been identified. Lhcb1 and Lhcb2 encode apoproteins of the major LHCII, whereas Lhcb3, Lhcb4, Lhcb5 and Lhcb6 encode apoproteins ofminor complexes, which serve to link the peripheral LHCII with the core complex ofPS II (Peter and Thornber, 1991; Jansson, 1994). C. reinhardtii contains three major groups of LHCII apoproteins, referred to as polypeptides 11 (ca. 31 kDa), 16 (ca. 26.5 kDa) and 17 (ca. 25.7 kDa). The trimeric form of LHCII from Chlamydomonas contains these three polypeptides plus polypeptide 13 but seems to be less stable than analogous complexes from plants (Bassi and Wollman, 1991). Monomeric LHCII purified from the alga also contains polypeptide 10, the apoprotein of CP26. In general, the complexes from Chlamy-

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domonas are very similar in composition to those in plants (Bassi and Wollman, 1991). The cloned Chlamydomonas Lhcb1 gene, from linkage group 6, apparently encodes a member of the polypeptide 11 group (Imbault et al., 1988). The Chlamydomonas Lhcb1 gene product is related phylogenetically to the Lhcb1, Lhcb2 and Lhcb3 groups from plants (Jansson, 1994; Green and Durnford, 1996). Current information is insufficient to correlate each algal apoprotein with one of the Lhcb1 through Lhcb6 species from plants. Results obtained with high resolution electrophoresis suggest that the total number of related LHCII apoproteins is perhaps as high as ten, with several of the major polypeptide fractions resolved into subfractions (Allen and Staehelin, 1994). Polyclonal antibodies raised against one of the major apoproteins from Chlamydomonas react also with the other species (Bassi and Wollman, 1991; Plumley et al., 1993). Light-harvesting complexes (LHCI) associated with PS I also contain a large family of apoproteins, with about ten proteins ranging in size from 31 kDa to 20 kDa (Bassi et al., 1992; Green and Durnford, 1996). A Lhca1 cDNA was obtained that encodes a 20 kDa apoprotein of LHCI (Hwang and Herrin, 1993). Consistent with findings of Bassi et al. (1992), that long wavelength fluorescence of LHCI at 77 K results from an aggregated state, fluorescence of PS I-minus C. reinhardtii CC–2341 cells at 708 nm and of y1 cells at 715 nm increased gradually during greening at 38 °C after a lag of about 15 min (White and Hoober, 1994). Apparently the longer wavelength maximum with y1 cells indicated a greater degree of aggregation facilitated by a functional PS I core. Analogous observations were made for PS I assembly in barley (Dreyfuss and Thornber, 1994b)

B. Synthesis of LHCII Apoproteins and Assembly of the Complex Because translatable mRNA accumulated in C. reinhardtii y1 cells during a brief incubation in the dark at 38 °C (Hoober et al., 1982), maximal arginine into LHCII apoproteins incorporation of occurred immediately when such cells were exposed was to light. The linear rate of incorporation of five- to ten-fold greater in the light than in the dark (Hoober et al., 1990). Accumulation of LHCII was monitored continuously by fluorescence of cells treated with chloramphenicol to inhibit synthesis of

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reaction center apoproteins on chloroplast ribosomes. Light of 650 nm, a wavelength optimal for reduction of Pchlide to Chlide and also near the absorption maximum of Chl was sufficient to support the process. Energy absorbed by Chl b was emitted from Chl a as fluorescence as the result of energy transfer within newly assembled LHCII (Booth and Paulsen, 1996). The kinetics of assembly (Fig. 2, curve labeled +CAP) were characterized by two phases, a 30- to 60-s period immediately after the start of illumination that involved photoreduction of the pre-existing pool of Pchlide, followed by a slightly slower phase limited probably by the rate of Chl synthesis (White et al., 1996). In the absence of chloramphenicol (Fig. 2, –CAP), fluorescence of newly formed LHCII was quenched, which indicated that a functional energy transduction system was also assembled within the first minute of illumination (White and Hoober, 1994; White et al., 1996). An increase in the amount of LHCII was not detected in the dark, consistent with the lack of synthesis of Chl. Geranylgeraniol esters of Chlides were the initial

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products of Chl synthesis (Maloney et al., 1989). Normally, the alcohol side chains are reduced stepwise to the phytol moiety. LHCII assembly at 38 °C captured newly synthesized Chl so rapidly that the side chains remained mostly unreduced, with geranylgeranyl Chlides as the predominant species. Because the extent of reduction of the side chains in Chl b was very similar to that in Chl a, either Chl a initiated partial assembly of LHCII, within which about half of the Chl a molecules were subsequently oxidized to Chl b, or LHCII assembly incorporated both Chl species from a common pool. Accumulation of Chl b did not occur when synthesis of LHCII apoproteins was inhibited. Whereas greening was supported by a high rate of synthesis of the LHCII apoproteins, amounts nearly equivalent to those in thylakoid membranes were recovered in the soluble fraction of cells treated with Because this procedure does not 0.1 M solubilize proteins integrated into a membrane structure, these results suggested that some LHCII apoproteins accumulated in a non-membrane environment. Apoproteins were not transferred from the alkali-soluble pool to membranes in pulse-chase experiments, nor were alkali-soluble apoproteins recovered in purified chloroplasts, which demonstrated that this fraction was not an intermediate in LHCII assembly. Apoproteins in the alkali-soluble and -insoluble fractions were all the size of the mature, processed forms (White et al., 1996; Park and Hoober, 1997). Proteases that cleave the targeting sequence are thought to be localized entirely within the chloroplast stroma (Su and Boschetti et al., 1993; VanderVere et al., 1995), and thus the N-termini of LHCII apoprotein precursors, even those in the alkalisoluble fraction, had apparently gained access to the stromal protease. In contrast to the processing of precursors synthesized in vitro to only about 50% completion by extracts containing processing proteases (Marks et al., 1985), the environment of the precursors in vivo apparently facilitated rapid and quantitative processing. These results suggested that precursors were processed as they reside in the envelope, with the N-terminus extending into the interior of the plastid (Schnell and Blobel, 1993). Recovery of a significant fraction of the apoproteins in a non-membrane form suggested that their synthesis was not a rate limiting factor in LHCII assembly.

III. Site of Assembly of LHCII During Initial Greening

A. Ultrastructure of the Developing Chloroplast Electron microscopy revealed that in cells greening at 25 °C, thylakoid membranes increased gradually with time in the chloroplast (Ohad et al., 1965b). In these early experiments, cells were chilled before treatment with fixative, and connections between thylakoid and envelope membranes were not detected. When degreened cells were exposed to light at 38 °C for several minutes and then injected directly into fixative in preparation for microscopy, numerous extensions of envelope membrane were observed in addition to accumulation of vesicles within the stroma (Hoober et al., 1991). These latter images suggested that expansion of the inner membrane was initiated immediately when cells were exposed to light. Although these micrographs cannot be interpreted in a dynamic sense, stromal vesicles appeared to be derived from invaginations of the inner membrane. Such structures were not generated by fixation of cells incubated under the same conditions but not exposed to light. The distribution of thylakoid membranes in degreened cells shortly after exposure to light was also examined by confocal scanning laser microscopy. Fluorescence of Chl in green cells, the control for this experiment, corresponded with the distribution of thylakoid membranes throughout the chloroplast (Fig. 3a). Fig. 3b shows cells of a PS II-deficient mutant generated by insertional mutagenesis (designated R25), in which energy absorbed by LHCII was released as fluorescence. During the early stage of greening, most Chl fluorescence was localized in a discontinuous pattern around the periphery of the chloroplast. These images supported the conclusion drawn from electron microscopy that during the early phase of greening, thylakoid membranes were associated with the envelope.

B. Immunolocalization of LHCII Apoproteins Crucial for understanding these findings was knowledge of the cellular distribution of LHCII apoproteins during the early stages of greening. For example, to discriminate whether folds in the chloroplast envelope were developing thylakoid domains or simply lipid structures, it was necessary to determine the location of newly synthesized

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apoproteins. Immunoelectron microscopy was performed with antibodies raised against polypeptide 11 (Hoober et al., 1982). The antibody preparation cross-reacts with polypeptides 10,11,13,16 and 17, which are all components of LHCII (Bassi and Wollman, 1991; Plumley et al., 1993). Experiments with each of the following mutant strains provided important information.

1. Strain y1 After degreened cells were exposed to 5 min of light of 38 °C, the shortest time in which a definitive pattern of labeling of the chloroplast was observed, binding of antibodies to LHCII apoproteins was detected along the inside surface of the envelope. The density of gold particles, conjugated to protein A to detect bound antibodies, along the envelope was greater after 15 min of light. Additional clusters of gold particles, which were not observed in darkgrown cells and perhaps marked vesicles within the stroma, occurred over areas deeper in the chloroplast at the longer time (White et al., 1996). To preserve antigenicity, the protocol for immunoelectron microscopy did not provide definition of membrane structures. Concomitant with immunolabeling of the chloroplast, binding of antibodies was detected over small

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vacuoles outside of the chloroplast. These vacuoles were discovered during the first electron microscopic studies of C. reinhardtii by Sager and Palade (1957), who noted their dense, polymorphic content. A quantitative analysis of the labeling showed that LHCII apoproteins accumulated in the vacuoles in parallel with the increase in the chloroplast (White et al., 1996). However, labeling of the vacuoles was not detected with sections of dark-grown y1 cells or when preimmune serum was used.

2. Strain cw15 arg7A Further studies showed that large, electron-opaque granules within vacuoles of cell wall-deficient (cw) mutant strains, first described by Davies and Plaskitt (1971), were immunolabeled in cells grown in the light or dark at 28 °C (Komine et al., 1996; Wolfe et al., 1997). As shown in Fig. 4, immunogold particles were detected only over thylakoid membranes within the chloroplast and over granules within vacuoles. Immunoblot analysis after electrophoresis of purified granules showed that the immunoreactive components were mature-sized LHCII apoproteins, which were recovered at higher abundance in granules from cells incubated at 38 °C than at 28 °C (White et al., 1996; Komine et al., 1996). The presence of LHCII apoproteins in vacuoles in the light or dark when

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cells were grown at 28 °C demonstrated that accumulation outside of the chloroplast was not an effect of the higher temperature.

3. Strain cbn1 -113 arg2 y A y derivative of strain cbn1-113 arg2, which does not make Chl b (Michel et al., 1983; Chunaev et al., 1991), was isolated for use in greening experiments. These cells synthesized LHCII apoproteins at nearly the same rate in the dark as in the light, yet the proteins did not accumulate in the dark. With immunoelectron microscopy, binding of antibodies was not detected over the chloroplast in dark-grown, yellow cells or to a significant extent over vacuoles (Fig. 5a). Conversely, in cells incubated at 38 °C in the dark, vacuoles outside of the chloroplast were heavily labeled and the apoproteins seemed also to be dispersed within the cytosol (Fig. 5c). When these cells were exposed to light, labeling ofthe chloroplast and cytosol increased with time (Fig. 5d). These data indicated that Chl a was necessary for accumulation of the apoproteins in the chloroplast. In the absence of Chl synthesis, the apoproteins were detected but not inside the organelle. The cbn1-113 arg2 y cells were pale-green when grown in the light at 28 °C in medium lacking acetate. The density of immunogold particles over the chloroplast in sections of such cells was 48% of that in cells grown with acetate. Concomitantly, granules

in cytoplasmic vacuoles became more heavily labeled (Park and Hoober, 1997). As in the chlorina f2 mutant of barley (Bellemare et al., 1982), the rate of synthesis of LHCII apoproteins in cbn1-113 arg2 y cells was similar to that in wild-type. Yet the amount of these proteins in the chloroplast was related to the level of Chl. Rather than degradation in the chloroplast, a process implied from results with the barley mutant, the apoproteins in the algal cells were collected in cytoplasmic vacuoles.

4. Strain MC9 A Chl b-deficient, metronidazole-resistant mutant (designated MC9) was selected after insertional mutagenesis of cw15 arg7A. Mutant cells grown at 28 °C have a high Chl a/b ratio (8 to 10), are deficient in LHCII apoproteins when grown in the light, and green slowly after growth in the dark (Wolfe et al., 1997). Fluorescence spectra at 77K showed a deficiency in LHCII and LHCI. The chloroplast contained a paucity of thylakoid membranes, arranged as single lamellae, and relatively few antibodies to LHCII apoproteins bound to sections over the organelle. Although the rate of synthesis of LHCII apoproteins was well below the expected capacity of the chloroplast to import the proteins, granules in cytoplasmic vacuoles (Fig. 6a) were immunoreactive with antibodies against LHCII apoproteins (Fig. 6b). Our findings suggest that this mutant is deficient in a

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factor required for LHCII assembly and that apoproteins synthesized in excess ofthe reduced rate of LHCII assembly were shunted into vacuoles.

C. Fate of Cytoplasmic LHCII Apoproteins LHCII apoproteins did not normally accumulate in dark-grown cells even though they were synthesized at detectable rates. The absence of labeling of the chloroplast in such cells could be explained by rapid proteolysis of apoproteins imported into the organelle, although this possibility does not account for binding of antibodies to vacuoles. Davies and Plaskitt (1971) and our work (Komine et al., 1996; Wolfe et al., 1997) demonstrated that vacuolar granules were released from wall-deficient cwl 5 cells by exocytosis (see Fig. 6a), a process perhaps related to the normal delivery of cell wall precursors. This process may also be responsible for maintaining a low level of the LHCII apoproteins in dark-grown cells. Because

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growth occurs in the dark, and thus cell wall constituents would be needed outside the plasmalemma, exocytosis may be commensurate with growth. Impairment of cell wall formation in mutant strains apparently leads to greater accumulation and/ or retention of vacuolar material within the cell. It is notable that numerous granules appeared between the cell membrane and cell wall about 12 h after wild-type cells grown in nitrogen-deficient medium were shifted to nitrogen-sufficient medium (Plumley et al., 1989). It is not known how LHCII apoproteins enter vacuoles.

IV. Conclusions Greening cells of C. reinhardtii y 1 synthesize Chl at a rate of molecules per min per cell at 38 °C. From the Chl a/b ratio of 2.3 to 2.5, and assuming that most of the Chl b resides within LHCII, the rate

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of total import of LHCII apoproteins can be calculated as at least molecules per min per chloroplast, based on the composition of crystallized LHCII (Kühlbrandt et al., 1994). Because nearly half of the mature LHCII apoproteins accumulate in an alkalisoluble fraction and thus are not integrated into thylakoid membranes, the total rate of synthesis and import of these proteins during greening at 38 °C molecules per min per may be as high as 3 to cell. This rate provides an amount of protein that is sufficient for localization by immunoelectron microscopy during the early stage of chloroplast development. Experiments with immature chloroplasts purified from pea seedlings described a flux of precursors of LHCII apoproteins into the organelle. Whereas proteins targeted to the stromal or luminal compartments were imported into purified chloroplast at a rate of molecules per minute, near the estimated maximal rate in vivo (Cline and Henry, 1996), LHCII apoproteins were imported at a rate of only a few hundred per minute per chloroplast (Reed et al., 1990; Yuan et al., 1993). Rapid-stopping techniques demonstrated that soluble intermediates accumulated with kinetics typical for a precursorproduct relationship with the membrane-integrated form (Reed et al., 1990). The soluble form consisted

of the apoproteins in a complex with stromal proteins, a state competent for insertion into the membrane (Cline et al., 1989; Reed et al., 1990; Payen and Cline, 1991). These observations formed the basis of the ‘soluble intermediate’ model (Fig. 1). A stromal protein involved in this process is CP54, a homologue of a subunit of the signal recognition particle (Li et al., 1995). Stromal extracts depleted immunologically of CP54 were unable to form the soluble complex or facilitate integration ofthe apoproteins into thylakoid membranes (Cline and Henry, 1996). CP54 bound nonpolar sequences preferentially, and interacted strongly with the third membrane-spanning helix of LHCII apoproteins (High et al., 1997). Chl synthesis was not required for import under the conditions of these experiments, although Kohorn et al. (1986) showed that addition of 5-aminolevulinate and Sadenosyl methionine, precursors of Chl, markedly enhanced recovery of LHCII apoproteins in chloroplasts. A common means to evaluate correct integration of LHCII apoproteins into thylakoid membranes is the extent ofprotection the membrane provides to proteolysis. Treatment of the stromal side of thylakoid membranes with proteases such as trypsin or thermolysin produced fragments lacking 1 to 2 kDa from the N-terminus (Hoober et al., 1990; Plumley and Schmidt, 1995). Most ofthe apoproteins

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integrated directly into the membrane in vitro were similarly protected from proteolysis and were recovered in complexes with Chl after solubilization of the membrane and electrophoresis. However, LHCII reconstituted in vitro also showed the same resistance to proteolysis as the complex in membranes (Paulsenetal., 1993). Some of the LHCII apoproteins recovered from membranes after in organello import or direct in vitro insertion into thylakoid membranes were digested to a smaller fragment, which lacked an additional 5 to 6 kDa. The abundance of the smaller proteolytic fragment at early stages of import was interpreted as evidence for an intermediate stage in assembly of the complex, in which more of the Nterminus was exposed on the stromal surface of the membrane (Reed et al., 1990; Yuan et al, 1993; Kuttkat et al., 1996). Alternatively, the smaller fragment may result from cleavage near the Cterminus, which is normally on the luminal surface of the membrane, and would suggest incorrect orientation of at least a portion of the protein. Extensive data obtained from in vitro systems support the ‘soluble intermediate’ model (Cline and Henry, 1996). However, these systems contain only the purified chloroplast without the cytosol that surrounds the organelle in vivo. In experiments in which cytosolic proteins from wheat germ were added to isolated pea chloroplasts, import of LHCII precursors was almost completely inhibited (Yuan et al., 1993), which may indicate the importance of the in vivo environment. Further distinction between in vitro and in vivo import was obtained by Silva-Filho et al. (1997) with a chimeric protein that included the transit peptide and a few amino acids of the mature triose phosphate 3-phosphoglycerate-phosphate translocator (an inner envelope membrane protein) linked to chloramphenicol transacetylase. Whereas the protein containing 5 amino acids of the mature protein was imported into the chloroplast in vitro, a protein the size of the processed form was recovered in the cytosol rather than the chloroplast from intact leaves of tobacco transformed with the gene encoding this protein. These results suggest that import in vivo is more stringent than in in vitro experiments. Because inclusion of longer lengths of the mature translocator led to chloroplast localization in vivo, specific interaction with components within the chloroplast apparently is normally involved during import. Our results on LHCII assembly in vivo suggest that bulk import of the apoproteins requires synthesis of Chi, and that in its absence the partially imported

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and processed proteins are retracted into the cytosol. Chl a and xanthophylls possibly trigger folding of apoproteins, as described by Paulsen et al. (1993), as they reside within the import apparatus in the envelope. The formation of folds or loops within the proteins, which can occur on a microsecond time scale (Hagen et al., 1996), may sequester Chl a molecules from their environment. Because synthesis of Chl b seems to depend upon the presence of LHCII apoproteins, an intermediate form of LHCII may be assembled, within which about half of the Chl a molecules are oxidized to Chl b (Plumley and Schmidt, 1995). This pathway would explain the similarities in side chain composition of the two forms of Chl. Either cytosolic proteins remain temporarily bound to LHCII apoproteins and abort import when Chl is not made, or the apoproteins slip into the intermembrane space when import is incomplete, as suggested by Scott and Theg (1996). Although our work has provided an outline of LHCII assembly in vivo, with transfer of LHCII from the envelope to thylakoid membranes as described by the ‘membrane vesicle intermediate’ model, studies with cell-free systems are required to identify cytosolic and additional stromal factors that are involved with integration of LHCII apoproteins into membranes. Further work is also required to understand the transfer of LHCII apoproteins into vacuoles and the roles these structures play in the physiology of Chlamydomonas.

Acknowledgments This is publication number 331 from the Arizona State University Center for the Study of Early Events in Photosynthesis. L. Eggink was supported by Graduate Research Training Grant DGE9553456 from the National Science Foundation.

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Henningsen KW and Boynton JE (1969) Macromolecular physiology of plastids. VII. The effect of a brief illumination on plastids of dark-grown barley leaves. J Cell Sci s5: 757–793 Henningsen KW and Boynton JE (1970) Macromolecular physiology of plastids. VIII. Pigment and membrane formation in plastids of barley greening under low light intensity. J Cell Biol 44: 290–304 Henningsen KW and Boynton JE (1974) Macromolecular physiology of plastids. IX. Development of plastid membranes during greening of dark-grown barley seedlings. J Cell Sci 15: 31–55 High S, Henry R, Mould RM, Valent Q, Meacock S, Cline K, Gray JC and Luirink J (1997) Chloroplast SRP54 interacts with a specific subset of thylakoid precursor proteins. J Biol Chem 272:11622–11628 Hobe S, Prytulla S, Kühlbrandt W and Paulsen H (1994) Trimerization and crystallization of reconstituted lightharvesting chlorophyll a/b complex. EMBO J 13: 3423–3429 Hobe S, Förster, Klinger J and Paulsen H (1995) N-proximal sequence motif in light-harvesting chlorophyll a/b-binding protein is essential for trimerization of light-harvesting chlorophyll a/b complex. Biochemistry 34: 10224–10228 Hoober JK( 1972) A major polypeptide of chloroplast membranes of Chlamydomonas reinhardi. J Cell Biol 52: 84–96 Hoober JK (1987) The molecular basis of chloroplast development. In: Hatch MD and Boardman NK (eds) The Biochemistry of Plants, Vol 10, Photosynthesis, pp 1–74. Academic Press, San Diego Hoober JK and Stegeman WH (1976) Kinetics and regulation of synthesis of the major polypeptides of thylakoid membranes in Chlamydomonas reinhardtii y-1 at elevated temperatures. J Cell Biol 70: 326–337 Hoober JK, Marks DB, Keller BJ and Margulies MM (1982) Regulation of accumulation of the major thylakoid polypeptides in Chlamydomonas reinhardtii y-1 at 25 °C and 38 °C. J Cell Biol 95: 552–558 Hoober JK, Boyd CO and Paavola LG (1991) Origin of thylakoid membranes in Chlamydomonas reinhardtii y-1 at 38 °C. Plant Physiol 96: 1321–1328 Hoober JK, Maloney MA, Asbury LR and Marks DB (1990) Accumulation of chlorophyll a/b-binding polypeptides in Chlamydomonas reinhardtii y-1 in the light or dark. Plant Physiol 92: 419–426 Hoober JK, White RA, Marks DB and Gabriel JL (1994) Biogenesis of thylakoid membranes with emphasis on the process in Chlamydomonas. Photosynth Res 39: 15–31 Hwang S and Herrin DL (1993) Characterization of a cDNA encoding the 20-kDa Photosystem I light-harvesting polypeptide of Chlamydomonas reinhardtii. Curr Genet 23: 512–517 Imbault P, Wittemer C, Johanningmeier U, Jacobs JD and Howell SH (1988) Structure of the Chlamydomonas reinhardtii cabII­ 1 gene encoding a chlorophyll-a/b-binding protein. Gene 73: 397–407 Jansson S (1994) The light-harvesting chlorophyll a/b-binding proteins. Biochim Biophys Acta 1184: 1–19 Kohorn BD, Harel E., Chitnis PR, Thornber JP and Tobin EM (1986) Functional and mutational analysis of the lightharvesting chlorophyll a/b protein of thylakoid membranes. J Cell Biol 102: 972–981 Komine Y, Park H, Wolfe GR and Hoober JK (1996) Secretory

Chapter 19 Assembly of Light-Harvesting Systems

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organization of higher plant Photosystem II light-harvesting pigment-proteins. J Biol Chem 266: 16745–16754 Plumley FG and Schmidt GW (1987) Reconstitution of chlorophyll a/b light-harvesting complexes: Xanthophylldependent assembly and energy transfer. Proc Natl Acad Sci USA 84: 146–150 Plumley FG and Schmidt GW (1995) Light-harvesting chlorophyll a/b complexes: Interdependent pigment synthesis and protein assembly. Plant Cell 7: 689–704 Plumley FG, Douglas SE, Branagan-Switzer A and Schmidt GW (1989) Nitrogen-dependent biogenesis of chlorophyll-protein complexes. In: Briggs WR (ed) Photosynthesis, pp 311–329. Alan R Liss, New York Plumley FG, Martinson TA, Herrin DL, Ikeuchi M and Schmidt GW (1993) Structural relationships of the Photosystem I and Photosystem II chlorophyll a/b and a/c light-harvesting apoproteins of plants and algae. Photochem Photobiol 57: 143–151 Pogson B, McDonald DA, Truong M, Britton G and DellaPenna D (1996) Arabidopsis carotenoid mutants demonstrate that lutein is not essential for photosynthesis in higher plants. Plant Cell 8: 1627–1639 Preiss S and Thornber JP (1995) Stability of the apoproteins of light-harvesting complex I and II during biogenesis of thylakoids in the chlorophyll b-less barley mutant chlorina f2. Plant Physiol 107: 709–717 Rawyler A, Meylan-Bettex M and Siegenthaler PA (1995) (Galacto)lipid export from envelope to thylakoid membranes in intact chloroplasts. II A general process with a key role for the envelope in the establishment of lipid asymmetry in thylakoid membranes. Biochim Biophys Acta 1233: 123–133 Reed JE, Cline K, Stephens LC, Bacot KO and Viitanen PV (1990) Early events in the import/assembly pathway of an integral thylakoid protein. Eur J Biochem 194: 33–42 Sager R (1955) Inheritance in the green alga Chlamydomonas reinhardi. Genetics 40: 476–489 Sager R and Palade GE (1957) Structure and development of the chloroplast in Chlamydomonas, I. The normal green cell. J Biochem Biophys Cytol 3: 463–487 Sato N, Sonoike K, Kawaguchi A and Tsuzuki M (1996) Contribution of lowered unsaturation levels of chloroplast lipids to high temperature tolerance of photosynthesis in Chlamydomonas reinhardtii. J. Photochem Photobiol B: Biol 36: 333–337 Schnell DJ and Blobel G (1993) Identification ofintermediates in the pathway of protein import into chloroplasts and their localization to envelope contact sites. J Cell Biol 120:103–115 Scott SV and Theg SM (1996) A new chloroplast protein import intermediate reveals distinct translocation machineries in the two envelope membranes: Energetics and mechanistic implications. J Cell Biol 132: 63–75 Silva-Filho M de C, Wieërs M-C, Flügge U-I, Chaumont F and Boutry M (1997) Different in vitro and in vivo targeting properties of the transit peptide of a chloroplast envelope inner membrane protein. J Biol Chem 272: 15264–15269 Su Q and Boschetti A (1993) Partial purification and properties of enzymes involved in the processing of a chloroplast import protein from Chlamydomonas reinhardtii. Eur J Biochem 217: 1039–1047 Terao T and Katoh S (1989) Synthesis and breakdown of the apoproteins of light-harvesting chlorophyll a/b proteins in

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chlorophyll b-deficient mutants of rice. Plant Cell Physiol 30: 571–580 VanderVere PS, Bennett TM, Oblong JE and Lamppa GK (1995) A chloroplast processing enzyme involved in precursor maturation shares a zinc-binding motif with a recently recognized family of metalloendopeptidases. Proc Natl Acad Sci USA 92: 7177–7181 Wei N and Deng X-W (1996) The role of the COP/DET/FUS genes in light control of Arabidopsis seedling development. Plant Physiol 112: 871–878 Whatley JM, Hawes CR, Home JC and Kerr JDA (1982) The establishment of the plastid thylakoid system. New Phytol 90: 619–629 White RA and Hoober JK (1994) Biogenesis of thylakoid

membranes in Chlamydomonas reinhardtii y1. A kinetic study of initial greening. Plant Physiol 106: 583–590 White RA, Wolfe GR, Komine Y and Hoober JK (1996) Localization of light-harvesting complex apoproteins in the chloroplast and cytoplasm during greening of Chlamydomonas reinhardtii at 38 °C. Photosynth Res 47: 267–280 Wolfe GR, Park H, Sharp WP and Hoober JK (1997) Lightharvesting complex apoproteins in cytoplasmic vacuoles in Chlamydomonas reinhardtii (Chlorophyta). J. Phycol 33: 377– 386 Yuan J, Henry R and Cline K (1993) Stromal factor plays a essential role in protein integration into thylakoids that cannot be replaced by unfolding or by heat shock protein Hsp70. Proc Natl Acad Sci USA 90: 8552–8556

Chapter 20 Pigment Biosynthesis: Chlorophylls, Heme, and Carotenoids Michael P. Timko Department of Biology, University of Virginia, Charlottesville, VA 22903, U.S.A.

Summary I. Introduction

II. Tetrapyrroles and Their Derivatives—An Overview

III. Formation of ALA

A. B. Glutamyl-tRNA Synthetase C. Glutamyl-tRNA Reductase D. Glutamate-1-Semialdehyde Aminotransferase IV. The Pathway from ALA to Protoporphyrin IX A. Porphobilinogen Biosynthesis B. Porphobilinogen Deaminase C. Uroporphyrinogen III Synthase D. Uroporphyrinogen III Decarboxylase E. Coproporphyrinogen III Oxidase F. Protoporphyrinogen IX Oxidase V. The Magnesium Branch—Chlorophyll Formation A. Mg Chelatase B. Mg-Protoporphyrin IX Methyltransferase C. Isocyclic Ring Formation D. Vinyl Reduction E. Reduction of Protochlorophyllide 1. The Light-Dependent Reaction 2. Light-Independent Reaction F. Phytylation G. Reaction Center Chlorophylls and Pheophytin H. Formation of Chlorophyll b VI. The Iron Branch—Formation of Heme A. Ferrochelatase B. Plastidic and Non-plastidic Heme and Heme Derivatives VII. Light and Metabolic Regulation of Chlorophyll Formation VIII. Carotenoids A. The Function and Distribution of Carotenoids B. Pathway for Carotenoid Formation 1. Formation of Isoprene Units and Synthesis of GGPP 2. Phytoene Synthase 3. Phytoene Desaturation 4. Cyclizations of Lycopene 5. Xanthopnyll Biosynthesis C. Pathway Regulation Acknowledgments References

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Michael P. Timko

Summary The two major pigments found in photosynthetic eukaryotic cells are the tetrapyrroles (including chlorophylls, heme, and their derivatives) and the carotenoids. Both of these classes of molecules play an important role in the light absorption and energy transduction processes ofphotosynthesis and they also participate in numerous other metabolic activities in the cell. Over the past several years there has been a considerable advancement in our understanding of the biochemistry and genetic regulation of tetrapyrrole and carotenoid formation. Genes encoding many of the enzymes of the two biosynthetic pathways have been isolated and their nucleotide and encoded primary protein structures determined. Such molecular analysis has greatly facilitated the examination of how enzyme synthesis and activity are regulated throughout development and under a variety of different growth and environmental conditions. It has also led to new insights into the reaction mechanisms and specificities of several key enzymes in these processes. In this chapter, I present an overview of the recent developments in tetrapyrrole and carotenoid biosynthesis, drawing upon work carried out in a wide range of organisms in order to better illustrate certain features of the biosynthetic process in question or to highlight particularly important differences among species. I. Introduction The two major pigments found in photosynthetic organisms are the tetrapyrroles and carotenoids. Both of these classes of molecules play an important role in photosynthesis and they also participate in numerous other metabolic processes in the cell. This chapter focuses primarily on the biochemistry and genetic regulation of tetrapyrrole and carotenoid formation, with particular emphasis placed upon our current understanding of the physical characteristics of the enzymes involved, their reaction mechanisms, substrate specificities and cofactor requirements. How the synthesis and activity of the enzymes comprising these two pathways are coordinated and regulated by various endogenous and exogenous factors (e.g., metabolite availability, light) during growth and differentiation are also discussed. By and large, our current understanding of tetrapyrrole and carotenoid biosynthesis comes from experimental work carried out in a wide range of Abbreviations: ALA– -aminolevulinicacid; CoA–coenzyme A; CPOX – coproporphyrinogen III oxidase; EDTA– ethylene dinitrilo-tetraacetic acid; EGTA – ethyleneglycol-bis-( ether) N,N´-tetraacetic acid; gabaculine – 3-amino2,3- dihydrobenzoic acid; GGPP–geranylgeranylpyrophosphate; GluRS – glutamyl-tRNA synthetase; GluTR – glutamyl-tRNA reductase; GSA – glutamate-1-semialdehyde; GSA-AT – glutamate-1-semialdehyde aminotransferase; PAGE – polyacrylamide gel electrophoresis; PBGD – porphobilinogen deaminase; PBGS – porphobilinogen synthase; PDS – phytoene synthase; POR – NADPH:protochlorophyllide oxidoreductase; PSY – phytoene synthase; S. cerevisiae – Saccharomyces cerevisiae; SAM – S-adenosyl-L-methionine; SDS – sodium lauryl sulfate; UROS – uroporphyrinogen III synthase; X – any amino acid

organisms, notably photosynthetic bacteria and vascular plants. Where information was available specifically dealing with pigment formation in Chlamydomonas spp., this work has been emphasized. Because of the breadth of information available on tetrapyrrole and carotenoid biosynthesis, the scope of treatment of some topics had to be limited. Processes not documented in Chlamydomonas spp. or aspects of regulation specific to multicellular organisms with complex developmental patterns were essentially omitted. A number of reviews reporting recent progress in various aspects of tetrapyrrole (Beale, 1993, 1995; Biel, 1995; von Wettstein et al., 1995; Fujita, 1996; Reinbothe and Reinbothe, 1996) and carotenoid (Bartley et al., 1994; Hirschberg and Chamovitz, 1994; Armstrong, 1995; Bartley and Scolnik, 1995; Armstrong and Hearst, 1996; Yamamoto and Bassi, 1996) formation in photosynthetic organisms have been published within the past few years. The reader is referred to these articles for additional information on aspects ofthe respective pathways not specifically addressed in this chapter. II. Tetrapyrroles and Their Derivatives—An Overview At least three major structurally and functionally distinct groups of tetrapyrroles are found in photosynthetic eukaryotes. These are the porphyrins, the - porphyrins, and the bilins. The three groups are distinguished by the macrocyclic or linear arrangement of their pyrrole components and in the case of the macrocyclic tetrapyrroles by the metal ion which they chelate. The - porphyrins,

Chapter 20 Pigment Biosynthesis consisting predominantly of the chlorophylls and their derivatives, are the most abundant class of tetrapyrroles found in most vascular and non-vascular land plants and algae. Bound to their apoproteins, chlorophylls and their derivatives form the lightharvesting and reaction center complexes of PS I and PS II and are intricately involved in the trapping of light energy and its conversion into stored chemical energy in the process of photosynthesis. The porphyrins, typified by protoheme and heme, comprise the second class of cyclic tetrapyrroles found in photosynthetic organisms. Protoheme and heme are generally present in all cell types where they function as cofactors or prosthetic groups for various soluble oxidative enzymes (such as catalases, peroxidases, and cytochrome P450s) and membranelocalized cytochromes comprising the mitochondrialand plastid-localized electron transfer chains. In nonphotosynthetic eukaryotes, protoheme and its derivatives have also been shown to function as effector molecules capable of regulating cellular processes such as transcription, translation, and protein translocation into organelles (Lathrop and Timko, 1993; Zhang and Guarente, 1995). Although similar regulatory functions for (proto)heme have yet to be demonstrated in photosynthetic organisms, chlorophyll and several of its biosynthetic intermediates have been shown to be involved in the regulation of translation on plastid ribosomes and import into plastids (Reinbothe et al., 1995, 1996a; Rochaix, 1996). The linear, non-metal chelating bilins are a structurally and functionally diverse group of molecules that comprise the third class oftetrapyrroles found in photosynthetic cells. Within this group, the phycobilins and phytochromobilin are of particular importance. The phycobilins, principally phycocyanins and phycoerythrins, are the major lightharvesting pigments found in the photosynthetic membranes of cyanobacteria and red algae (Beale, 1993, 1994). Phytochromobilin is covalently attached to the phytochrome apoprotein forming a holocomplex that serves as a major red light photoreceptor involved in regulating photomorphogenesis in plants (Quail et al., 1995). While phytochrome has been found in some algae (Lagarias et al., 1995) and recently in cyanobacteria (Kaneko et al., 1996), its exact role in these organisms is still unresolved. Other tetrapyrrolic compounds, such as siroheme, the prosthetic group of the nitrite and sulfite reductases (Crawford, 1995) and corrins (e.g., vitamin

379 )(Battersby, 1994), are also found in photosynthetic organisms. However, the distribution of the corrinoids appears to be quite restricted, with their formation primarily found in photosynthetic prokaryotes. Figure 1 shows schematically the general pathway for the formation of the major tetrapyrroles found in photosynthetic eukaryotes. All tetrapyrroles are derived from a common biosynthetic precursor, ALA, formed in photosynthetic organisms by one of two possible biosynthetic routes (discussed below). The biosynthetic steps from ALA to protoporphyrinogen IX constitute the major, common portion of the pathway. Starting at protoporphyrinogen IX, intermediates in the pathway may be channeled to different locations within the plastid (i.e., thylakoid or envelope membranes) or transported outside the plastid for use by enzymes in other subcellular compartments (i.e., mitochondrion). In plastids, protoporphyrin IX is the last common intermediate -branch of the pathway that gives between the rise to chlorophylls a and b and their derivatives and - branch of the pathway that yields protoheme, the heme, and in some cell types the linear tetrapyrroles such as phytochromobilin. Siroheme and the corrinoids are formed in a branch of the pathway that originates with uroporphyrinogen III. After decades of intensive study, nearly all of the enzymes involved in chlorophyll and heme formation have been identified and characterized to various extents with respect to their physical properties, catalytic requirements, substrate and cofactor specificity, and intracellular distribution. In addition, genes encoding many of the enzyme components of chlorophyll and heme biosynthesis have been characterized from at least one photosynthetic organism, and in many cases from a variety of photosynthetic prokaryotic and eukaryotic cells. These findings are summarized below.

III. Formation of ALA ALA is considered the first committed precursor of tetrapyrrole biosynthesis and its formation is known to occur by two distinct biosynthetic routes, differing significantly in their enzymatic machinery and substrate requirements. In animal cells, nonphotosynthetic lower eukaryotes (e.g., S. cerevisiae and other fungi), and the -group of purple nonsulfur bacteria (e.g., Rhodobacter, Rhodopseudo­ monas, Rhizobium), ALA is formed from glycine

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and succiny l-CoA in a condensation reaction catalyzed by the pyridoxal phosphate-requiring enzyme ALA synthase (EC 2.3.1.37, ALAS) (Beale 1995). ALAS has been studied extensively and the

Michael P. Timko

biochemical and genetic factors that regulate its formation and activity have been described in considerable detail (Dailey, 1990; Jordan, 1991; Beale, 1993, 1995).

Chapter 20 Pigment Biosynthesis

In most photosynthetic organisms, including plants, red and green algae, cyanobacteria and other photosynthetic prokaryotes (i.e., prochlorophytes, the purple and green sulfur bacteria, the green nonsulfur bacteria, and heliobacteria), ALA is formed by an alternative pathway starting with glutamate (Beale, 1995). This pathway (Fig. 2), referred to as the C5 pathway, requires four separate components: three different enzymes (glutamyl-tRNA synthetase (GluRS), glutamyl-tRNA reductase (GluTR), and glutamate-1-semialdehyde aminotransferase (GSAThe formation of AT)) and an activated ALA via the pathway begins with the ATP-dependent activation of glutamate by its esterification to a plastid (Kumar et al., 1996a). This reaction is catalyzed by GluRS. The activated, tRNA-bound glutamate is then reduced by the pyridine nucleotide requiring enzyme GluTR to form glutamate-1-semialdehyde (GSA). Finally, GSA is converted to ALA by transfer of the C4 amino group to the C5 carbon in a reaction catalyzed by GSA-AT. The basic features of pathway appear to be similar or identical in the various plants, algae, and bacteria examined and it has been shown that in some cases reaction components from different sources can be combined to reconstitute ALA biosynthetic activity (see Beale, 1993; 1995).

A. In all organisms using the pathway examined thus far, the RNA required for ALA formation is a containing the UUC anticodon for glutamate (Huang et al., 1984; Schön et al., 1986; Schneegurt and Beale, 1988). While it was suggested initially that involved in ALA formation may be a the special tRNA species, several lines of evidence have

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shown that this is not the case and that the same used for ALA formation is also used for protein synthesis (Kumar, et al., 1996a). In plants and algae, including Chlamydomonas spp., the is encoded in the chloroplast DNA (Sugiura, 1992) and its nucleotide sequence is highly conserved from E. coli (Kumar et al., 1996a). The and barley chloroplasts contain several modified bases, including a 5-methylaminomethyl-2-thiouridine in the first anticodon position shown to be necessary in the charging of the tRNA with glutamate by GluRS (Sylvers et al., 1993). Sequence comparison genes from an evolutionarily diverse of pathway and group of organisms possessing the chemical modification studies of the tRNA have also identified other conserved bases thought to be to GluRS and required for binding of the GluTR (Rogers and Söll, 1993; Willows et al., 1995) or to elongation factor EF-Tu (Stange-Thomann et al., 1994).

B. Glutamyl-tRNA Synthetase The same glutamyl-tRNA synthetase (EC 6.1.1.17, GluRS) used for protein synthesis catalyzes the esterification of glutamate onto for use in ALA formation (Kumar et al., 1996a). Like other well characterized aminoacyl-tRNA synthetases, the GluRS involved in ALA formation requires ATP and for activity. GluRSs have been isolated and characterized from C. reinhardtii (Chang et al., 1990; Chen et al., 1990a), Chlorella vulgaris (Weinstein et al., 1987), and barley (Bruyant and Kannagara, 1987), as well as from a number ofphotosynthetic and nonphotosynthetic prokaryotes (Beale, 1995). The Chlamydomonas GluRS was analyzed by both denaturing SDS-PAGE and sedimentation on glycerol

382 gradients and determined by Chen et al. (1990a) to be a monomer of 62 kDa. In contrast, Chang et al. (1990) reported that the Chlamydomonas GluRS was a 60 kDa dimer that could be denatured into identical subunits of approximately 32.5 kDa. Only a single GluRS was detected in barley chloroplasts for use in both protein synthesis and ALA formation (Bruyant and Kannangara, 1987). In Chlorella, several GluRSs were reported to be present, but only one produced a glutamyl-tRNA used in ALA formation (Avissar and Beale, 1988). Similarily, in Synechocystis strain PCC 6803, the same GluRS for use in ALA formation is used to charge used to charge the tRNA for protein synthesis (Rieble , and Beale, 1991a). Unlike its substrate, GluRS is likely to be encoded in nuclear DNA (Kumar et al., 1996a). However, a gene encoding the enzyme has yet to be isolated from plants or algae.

C. Glutamyl-tRNA Reductase Glutamyl-tRNA reductase (GluTR) catalyzes the and NADPH-dependent reduction ofglutamylto form glutamate-1-semialdehyde (GSA), a hydrated hemiacetal form of GSA, or a cyclized form of the compound (Houen et al., 1983; Hoober et al., 1988, Jordan et al., 1993)(Fig. 2).The is released in the reaction becoming available for another round of aminoacylation by GluRS. Despite their relatively low abundance within cells, GluTRs have now been purified and characterized from a number of photosynthetic and non-photosynthetic organisms and found to differ considerably with respect to their molecular masses, subunit compositions, and catalytic activities (Weinstein et al., 1987; Chen et al., 1990b; Rieble and Beale, 1991b; Pontoppidan and Kannangara, 1994; Kumar et al., 1996a). The enzyme from C. reinhardtii has been reported to be a monomer with a molecular weight of 130 kDa (Chen et al., 1990b). This contrasts with the purified enzyme from barley which has a molecular mass of 270 kDa and is composed of four to six identical subunits of 54 kDa (Pontoppidan and Kannangara, 1994). Genes encoding GluTR have been isolated and characterized from a number of photosynthetic organisms (Beale, 1993, 1995; Kumar et al., 1996a). Initially, sequences encoding the GluTR activity in plants were identified by their ability to complement hemA mutants of E. coli, which are defective in ALA formation (Ilag et al., 1994). Plant genes encoding

Michael P. Timko GluTR (designated HEMA) have now been isolated from Arabidopsis (Ilag et al., 1994; Kumar et al., 1996b), cucumber (Tanaka et al., 1996), and barley (Bougri and Grimm, 1996) and shown to encode approximately 50 kDa proteins with conserved amino acid sequences. Like many other plastid localized proteins, the plant GluTRs are synthesized outside the plastid as higher molecular weight precursors that are post-translationally imported (Ilag et al., 1994). In Arabidopsis and cucumber two differentially expressed HEMA genes were reported to be present in the nuclear genome (Kumar et al., 1996b; Tanaka et al., 1996), whereas three to four genes are reported to be present in barley (Bougri and Grimm, 1996). The functional significance of having multiple expressed HEMA genes remains to be elucidated. A number of studies have shown that GluTRs recognize the tRNA cofactor in a nucleotide sequencespecific fashion. For example, the E. coli could serve as substrate for the GluTRs from Chlamydomonas and Arabidopsis, but was not recognized by the purified GluTR from barley (Huang and Wang, 1986; Ilag et al., 1994; Pontoppidan and Kannangara, 1994). Specific residues important in with GluTR controlling the interaction of have recently been determined for Euglena gracilis was found where a single point mutation in to uncouple chlorophyll and plastid protein synthesis (Stange-Thomann et al., 1994). The could still be charged by GluRS, but was no longer able to participate in the GluTR reaction. Affinity purified GluTRs from Synechocystis strain PCC 6803 and several other oxygenic prokaryotes were reported to be capable of catalyzing GSA in the absence of formation fromglutamylGluRS and GSA-AT (Rieble and Beale, 1991b). On the other hand, there is evidence that the GluRS and GluTR of Chlamydomonas form a stable complex capable of co-sedimenting during centrifugation through glycerol gradients (Jahn, 1992). Stable complexes were formed only in the presence of or when appropriate combinaglutamyltions of ATP, glutamate, and were included in the assay mixture, indicating that aminoacylatedwas required for complex formation. Addition of NADPH resulted in the reduction of tRNA-bound glutamate and complex dissociation. The formation of such a ternary complex was suggested to be a possible mechanism for channeling specifically to ALA biosynthesis rather than to protein synthesis (Jahn, 1992).

Chapter 20 Pigment Biosynthesis

D. Glutamate-1-Semialdehyde Aminotransferase Glutamate-1 -semialdehyde aminotransferase (EC 5.4.3.8, GSA-AT) catalyzes the final step in ALA formation, the transamination of GSA to ALA (Fig. 2). GSA-ATs have been purified from a variety of organisms and their biochemical properties and catalytic mechanisms have been characterized in considerable detail. It is now generally accepted that GSA-AT converts GSA to ALA in a ping-pong bi-bi group from the C4 mechanism in which the carbon of GSA is transferred to the C5 carbon of ALA, with pyridoxal 5´-phosphate as a cofactor and 4,5-diaminovaleric acid as an intermediate (Kannangara et al., 1994). The enzymes isolated from plants (Grimm et al., 1989), Chlorella (Avissar and Beale, 1989), Chlamydomonas (Wang et al, 1984; Jahn et al., 1991) and Synechococcus strains PCC 6301 (Grimm et al., 1989) and PCC 6803 (Rieble and Beale, 1991a) appear to function as homodimers consisting of 43–46 kDa subunits. The purified enzyme contains a pyridoxal-phosphate cofactor (Avissar and Beale, 1989) bound at a conserved Lys within the proposed active site domain (Grimm et al., 1992) and requires no added substrates other than GSA for activity. The presence of abound pyridoxalphosphate cofactor on the enzyme is consistent with the observation that ALA formation is readily inhibited by gabaculine and aminooxyacetate, known pyridoxal antagonists (Avissar and Beale, 1989). Cloned cDNAs and genomic sequences encoding GSA-AThave now been isolated from barley (Grimm, 1990), soybean (Sangwan and O’Brian, 1993), Arabidopsis (Ilag et al., 1994), and tobacco (Höfgen et al., 1994). At least two distinct genes encoding

383 GSA-AT are present in the nuclear genome in plants, whereas only a single gene appears to be present in Chlamydomonas (Matters and Beale, 1994). The GSA-ATs from the various plants and algae display a high degree of sequence similarity and are similar in primary structure to other members ofthe aspartate aminotransferase enzyme family (Ilag et al., 1994; Matters and Beale, 1994).

IV. The Pathway from ALA to Protoporphyrin IX All available information indicates that the biosynthetic steps of tetrapyrrole formation between ALA and protoporphyrin IX are absolutely conserved among prokaryotic and eukaryotic cells regardless of whether they are photosynthetic. While some characterization of this portion of the tetrapyrrole biosynthetic pathway has been done in Chlamydomonas, a larger part of the information now available on these processes comes from studies involving other organisms.

A. Porphobilinogen Biosynthesis Porphobilinogen synthase (EC 4.2.1.24, PBGS; also known as ALA dehydratase, ALAD) catalyzes the asymmetric condensation of two ALA molecules to form porphobilinogen (PBG), the first pyrrole in the pathway. One proposed reaction scheme for PBG formation (Spencer and Jordan, 1995) is presented in Fig. 3. While the basic catalytic properties of PBGSs isolated from various prokaryotic and eukaryotic organisms are the same, differences in enzyme structure, metal ion requirements for

384 catalysis, and oxygen/sulfhydryl sensitivity have been reported (Jordan, 1991; Jaffe, 1995). All PBGSs are oligomeric proteins ranging in size between 250– 340 kDa (Jordan, 1991; Jaffe, 1995). Although early descriptions of the enzyme isolated from vascular plants reported a homohexameric protein with subunits of40–50 kDa, recent studies ofthe physical properties of the pea PBGS purified from an E. coli strain engineered to overproduce the enzyme showed that the plant enzyme was an octomer, similar to its animal and bacterial counterparts (Cheung et al., 1997). Almost all PBGSs characterized thus far require a or , for activity. In divalent metal ion, either general, mammalian, yeast, and most prokaryotic PBGSs require ions for full activity, while those ofplants and algae require (Jordan, 1991; are found to be Jaffe, 1995). PBGSs utilizing are not. oxygen sensitive, while those utilizing Based upon available biochemical and structural analyses, Jaffe (1995) has proposed that the metal ion selectivity and oxygen sensitivity of the various PBSGs is determined by the three possible metal binding sites (termed A, B and C) found in the enzyme. All mammalian PBGSs use only the A and at each position. Four B sites and coordinate one at the BCys residues contribute ligands to bind site ( ), and at least one His, one Tyr, and one Asp . E. coli, cyanobacteria, residue are ligands to and and most other bacterial ALADs have a site, and also coordinate at a C site whose bound at the C location has yet to be defined. site of the E. coli PBGS functions allosterically to stimulate enzyme activity when the A and B sites are occupied (Mitchell and Jaffe, 1993). In contrast, plant, algal, and some photosynthetic bacteria (e.g., . In these Rhodobacter spp.) coordinate only organisms the four Cys residues comprising the

Michael P. Timko site have been replaced by Asp residues, creating binding (Boese et al., potential ligands for are 1991). In the case of pea PBGS, only two bound per subunit (Cheung et al., 1997). Which sites are occupied by the bound metal in the plant enzyme is not known at this time. Nucleotide and deduced amino acid sequence information is now available for about 16 different PBGSs from various plants, algae, and photosynthetic bacteria (Jaffe, 1995). Only a single gene encoding PBGS appears to be present in the Chlamydomonas nuclear genome (Matters and Beale, 1995a). This gene, designated Alad, encodes a 43 kDa protein with 50–60% sequence identity to the enzymes found in plants. Like the plant enzymes, the Chlamydomonas PBGS contains the highly conserved Asprich domain proposed to be involved in binding.

B. Porphobilinogen Deaminase Porphobilinogen deaminase (EC 4.1.3.8, PBGD; also known as hydroxymethylbilane synthase) catalyzes the deamination and polymerization offour molecules of PBG resulting in the formation of the first tetrapyrrole in the pathway, 1 -hydroxymethylbilane (preuroporphyrinogen), an unstable linear molecule (Fig. 4). During or immediately after its release from PBGD, 1-hydroxymethylbilane is cyclized and isomerized by the enzyme uroporphyrinogen III synthase (UROS) to yield uroporphyrinogen III. Uroporphyrinogen III is the only isoform of the tetrapyrrole capable of being used in subsequent steps of the biosynthetic pathway. In the absence of UROS, the unstable 1 -hydroxymethylbilane cyclizes spontaneously to form uroporphyrinogen I, a biologically irrelevant isomer (Battersby et al., 1979). PBGDs have been purified from various photosynthetic bacteria, algae, and vascular plants, and in

Chapter 20 Pigment Biosynthesis all cases, the enzyme has been found to be a monomer ranging in molecular weight between 34–44 kDa (Spano and Timko, 1991; Jones and Jordan, 1994; Shoolingin-Jordan, 1995). The reaction catalyzed by PBGD is unique in that a dipyrromethane cofactor covalently attached to the enzyme is used as a primer for the subsequent addition of the four substrate PBG molecules of the growing mono- through tetrapyrrole chain (Jordan and Warren, 1987; Hart et al., 1987). The dipyrromethane is covalently attached to the enzyme at a Cys universally conserved in all PBGDs characterized to date (Shoolingin-Jordan, 1995). After completion of tetrapyrrole assembly, the bond between the dipyrromethane and the tetrapyrrole chain is cleaved, releasing the 1hydroxymethylbilane, and leaving the dipyrromethane attached to the enzyme. There does not appear to be any additional metals or cofactors required for enzyme activity. The important structural features in PBGDs, the reaction mechanism, and the molecular factors controlling enzyme formation and activity have been the subject of a recent review (Shoolingin-Jordan, 1995). Genes encoding PBGD have been cloned from a wide variety of species. However, up to this point only a very limited number of genes have been isolated from photosynthetic organisms (i.e., E. gracilis (Sharif et al., 1989), pea (Witty et al., 1993), and Arabidopsis (Lim et al., 1994)). When the deduced primary structure of the plant and algal PBGDs are compared to those from animals and bacteria, only 15–20% overall sequence similarity is observed (Shoolingin-Jordan, 1995). The majority of the conserved residues are involved in catalysis and substrate binding. The crystallization of the E. coli PBGD and determination ofits three dimensional structure at 1.7 Å resolution has greatly assisted in understanding enzyme function and has led to the identification of residues required for assembly of the dypyrromethane cofactor, PBG polymerization, and catalysis (Louie et al., 1996).

C. Uroporphyrinogen III Synthase In the presence of uroporphyrinogen III synthase (co-synthase) (EC 4.2.1.75, UROS), 1-hydroxymethylbilane is cyclized and the D ring of the tetrapyrrole is inverted, forming the uroporphyrinogen III molecule (Fig. 4). The mechanism of cyclization and the nature of the rearrangement that occurs to produce the uroporphyrinogen III isomer containing

385 the ring D inversion has been studied extensively, and it is now generally believed that it is not just the acetate and propionate side chains, but the whole pyrrole ring, that has been turned around (Leeper, 1991; Shoolingin-Jordan, 1995). UROS has been purified from Euglena (Hart and Battersby, 1985), spinach (Higuchi and Bogorad, 1975), and E. coli (Jordan et al., 1988a). The native enzyme purified from these various sources is a monomer of approximately 30 kDa. There is no apparent metal requirement for enzyme activity and upon purification UROS does not appear to contain any bound cofactors. Genes encoding UROS have been isolated from Synechococcus strain PCC 7942 (Jones et al., 1994) and several non- photosynthetic microbes (see Beale, 1995), but not from plants or algae. The cyanobacterial gene encodes a 28 kDa protein that has only 26% sequence identity with the enzyme of E. coli (Jordan et al., 1988b). However, this is not unusual since it has been previously noted that a low conservation of primary protein structure exists among UROSs from other sources (Jones et al., 1994). To ensure that the formation of uroporphyrinogen III is not rate limiting, UROS must be present in excess in the cell in order to cyclize and isomerize the 1-hydroxymethylbilane as rapidly as it is formed, and some type of close physical association must exist between PBGD and UROS. Evidence for complex formation between PBGD and UROS has been obtained by enzyme co-sedimentation studies on sucrose density gradients (Higuchi and Bogorad, 1975) and by affinity chromatography (Frydman and Feinstein, 1974). In both cases, mobility of PBGD was influenced by the presence of UROS. Further evidence for complex formation by the two enzymes comes from the observation that inclusion of UROS for PBG of the in the reaction mixture altered the Euglena PBGD (Battersby et al., 1979) and facilitated the release of 1-hydroxymethylbilane from the Rb. sphaeroides enzyme (Rosé et al., 1988). The first branch point in the tetrapyrrole biosynthetic pathway occurs after uroporphyrinogen III formation and leads to the formation ofsirohemes and corrins. Like other photosynthetic eukaryotes, Chlamydomonas spp. cells contain siroheme as the prosthetic group of the nitrite and sulfite reductases (Crawford, 1995). The conversion of uroporphyrinogen III to siroheme requires the methylation of the tetrapyrrole ring at positions 1 and 3 to form precorrin 2, oxidation of this intermediate to

386 tetrahydroporphyrin (sirodihydrochlorin), and the into the molecule subsequent insertion of (reviewed in Beale, 1993). In some organisms, precorrin 2 is converted to corrins. Corrin biosynthesis (including the formation of vitamin ) has not been described in Chlamydomonas spp., but has been studied extensively in other organisms (Battersby, 1994).

Michael P. Timko identity to the enzymes isolated from nonphotosynthetic organisms (Mock et al., 1995). Like most of the enzymes of tetrapyrrole biosynthesis in photosynthetic eukaryotes, the URODs of tobacco and barley are encoded in the nucleus, synthesized as higher molecular weight precursors in the cytosol, and post-translationally imported into the plastid.

E. Coproporphyrinogen III Oxidase D. Uroporphyrinogen III Decarboxylase The subsequent formation of protoheme and chlorophylls requires that uroporphyrinogen III is decarboxylated at all four acetate side chains. The enzyme catalyzing this decarboxylation reaction is uroporphyrinogen III decarboxylase (EC 4.1.1.37, UROD). The product of the reaction, coproporphyrinogen III, contains four methyl groups in place of the four acetate side chains. In mammalian cells, when substrate concentrations are low, decarboxylation of uroporphyrinogen III takes place beginning with the acetate side chain present on ring D and proceeds in an ordered manner through the acetate side chains of ring A, ring B, and finally ring C (Luo and Lim, 1993). At high substrate concentration the pattern of decarboxylation is random. A similar reaction sequence is expected to take place in photosynthetic organisms. UROD has been purified to near homogeneity from Rb. sphaeroides (Jones and Jordan, 1993) and partially purified and characterized from Euglena (Juknat et al., 1989) and tobacco (Chen and Miller, 1974). Under both denaturing and non-denaturing conditions the enzyme from Rb. sphaeroides behaved as a monomer of approximately 41 kDa, whereas the algal enzyme appeared slightly larger with a molecular weight of about 54 kDa. The tobacco and Euglena enzymes require no metals for activity and are inhibited by sulfhydryl-modifying reagents. The Euglena UROD is reported to be capable of using either uroporphyrinogen III or uroporphyrinogen I as substrate, with uroporphyrinogen III being a much better substrate (Juknat et al., 1989). Genes encoding UROD have been isolated and characterized from Synechococcus strain PCC 7942 (Kiel et al., 1991), tobacco and barley (Mock et al., 1995), as well as from a variety of non-photosynthetic organisms. The deduced sequences of the tobacco and barley URODs are very similar to each other and to the cyanobacterial protein (86% and 73% identical, respectively), but have only between 33% and 50%

Coproporphyrinogen III oxidase (EC 1.3.3.3, CPOX) oxidatively decarboxylates the propionic acid side chains on rings A and B of coproporphyrinogen III, forming vinyl groups at these positions and generating protoporphyrinogen IX as product (Leeper, 1991). In most photosynthetic eukaryotic cells and aerobically-grown prokaryotes, CPOX requires molecular oxygen for its activity. The purified enzyme consists of two. identical 35–37 kDa subunits (Hsu and Miller, 1970; Bogard et al., 1989;Camadro et al., 1986). In mammalian cells, CPOX is associated with the intermembrane space of mitochondria (Grandchamp and Nordmann, 1978), while in S. cerevisiae it is a cytosolic enzyme (Camadro et al., 1986). In plants, CPOX appears to be localized exclusively in the plastid stroma (Smith et al., 1993). CPOXs isolated from plants and algae are reported to be stimulated by metals such as , and , and inhibited by treatment with EDTA. This contrasts with the enzymes isolated from animals and yeast which either require no metal for activity or are stimulated by the addition of or (Medlock and Dailey, 1996). Cloned cDNAs and nuclear genomic fragments encoding CPOX have been isolated and sequenced from C. reinhardtii (Hill and Merchant, 1995) and several plants, including soybean, tobacco, and barley (Madsen et al., 1993; Kruse et al., 1995). The encoded proteins show significant similarity to each other and to other prokaryotic and mammalian CPOXs. The Chlamydomonas enzyme, like those of vascular plants, is synthesized in the cytosol as a higher molecular weight precursor, which is efficiently imported into plastids, processed and localized to the stroma (Madsen et al., 1993; Kruse et al., 1995; Hill and Merchant, 1995). In Chlamydomonas, the expression of CPOX is regulated by the availability of copper in the growth medium (Chapter 31, Merchant). If the amount of copper is not sufficient to support plastocyanin accumulation at the stoichiometry required for photosynthesis, the cell

Chapter 20 Pigment Biosynthesis uses a c-type cytochrome (Cyt ) as an alternative carrier. Formation of this cytochrome de novo requires additional heme formation by the cell and thus induction of CPOX activity.

F. Protoporphyrinogen IX Oxidase Protoporphyrinogen oxidase (EC 1.3.3.4, PPOX) catalyzes the oxidation of protoporphyrinogen IX to protoporphyrin IX, a fully aromatic molecule and the only metal-free porphyrin occurring in the biosynthetic pathway. The oxidation involves the removal of six hydrogen atoms, two from two of the nitrogen atoms and one from each meso-carbon (i.e., C5, C10, C15, and C20) (Leeper, 1991). Although protoporphyrinogen IX undergoes autoxidation when exposed to molecular oxygen, the reaction is enzymemediated in photosynthetic cells. PPOX has been purified to various extents from a number of photosynthetic organisms. The enzyme was first shown to be associated with organellar fractions (a mixture ofplastids and mitochondria) prepared from barley leaves (Jacobs and Jacobs, 1987), and later demonstrated to be present in detergent-solubilized mitochondrial membrane fractions (Jacobs et al., 1989). The enzyme was unequivocally demonstrated to be present in both mitochondria and etioplasts prepared from pea leaves and Arum spadices by Smith et al. (1993), indicating that this step of tetrapyrrole formation, the last reaction in common to chlorophyll and heme biosynthesis, occurs in parallel in these two organellar locations. The mitochondria-associated PPOX from plants was reported to be a 210 kDa oligomer, consisting of 36 kDa subunits (Jacobs and Jacobs, 1987). The subunit molecular weight of the plant PPOX reported by these researchers is considerably smaller than that reported for PPOXs in other organisms (Dailey, 1990). Further conformation of duplicate pathways for protoporphyrin IX formation located in the plastids and mitochondria ofplants came from the cloning of cDNAs encoding organelle-specific forms of PPOX. A cDNA encoding a PPOX from Arabidopsis was isolated by functional complementation of a hemG mutant of E. coli (Narita et al., 1996). The Arabidopsis cDNA encodes a 57.7 kDa protein containing a 60– 70 amino acid residue segment at the amino-terminus, thought to function as the transit sequence required for organelle import. The Arabidopsis PPOX is about 27% identical to the PPOX encoded by the hemY gene in B. subtilis and the Hem Y-like enzymes present

387 in various mammalian cells, but surprisingly shows no similarity to the hemG-encoded enzyme for E. coli which it complements. The lack of similarity between the plant and bacterial PPOXs is thought to reflect the fact that the electron acceptors for the two classes of enzymes are different (Narita et al., 1996). Two distinct cDNAs were isolated recently from tobacco, also by functional complementation of the E. coli hemG mutant (Lermontova et al., 1997). One cDNA encodes a 548 amino acid protein designated PPOX I that contains a 50 amino residues aminoterminal segment potentially required for translocation into the plastid. Upon import into plastids, PPOX I is processed to yield a mature protein of approximately 53 kDa that is equally distribed between the stroma and thylakoid fractions. The second tobacco cDNA encodes a protein (PPOX II) consisting of 504 amino acid residues that is translocated into the mitochondrion with no apparent change in size after import. The plastid- and mitochondria-localized PPOXs from tobacco contain only27.2% identical amino acid residues. The plastidlocalized PPOX I of tobacco is 71.2% identical to the Arabidopsis PPOX, suggesting that the latter is likely a plastid localized form of the enzyme. PPOX is the cellular target for the p-nitrosubstituted diphenyl ether class of photobleaching herbicides, known to cause rapid photooxidative damage in plants (Matringe et al., 1989). A C. rein­ hardtii strain resistant to N-phenylimide and diphenyl ether herbicides has been isolated and shown to contain a dominant nuclear mutation (rs3) thought to affect the interaction between PPOX and the photobleaching herbicides (Oshio et al., 1993; Sato et al., 1994). A 10 kb fragment of genomic DNA containing the entire Rs3 gene has been cloned from C. reinhardtii and shown to confer herbicide resistance upon transformation into wild-type cells (Randolph-Anderson et al., 1998). The Rs3 gene contains a G A base change that results in aVal Met substitution in the PPOX present in the mutant versus wild-type strains. The Chlamydomonas PPOX has approximately 56% identity to the Arabidopsis PPOX within the carboxy-terminal 187 amino acid residues of protein. Following the formation ofprotoporphyrin IX, the tetrapyrrole biosynthetic pathway separates into two main branches that give rise to the (i.e., protoheme and heme) and the (i.e., chlorophyll and its derivatives). The flow of biosynthetic intermediates beyond this point in the

388 pathway is controlled by two enzymes: ferrochelatase, branch, and magnesium operating for the branch. Since chlorophylls chelatase, for the are the predominant porphyrin formed by Chlamy­ domonas spp., their formation will be discussed first.

V.The Magnesium Branch—Chlorophyll Formation The pathway from protoporphyrin IX to chlorophyll a consists of six steps, beginning with the insertion ion into protoporphyrin IX. The of a IX is then esterified to form IX monomethyl ester (Mg-PME). An isocyclic ring (ring E) is formed on the molecule, and there is a subsequent reduction of the 8-vinyl group. This is followed by the reduction of ring D and esterification of the 17-propionate side chain. The primary product of this reaction series is chlorophyll a.

A. Mg Chelatase Insertion of into protoporphyrin IX is catalyzed in an ATP-dependent reaction by the oligomeric IX chelatase (Mg enzyme chelatase). Until recently, little was known about the physical properties ofthe enzyme responsible for the insertion of into protoporphyrin IX or the gene(s) that encode it. Biochemical analyses and cell fractionation studies established early on that the Mg chelatase activity present in vascular plant chloroplasts was composed of at least two separate components, one soluble and one membraneassociated, and that ATP was required for activity (Walker andWeinstein, 1991). Additional information on the physical properties of the enzyme and the subsequent cloning of genes encoding its subunits was provided by studies involving the biochemical and genetic analysis of chlorophyll biosynthesis mutants in plants (von Wettstein et al., 1995) and photosynthetic bacteria (Bauer et al., 1993). We know now that Mg chelatase consists of three distinct protein subunits encoded by three separate genes or gene families. Genes encoding the three subunits of Mg chelatase were initially identified by insertional mutagenesis studies in Rhodobacter spp., where loss of expression of either bchD, bchI, or bchH was correlated with the accumulation of protoporphyrin IX and absence of enzymatic activity

Michael P. Timko (Bollivar et al., 1994a). Unambiguous proof that the bchD, bchI, and bchH gene products constitute a functional Mg chelatase was provided by Gibson et al. (1995), who showed that it was possible to reconstitute enzyme activity in vitro by mixing extracts of E. coli strains independently overexpressing the bchD, bchI, and bchH gene products. Based upon subsequent biochemical studies, the 140kDa bchH gene product has been proposed as the subunit responsible for binding protoporphyrin IX , while a complex prior to the insertion of formed by the 40 kDa bchD and 70 kDa bchI gene products is thought to be involved in ATP binding and metal insertion (Willows et al., 1996). Homologs of the Rhodobacter spp. genes encoding the three Mg chelatase subunits have been identified in cyanobacteria, algae, and plants, and have received a variety of designations. In Synechocystis strain PCC 6803 the bchD, bchI, and bchH homologs are referred to as chlD, chlI and chlH (Jensen et al., 1996a). In barley, they have been designated as Xantha-g, Xantha-h, and Xantha-f (von Wettstein et al., 1995; Jensen et al., 1996b). The ch42 mutation in Arabidopsis(Koncz et al., 1990; Gibson et al., 1996), the CHL1 gene from soybean (Nakayama et al., 1995), and ccsA gene in Euglena and other algae (Orsat et al., 1992; Reith and Munholland, 1993; Jensen et al., 1996b) encode BchI homologs, whereas the olive mutant in Antirrhinum majus (Hudson et al., 1993) encodes a BchH homolog that cross-reacts immunologically with the 150 kDa protein encoded by Xantha-h. While the BchI homologs of A. majus, Arabidopsis, and soybean are encoded in the nucleus, the ccsA genes of E. gracilis, Porphyra purpurea, Olisthodiscus luteus and Cryptomonas are i n the chloroplast genome (Orsat et al., 1992; Reith and Munholland, 1993; Jensen et al., 1996b). In Chlamydomonas, two mutants blocked in their ability to form have been described (Wang et al., 1974). The brc1 mutant is unable to form chlorophyll in the dark, but when grown in the light, accumulates protoporphyrin IX and synthesizes chlorophyll. In contrast, the brs1 strain cannot synthesize chlorophyll in the light or dark. To account for these observations, Wang (1978) reactions proposed that two separate occur in Chlamydomonas, one light-mediated and the other operating in the dark. The mutation in brc1 was proposed to affect only the dark reaction, whereas brs 1 lacks both the light and dark activities. Such a model predicts that mutations affecting only the light

Chapter 20 Pigment Biosynthesis pathway should also be found. The strain grc1 contains such a mutation. This strain is indistinguishable from wild-type when grown in the dark, but accumulates protoporphyrin IX in the light (Wang, 1978). Lightgrown brc1 and grc1 contain less chlorophyll than wild-type cells, consistent with the fact that the cells activity. lack part of their Several other mutants have been described that affect IX formation in C. reinhardtii. These include chl1 (Stolbova, 1971), which most likely represents an allele of brs1, and y-y (Nicholson-Guthrie and Guthrie, 1987), which accumulates large amounts of protoporphyrin IX and is incapable of chlorophyll formation in light or darkness. The nuclear mutation r1 produces no phenotype by itself, but in combination with other mutations blocking chlorophyll formation (e.g., brcl, brs1, y1) raises the level of the accumulated intermediate (i.e., protoporphyrin IX or protochlorophyllide) approximately 20-fold (Wang et al., 1975). This led to the suggestion that r1 may affect ALA formation by reducing the sensitivity of one or more of the pathway enzymes to feedback inhibition (Wang et al., 1975; Wang, 1978).

B. Mg-Protoporphyrin IX Methyltransferase Insertion of into the porphyrin ring is followed by methylation of the 13-propionic group of IX IX forming monomethyl ester (Mg-PME). The enzyme SIX adenosyl-L-methionine: methyltransferase (EC 2.1.1.11, PPMT) catalyzes this reaction using SAM as the methyl donor (Fig. 5). PPMTs have been characterized from several plants,

389 E. gracilis, and the photosynthetic bacterium Rb. sphaeroides, and have been shown to have distinctly different kinetic mechanisms (see Richards, 1993). The PPMTs isolated from plants use a ping-pong mechanism, with the SAM binding first, followed by release of S-adenosyl-homocysteine, before binding IX. The of the second substrate Euglena enzyme is reported to have a random mechanism, whereas the PPMT isolated from Rb. sphaeroides has an ordered mechanism in which the IX is bound first. A single gene, designated bchM, encodes PPMT in Rb. capsulatus and Rb. sphaeroides (Bollivar et al., 1994a; Gibson and Hunter, 1994). Based on mutational studies and biochemical analysis of bchM disruption mutants, it was concluded that the bchM gene product was involved in Mg-MPE formation. However, confirmation of the exact role of the bchM gene product in bacteriochlorophyll formation was obtained only after it was shown that extracts of E. coli over expressing the Rb. capsulatus bchM gene product were able to form a MgPME product with the correct spectroscopic IX and properties using SAM as substrate (Bollivar et al., 1994b). A homolog of the bacterial PPMT gene was identified in Synechocystis strain PCC 6803 by functional complementation of a bchM mutant of Rb. capsulatus (Smith et al., 1996). The cyanobacterial gene, designated chlM, encodes a PPMT that has 29% sequence identity to the Rb. capsulatus enzyme, possibly reflecting the fact that the PPMTs of higher plants, algae and photosynthetic bacteria possess unique reaction mechanisms.

Michael P. Timko

390

C. Isocyclic Ring Formation In the next step of the biosynthetic pathway, IX monomethyl ester is converted ,8-divinylpheoporphyrin (divinyl into protochlorophyllide) by the enzyme IX monomethyl ester (oxidative) cyclase (Mg-PME cyclase) (Fig. 5). In aerobic chlorophyllsynthesizing organisms, the formation of the isocyclic ring involves at least three separate steps in which and both are reaction intermediates in the cyclization process (Castelfranco et al., 1994). In studies have shown that the addition, oxygen atom in ring E is derived from atmospheric (Walker et al., 1989). Much of the initial characterization of Mg-PME cyclase activity was performed using extracts of higher plant chloroplasts (Castelfranco et al., 1994). More recently, the Mg-PME cyclase activity has been examined in Chlamydomonas (Bollivar and Beale, 1995, 1996) and Synechocystis strain PCC 6803 (Bollivar and Beale, 1996). The enzyme from higher plants can be fractionated into two components : a soluble component with a molecular weight of greater than 30 kDa and a labile, membranebound component (Walker et al., 1991; Castelfranco et al., 1994). The Chlamydomonas Mg-PME cyclase is tightly associated with the thylakoid membrane fraction of lysed chloroplasts, and attempts to solubilize it by physical disruption or detergent treatment have been unsuccessful (Bollivar and Beale, 1996). In contrast, the cyanobacterial enzyme could be separated into soluble and membrane-bound components and like the plant enzyme its activity could be reconstituted in vitro by combining the two fractions. The enzymes from Chlamydomonas, Synechocystis strain PCC 6803, and plants require both NADPH (or NADH) and foractivity. Inhibitor studies carried out using the Chlamydomonas enzyme indicated that the algal Mg-PME cyclase does not require a heme or flavin cofactor for activity. This is in contrast to what has been observed for the plant enzyme, that was reported to require ferric heme for activity (Whyte and Castelfranco, 1993). The algal enzyme was found to be sensitive to suggesting a possible role for non-heme iron in the were reported to inhibit reaction. While the in vitro activity of the plant Mg-PME cyclase, a for activity was not tested (Walker requirement of et al., 1991). At the present time, no information is available on

the structure or expression of genes encoding the Mg-PME cyclase from photosynthetic eukaryotes. In Rb. capsulatus, mutagenesis studies showed that insertional disruption of bchM and bchE resulted in the accumulation of Mg-PME by the mutant strains (Bollivar et al., 1994a). The bchM gene has been shown recently to encode the PPMT activity for bacteriochlorophyll formation (Bollivar et al., 1994b), suggesting that bchE may encode the enzyme activity required for isocyclic ring formation. The enzymatic function of the 66 kDa protein encoded by bchE has yet to be determined.

D. Vinyl Reduction In most plants, algae, and photosynthetic bacteria, (bacterio)chlorophyll a exists as a heterogeneous mixture of monovinyl and divinyl forms of the molecule with the final composition of the chlorophyll pool dependent upon the species, growth conditions, and stage of development (Rebeiz et al., 1994). The divinyl forms of chlorophyll a (or its biosynthetic intermediates after Mg-PME) contain vinyl groups at the 3- and 8-positions in rings A and B, respectively, whereas the monovinyl forms contain a vinyl group in the 3-position of ring A and an ethyl group at the 8-position on ring B (Fig. 5). The exact timing of the reduction of the 8-vinyl group and the nature of the enzyme involved are a matter of some debate. It was originally thought that vinyl reduction occurred at the protochlorophyllide stage, with 3,8divinyl protochlorophyllide being reduced to 3monovinyl protochlorophyllide by an 8-vinyl reductase (Whyte and Griffiths, 1993). However, the results from a number of studies showed that other chlorophyll biosynthetic intermediates, including 1X, Mg-PME, and 3,8-divinyl chlorophyllide, can also serve as substrates for vinyl reduction (Richards et al., 1993; Rebeiz et al., 1994). Whether one 8-vinyl reductase with broad specificity or several 8-vinyl reductases are involved in the conversion of 3,8-divinyl tetrapyrroles to 3monovinyl tetrapyrroles is not known. It is also not known ifthe recognized variability in the 3-monovinyl and 3,8-divinyl protochlorophyllide levels in different species reflects species-specific differences in the distribution of putative 8-vinyl reductase activities or results from differences in substrate and/or cofactor availability during development. Some insight into the function of 8-vinyl reductases and possible factors contributing to the size of the monovinyl- and divinyl-chlorophyll pools found in

Chapter 20 Pigment Biosynthesis some cell types comes from studies of bacteriochlorophyll biosynthesis mutants in Rb. capsulatus. Suzuki and Bauer (1995) have shown that mutations in the bchJ locus of Rb. capsulatus cause cells to accumulate 3,8-divinyl protochlorophyllide preferentially, suggesting that bchJ may encode an 8-vinyl reductase. Cells with the bchJ mutation also have reduced total bacteriochlorophyll levels relative to wild-type cells, with the level defined by the amount of 3,8-divinyl protochlorophyllide present. Strains blocked in their ability to reduce protochlorophyllide in a light-independent manner (e.g., bchL, bchN, and bchB mutants) still accumulate a mixed pool of 3monovinyl- and 3,8-divinyl protochlorophyllide; however, double mutants containing defects in bchJ and either bchL, bchN, or bchB, accumulate greater amounts of 3,8-divinyl protochlorophyllide, suggesting that the light-independent protochlorophyllide reductase in Rb. capsulatus may discriminate between substrates, favoring 3-monovinyl protochlorophyllide. One cannot exclude the possibility that a second 8-vinyl reductase is present that uses an earlier divinyl intermediate, or that the product of the bchJ gene interacts with other factors that condition its substrate specificity.

E. Reduction of Protochlorophyllide Two distinct enzyme mechanisms for the reduction of protochlorophyllide to chlorophyllide have evolved in nature. One mechanism is completely dependent upon light for its activity, whereas the second mechanism is capable of operating independently of light availability. Both the lightdependent and light-independent mechanisms for protochlorophyllide reduction are present in wildtype Chlamydomonas cells. The two enzymes presumably share a common pool of 3-monovinyland 3,8-divinyl protochlorophyllide substrates and give rise to indistinguishable reaction products. This, however, remains to be unequivocally determined. A significant amount of information is now available on the structure and expression characteristics of the light-dependent enzyme for protochlorophyllide reduction (Reinbothe and Reinbothe, 1996; Reinbothe et al., 1996a; Fujita, 1996). In contrast, little is known about the enzyme(s) mediating lightindependent protochlorophyllide reduction and its catalytic requirements (Reinbothe et al., 1996a; Fujita, 1996).

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1. The Light-Dependent Reaction The light-dependent reduction of protochlorophyllide is catalyzed by the enzyme NADPH: protochlorophyllide oxidoreductase (EC 1.6.99.1, POR), and involves the overall trans addition of hydrogen to the C17 and C18 positions in ring D of the protochlorophyllide molecule, forming chlorophyllide (Fig. 6). Although the exact role of light in the reaction mechanism is unclear, it is known that the action spectrum for reduction is almost identical to the absorption spectrum of phototransformable protochlorophyllide, indicating that the pigment itself acts as the photoreceptor in this mechanism (Griffiths, 1991). Protochlorophyllide photoconversion most likely involves a single photochemical event in which, after illumination of the pigment-protein complex, the absorbed quantum of light energy brings about the direct transfer of two electrons, possibly as hydride, from the pro-S or ‘B’ face of NADPH to protochlorophyllide (Griffiths et al., 1996). Despite initial reports of a flavin associated with POR (Walker and Griffiths, 1988) and studies suggesting that PORcatalyzed protochlorophyllide reduction involves a flavin intermediate (Nayar and Begley, 1996), recent studies by Martin et al. (1997) have shown that protochlorophyllide photoconversion takes place in the absence of flavin. Both 3-monovinyl- and 3,8divinyl protochlorophyllide can be used equally well as substrates by the plant enzyme, whereas in most cases, protochlorophyll does not appear to be a suitable substrate (Griffiths, 1991; Knaust et al., 1993; Whyte and Griffiths, 1993; Schoch et al., 1995). A single nuclear gene, which we suggest be designated Por1, encodes the POR in C. reinhardtii (Li and Timko, 1996). The Chlamydomonas Por1 gene encodes a 397 amino acid protein, of which the amino-terminal 57 residues comprise the transit peptide required for import into the chloroplast. The Chlamydomonas POR has approximately 65–70% sequence identity to the PORs from higher plants and 52–56% identity to the enzymes from cyanobacteria (Li and Timko, 1996; Fujita, 1996). The greatest amount of variation in primary protein structure among the various PORs is found within the transit peptide of the prePOR and at the aminoterminus of the mature protein. The Chlamydomonas POR, like other higher plant and algal PORs, belongs to an extended superfamily of NAD(P)(H)-dependent dehydrogenases/reductases

392

that share many common structural and mechanistic properties (Wilks and Timko, 1995). These include a highly conserved pentapeptide, Tyr-X-X-X-Lys, that forms part of the active site domain, and a canonical nucleotide binding domain, Gly-X-X-X-Gly-X-Gly, near its ammo-terminal end. The Chlamydomonas FOR also contains the four Cys residues conserved in almost all PORs characterized to date (Li and Timko, 1996). Chemical modification studies (Oliver and Griffiths, 1981) and analysis of POR proteins containing site-directed mutations (H. M. Wilks and M. P. Timko, unpublished) have shown that these Cys residues are involved in catalysis, and that at

Michael P. Timko

least one Cys located within the active-site fold of the protein is absolutely essential for enzyme activity. Seven allelic mutations, mapping to a single nuclear locus designated PC1, were isolated and shown to cause a complete loss of light-dependent protochlorophyllide reduction in C. reinhardtii (Ford et al., 1981, 1983). Although defective in lightdependent protochlorophyllide reduction, pc1 cells retain their capacity for light-independent protochlorophyllide reduction and are capable of synthesizing approximately 52% of wild-type levels of chlorophyll in the dark and about 36% of the wildtype chlorophyll levels in the light. Cells containing

Chapter 20

Pigment Biosynthesis

both pc1 and one of the y mutations (i.e., y5, y7) which block light-independent protochlorophyllide reduction are incapable offorming chlorophyll in the light or dark. Based on the results of cell fractionation experiments showing that a 320 kDa membrane protein aggregate from y7 cells was capable of lightdependent POR activity, whereas similar fractions from pc1 y7 cells were inactive, Ford et al. (1983) concluded that pc1 must encode a defective POR protein. That pc1 encodes a mutant Por1 gene was demonstrated by Li and Timko (1996). These investigators isolated and characterized the genes encoding POR from wild-type and pc1 cells and showed that the Por1 gene in pc1 contained a frame shift mutation in the coding region of the gene. Furthermore, transformation of the wild-type Por1 gene into pc1 y7 cells was sufficient to restore protochlorophyllide photoconversion.

2. Light-Independent Reaction Among the earliest observations leading to the finding that two separate mechanisms for chlorophyll formation exist, was the identification of nuclear mutants in Chlamydomonas and other algae that failed to accumulate chlorophyll in the dark, but were still able to synthesize chlorophyll in the light. The first mutant of this type to be isolated and fully characterized was y1 in C. reinhardtii, given its designation based upon the ‘yellow-in-the-dark’ phenotype of the cells (Sager and Palade, 1954). Grown in the dark, y1 mutants accumulate protochlorophyllide, suggesting that the block in chlorophyll biosynthesis occurs at the protochlorophyllide reduction step of the pathway. However, since y1 mutants are still able to reduce protochlorophyllide and synthesize chlorophyll when grown in the light, it suggests that a separate mechanism for chlorophyll formation exists in wildtype cells that operates concurrent to and separate from the light-dependent mechanism for chlorophyll formation. Furthermore, the light-independent pathway contributes significantly to the total chlorophyll pool in Chlamydomonas, since cells that lack the light-dependent mechanism of protochlorophyllide reduction (e.g., pc1) still accumulate nearly one-half of their wild-type levels of chlorophyll (Ford et al., 1983). The y1 mutation is the most frequently encountered of the ‘yellow-in-the-dark’ mutations in

393 C. reinhardtii, and a number of alleles of this gene have been identified, including y1-a (Wang et al., 1977) and y 1-4, a genetically stable temperaturesensitive allele (Ford and Wang, 1980b, 1982). At least seven other y mutants not allelic with y1 have been identified and have been designated y5 through y10, spy6 and spy200 (Ford and Wang, 1980a,b, 1982). In complementation tests, all of the heterozygous diploids produced from crosses of these various strains have a wild-type phenotype indicating that the mutations are recessive (Ford and Wang, 1980a,b). The exact role of the various y genes and their encoded products in light-independent protochlorophyllide reduction remains uncertain. In addition to the various nucleus-encoded y-genes, we now know that light-independent protochlorophyllide reduction in Chlamydomonas involves the products of three distinct chloroplast-encoded genes, designated chlL, chlN, and chlB. Several independent lines of investigation led to the identification of these genes and their role in chlorophyll formation in the dark (Bauer, et al., 1993; Fujita, 1996). Insertional mutagenesis studies using wild-type Rb. capsulatus cells identified three different loci required for lightindependent bacteriochlorophyll formation in the bacterium. Disruption of any one of these genes, designated bchL, bchN, and bchB, resulted in the loss of bacteriochlorophyll formation and the accumulation of protochlorophyllide by the mutant cells, suggesting that the products of these three genes function at the protochlorophyllide reduction step of the bacteriochlorophyll biosynthesis pathway (Bauer et al., 1993). Subsequently, homologs of all three Rb. capsulatus genes (bchL, bchN, and bchB) were found in either the chromosomal DNA of anoxygenic photosynthetic bacteria and cyanobacteria or chloroplast genomes of non-vascular plants, gymnosperms, and algae capable ofdark chlorophyll formation (Fujita, 1996). In Chlamydomonas spp., the bchL, bchN, and bchB homologs have been designated as chlL (or frxC) (Suzuki and Bauer, 1992; Huang and Liu, 1992), chlN (or gidA) (Roitgrund and Mets, 1990; Choquet et al., 1992) and chlB (Li et al., 1993; Liu et al., 1993; Richard et al., 1994), respectively. No homologs of these genes have yet to be identified in the plastid genomes of angiosperms or the Euglenophyta, consistent with the inability of these organisms to form chlorophyll in the dark. The only exceptions reported thus far are the gymnosperms Ginko biloba and Larix kaempferi,

Michael P. Timko

394 which have all three plastid genes, but do not green in the dark (Fujita, 1996). Whether the chlL, chlN, and chlB genes present in these species are actively transcribed and encode functional proteins has not been determined. While direct biochemical evidence for the involvement of the chlN, chlL, and chlB gene products in light-independent protochlorophyllide reduction in Chlamydomonas has yet to be obtained, a functional role for the chlL and chlB genes has been demonstrated by mutagenesis experiments in which targeted disruption of the genes resulted in the loss of chlorophyll formation in the dark and the accumulation of protochlorophyllide (Suzuki and Bauer, 1992; Li et al., 1993; Liu et al., 1993). In Marchantia, the chlL-encoded protein (ChlL) was immunolocalized to the stromal fraction of chloroplasts (Fujita et al., 1989), whereas the chlB­ encoded protein (ChlB) was reported to be associated with the membrane fraction in cyanobacteria (Fujita et al., 1996). ChlB was also immunolocalized to the thylakoid membranes in synchronized C. eugametos cells, sampled early in the dark phase of the lightdark cycle (Richard et al., 1994). Taken together, these data suggest that the light-independent enzyme may consist of both soluble and membrane-associated components. In Synechocystis strain PCC 6301, lightindependent protochlorophyllide reductase activity was associated with the plasma membrane fraction (Pescheck et al., 1989). The enzymatic activity , was enhanced by required NADPH and menadione, a mediator of electron transport between NADPH and membrane-bound enzymes, and was inhibited by treatment with EGTA. The proteins encoded by chlN, chlL, and chlB (designated ChlN, ChlL, and ChlB, respectively) and their homologs share no sequence similarity to the known light-dependent PORs. However, ChlN, ChlL, and ChlB show significant similarity to the bchX, bchY, and bchZ gene products encoding the chlorophyllide reductase of Rb. capsulatus, an enzyme responsible for reduction of the chlorin ring B in bacteriochlorophyll formation, and to the three subunits of nitrogenase (Fujita et al. 1991, 1993; Burke et al., 1993), a multisubunit enzyme that catalyzes the reduction of to ammonia (Peters et al., 1995). The nitrogenase enzyme consists of two separable components: a MoFe-protein with an structure encoded by the nifD and nifK. genes, respectively, and the Fe-protein, which is a dimer encoded by the nifH gene. The Fe-protein is thought

to be responsible for the transfer of electrons to the MoFe-protein in a reaction that is coupled with the hydrolysis of whereas the MoFe-protein catalyzes the reduction of directly. ChlL and ChlB are approximately 35% and 19% identical to the nifH and nifK gene products, respectively (Fujita et al., 1991; Suzuki and Bauer, 1992; Li et al., 1993), whereas ChlN is approximately 19% identical to the nifD encoded protein (Fujita et al., 1993). ChlL is reported to chromatograph as a homodimer and, like its nifH analog, contains four conserved Cys residues capable of forming a single [4Fe-4S] cluster and two conserved ATP binding domains per homodimer (Suzuki and Bauer, 1992; Fujita et al., 1996). These similarities suggest that the light-independent protochlorophyllide reductase may form a multisubunit complex similar to nitrogenase in which ChlL might function as a specific donor of electrons to a ChlN-ChlB component responsible for protochlorophyllide reduction (Fujita, 1996). As noted previously, the exact role(s) played by the nucleus-encoded y gene counterparts in the process of light-independent protochlorophyllide reduction remains to be determined. The y genes could potentially encode additional subunits of the light-independent protochlorophyllide reductase complex, be proteins required for substrate or cofactor formation, or serve as transcriptional or posttranscriptional regulators of plastid gene expression (e.g., chlN, chlL, and chlB). The involvement of nucleus-encoded factors in the regulation of various aspects of Chlamydomonas chloroplast gene expression is well documented (Rochaix, 1996). Elucidating the function of these genes will likely require their isolation and full characterization.

F. Phytylation The final step of chlorophyll synthesis is the esterification of a long chain alcohol to the propionic acid side chain of ring D. This reaction, referred to as phytylation, is catalyzed by the enzyme chlorophyll synthetase (Fig. 7). The chlorophyll synthetase reaction has been studied extensively in plastids of vascular plants (Rüdiger, 1993) and more recently in Rb. capsulatus (Bollivar et al., 1994c). Both chlorophyllides a and b are suitable substrates for chlorophyll synthetase, whereas protochlorophyllide is not (Helfrich and Rüdiger, 1992; Rüdiger, 1993). In etioplasts, chlorophyllide is esterified initially

Chapter 20 Pigment Biosynthesis

with geranylgeraniol, followed by the stepwise reduction of the geranylgeranyl group to dihydrogeranylgeranyl, tetrahydrogeranylgeranyl, and finally to phytol (hexahydrogeranylgeranyl). In mature chloroplasts, phytol pyrophosphate appears to be the preferred substrate for the enzyme (Rüdiger, 1993). The hydrogenation of geranygeranyl to phytol (Schoch et al., 1980), and requires NADPH and can take place either using pyrophosphates (GGPP phytol pyrophosphate) or pigments (geranylgeranylchlorophyll) as substrates. While chlorophyllide hydrogenation of GGPP occurs in the chloroplast envelope membrane, the hydrogenation of geranylgeranylchlorophyllide most likely takes place at the site of chlorophyllide formation, either in the thylakoid membrane of mature chloroplasts or the prolamellar body and prothylakoid membrane of etioplasts (Soil et al., 1983; Rüdiger, 1993). Two open reading frames have been identified in the photosynthetic gene cluster in Rb. capsulatus

395

that when disrupted result in defects in esterification of chlorophyllide and the subsequent hydrogenation of esterified geranylgeranyl to phytol (Bollivar et al., 1994c). These genes, designated bchG and bchP, are proposed to encode the bacteriochlorophyll synthetase and geranylgeranyl hydrogenase, respectively. A homolog of bchG, designated G4, was isolated from Arabidopsis and shown to encode a 42 kDa polypeptide with 61 % similarity to the Rb. capsulatus protein (Gaubier et al., 1995). Definitive proof that the G4-encoded protein is a functional chlorophyll synthetase remains to be provided.

G. Reaction Center Chlorophylls and Pheophytin The bulk of chlorophyll a and b found in photosynthetic membranes of higher plants and green algae is associated with a family of nuclear-encoded proteins known as the light-harvesting chlorophyll a/b-binding polypeptides (or LHCP) (Hoober et al.,

Michael P. Timko

396 1994). A much smaller fraction of the chlorophyll a pool is used in the assembly of the PS I and PS II reaction centers. From various biochemical and spectroscopic analyses we know that a number of specialized chlorophyll a derivatives are present within the PS I or PS II reaction centers. In the PS I reaction center, a special pair of chlorophyll molecules, termed are present. Attempts to define the nature of these chlorophylls led to the identification of RC I, a 13(S)-hydroxy-20-chloroderivative of chlorophyll a found in PS I preparations from various plants and algae (Dörnemann and Senger, 1986). Whether RC I is the chromophore for or the primary acceptor is not known. However, its role as the primary electron donor in PS I seems unlikely since the amounts of RC I detectable in highly purified PS I preparations appears to be too low relative to levels of observed (Watanabe et al., 1987). Several groups (Watanabe et al., 1985; Kobayashi of et al., 1988) have identified a chlorophyll a, termed chlorophyll a´, in PS I preparations from higher plants, algae, and cyanobacteria. Unlike RC I, chlorophyll a´ and are present in equimolar amounts in these reaction center preparations. More importantly, upon addition of purified chlorophyll a´ to cell-free preparations containing the 65–66 kDa PS I apoprotein, spectral shifts are observed that are similar to those seen for in response to redox state changes and illumination (Watanabe et al., 1987). Although the origin of chlorophyll a´ is not known, it is assumed to be formed from chlorophyll a by spontaneous epimerization during the incorporation of the pigment into the PS I reaction center. Consistent with this, Helfrich et al. (1994) have shown that chlorophyllide a´ is not a substrate for chlorophyll synthetase and therefore chlorophyll a´ formation must occur only after esterification has taken place. The primary electron acceptor of PS II is pheophytin a, a chlorophyll a derivative lacking the atom (Kobayashi et al., 1988). It is chelated generally assumed that the pheophytin a associated with PS II reaction centers is formed by the during insertion of spontaneous loss of the chlorophyll a into the reaction center complex. However, recent studies in higher plants (Litvin et al., 1993; Ignatov and Litvin, 1994) suggest that pheophytin a formation may proceed directly from minor, long-wavelength chlorophyll precursors (e.g., the chlorophyllide precursor, P650).

H. Formation of Chlorophyll b In addition to chlorophyll a, most plants and green algae contain smaller amounts of chlorophyll b. Chlorophyll b differs structurally from chlorophyll a only at the C7 position in ring B, where a formyl group has replaced the methyl group present in chlorophyll a. At least three different types of enzyme(s) have been proposed to be involved in the formation of chlorophyll b: a mixed-function oxygenase with broad specificity, a dioxygenase, or a monooxygenase and dehydrogenase working jointly (Porra et al., 1993). Although the exact reaction sequence leading to chlorophyll b formation is still in question, it is generally believed that chlorophyll b is derived from chlorophyll a or an earlier biosynthetic precursor by oxidation of the methyl group at C7, most likely using molecular oxygen (Schneegurt and Beale, 1992; Porra et al., 1993). Ambiguity surrounding the mechanism for chlorophyll b biosynthesis arises from the inability of numerous in vivo and in vitro studies to resolve whether oxidation of the ring B methyl group occurs before or after phytylation (Beale and Weinstein, 1990). Early in vivo studies clearly indicated that radiolabeled ) chlorophyll biosynthetic precursors (e.g., were rapidly incorporated into chlorophyll a during greening of etiolated plant tissues (Akoyunoglu et al., 1967). In these tissues, chlorophylls a and b accumulated proportionally, with chlorophyll b being formed only after a sufficient pool of chlorophyll a precursors accumulated (Oelze-Karow and Mohr, 1978). Light was not specifically required for chlorophyll b formation, since etiolated seedlings given a brief initial exposure to light continued to form chlorophyll b in the dark. The stimulation of chlorophyll b formation observed upon illumination in some plant species was thought to reflect the fact that light was required for the continued synthesis of chlorophyll a precursors (Oelze-Karow and Mohr, 1978). Several laboratories have also shown that chlorophyll b can be formed from 3-monovinyl protochlorophyllide a, 3,8-divinyl protochlorophyllide a, or chlorophyllide a (Kotzabasis and Senger, 1989; Shedbalker et al., 1991; Rüdiger, 1993). These observations support the earlier findings of Bednarik and Hoober (1985a,b) who showed that Chlamydomonas y1 cells grown at elevated temperatures (38 °C) in the dark in the presence of either o- or m -phenanthrolines secreted chloro-

Chapter 20 Pigment Biosynthesis phyllide b, the unesterified precursor of chlorophyll b, into the surrounding medium at a rate equal to that of total chlorophyll production in the light. Green cells treated in the same manner excreted 3,8-divinylchlorophyllide b, whereas cells grown at 38 °C in the dark without phenanthroline treatment excreted protochlorophyllide. The role of phenanthrolines in chlorophyllide b synthesis is not known. However, Hoober and his colleagues have suggested that phenanthroline may be functioning in place of an endogenous polyaromatic effector molecule, possibly chlorophyll a, required by the chlorophyllide b forming system (Bednarik and Hoober, 1985b; Hoober et al., 1994). Consistent with this suggestion, plastid membrane fractions prepared from darkadapted (yellow) cells were able to catalyze the conversion of exogenously supplied protochlorophyllide to chlorophyllide b when supplemented with phenanthrolines (Bednarik and Hoober, 1985b). Although conversion of chlorophyll b to chlorophyll a was thought to be unlikely because reduction of the formyl group to a methyl group is a thermodynamically difficult reaction, several articles have appeared recently reporting chlorophyll b conversion to chlorophyll a via a 7- hydroxymethyl chlorophyll intermediate (Ito et al., 1996; Ohtsuka et al., 1997). Variations in both chlorophyll a and chlorophyll b structure resulting from modifications introduced at different locations within the porphyrin ring or phytol tail have also been observed in some species (Rebeiz et al., 1994). It is not known at which steps during chlorophyll b biosynthesis that the observed modifications to the porphyrin ring or phytol tail take place, nor what possible functional significance these modifications may have. Genetic analysis of chlorophyll b-less mutants of Chlamydomonas has provided strong evidence that only a single gene product is involved specifically in the conversion of protochlorophyllide b to chlorophyllide b. Analysis of 54 independently-induced chlorophyll b-less mutants revealed that all carried allelic mutations of a single gene, designated cbn1 (Chunayev et al., 1991). Subsequently, 13 linked (e.g., sub1, sub2, sub8) and several unlinked (e.g., sub9) nuclear suppressors of the cbn1 mutation were isolated that resulted in complete recovery of chlorophyll b and neoxanthin formation to wild-type levels (Nikulina et al., 1997). One such suppressor mutation, designated sub9, was reported to confer an enhanced capacity for chlorophyll b formation, suggesting that it directly affected a component

397 involved in the regulation of the chlorophyll b biosynthesis. VI. The Iron Branch—Formation of Heme In Chlamydomonas spp., as in other photosynthetic organisms, the large amounts of chlorophylls a and b required for the assembly and function of the photosynthetic apparatus likely result in a preferential of shuttling of protoporphyrin IX to the the pathway. The remaining portion of this intermediate is used to form protoheme, heme, and their derivatives utilized in the various other porphyrin-requiring processes throughout the cell.

A. Ferrochelatase Ferrochelatase (EC 4.99.1.1) catalyzes the insertion ) into protoporphyrin IX to form of ferrous iron ( protoheme. The enzyme has been purified and characterized from a wide variety of organisms. In most non-photosynthetic eukaryotes, ferrochelatase is a monomeric protein of 40–42 kDa (Beale, 1993, 1995). In S. cerevisiae and mammalian cells, the enzyme is associated with the inner mitochondrial membranes where its activity is regulated by lipids, metals, and porphyrins (see Dailey, 1990). Ferrochelatase activity has been measured in cell-free extracts, and partially-purified membrane fractions from proplastids, chloroplasts, and mitochondria (Little and Jones, 1976). The enzymes from plastid and mitochondrial membrane fractions have similar requirements for activity, but differ slightly in their pH optimum (Porra and Lacelles, 1968; Little and Jones, 1976). Both enzymes are inhibited by Fe-, Mg-, and Zn-protoporphyrins and N-methylporphyrins. Fractionation studies have shown that all of the ferrochelatase activity present in mature chloroplasts is associated with the thylakoid membranes (Matringe et al., 1994). A cDNA encoding ferrochelatase was isolated from Arabidopsis by Smith et al. (1994) by functional complementation of a yeast mutant defective in this enzyme activity. The cDNA encodes a 52 kDa protein with 25–35% sequence identity to ferrochelatases from other organisms. The Arabidopsis enzyme contains a 65 amino acid transit peptide, which was demonstrated to mediate chloroplast, but not mitochondrial, targeting and import. Subsequently, cDNAs have been isolated from barley and cucumber

398 that encode ferrochelatases slightly larger in molecular mass (53.5 kDa and 57.2 kDa, respectively) than that reported to be present in Arabidopsis (Miyamoto et al., 1994). The organellar location of the proteins encoded by these two cDNAs was not determined in this study.

B. Plastidic and Non-plastidic Heme and Heme Derivatives Isolated chloroplasts readily synthesize chlorophyll indicating and heme from or that the enzymes required for both branches of the tetrapyrrole biosynthetic pathway are present and active in these organelles (Fuesler et al., 1984; GomezSilva et al., 1985). Heme is also found elsewhere in the cell, notably in the mitochondrion, endoplasmic reticulum, peroxisomes and glyoxysomes. Since there is no evidence for a mitochondrial ALA synthase activity in photosynthetic eukaryotes except in Euglena, non-plastidic heme found in Chlamydomonas cells and those of other plants and algae must originate from intermediates made either in the plastid and converted to protoheme in the cytosol or mitochondrion, or are formed directly from protoheme exported from the plastid (Thomas and Weinstein, 1990). There is now substantial evidence that the plastids are the sole source of porphyrin intermediates up to coproporphyrinogen III and that protoporphyrinogen IX or a later pathway intermediate is exported from the plastid for the formation of non-plastidic heme (Smith, 1988; Smith et al., 1993). Both PPOX and ferrochelatase activities have been found in mitochondria, suggesting that a portion of the nonplastidic heme pool may be formed in this organelle. However, the location of non-plastidic heme formation remains an open question. Protoheme (heme b) formed within the plastid and retained there for use in various plastid-localized functions is either incorporated directly into various b-type cytochromes (e.g., cytochrome ) present in the photosynthetic electron transport chain or serves as the precursor to heme c, the covalently attached prosthetic group of cytochromes c and f (Beale, 1995; Howe and Merchant, 1995). Attachment of protoheme to the cytochrome occurs by ligation of the vinyl groups of protoheme to cysteine residues on the apoprotein catalyzed by specific cytochrome c-heme lyases. A chloroplast gene, designated ccsA, that has limited sequence identity to bacterial genes

Michael P. Timko required for the biogenesis of c-type cytochromes has been identified in C. reinhardtii and shown by targeted inactivation and complementation studies to be necessary for protoheme attachment to cytochromes and f in this alga (Xie and Merchant, 1996). Protoheme exported from the plastid (or formed in reactions localized in other cellular compartments) has a variety of possible functions. It can be used to form respiratory cytochromes soluble in the cytosol or associated with various intracellular membranes. It can be converted to heme a and used as the prosthetic group of mitochondrial cytochrome oxidases, or it can be covalently attached to proteins, such as the cytosolic-, mitochondrial-, and microbody-localized c-type cytochromes. (Proto)heme may also be used to regulate a variety of processes within the cell, including transcription, translation, import into organelles, the activity of soluble and endoplasmic reticulum-bound cytochrome P450s, and catalase function in the peroxisomes. Unfortunately, there is currently little known about the regulatory functions played by (proto)heme in Chlamydomonas and other photosynthetic eukaryotic organisms.

VII. Light and Metabolic Regulation of Chlorophyll Formation Tetrapyrrole biosynthesis and subsequent chlorophyll accumulation are regulated in Chlamydomonas spp. and other photosynthetic eukaryotic cells by a combination of factors, that include light and metabolism. One of the first recognized control points in these processes is ALA formation. ALA formation is thought to be rate limiting in plants since the exogenous application of ALA to cotyledons of etiolated seedlings leads to a dramatic increase in protochlorophyllide, the only intermediate in chlorophyll formation that normally accumulates to detectable levels in these tissues. Feeding ALA to dark-grown Chlamydomonas y1 cells also results in increased levels of protochlorophyllide (Wang et al., 1977). Such an increase would only be expected if ALA formation was feedback inhibited by one or more of the end-products of the pathway, either protochlorophyllide or protoheme (Beale and Weinstein, 1990). Which enzyme ofALA formation is the likely target for metabolite repression of ALA formation is not known. Purified GluRS from Chlamydomonas was inhibited in vitro over 90% by

Chapter 20 Pigment Biosynthesis the presence of 5 M heme in the assay mixture (Chang et al., 1990), whereas the GluRS from Scenedesmus obliquus was reported to be inhibited by protochlorophyllide and to a lesser extent by chlorophyll a and protoporphyrin IX (Dörnemann et al., 1989). In contrast, physiologically relevant concentrations ofeither heme or protochlorophyllide had no effect on GluRS activity from Synechocystis (Rieble and Beale, 1991a). Feedback inhibition of ALA formation at the GluRS step seems unlikely, since in most organisms the GluRS used for ALA formation is also used for protein synthesis, and the available in the cell level of fully acylated does not change under various growth conditions (O’Neill and Söil, 1990; Kumar et al., 1996a). A more likely target site for feedback control of ALA formation is GluTR. GluTRs have been purified from a variety of photosynthetic organisms and enzyme activity has been shown to be inhibited by physiologically relevant heme concentrations (Rieble and Beale, 1991b; Weinstein et al., 1993; Pontoppidan and Kannangara, 1994). Furthermore, multiple isoforms of the enzyme are reported to be present in plants, at least one of which appears to be specific for photosynthetic cells. This suggests that GluTR may have an important role in controlling ALA formation in response to changing cellular needs for tetrapyrrole precursors (Bougri and Grimm, 1996; Tanaka et al., 1996). , the ChlamyIn the presence of acylated domonas GluRS and GluTR are reported to form a stable complex (Jahn, 1992). Since GluTR competes with EF-Tu for binding of complex formation between GluRS and GluTR may be a possible mechanism for channeling specifically to ALA formation rather than to protein synthesis (Bougri and Grimm, 1996). The binding affinity ofGluTR to GluRS may be influenced by the presence of heme through specific sites in GluTR. Such heme response sites are known to exist in other proteins that use heme as an effector to regulate their activity (Zhang and Guarente, 1995). In Chlamydomonas, neither transcription of trnE1, (Jahn, 1992), nor the plastid gene encoding the level of GluRS (Mau et al., 1992) is significantly altered by light during greening. This is similar to what has been observed in higher plants, where the (Berry-Lowe, 1987) and levels of both GluRS (Bougri and Grimm, 1996) do not appear to change in etiolated seedlings following illumination. Increased levels of GluTR and GSA-AT activity

399 were observed in dark-grown Chlamydomonas cells transferred into the light, with GSA-AT exhibiting the largest increase in enzyme activity (Mau et al., 1992). Light treatment has also been shown to bring about an increase in GSA-AT mRNA levels in light:dark synchronized-cell cultures, with a twofold increase in mRNA abundance observed in the first 0.5 h in the light and a 26-fold increase in message levels found after 2 h of illumination (Matters and Beale, 1994). The observed increase in GSA-AT mRNA in the synchronized cells was comparable to the previously reported increases in GSA-AT activity in dark-grown Chlamydomonas following transfer into the light. This differs slightly from what has been reported to occur during the greening of etiolated plants, where light-induced increases in GluTR and GSA-AT activity were not always coupled to increased levels of GluTR and GSA-AT mRNAs (Grimm, 1990; Sangwan and O’Brian, 1993; Ilag et al., 1994; Bougri and Grimm, 1996). The observed lack of correlation between message abundance and enzyme activity levels was suggested to be indicative of the involvement of post-transcriptional events in the control ALA synthesis (Bougri and Grimm, 1996). The differences observed between greening angiosperm seedlings and synchronized Chlamydomonas cells in the regulation of GluTR and GSA-AT expression in response to light may also reflect physiological differences between the two cell types (Matters and Beale, 1994). Since the cells in etiolated tissues are poised for greening, GluTR and GSA-AT mRNA levels would be expected to be high in order to accommodate the need for rapid ALA formation upon illumination, whereas in cells growing synchronously under light/dark cycles, new chlorophyll synthesis would only be required upon cell division, which would occur only once during the dark phase. It is worth noting that Arabidopsis seedlings grown in dark/light cycles exhibit more dramatic changes in GluTR and GSA-AT mRNA levels, with the abundance of both mRNAs low during the dark period and high in the light (Ilag et al., 1994). The mechanism by which light regulates gene expression involved in ALA formation is not known. In higher plants, ALA formation is thought to be regulated in part by a low fluence phytochrome response (Huang et al., 1989). Both red and blue light treatments were found to be effective inducers of ALA formation in dark-grown wild-type E. gracilis cells (Mayer and Beale, 1990), but only blue light

400 treatment was required to induce the enzymes involved in ALA formation in an achlorophyllous mutant of this alga (Mayer and Beale, 1991). In Chlamydomonas, expression of the nuclear genes encoding GSA-AT (designated Gsa) and PEGS (designated Alad) were shown to be regulated by a carotenoid-type blue-light photoreceptor system rather than by phytochrome, rhodopsin, or a protochlorophyllide-based photoreceptor (Matters and Beale, 1995b). Subsequent studies aimed at defining more precisely the molecular mechanism of blue-light induction of Gsa expression showed that and calmodulin are essential components both of the signal transduction chain leading to increased Gsa transcription (Im et al., 1996). It was also shown that blue-light induction of Gsa transcription likely involved an acetate-dependent mobilization of pools, with acetate most likely acting as internal a source of metabolic energy for the mobilization process. The steps between ALA formation and metal chelation are considered not to be rate limiting. In Chlamydomonas, the level of mRNA encoding PBGS increased about seven-fold upon transfer of synchronously growing cells from dark to light, with a corresponding three-fold increase in PBGS activity (Matters and Beale, 1995ab). By comparison, PBGS, PBGD, and UROS activities have been reported to increase three- to six-fold in etiolated pea leaves during the first 48–60 h of greening (Smith, 1986; Spano and Timko, 1991; Witty et al., 1993). In addition, expression of PBGS, PBGD, and CPOX appears to be regulated to a greater extent by developmental age rather than light, with higher levels of immunodetectable protein and mRNA found early in leaf development in the light or dark compared to later stages when the tissues are completely differentiated (He et al., 1994; Kruse et al., 1995). The chelation of metal by protoporphyrin IX is also likely to be an important regulatory point controlling the flow of intermediates through the pathway. To what extent the plastid-localized ferrochelatase and Mg chelatase compete for available substrate is not known. Based solely upon the abundance of end-product, the majority of protoporphyrin IX appears to be shuttled to the branch of the pathway during chlorophyll formation. Consistent with this observation, Mg chelatase activity is low in etiolated seedlings and increases significantly in the light (Walker and Weinstein, 1991; von Wettstein et al., 1995). This may be the

Michael P. Timko result of altered availability of one or more of the enzyme subunits. For example, the abundance of Xantha-f and Xantha-h transcripts have been shown to increase in etiolated barley seedlings upon illumination (Jensen et al., 1996b). While the Xantha-f transcript increased 18–20-fold upon illumination and remained high for approximately 20 h, the Xantha-h transcript increased only two- to three-fold, peaking at 6 h post-illumination and then gradually declining to dark levels. Both the Xantha-f gene product and Mg chelatase activity exhibited a diurnal fluctuation. Since the protein encoded by Xantha-f is thought to be responsible for insertion binding protoporphyrin IX while the step appears to be catalyzed by other components of the enzyme complex, it is possible that the Xantha-f gene product may function to regulate the flow of branch of the protoporphyrin IX through the pathway (Hudson et al., 1993; Gibson et al., 1995). By comparison, ferrochelatase mRNA was found to be induced only five-fold upon illumination of etiolated Arabidopsis (Smith et al., 1994). This low level of induction matches the rather static production of protoheme by the plastid during this time frame (Beale and Weinstein, 1990). In angiosperms, the light-dependent reduction of protochlorophyllide is the last major control point in the chlorophyll branch of the pathway. Two forms of POR are known to exist in most monocot and dicot species. These two forms, termed PORA and PORB, differ in their pattern of expression, abundance, and activity during light-induced development (Holtorf et al., 1995; Armstrong et al., 1995). PORA is the predominant form of the enzyme present in etiolated tissues and along with its substrates, protochlorophyllide and NADPH, accumulates to high levels in the prolamellar bodies of etioplasts. Since the amounts of PORA protein and the mRNA that encodes it decrease dramatically upon illumination, it is thought that PORA functions only during the very early stages of greening. PORB is thought to be responsible for the reduction of protochlorophyllide during the later stages ofgreening and for chlorophyll formation in mature, green tissues. PORB is present only in minor amounts in the thylakoid membranes of developing and mature chloroplasts and its expression appears to be constitutive throughout development. In contrast, transcription of the PorA gene is negatively regulated by phytochrome (Holtorf et al., 1995; Armstrong et al., 1995). In addition to regulation at the level of transcription, in vitro import

Chapter 20 Pigment Biosynthesis studies performed with radioactively-labeled PORA and PORB precursor polypeptides have shown that import of prePORA is regulated by substrate (protochlorophyllide) availability at the plastid envelope membrane, whereas prePORB import occurs in the absence of protochlorophyllide (Reinbothe et al., 1995). In wild-type Chlamydomonas, the levels of POR protein and the mRNA that encodes it are high in dark-grown cells, but absent or very low in cells grown in the light (Li and Timko, 1996). What factors control Por1 expression and the abundance and activity of its encoded protein remain to be determined. Interestingly, the three plastid genes encoding subunits of the light-independent proto­ chlorophyllide reductase in Chlamydomonas (i.e., chlL, chlN, and chlB) appear to be expressed both in dark- and light-grown cells, with at least one subunit reported to have greater abundance in the light (Li et al., 1993; Fujita, 1996). Further characterization of this enzyme and the abundance and distribution of its subunit components is clearly necessary. Furthermore, how the two different routes for chlorophyll formation are integrated in organisms such as Chlamydomonas and whether they serve specific roles in the formation and assembly of the photosynthetic apparatus and overall plastid development remains an open question. One of the primary roles for PORA in higher plants has been suggested to be photoprotection against photooxidative damage during the initial stages of greening (Reinbothe et al., 1996b). Given that wild-type Chlamydomonas cells have the capacity for light-independent chlorophyll formation, whether the Por1 gene product in Chlamydomonas serves a similar protective function is not known. Cells lacking the ability to photoconvert proto­ chlorophyllide (i.e., pc1) grow more slowly and accumulate chlorophyll to a lesser extent than wildtype cells, possibly reflecting the adverse effects of photooxidative damage. This is certainly the case in pc1 y7 and similar double mutants that completely lack the ability to reduce protochlorophyllide and are incapable of growth in the light (Ford et al., 1983). The reduced levels of chlorophyll formed in pc1 may also reflect the inability of the cells to utilize a portion of their protochlorophyllide pool if, as suggested by Suzuki and Bauer (1995), the lightindependent enzyme is discriminatory for the 3­ monovinyl protochlorophyllide substrate.

401 VIII. Carotenoids

A. The Function and Distribution of Carotenoids Carotenoids are hydrophobic, long-chain isoprenoid compounds synthesized in the chloroplast by the condensation of eight isoprene units. In Chlamy­ domonas spp., as in other photosynthetic organisms, carotenoids have two recognized primary functions. First, they serve as accessory pigments involved in the harvesting of light energy for use in photo­ synthesis. Most carotenoids absorb light energy primarily in the blue region of the visible light spectrum (400–600 nm) and the energy they absorb is readily transferred to chlorophylls. In this capacity, carotenoids act as true accessory pigments to the major light-harvesting and reaction center chlorophyll-protein complexes by trapping radiant light energy in the regions of the spectrum not absorbed by chlorophylls a and b (Armstrong, 1995; Bartley and Scolnik, 1995). Second, carotenoids function as protective agents preventing photooxidative damage of cellular membranes. The protective function of carotenoids derives from the unique ability of colored carotenoids to dissipate excess energy via the xanthophyll cycle and to quench chlorophyll triplet states that arise during light absorption that would otherwise lead to the generation of reactive singlet oxygen. Such compounds are potentially harmful to photosynthetic cells because of their ability to cause membrane peroxidation, protein denaturation, and pigment (porphyrin) bleaching, all of which can lead to cell death (Bartley and Scolnik, 1995; Reinbothe et al., 1996c). The protective role of carotenoids in vivo has been examined primarily through the use of bleaching herbicides, such as norflurazon, that block carotenoid formation. This topic has been discussed in considerable detail elsewhere (Bramley, 1994). Carotenoids and intermediates in their biosynthesis (e.g., FPP, GGPP) are also used in the formation of other important compounds within the cells. These include the phytol tail of chlorophyll, the side chain of heme a, various prenylquinones essential for photosynthesis (e.g., plastoquinone and ), sterols, mating factors, cytokines, and photoreceptor pigments (Britton, 1993; Sandmann, 1994). The mechanism for their formation will not be considered at this time. Sager and Zalokar (1958) published the first chromatographic characterization of carotenoids from wild-type C. reinhardtii (strain 21gr). Among the

402 carotenoids identified were various and and lutein. During subsequent decades, this list has been expanded and refined and we now recognize that Chlamydomonas spp., as well as other members of the green algae (Chlorophyceae), generally have carotenoid compositions similar to those of higher plants with the major components lutein, violaxanthin, and neo­ being xanthin. Smaller amounts of crypto­ xanthin, luteoxanthin, and zeaxanthin are also found along with many other minor components that vary from species to species. The carotenoid composition of C. reinhardtii and the ratio of chlorophyll/ carotenoid is known to be dependent upon a number of factors, including light availability and intensity and nutritional status, and is subject to species specific differences (Francis et al., 1975; Radchenko, 1977a,b). The largest fraction of carotenoids found in plastids is associated with the thylakoid membranes, with the remainder associated with the chloroplast envelope (Britton, 1993; Yamamoto and Bassi, 1996). This localization is consistent with the observation that all of the committed steps for carotenoid formation are localized to the plastid (McGarvey and Croteau, 1995). It appears that all of the enzymes of the pathway are encoded by nuclear genes and imported post-translationally into the plastids (Bartley et al., 1994; Bartley and Scolnik, 1995; Armstrong and Hearst, 1996). It is likely that the overall functioning of the pathway is critically dependent upon the organization of the enzymes into a functional membrane-bound or membrane-associated multi­ enzyme complex localized in the plastid thylakoid or envelope membranes, the former being the more likely location (Britton, 1993; Yamamoto and Bassi, 1996).

B. Pathway for Carotenoid Formation Within the past few years there has been a tremendous increase in our understanding of the processes of carotenoid formation and the factors that control the accumulation and intracellular distribution of the enzymes and substrates involved. The pathway for carotenoid biosynthesis can be considered in three parts (Fig. 8). The first part of the pathway provides the isoprene units that are the biosynthetic building blocks of carotenoids and other important cellular compounds. It begins with the formation of mevalonic acid (MVA), a molecule used in many biosynthetic

Michael P. Timko processes, and leads to the production of GGPP, the direct precursor of carotenoids. The second part of the biosynthetic pathway involves the conversion of GGPP to lycopene. The first committed step in this portion of the pathway is the head to head condensation of two molecules of GGPP to yield phytoene. The subsequent steps are desaturation reactions, that lead to the formation of lycopene. The third part of the pathway consists of a series of cyclization reactions that form the cyclic and and their derivatives (e.g., xanthophylls). Numerous branch points occurring within the pathway, and a variety of enzymes capable of introducing structural modifications, create the wealth of diversity recognized within the total carotenoid pool of the cell.

1. Formation of Isoprene Units and Synthesis of GGPP The biosynthesis of carotenoids begins with the conversion of acetyl-CoA to the active isoprene unit, isopentenyl pyrophosphate (IPP). The initial steps of the pathway involve the condensation of three acetylCoA molecules to produce a compound, 3hydroxy-3-methylglutaryl- CoA (HMG-CoA). The formation of HMG-CoA in non-photosynthetic eukaryotic cells is catalyzed by two separate enzymes, acetyl-CoA acetyltransferase and HMG-CoA synthase (McGarvey and Croteau, 1995). In cyanobacteria, algae, and vascular plants there is considerable evidence indicating that the two reactions are catalyzed by a single enzyme that requires and a quinone as cofactors (Weber and Bach, 1994). HMG-CoA is then converted to MVA in a reaction involving two reduction steps that is catalyzed by the enzyme HMG-CoA reductase. In the reaction, two molecules of NADPH are consumed, one at each step of the reduction process. The MVA is then phosphorylated by the action of two separate kinases (mevalonate kinase and phosphomevalonate kinase) to form 5-pyrophosphomevalonate. The 5-pyrophosphomevalonate product is then decarboxylated by a pyrophosphomevalonate decarboxylase forming pyrophosphate (IPP). IPP is the five carbon isoprene unit used as the building blocks for the subsequent formation of all long-chain isopre­ noids, including carotenoids. To form long-chain molecules, IPP must be first isomerized to dimethylallyl pyrophosphate

Chapter 20 Pigment Biosynthesis

(DMAPP). This reaction is catalyzed by IPP isomerase, an enzyme whose activity was initially characterized from extracts of lysed chromoplasts (Dogbo and Camara, 1987). DMAPP and three IPP molecules are then assembled in a series of l´–4 compound condensation reactions that yield the GGPP. While it was initially thought that one enzyme,

403

a GGPP synthase, was responsible for carrying out all of the condensation reactions from DMAPP to GGPP, it is now clear that GGPP formation is catalyzed by a group of highly related prenyl­ transferases (Dogbo and Camara, 1987; Laferrière and Beyer, 1991). Each isoform existing in the plant appears to have a different substrate and chain length

Michael P. Timko

404 specificity (Croteau and Purkett, 1989). For example, geranylpyrophosphate synthase (GPP synthase) intermediate GPP from IPP and forms the DMAPP, whereas farnesylpyrophosphate synthase (FPP synthase) catalyzes the formation of FPP, a molecule, in a two-step reaction. The enzyme first forms GPP from IPP and DMAPP, and then another IPP is added to yield FPP. GGPP synthase is responsible for generating GGPP, which it does in a reaction involving three separate condensations steps beginning with DMAPP and IPP (McGarvey and Croteau, 1995). The prenyltransferases are all homodimers composed of approximately 37 kDa or subunits and require a divalent metal (either ) for activity. With the exception of HMG-CoA reductase, little is known about the structure of the genes encoding the enzymes involved in the conversion of MVA into IPP in plants and algae. Genes encoding HMG-CoA reductases have been isolated and characterized from a variety of vascular plant species and have been shown to be differentially expressed within various tissues of the plant and at different times during development (McGarvey and Croteau, 1995). The expression of the various HMG-CoA reductase genes has also been shown to be induced by a variety of biotic and abiotic factors, including temperature, light, pathogen attack and wounding (Stermer, et al., 1994). In plants where it has been carefully examined, HMG-CoA reductase activity appears to be regulated in a complex manner that involves differential rates of gene transcription and posttranslational modi­ fication of the enzyme (Stermer et al., 1994; Re et al., 1995). Factors such as intracellular calcium levels, calmodulin, and proteolytic degradation may be important components in the regulatory mechanisms. Both biochemical and immunological evidence exists for multiple isoforms of HMG-CoA reductasekinases in plants (Ball et al., 1994, 1995). These enzymes display a high degree of molecular and biochemical similarity to HMG-CoA reductasekinases found in other organisms.

2. Phytoene Synthase The first dedicated step in carotenoid biosynthesis is the head-to-head condensation of two molecules of GGPP to form phytoene. This reaction is catalyzed by the enzyme phytoene synthase (PSY). Phytoene formation occurs in a two-step reaction in which two molecules of GGPP are first condensed to form the

reaction intermediate, prephytoene pyrophosphate (PPPP). PPPP is then converted into phytoene (Camara, 1993). The PSY from bell pepper has been shown to be a monomeric protein of approximately for activity. The reaction 47 kDa that requires mechanism for this enzyme, including the possible steriochemistry that leads to the recognized cis-trans isomerizations, has been recently described in considerable detail by Sandmann (1994). Genes encoding PSY (designated Psy) have been isolated and characterized from a variety of vascular plants (cf. Bartley and Scolnik, 1995) and photo­ synthetic bacteria (Armstrong, 1994; Armstrong and Hearst, 1996). In tomato, and likely other plant species, PSY is encoded by a small gene family in the nucleus (Bartley and Scolnik, 1993). The deduced amino acid sequences of the PSY proteins from plants and photosynthetic bacteria are similar to each other and are similar to the deduced amino acid sequences ofsqualene synthase and dehydrosqualene synthetase, enzymes responsible for sterol and carotenoid biosynthesis, respectively (Hirschberg and Chamovitz, 1994; Sandmann, 1994). The structural similarity among these proteins is not surprising, since the biosynthetic intermediates used as substrates (i.e., prephytoene pyrophosphate and presqualene pyrophosphate) are analogous molecules (Britton, 1993).

3. Phytoene Desaturation Phyloene, a colorless molecule, is converted by a series of four desaturation reactions to the red colored carotenoid lycopene. In vascular plants, the conversion of phytoene to lycopene takes place via (yellow), and phytofluene (colorless), neurosporene (orange) intermediates. Beginning with phytofluene, each subsequent intermediate contains an additional two conjugated double bonds, such that the number of double bonds present in the molecule increases from three in phytoene to eleven in lycopene. In non-photosynthetic bacteria and fungi, a single desaturase enzyme, encoded by the crtI gene, carries out the four step desaturation of phytoene to neurosporene (Sandmann, 1994). In contrast, two enzymes, phytoene desaturase (PDS) and desaturase, catalyze the desaturation reactions in plants, algae, and cyanobacteria (Hirschberg and Chamovitz, 1994; Bartley and Scolnik, 1995). Phytoene desaturase (PDS) carries out the first two desaturation reactions, converting 15-cis phytoene

Chapter 20 Pigment Biosynthesis to (Beyer et al., 1989). Since the is mostly in the all-trans configuration, the enzyme is also thought to be responsible for the isomerization that occurs at this step as well. The cofactors required by PDS for activity are still unclear, although there is or may be evidence that either necessary. Hugueney et al. (1992) reported the isolation of a full-length cDNA from bell pepper, thought to encode a 56 kDa bifunctional phytoene/ phytofluene desaturase. When the 56 kDa protein was overexpressed in E. coli, the resulting enzyme activity was found to be capable of converting The purified, overexpressed lycopene into enzyme contained a bound cofactor. Genes encoding PDS, designated Pds, have now been characterized from a number of vascular plants species (Bartley and Scolnik, 1995). The produced by PDS is converted to lycopene through two additional desaturation reactions, catalyzed by desaturase (Linden et al., 1994). Little is known about the reaction mechanism for this enzyme. Desaturation of phytoene as a is known to be dependent upon molecular is not possible final electron acceptor, although thought to be directly involved in the reaction (Britton, 1993). Genes encoding two distinct types of desaturase have now been isolated from photosynthetic organisms. One gene, isolated from the cyanobacterium Anabaena strain PCC7120 (Linden et al., 1993), encodes a protein related to a crtI-type phytoene desaturase found in heterotrophic bacteria and fungi that are capable of completing the entire process of lycopene desaturation (Linden et al., 1993, 1994). The other genes, isolated from pepper (Albrecht et al., 1995) and Synechocystis strain PCC6803 (Hirschberg et al., 1997) encode desaturases related to pds-type phytoene desaturases.

4. Cyclizations of Lycopene Cyclization of the end groups found on various carotenoids is now generally accepted to take place at the lycopene level of desaturation. Cyclization is attack on the C2 carbon of a suitably initiated by folded acyclic end group. The alternative loss of a proton from the C6 or C4 carbon gives rise to the or respectively, and results in the production or (Britton, 1993). The of either enzymes responsible for the conversion of phytoene are termed lycopene cyclases into or

405 as a cofactor and (LYCs). The LYCs require are able to use either acyclic lycopene, or its precursor neurosporene, as substrate (Sandmann, 1994). Studies of carotenoid biosynthesis mutants of Arabidopsis thaliana have demonstrated that two different LYCs exist, which are involved in designated and the specific cyclization of lycopene to either or respectively (Cunningham et al., 1996). The and are related structurally, but differ in their enzymatic activity. The adds only a to lycopene forming whereas single adds two forming The the formation of allows for the subsequent formation of minor cyclic carotenoids such as zeaxanthin, violaxanthin, and neoxanthin, while the formation of is required for the production of xanthophylls such as lutein. Nucleotide and deduced amino acid sequences are of Arabidopsis now available for the and of pepper, (Cunningham et al., 1996) and the tomato, and tobacco (Hugueney et al., 1995; Pecker et al., 1996). The amino acid sequences of the plant LYCs are very similar to each other and to the isolated from other species. The significance of the observed conservation in protein structure and the relationship between enzyme structure and function have been discussed in detail elsewhere (Armstrong and Hearst, 1996; Cunningham et al., 1996).

5. Xanthophyll Biosynthesis The and serve as the substrates for the formation of a wide variety of compounds, including the xanthophylls. Xanthophylls are oxy-, hydroxy-, epoxy-, and furanoxy-derivatives formed by various hydroxylation and epoxidation/de-epoxidation reactions which take place within the cyclic end groups of the carotenes. Despite their importance in photosynthetic organisms, little is known about the physical properties of the enzymes involved in xanthophyll formation, their reaction mechanisms, and requirements for activity. Equally little is know about the nature of the gene families that encode these enzymes and how their expression is regulated (for discussion see Sandmann, 1994; Bartley and Scolnik, 1995; Armstrong and Hearst, 1996). With the recent cloning of several genes from vascular plants encoding enzymes involved in xanthophyll formation [e.g., capsathin-capsorubin synthase (Bouvier et al., 1994), hydrolase (Sun et

Michael P. Timko

406 al., 1996), zeaxanthinepoxidase(Marin et al., 1996), and violaxanthin de-epoxidase (Bugos and Yama­ moto, 1996)] considerably more insight into this important biosynthetic process should be forth­ coming.

C. Pathway Regulation Only a limited amount of information is currently available on how carotenoid biosynthesis is regulated at the molecular level in plants and algae. Within the early, non-committed portion of the pathway (i.e., the biosynthetic steps between MVA and GGPP formation), the synthesis and activity of HMG-CoA reductase likely plays a significant role is regulating the flow of metabolites available for carotenoid biosynthesis. The cellular and developmental factors that regulate the expression of HMG-CoA reductase expression and other enzyme activities prior to GGPP formation have been reviewed recently (Stermer et al., 1994; McGarvey and Croteau, 1995). Within the committed portion of the pathway (i.e., GGPP through lycopene), there is now substantial evidence indicating that differential gene expression plays a key role. For example, genes encoding PDS and PS Y in plants are known to be regulated both spatially and temporally during development, and to be under the control of various environmental cues, such as light and temperature (Bartley et al., 1994; Bartley and Scolnik, 1995). Recent studies employing transgenic plants expressing antisense Pds RNAs indicate that transcription of the Pds gene may be subject to endproduct regulation, and that in green tissues, carotenoid and chlorophyll levels are tightly coregulated (Corona et al., 1996). The coregulation of carotenoid and chlorophyll formation is well documented in the literature. In fact, not only are carotenoid and chlorophyll accumulation interdependent, but inhibition of either biosynthesis pathway dramatically affects the synthesis, assembly, and stability of the pigmentbinding proteins of the reaction centers and antennae complexes (Herrin et al, 1992; Plumley and Schmidt, 1995; Thomas 1997). Studies demonstrating that the formation and/or availability of specific carotenoids are important factors governing the overall synthesis and assembly of the photosynthetic apparatus during photomorphogenesis have also been reported (Britton, 1993; Hoober et al., 1994). For example, both biochemical studies and analysis of genetic mutants have established that chlorophyll b, and possibly xanthophylls, are required for integration

and stabilization of the LHCII polypeptides in Chlamydomonas (Hoober et al., 1994; Plumley and Schmidt, 1995). These aspects of photosynthetic membrane assembly are dealt with in other sections of this text. A number of mutants have been isolated and characterized in Chlamydomonas spp. that are deficient in either carotenoid and/or chlorophyll formation (c.f. Harris, 1989). Given the fact that one of the primary roles of carotenoids is to protect porphyrins from photobleaching, at least in some of these mutants, chlorophyll loss is a secondary defect. Considerably more needs to be learned about the synthesis and regulation of both tetrapyrrole and carotenoid formation in Chlamydomonas spp., not only with respect to the involvement of these molecules in assembly of the photosynthetic apparatus, but also their role in other cellular processes.

Acknowledgments I wish to thank Bruce Cahoon, Jeff Skinner, Nikolai Lebedev, Heather Brookman and Anthony Spano for their critical reading of the manuscript and Janel Sennewald for preparing the figures and checking literature citations. This work was supported by a grant from the US Department of Energy (DEFG05­ 94ER20144).

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Michael P. Timko T (1987) Chlorophyll a' in photosynthetic apparatus: Reinvestigation. In: Biggins J(ed), Progress in Photosynthesis Research, Vol I, pp 303–306. Martimis Nijhoff, Boston Weber T and Bach TJ (1994) Conversion of acetyl-coenzyme A into 3-hydroxy-3-methylglutaryl coenzyme A in radish seedlings: Evidence of a single monomeric protein catalyzing double condensation reaction. a Biochim Biophys Acta 1211: 85–96 Weinstein JD, Mayer SM and Beale SI (1987) Formation of from glutamie acid in algal extracts: Separation into an RNA and three required enzyme components by serial affinity chromatography. Plant Physiol 84: 244–250 Weinstein JD, Howell RW, Leverette RD, Grooms SY, Brignola PS, Mayer SM and Beale SI (1993) Heine inhibition of synthesis is enhanced by glutathione in cell-free extracts of Chlorella. Plant Physiol 101: 657–665 Whyte BJ and Castelfranco PA (1993) Further observations on the Mg-protoporphyrin IX monomethyl ester (oxidative) cyclase system. Biochem J 290: 355–359 Whyte BJ and Griffiths WT (1993) 8-Vinyl reduction and chlorophyll a biosynthesis in higher plants. Biochem J 291: 939–944 Wilks HM and Timko MP (1995) A light-dependent comple­ mentation system for analysis of NADPH:protochlorophyllide oxidoreductase. Identification and mutagenesis of two conserved residues that are essential for enzyme activity. Proc Natl Acad Sci (USA) 92: 724–728 Willows RD, Kannangara CO and Pontoppidan B (1995) involved in recognition by barley Nucleotides of chloroplast glutamyl-tRNA synthetase and glutamyl-tRNA reductase. Biochim Biophys Acta 1263: 228–234 Willows RD, Gibson LCD, Kannangara CG, Hunter CN and von Wettstein D (1996) Three separate proteins constitute the magnesium chelatase of Rhodobacter sphaeroides. Eur J Biochem 235: 438–443 Witty, M, Wallace-Cook ADM, Albrecht H, Spano AJ, Michel H, Shabanowitz J, Hunt DF, Timko MP and Smith AG (1993) Structure and expression of chloroplast-localized porpho­ bilinogen deaminase from pea (Pisum sativum L.) isolated by redundant PCR. Plant Physiol 103: 139–147 Xie Z and Merchant S (1996) The plastid-encoded ccsA gene is required for heme attachment to chloroplast c-type cytochromes. J Biol Chem 271: 4632–4639 Yamamoto HY and Bassi R (1996) Carotenoids: localization and function. In: Ort DR and Yocum CF (eds) Oxygenic Photosynthesis, pp 539–583. Kluwer Academic Publishers, Dordrecht Zhang L and Guarente L (1995) Home binds to a short sequence that serves a regulatory function in diverse proteins. EMBO J 14:313–320

Chapter 21

Glycerolipids: Composition, Biosynthesis and Function in Chlamydomonas Antoine Trémolières Institut de Biotechnologie des Plantes, Université Paris-Sud, Bâtiment 630, 91405 Orsay Cedex, France

Summary 415 Introduction 416 II. Glycerolipid and Fatty Acid Composition of Chlamydomonas. 417 A. Fatty Acid Composition 417 B. Lipid Class Composition and Sub-Cellular Localization 417 C. Fatty Acid Composition of Lipid Species 419 III. Lipid Metabolic Pathway in Chlamydomonas spp. 422 IV. In vivo Modifications of Lipid Composition in Chlamydomonas 425 V. Mutants Affected in Lipid Composition 426 VI. Involvement of Lipids in the Functional Organization and the Biogenesis of the Photosynthetic Apparatus. 428 Acknowledgments 429 References 429

Summary The lipid composition of Chlamydomonas spp. is described. These algae contain all the fatty acids characteristic of higher eukaryotic photosynthetic tissues and also some unusual fatty acids. The chloroplast membranes contain galactolipids, sulfolipid and phosphatidylglycerol as do all photosynthetic membranes. Phosphatidylethanolamine and phosphatidylinositol are found outside the chloroplast, but phosphatidylcholine is absent. Rather, an unusual betaine-containing lipid, diacylglycerol-trimethyl-homoserine, is present. The fatty acid distribution on the sn-1 and sn-2 carbons of the glycerol in different lipid classes is used to propose a scheme for the organization of glycerolipid metabolism in Chlamydomonas spp. in which most of the polyunsaturated fatty acids in the chloroplast are synthesized by the intrachloroplastic desaturase and acylase pathways. A set of experiments which demonstrate the ability of Chlamydomonas spp. to take up lipids from liposomes added to the medium, and consequently to change fatty acid and lipid composition is presented. Mutants affected in lipid composition are described. Four of them are affected in the biosynthesis of phosphatidylglycerol, one is affected in the synthesis of the sulfoquinovosyldiacylglycerol, while another shows a reduced level of chloroplastic poly-unsaturated fatty acids. Finally, evidence for the involvement of certain lipids such as phosphatidylglycerol or sulfoquinovosyl-diacylglycerol in some steps of thylakoid membrane biogenesis are presented.

J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 415–431. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

416 Introduction The lipid composition of Chlamydomonas spp. is characteristic of a typical eukaryotic photosynthetic organism. Therefore, it is a good model for the study of the organization of glycerolipid metabolism in eukariotic photosynthetic tissues, or the role of some specific lipids in membrane biogenesis or other cellular processes. However, several specific differences in lipid composition between Chlamy­ domonas spp. and other eukaryotic phototrophs present opportunities to study some unique aspects of lipid metabolism in Chlamydomonas spp. The most important difference is the absence of phosphatidylcholine (PC) and the presence (perhaps to replace PC) of diacylglycerol-trimethyl-homoserine (DGTS). DGTS contains betaine but no phosphoryl group and is found in several Chlorophyceae. Another difference is that monogalactosyldiacylglycerol (MGDG), which constitutes the main chloroplastic lipidic species in Chlamydomonas spp. contains a huge amount of hexadecatetraenoic acid — an unusual highly unsaturated fatty acid. Other differences are found in the extra-chloroplastic lipid species in that: 1) cis-vaccenic acid, an isomer of an octadecenoic fatty acid found in trace amounts in most eukaryotic organisms, is quite abundant in Chlamydomonas spp. and 2) an unusual octadecatrienoic acid, octadecatrienoic acid found only in pine trees (Gresti et al., 1996) is also found in Chlamydomonas spp. Although the lipid composition of Chlamydomonas spp. is now well documented, few metabolic, enzymatic or genetic studies of lipid metabolism in this organism have been undertaken.

Abbreviations: ACP – acyl carrier protein; sub-units of the ATP synthase complex; C16:0 – palmitic acid; or hexadecamonoenoic acid; – Trans- 3-hexadecenoic acid; C16:2 – hexadecadienoic acid; C16:3–hexadecatrienoic acid; C16:4–hexadecatetraenoic acid; C18:0–stearic acid; –oleic acid; –cis-vaccenic acid; C18:2–linoleic acid; – octadecatrienoic acid; -linolenic acid; C20:4 – eicosatetrienoic acid; DGDG – digalactosyldiacylglycerol; DGTS – diacylglycerol-trimethyl-homoserine; LHC I – Light harvesting chlorophyll protein complex associated with PS I; LHCII – Light harvesting chlorophyll a+b protein complex associated with PS II; MGDG – monogalactosyldiacylglycerol; PC–phosphatidylcholine; PE – phosphatidylethanolamine; PI – phosphatidylinositol; – polyphosphoinositols; PG – phosphatidylglycerol; PS I, PS II – Photosystem I and II, respectively; SQDG – sulfoquinovosyldiacylglycerol

Antoine Trémolières As a consequence, we are far from a full understanding of the organization of lipid metabolism in Chlamy­ domonas spp. and its genetic control. The metabolic pathways established for higher eukaryotic photosynthetic organisms are assumed to occur in Chlamydomonas spp. However, the desaturases leading to the synthesis of cis-vaccenate and octadecatrienoic acid, uncharacterized with respect to sequencing, remain to be characterized at the biochemical and genetic levels, and the pathway for the biosynthesis of DGTS has been only partly elucidated. A small set of C. reinhardtii mutants generated by random mutagenesis and affected in lipid composition has been isolated. These mutants constitute good tools to study the role of some lipids involved in different cellular processes. It is possible to manipulate the lipid composition in vivo by feeding Chlamydomonas spp. with liposomes. Despite the barrier of the cell wall, lipids enter the cells rapidly where they can be either metabolized or integrated into intra-cellular membranes. This technique is not only a powerful tool to study different aspects of lipid metabolism or function but is also a tool to study intracellular lipid trafficking. The ability of Chlamydomonas spp. to ingest lipids has never received any attention although it is possible that exogenous lipids could be used by the algae as a reduced carbon source. Lipid intracellular signaling pathways which exist in animal cells (and which probably are also present in plant cells) have been characterized in Chlamy­ domonas spp. One such path way starts from mediatorinduced cleavage of the poly-phosphoinositols by a phosphatidylinositol-specific phospholipase C, producing diacylglycerol and phosphatidylinositoltri-phosphate which acts as a second messenger in a complex set of interactions with several kinases (Musgrave et al., 1992). Another signaling pathway involves the cleavage of phospholipids by phospholipase D to produce phosphatidic acid which plays a regulatory function in intracellular calcium flux. Recently, a new lipid, diacylglycerolpyrophosphate, which is involved in cellular signaling has been discovered in Chlamydomonas spp. (Munnik et al., 1996). Hence, Chlamydomonas spp. constitute a fascinating model to study the metabolism and function of cellular lipids at both the biochemical and genetic levels.

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417

II. Glycerolipid and Fatty Acid Composition of Chlamydomonas.

A. Fatty Acid Composition The total fatty acid composition of C. reinhardtii strain 137c analyzed by Giroud et al. (1988), is given in Table 1. The components are C16:0; two isomers of C16:3

(tentative structures proposed: 7,10 and ,10,13); C16:4 (tentative structure proposed:

7,10,13); C18:0; and {The fatty acid was previously thought to be acid, which is generally found in animal cells but it has been demonstrated that it is a rare isomer of octadecatrienoic acid with its first double bond at the position (Giroud et al., 1988)}. Longer chains (not reported in the Tables) were also found at lower abundance. For example, in gametes of C. eugametos C22:l and C20:4 fatty acids were found. The structures of these long unsaturated fatty acids have not been elucidated but the C20:4 was shown to be distinct from arachidonic acid (Brederoo et al., 1991). 2-hydroxy fatty acids are also found in Chlamydomonas spp. The most abundant of these is 2-hydroxyhexadecanoic acids but longer-chain 2-hydroxy acids are also found, including substantial amounts of the C26 compound 2-hydroxyhexacosanoic acids (Matsumoto et al., 1984). Thus, all the fatty acids characteristic of higher eukaryotic photosynthetic tissues are found acid in Chlamydomonas spp. including which represents up to 70% in leaves of higher plants (James and Nichols, 1966) and which plays a crucial role in the stabilization of the major photosynthetic antenna LHC II in a trimeric state (Trémolières and Siegenthaler, 1997). Chlamydomonas spp. also contain quite unusual fatty acids such as C16:4, which is concentrated in MGDG, cis-vaccenic acid which is largely characteristic of bacteria and found only in trace which amounts in most higher plants, and is found otherwise only in pine seeds (Gresti et al., 1996).

B. Lipid Class Composition and Sub-Cellular Localization The structures of the glycerolipids found in Chlamydomonas spp. are shown in Fig. 1. Until 1988

it was not clear if PC was present in C. reinhardtii, but a careful analysis (Giroud et al., 1988) has proven that this phospholipid, found in almost all eukaryotic organisms, is absent. Among the Chlorophyceae studied so far, only C. reinhardtii (Eichenberger, 1982), C. eugametos (Brederoo et al., 1991) and Volvox carteri (Moseley and Thompson, 1980) are devoid of this phospholipid. The betaine lipids (a homoserine containing lipid-diacylglycerol-trimethyl-homoserine- and an alanine containing ) which have never been found in angiosperms, represent a prominent group ofunusual glycerolipids produced by almost all cryptogamic plants (Eichenberger, 1982; Sato and Furaya, 1984, 1985; Dembitsky and Rozensvet, 1989; Dembitsky et al., 1990; Eichenberger, 1990). In the Volvocales, all strains analyzed so far contain diacylglyceroltrimethyl-homoserine but lack (Janero and Barnett, 1982 a; Eichenberger, 1993). Since its identification, DGTS (Brown and Elovson, 1974), is presumed to substitute for PC, based on the zwitterionic structure which is characteristic of both lipids. All the lipid characteristic of photosynthetic

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membranes from cyanobacteria to land plants are also found in the thylakoid membranes of Chlamy­ domonas spp.—specifically, MGDG, DGDG, SQDG and PG. Outside the chloroplast, in addition to DGTS, PE and DPG are found. DPG is a lipid specifically associated with the mitochondrial inner membrane in all eukaryotic organisms so far analyzed. Janero and Barnett (1982 b) identified DPG in C. reinhardtii but they did not localize it to a specific cellular fraction. It is not known whether DGTS is totally extrachloroplastic or if some is localized in the chloroplast envelope. DGTS was detected in the chloroplast envelope of C. reinhardtii by MendiolaMorgenthaler et al. (1985), but it was difficult to insure that the chloroplast envelope fraction was fully devoid of any extra-chloroplastic membranes such as endoplasmic reticulum fragments. Finally, phosphatidylinositol (PI) and its polyphosphorylated derivatives have been found in Chlamydomonas spp. These lipids, associated with intra-cellular signaling pathways in animal cells, were found to be concentrated in the plasma membrane (Brederoo et al., 1991). The typical lipid class composition of Chlamydomonas spp. is given in Table 2.

C. Fatty Acid Composition of Lipid Species The fatty acid composition, their positional distribution and the composition of molecular species glycerolipids of Chlamydomonas spp. are given in Tables 3 and 4 (Giroud et al., 1988). MGDG is highly unsaturated with more than 70% polyunsaturated fatty acids. The sn-1 position is esterified mainly with while almost all (95%) of the C16:4 is esterified at the sn-2 position. In DGDG, two fatty esterified at the sn-1 acids arepredominant: position and C16:0 found to be the main fatty acid at both positions (83%). PG contains a high percentage of a special fatty acid found exclusively in eukaryotic photosynthetic organisms (Dubacq and Trémolières, 1983): is found specifically esterified at the sn-2 position. and are concentrated in DGTS and in PE where they are esterified preferentially at the sn-2 position. is found mainly at the sn-1 position of PE and PI. Nevertheless, significant amounts of this fatty acid can be found in PG, SQDG, and traces are present in the two galactolipids. It is interesting to note that and C16:4 are localized almost exclusively in the two chloroplastic galactolipids, while both and C18:4 are virtually limited

419

to the extra-chloroplastic lipids, DGTS and PE. The distribution of fatty acids between the sn-1 and sn-2 positions of a glycerolipid in plants is generally considered to reflect its biosynthetic origin (Frentzen, 1986; Roughan and Slack, 1982, 1984). Lipids in which the sn-2 position is occupied by a C16 acid are assumed to have a chloroplastic (prokaryotic) origin, those with C18 acid at this sn-2 position have an extra-plastidic (eukaryotic) origin. Application of this principle to the lipids of Chlamydomonas spp. has led to the proposition that MGDG, DGDG and SQDG are synthesized almost exclusively by the chloroplast. When the same principle is applied to PG, it appears that only 88% of this lipid is produced by the chloroplast. This agrees with the fact that PG has been shown to be synthesized also outside the chloroplast (Marshall and Kates, 1972). But the question of whether PG is located outside the chloroplast and if some chloroplastic PG could be imported from the cytosol remains open. Recent results obtained with a mutant of C. reinhardtii impaired in the chloroplastic synthesis of PG will be described later in Section IV. The results suggest that a fraction of PG found in the chloroplast comes from the cytosol. On the other hand, both DGTS and PE are supposed to be produced almost exclusively outside the chloroplast. The cytosolic origin of DGTS has been reported already for the fern Adiantum capillus veneris by Sato and Furuya (1983). The ‘prokaryotic’ proportion of these lipids can be explained by a less pronounced fatty acid specificity of the microsomal monoacylglycerol-3-P-acyltransferase (Frentzen et al., 1984). Nevertheless, the possibility that some synthesis of DGTS occurs in the chloroplast envelope cannot be ruled out. It is interesting to note that in Chlamydomonas spp., 86% of PI displays a ‘prokaryotic structure,’ suggesting that it is synthesized in the chloroplast.

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421

422 Since PI of Arabidopsis has been shown to be mainly ‘eukaryotic’ (Browse et al., 1986), and the enzymes synthesizing this lipid are situated in the endoplasmic reticulum of castor bean endosperm (Moore et al., 1973), the site of its synthesis in Chlamydomonas spp. could differ from that of land plants. The (a fatty acid found only in distribution of trace amounts in most higher plants) is also interesting to examine. This isomer is almost completely absent in the two chloroplast galactolipids and concentrated in the extra-chloroplastic PE, where it is esterified preferentially at the sn-1 position. This suggests that the desaturase leading to the formation of this fatty acid functions in the endoplasmic reticulum. The is found also in PG and SQDG fact that suggests that a small fraction of these lipids could be is also produced outside the chloroplast. concentrated in PI ( at the sn-1 position) and this fact can be explained ifwe imagine the following scheme for the biosynthesis of PI: diacylglycerol with a prokaryotic structure is first formed in the chloroplast and then exported into the cytosol where the final steps of the biosynthetic pathway occurs. This consists of the formation of CDP-diacylglycerol, the replacement of the CDP group by an inositol, and the desaturation ofthe 18:0 fatty acid at the sn-1 position to yield

III. Lipid Metabolic Pathway in Chlamydomonas spp. Although the lipid composition of Chlamydomonas spp. is very well documented, metabolic, enzymatic and genetic studies of lipid metabolism in this organism are in their infancy. As a consequence, we do not have a full understanding of the organization of lipid metabolism and its genetic control. In general, the metabolic pathways established for higher eukaryotic photosynthetic organisms have been assumed to exist in Chlamydomonas spp. as well. Based on the extensive analysis of lipid molecular species structure and sub-cellular localization (described in the previous paragraph), the results of a number ofmetabolic and enzymatic studies of fatty acid metabolism in Chlamydomonas spp., plus our present understanding of fatty acid metabolism in other eukaryotic tissues, we can propose a hypothetical but reasonable picture of the organization of glycerolipid metabolism in Chlamydomonas spp. This model provides a useful framework for future genetic studies.

Antoine Trémolières The organization of the glycerolipids in leaf tissues of higher plants can be summarized as follows (for a review, see Roughan and Slack, 1982): in plants, de novo fatty acid synthesis occurs in the plastid by a type II fatty acid synthetase (a prokaryotic type of fatty acid synthetase which uses ACP as the acyl carrier). Therefore, the plastid is the only source of fatty acids (in contrast to yeast and mammalian cells where a type I fatty acid synthetase is located in the cytosol). Plastid fatty acid synthetase produces mainly palmitoylACP and stearoylACP. The latter is largely desaturated by the stromal stearoylACPdesaturase to give oleoylACP. A fraction of the acyl-ACP produced by the plastidic fatty acid synthetase is used for acylation of glycerol-3-phosphate through the action of two acylases. The sn-1 glycerol-3-phosphate acyltransferase located in the stroma acylates the sn­ 1 position with C16 and C18 acyl chains producing lysophosphatidic acid, while an sn-2-lysophosphatidic acid acyltransferase located in the chloroplast envelope, which shows a high selectivity for C16 acyl chains, esterifies the sn-2 position. The phosphatidic acid so produced is converted into diacylglycerol and these diacylglycerols are used either for galactolipid synthesis through the action of UDP-galactosyl transferase or for the synthesis of SQDG and PG. PG is synthesized through the CDPdiacylglycerol pathway. Lipids formed inside the chloroplast are used as substrates for chloroplastic and desaturases located in the chloroplast envelope to produce poly-unsaturated lipid species. A fraction of the palmitoylACP and OleoylACP formed by the chloroplast fatty acid synthetase is converted into AcylCoA and exported to the cytoplasm. OleoylCoA is the preferentially exported species, but the mechanism of export is not known. AcylCoA exported from the chloroplast is used in the endoplasmic reticulum for the synthesis of extrachloroplastic membrane lipids. Cytosolic acylases show a markedly distinct acyl chain selectivity compared to chloroplastic acylases; the former acylate the sn-2 position with C18 acyl chains (while the sn-1 position can be acylated with both C18 and C16 acyl chains). A distinct set of extrachloroplastic desaturases, located in the endoplasmic reticulum, use mainly PC as a substrate to produce extrachloroplastic poly-unsaturated lipids. The acyl chain length selectivity of the two chloroplastic acylases is the same as that found in cyanobacteria. They are responsible for the synthesis (through the intrachloroplastic pathway) of lipids with either C18 or C16 acyl chains at the sn-1 position, but with only

Chapter 21 Glycerolipids C16 acyl chains at the sn-2 position. However, most lipids (and especially galactolipids) of land plant chloroplasts contain C18 acyl chains at both the sn-1 and sn-2 positions. A cooperative pathway between cytosol and chloroplasts is involved in the formation of these lipid species. PC is likely the C18 sn-1- C18 sn-2 diacylglycerol donor for the UDP-galactosyldiacylglyceroltransferase in the chloroplast envelope of land plants (Oursel et al., 1987). The cooperative organization of the chloroplastic (prokaryotic) and extra-chloroplastic (eukaryotic) pathways for the synthesis of glycerolipids assures the correct accumulation of all lipid classes while allowing great flexibility. Nevertheless, the lack of PC in Chlamydomonas spp. raises an important question. PC plays a crucial role in plant lipid metabolism. It is the main substrate for extrachloroplastic desaturases (Roughan and Slack, 1982), and it is probably a key player in the chloroplast envelope for both export and import of fatty acids from the cytosol to the chloroplast. The function of PC in fatty acid transport is well evidenced by the exceptionally high turnover rate of its fatty acids. A study of the incorporation of labeled acetate, palmitate and oleate in Chlamydomonas spp. by Giroud and Eichenberger (1989) led these authors to propose that DGTS could play the same role in Chlamydomonas spp. as does PC in land plants. This hypothesis is supported by several other studies of lipid metabolism in Chlamydomonas spp. (Seras et al., 1989; Grenier et al., 1991). Nevertheless, it remains to be proven that DGTS is present in the purified chloroplast envelope, as is the case for PC in vascular plants. Sirevåg and Levine (1972) identified a fatty acid synthetase activity in extracts of Chlamydomonas. In contrast to Euglena for which two distinct fatty acid synthetase activities were reported (Delo et al., 1971), only a single activity, dependent on added acyl carrier, was detected in C. reinhardtii extracts. Therefore, it can be concluded that in Chlamy­ domonas spp., as in land plants, all de novo fatty acid synthesis occurs in the plastid. Glycerol-3-Phosphate acyltransferase and lysophosphatidate acyltransferase have been localized to the chloroplast envelope, in association with the thylakoid membrane and pyrenoid tubules (Jeselma et al., 1982; Michaels et al., 1983). Galactosyl transferase was also shown to be associated also with the chloroplast envelope (Mendiola-Morgenthaler et al., 1983) while Hoppe and Schwenn (1981) reported the association of

423 SQDG synthesis with thylakoid membranes. Together all these observations strongly suggest that, in Chlamydomonas spp., the scheme of glycerolipid metabolism organization is similar to that found in land plants. Examination of Tables 2 and 3 suggest that the prokaryotic pathway for lipid synthesis by the intrachloroplastic enzymes has a more prominent role in Chlamydomonas spp. in comparison to most land plants. Therefore, with respect to acylase selectivities, it can be concluded that almost all galactolipids and sulfolipids are built in this way. Nevertheless, the fact that 1) a significant amount of (produced probably in the cytosolic compartment) is found in the sulfolipid, and 2) traces of this fatty acid are always detected in galactolipids indicates that Chlamydomonas spp. have the same ability as land plant photosynthetic tissues to use cytosolic fatty acids, via the eukaryotic pathway, for the synthesis of chloroplastic lipids. The PG also appears to be produced primarily in the chloroplast. Only about 10% of the PG is attributed to the eukaryotic pathway. The pathway of PI biosynthesis is highly unusual because although it is located mainly outside the chloroplast, the lipid has a pronounced prokaryotic structure. DGTS and PE have a largely eukaryotic structure, which is in good agreement with their roles in lipid metabolism in the cytosolic compartment. Metabolic radiolabeling studies suggest strongly that MGDG is the main substrate for chloroplastic desaturases while DGTS and PE are substrates for extra-chloroplastic desaturases and desaturases). A (especially for the hypothetical scheme of lipid metabolic pathways and their sub-cellular organization in Chlamy­ domonas spp. is proposed in Fig. 2. The proposed scheme results primarily from the analysis of fatty acid distribution in each lipid class and at each position of the glycerol backbone (Giroud and Eichenberger, 1988) and from the few studies of enzymes of lipid metabolism. It is clear that many steps in this pathway remain to be confirmed. One technical difficulty is the problem of isolating the chloroplast envelope in a highly purified state from Chlamydomonas spp. Once this is achieved, it would allow the exact localization of enzymes involved in lipid metabolism. Nevertheless, the proposed scheme should be useful as a basic pathway for the design of genetic approaches to the study of this problem.

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Glycerolipids

IV. In vivo Modifications of Lipid Composition in Chlamydomonas

Despite the barrier that the cell wall of Chlamy­ domonas spp. presents, it was shown by Grenier et al. (1991) that it is possible to increase significantly the lipid content and to change the fatty acid composition of the cells by growing them in the presence of liposomes. In that study, the authors used liposomes of PC, a lipid which is absent in Chlamydomonas spp., in order to follow the fate ofthe lipid inside the cell. Surprisingly, under most conditions, PC was detected only in trace amounts. In fact PC only accumulated at quite high concentrations when dipalmitoyl-PC liposomes were presented. Cell growth was strongly inhibited under these conditions. With unsaturated PC, such as dioleoyl or dilinoleoylPC, no inhibition of growth was observed at concentrations of up to 90 mg of liposomes per ml of culture, and PC did not accumulate in the cells. With a liposome concentration of 90 mg/ml, the lipid content ofthe cells is modified as follows: an increase of 160% is observed with dipalmitoyl-PC liposomes, 130% with dioleoyl-PC and up to 265% with dilinoleoyl-PC. Examination of lipid class composition showed that membrane polar lipid content remained quite stable while a very significant increase in the triacylglycerol content was observed. A four, five and ten-fold increase in triacyglycerol was observed in cells fed with dipalmitoyl-PC, dioleoylPC and dilinoleoyl-PC, respectively. The massive synthesis of triacylglycerol probably constitutes a regulatory mechanism which allows the algae to store the excess of exogenous fatty acids and to regulate membrane polar lipid composition. Even if the content of the polar lipid classes constituting the hydrophobic matrix of cell membranes remained quite stable, important changes in fatty acid composition were observed. This demonstrates that exogenous fatty acids from PC can be actively integrated into these lipid classes. With dioleoyl-PC liposomes, the C18:1 percentage increased mainly in DGTS. A big increase in percentage of C18:2 in this lipid was also observed. This suggests that DGTS is the preferred substrate extra-chloroplastic desaturase of C18:1. for the The percentage of C18:1 also was increased in PE but the percentage of C18:2 did not. However, a large increase in the percentage of was noted suggesting that PE is probably the preferential substrate for the desaturase. Curiously, no increase

425 in C18:1 was seen in chloroplastic lipid species but increased the percentage of C18:2 and markedly in DGDG (while the fatty acid composition of MGDG remained unaffected). This result can be interpreted either by proposing that oleic acid is imported from the cytosol to the chloroplast followed by efficient desaturation by the chloroplast desaturases, or by proposing that both C18:2 and are imported and incorporated directly into DGDG. With dilinoleoyl-PC liposomes, C18:2 increased mainly in DGTS suggesting a preferential acylation of the C18:2 fatty acid to DGTS. However, chloroplast lipid species also significantly accumulated this fatty acid. Together these results support strongly the cooperative pathway described above. It should be noted that such observations demonstrate that even if the import of C18 acyl chains from the cytoplasm to the chloroplast is very low in Chlamydomonas spp. under normal growth conditions (where the intra-chloroplastic desaturation is largely capable of synthesizing the poly-unsaturated fatty acids of the chloroplast), it is nevertheless obvious that all the machinery involved in the process is present and has the potential to work. For instance, it can be seen that when Chlamydomonas spp. is grown in the presence of dioleoyl or dilinoleoylphosphatidylcholine liposomes the C18/C16 ratio in the DGDG increases from around 54% to up to 70%. This shows that up to 20% of the molecules of this lipid have a C18:C18 structure and therefore indicates that they are imported from the cytosol. The mechanism of import remains unknown but the most probable explanation (taking into consideration the metabolic studies performed by Giroud and Eichenberger) is that it is due to DGTS at the level of the chloroplast envelope. This study gives a good picture of the most likely distribution of the different desaturases in the chloroplast and in the cytosol and of their preferential substrates: as in all other eukaryotic photosynthetic organisms the desaturase is exclusively located in the chloroplast. The chloroplast is also able to desaturate the C16:0 It possesses a full set of desaturases which in most probably use MGDG as substrate, the chloroplastic and desaturases leading to the synthesis of C18:2 and but also and 13 desaturases leading to the synthesis of C 16:4, which is a major fatty acid in MGDG. The -trans desaturase is probably also located in the chloroplast. In the endoplasmic reticulum, the most active desaturase (which clearly is not desaturase is a

426 regulated in the same way as is the equivalent chloroplast desaturase). The cytosolic uses preferentially DGTS as a substrate in the cytosolic compartment. This compartment also contains a desaturase which prefers PE (and also desaturase which probably PI) as substrate and a accepts preferentially PE (but also DOTS) as substrate. Finally, it should be noted that even if the extrachloroplastic desaturase seems to be only weakly active in Chlamydomonas spp., this enzyme is certainly present as it is needed for the synthesis of C18:4 which is concentrated in cytosolic lipid species. In addition to supporting the hypothesized organization of glycerolipid metabolism in Chlamy­ domonas spp., the ability to ‘capture’ exogenous lipids (by a mechanism which remains unknown) opens up the possibility to manipulate membrane lipid composition. For example, in Chlamydomonas spp. cultivated in the presence of dioleoyl or dilinoleoyl-PC liposomes, major changes in cell membrane ultrastructure of the thylakoid membranes were observed (Grenier et al., 1991).Furthermore, when C. reinhardtii mutants lacking the PG species (found only in with the special fatty acid the thylakoids membranes of eukaryotic photosynthetic organisms and which will be described in the next paragraph), were grown in the presence of liposomes of this lipid, it was possible to target the missing lipid efficiently to the thylakoid membranes. Only a part of the foreign PG was deacylated and the fatty acids were redistributed into the endogenous lipidic classes or degraded but a significant amount of the foreign lipids were stabilized in the thylakoid membranes (and nowhere else), allowing large compensation of the deficiency of this lipid in the mutants. An interesting observation was that the only molecular lipid classes which could be integrated into the membrane in a stable manner were the species found normally in the wild-type membrane. This observation suggests that this may be a specific interaction between the correct protein and the correct lipid species in the membrane which assures the stabilization of the lipid.

V. Mutants Affected in Lipid Composition A few mutants of C. reinhardtii affected in lipid composition have been isolated. In 1987, Garnier et al. isolated two nuclear mutants that lacked PS II activity and showed intriguing chlorophyll fluores-

Antoine Trémolières cence properties. Compared to classical mutants lacking PSII which show an abnormally high fluorescence level, these two mutants had a low level of fluorescence and were therefore named mf1 and mf2 (mf for minimum fluorescence). As shown in Table 5, these two mutants strains were fully devoid of the PG molecular species containing (Maroc et al., 1987; Dubertret et al., 1994). is a special fatty acid found specifically at the sn-2 position of PG in all eukaryotic photosynthetic organisms (Dubacq and Trémolières, 1983). As a consequence, these mutants contained only 30% of the normal PG content. In addition, the mutant mf2 contained an unusually high level of lyso-PG. It was first proposed by Seras et al. (1989), from results of a study of radioactive acetate incorporation into lipids and fatty acids in the wild-type and mutant strains, that the site ofthe mutation could be the gene coding for the desaturase. Thus, it was proposed that the mf1 and mf2 mutants contained two mutations: one affecting PSII activity and the desaturase. Unfortunately, attempts other the to separate the two mutations by crossing the mutants with wild strains followed by the analysis of their progeny was unsuccessful. Another strategy was to select suppressors of the mutation affecting PS II activity. Mutants with recovered PS II activity were selected, but no mutants with a wild-type level of PS containing PG II activity and lacking the were obtained. However, two new phenotypes were obtained through this study: the first class ofmutants called Pmf1 (P for ‘photosynthesis’, mf1 indicating that it was derived from the mf1 mutant) recovered around 10% ofthe wild-type level ofoxygen evolution and also about 10% of the normal level of containing PG. The second one called Pmf2, recovered 50% of both the photosynthetic activity and the normal level ofthe lipid. No segregation between the photosynthetic deficiency and the lipid deficiency were obtained. However, a careful examination of the lipid phenotypes of the four mutants so obtained indicated a more complex relationship than imagined originally. In fact, in addition to the most obvious change, which was the disappearance (in the two non-photosynthetic mutants) or the decrease (in the containing PG, two photosynthetic strains) of alteration of fatty acid composition in several lipid classes both inside and outside of the chloroplast was observed. Analysis of the fatty acid pattern of the mutants showed that in addition to the deficiency in desaturase activity, they were also affected

Chapter 21 Glycerolipids

in the chloroplastic -desaturase activity while all and other desaturase activities (the chloroplastic -desaturases and all the endoplasmic reticulum desaturases) were normal. Even if the target(s) of the mutation(s) affecting these four mutants has not yet been established, studies of their lipid metabolism have largely confirmed the scheme proposed for the organization of glycerolipid biosynthesis in Chlamydomonas spp. (El Maanni, 1996). An increase in the amount of fatty acids of cytosolic origin was

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observed in the four mutants. It should be noted that indicates that even the observed change in if the return of fatty acids from the cytosol to the chloroplast through the cooperative pathway plays a minor role in this alga, it does nevertheless occur and can operate, when necessary, according to physiological conditions or genetic conditions imposed by the background. Indeed, in these mutants it is this cooperative pathway which probably compensates for the deficiency of the desaturase activity and

428 allows the cells to keep a perfectly normal level of unsaturation in the chloroplastic lipids. A nuclear mutant containing only 5% of the SQDG was isolated by Sato et al. (1995a) and named hf2 because it had a very high level of chlorophyll fluorescence. Compared to the mf mutants already described, no other significant change in the lipid composition was detected suggesting that the mutation affected some steps in the synthesis of SQDG. Another mutant showing an abnormally high fluorescence level and called hf9 was isolated by the same group (Sato et al., 1995). This mutant showed a normal lipid composition but the chloroplast lipids were strongly reduced in polyunsaturated fatty acids (C16:2; C16:3; 16:4; C18:2; and increased in the percentage of and The poly-unsaturated fatty acid level remained normal in extra-chloroplastic lipid species. Interestingly and increased in DGTS but not in PE. This fatty acid profile is interpreted by the authors as resulting from a mutation affecting the chloroplastic desaturase. As for the mf mutants, the pleiotropic effect of the mutation could be explained through the cooperative relation between the chloroplastic and cytosolic pathways. In Table 5 the lipid characteristics of the six Chlamydomonas mutants isolated so far are summarized and the putative direct or indirect targets of the mutations are indicated.

VI. Involvement of Lipids in the Functional Organization and the Biogenesis of the Photosynthetic Apparatus Mutants of Chlamydomonas affected in lipid or fatty acid composition have been used with success to demonstrate a role for some lipids in the biogenesis and the functional organization of the photosynthetic membrane. The mutants mf1 and mf2, lacking the PG were found to be almost containing completely unable to form the trimeric LHC II which is believed to be the stable and functional form ofthe main light harvesting pigment-protein antenna complex normally associated with PS II, but also with PS I under conditions which reduce the plastoquinone pool (Maroc et al., 1987; Dubertret et al., 1994). Thylakoid membranes of these mutants were almost totally lacking in grana stacks and they present a very unusual ultrastructure with a normal amount of membranes but which are largely

Antoine Trémolières unappressed. Reincorporation of the missing lipid by incubating the mutants strains in the presence of liposomes (as described above) largely restored the ability to form the trimeric LHC II and to develop grana stacks. The fact that this restoration only occurred under conditions where de novo synthesis of LHC II apoproteins occurred shows that is involved in some step in the biogenesis of the trimeric LHC II (Garnier et al., 1990; Trémolières et al., 1991). This approach, using mutants of C. reinhardtii supplemented with liposomes, was the first demonstration in vivo of the important role played by PG in the initial step(s) of the trimerization of LHC II. This work also provided evidence correlating the stabilization of LHC II in a trimeric state and the formation of appressed membranes. It was observed that in the mutants lacking containing PG, the apoproteins of LHC II which cannot be stabilized in the trimeric form and are degraded at the N-terminal extremity. These results are in good agreement with those obtained by in vitro reconstitution experiments which showed that lipids, and more precisely PG, induce trimerization of monomeric LHC II. Furthermore, it has been shown, using mutated forms of recombinant LHC II expressed in E. coli, that the lipid probably interacts with a domain at the N-terminal end of the LHC II to induce a conformational change of the complex (including changes in the orientation of some chlorophylls) which allows the trimerization process to occur. This domain is situated 16 amino acids away from the N-terminal extremity and is constituted by a WYGPDR sequence in pea LHC II, (replaced in C. reinhardtii by a similar sequence FYGPNR). The sequence is called the ‘trimerization motif’ (Remy et al., 1984; Hobe et al., 1994; Hobe et al., 1995). It is a rare example ofa highly specific interaction between a lipid and a protein which induces first a change in the configuration of the chlorophyll-protein complex and then a cascade of reactions leading to the formation of complex membrane interactions which in turn leads to grana stacks. SQDG is found in all photosynthetic organisms. It would be interesting to know if this widespread occurrence results from the common phylogenetic origin of these organisms or because this lipid is important for some steps of the photosynthetic process. Sato et al. (1995a,b) isolated a C. reinhardtii mutant almost fully devoid of SQDG (around 5% of the wild type level), but otherwise having a normal lipid composition. The photosynthetic characteristics

Chapter 21 Glycerolipids of this mutant were examined and a 17% decrease in the growth rate was found, resulting from a decrease in PS II activity while PS I activity was unaffected. Genetic analysis showed that the lesion in PS II activity was associated with SQDG deficiency. An extensive analysis of the chlorophyll/protein complexes of the thylakoid membranes was performed including the PS II core, LHC II, PS I and LHC I as well as No significant differences in the polypeptide patterns were noticed, suggesting that if SQDG was not involved in the biogenesis and targeting of the PS II apo-proteins into the photosynthetic membrane, this lipid is nevertheless indispensable for the function of PS II. It remains to be seen if the trace amounts (5% of the wild-type level) of lipid, remaining in the mutant can nevertheless play some other role in the thylakoid membrane. SQDG has been found linked tightly to the purified LHC II of Chlamydomonas spp. (Sigrist complex of et al., 1988) and to the chloroplasts of land plants (Pick et al., 1985), although in very small amounts. Concerning the effect of the lipid deficiency on chloroplast ultrastructure, it was observed that the hf 2 mutant contained abnormal extremely curved thylakoid membranes showing that the virtual absence of SQDG has a strong effect on thylakoid membrane ultrastructure. It should be noted that the thylakoid membranes were normally appressed. Another mutant isolated by the same group (Sato et al., 1995b) was supposed to be affected in chloroplastic desaturase activity. In this mutant, when compared to the wild type, the percentage of poly-unsaturated fatty acids was reduced from 74% to 18% for MGDG, from 45% to 5% for DGDG and from 20% to a few % for PG. An increase from 20% in PG. Extrato 28% was observed for chloroplastic lipids were not affected. This mutant was strongly affected in its photosynthetic properties with a reduction of about 80 % in photosynthetic oxygen production and a concomitant decrease in the growth-rate under phototrophic conditions. These are very different results from those obtained with mutants of A. thaliana affected in the same desaturase and which are still capable of carrying out photosynthesis (Mc Court et al., 1987). This result can be explained by the fact that in Arabidopsis, contrary to Chlamydomonas spp., the participation of the cooperative pathway to the synthesis of chloroplastic lipids is very important and thus can compensate for the chloroplastic desaturase

429 deficiency. Despite this dramatic inhibition of the photosynthetic process in the hf9 mutant, the ultrastructure of the thylakoid membranes was not significantly affected. In fact, hyper-stacking was observed which could result from a pleiotropic effect of the mutation which leads to an increase in the percentage of PG. The photosynthetic properties and ultrastructural characteristics of the various mutants of Chlamydomonas spp. affected in lipid composition are summarized in Table 4.

Acknowledgments The author wishes to express his gratitude to W. Eichenberger and A. Boschetti from the University of Bern, in Switzerland, for kindly providing certain data included in this review. He is indebted to M. Hodges for carefully reading the manuscript. We wish also to acknowledge M. H. Trémolières for typing and L. Orr and Y. Bertrand for figures.

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430 Eichenberger W (1976) Lipids of Chlamydomonas reinhardtii under different growth conditions. Phytochemistry 15: 459– 463 Eichenberger W (1982) Distribution of diacylglycerol O-4´(N,N,N-trimethyl)homoserine in different algae. Plant Sci Lett 24: 459–463 Eichenberger W (1990) Identification ofnew plant lipids: structure of a second betaine lipid from algae. In: Quinn PJ and Harwood JL (eds) Plant Lipid Biochemistry, Structure and Utilization, pp 9–16. Portland Press, London Eichenberger W (1993) Betaine lipids in lower plants. Distribution of DGTS, DGTA and phospholipids and the intracellular localization and site of biosynthesis of DGTS. Plant Physiol Biochem 31: 213–221 Eichenberger W, Boschetti A and Michel HP (1986) Lipid and pigment composition of a chlorophyll b-deficient mutant of Chlamydomonas reinhardtii. Physiol. Plant 66: 589–594 El Maani A (1996) Etude du rôle du phosphatidylglycérol dans la biogenèse et l’organisation fonctionnelle dc la membrane photosynthétique chez quatre mutants de Chlamydomonas reinhardtii affectés dans le métabolisme des lipides. Thèse de Doctorat d’Etat, Université Paris-Sud, France. Frentzen M (1986) Biosynthesis and desaturation of different diacylglycerol moieties in higher plants. J Plant Physiol 124: 193–209 Frentzen M, Hares W and Schiburr A (1984) Properties of the microsomal glyceroI-3-P and monoacylglycerol-3P- acyltransferases from leaves. In: Siegenthaler PA and Eichenberger W (eds) Structure Function and Metabolism of Plant Lipids. pp 105–110. Elsevier Science Publishers, Amsterdam. Gamier J, Wu B, Maroc J, Guyon D and Trémolières A (1990) Restoration of both an oligomeric form of the light-harvesting antenna CP II and a fluorescence state II-state I transition by -trans-hexadecenoic acid containing phosphatidylglycerol, in cells of a mutant of Chlamydomonas reinhardtii. Biochim Biophys Acta 1020: 153–162 Giroud C, Gerbert A and Eichenberger W (1988) Lipids of Chlamydomonas reinhardtii. Analysis of molecular species and intracellular site(s) of biosynthesis. Plant Cell Physiol 29: 587–595 Grenier G, Guyon D, Roche O, Dubertret G and Trémolières A ( 1 9 9 1 ) Modification of fatty acid composition of Chlamy­ domonas reinhardtii cultured in the presence of liposomes. Plant Physiol Biochem 29: 429–440 Gresti J, Mignerot C, Bezard J and Wolff RL (1996) Distribution of olefinic acid in the triacylglycerols from Pinus coraiensis and Pinus pinaster seeds oil. J A O C S 73: 1539–1548 Hobe S, P r y t u l l a S, Kühlbrandt W and Paulsen H (1994) Trimerization and crystallization of reconstituted lightharvesting chlorophyll a/b protein complex. EMBO J 13: 3423–3429 Hobo S, Förster R, Klinger J and Paulsen H (1995) N-proximal sequence motif in light-harvesting a/b binding protein is essential for the trimerization of light-harvesting chlorophyll a/b complex. Biochemistry 43: 10224–10228 Hope W and Scwhenn JD (1981) In vitro biosynthesis of the plant sulfolipid: On the origin of the sulfonate group. Z Natürforch c36c: 820–826 James AT and Nichols BW (1966) Lipids of photosynthetic systems. Nature 210: 372–375 Janero DR and Barnett R (1982 a) Isolation and characterization

Antoine Trémolières of an ether-linked homoserine from the thylakoid membrane J Lipid Res 23: 307–316 of Chlamydomonas reinhardtii Janero DR and Barnett R (1982 b) Cardiolipin of Chlamydomonas reinhardtii 137+. Phytochemistry 21: 1151–1152 Jeselma CLAS, Michaels DR, Janero DR and Barnett RJ (1982) Membrane lipid metabolism in Chlamydomonas reinhardtii 137+ and y-1.I. Biochemical localization and characterization of acyltransferase activities. J Cell Sci 58: 469–488 Mac Court P, Kunstt L, Browse J and Somerville CR (1987) The effects of reduced amount of lipid unsaturation on chloroplast ultrastructure and photosynthesis in a mutant of Arabidopsis thaliana. Plant Physiol 84: 353–360 Maroc J, Trémolières A, Gamier J and Guyon D (1987) Oligomeric form of the light-harvesting chlorophyll a+b protein complex CP I I , phosphatidylglycerol; -trans-hexadecenoic acid and energy transfer in Chlamydomonas reinhardtii wild type and mutants. Biochim Biophys Acta 893: 91–99 Matsumoto GI, Shioya M and Nagashima H (1984) Occurrence of 2-hydroxy acids in microalgae. Phytochemistry 23: 1421– 1423 Marshall MO and Kates M (1972) Biosynthesis of phosphatidylglycerol by cell-free preparation from spinach leaves. Biochim Biophys Acta 260: 558–570 Mendolia- Morgenthaler LW, Eichenberger W and Boschetti A (1985) Isolation of chloroplast envelopes from Chlamydomonas. Lipid and polypeptide composition. Plant Sci 41: 97–10 Michaels ASCL, Jeselma CL and Barnett RJ (1983) Membrane lipid metabolism in Chlamydomonas reinhardtii 137+ and y-1 II Cytochemical localization of acyltransferases activities. J Ultrastr Res 82: 35–51 Moore TS, Lord JM, Kagawa T and Beevers H (1973) Enzymes of phospholipid metabolism in the endoplasmic reticuluin of castor bean endosperm. Plant Physiol 52: 50–53 Moseley FR and Thompson CA jr (1980) Lipid composition of Volvox carteri. Plant Physiol 65: 260–265 Oursel A, Escoffier A, Kader JC, Dubacq JP and Trémolières A (1987) Last step in the cooperative pathway for galactolipids synthesis in spinach leave: Formation of monogalactosyldiacylglycerol with C18 polyunsaturated acyl groups at both carbon atoms of the glycerol. FEBS Lett. 219: 393–399 Pick U, Gounaris K, Weiss M and Barber J (1985) Tightly bound Biochim Biophys Acta sulpholipid in chloroplast 808: 415–420 Remy R, Trémolières A and Ambard-Breteville F (1984) Formation of oligomeric light-harvesting chlorophyll a/b protein by i n t e r a c t i o n w i t h liposomes. Photobiochem Photobiophys 7: 267–276 Roughan PG and Slack CR (1982) Cellular organization of glycerolipid metabolism. Ann. Rev. Plant Physiol. 33: 97–132 Roughan PG and Slack CR (1984) Glycerolipid synthesis in leaves. Trends Biochem Sci 9: 383–386 Seras M, Gamier J, Trémolières A and Guyon D (1989) Lipid biosynthesis in cells of the wild type and two photosynthesis mutants of Chlamydomonas reinhardtii. Plant Physiol Biochem 27: 393–399 Sato N and Furuya M (1984) Distribution of diacylglyceroltrimcthylhomoscrinc in selected species of vascular plants. Phytochemistry 23: 1625–1627 Sato N and Furuya M (1985) Distribution of diacylglyceroltrimethylhomoserine and phosphatidylcholine in non-vascular

Chapter 21 Glycerolipids green plants. Plant Sci 38: 81–85 Sato N, Sonoike K, Tsuzuki M and Kawaguchi A (1995a) Impaired Photosystem II in a mutant of Chlamydomonas reinhardtii defective in sulfoquinovosyldiacylglycerol. Eur J Biochem 234: 16–23 Sato N, Tsuzuki M, Matsuda Y, Ehara T, Osafune T and Kawaguchi A (1995b) Isolation and characterization ofmutants affected in lipid metabolism of Chlamydomonas reinhardtii. Eu J Biochem 230: 987–993 Sigrist M, Zwillenberg C, Giroud CH, Eichenberger W and Boschetti A (1998) Sulfolipid associated with the lightharvesting complex associated with Photosystem II apoproteins of Chlamydomonas reinhardtii. Plant Science 58: 15–23 Trémolières A (1991) Lipid-protcin interactions in relation to light energy distribution in photosynthetic membrane of cukaryotic organisms. Role of acid containing phosphatidylglyccrol. Trends in Photochcm Photobiol 2: 13–32

431 Trémolières A and Siegenthaler PA (1997) Reconstitution of photosynthetic structures and activities with lipids. In: Siegenthaler PA and Murata N (eds) Lipids in Photosynthesis: Structure, function and Genetics, pp 175–189. Kluwer Academic Publishers, Dordrecht Trémolières A, Roche O, Dubcrtrct G, Guyon D and Gamier J (1991) Restoration of thylakoid appressions by acid containing phosphatidy Iglycerol in a mutant of Clilamydonionas reinhardtii. Relationship with the regulation of excitation energy distribution. Biochim Biophys Acta 1059: 286–292 Trémolières A, Dainese P and Bassi R (1994) Heterogenous lipid distribution among chlorophyll-binding proteins of Photosystem II in maize mesophyll chloroplasts. Eur J Biochem 22: 721–730

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Chapter 22 In vivo Measurements of Photosynthetic Activity: Methods Pierre Joliot, Daniel Béal and René Delosme Institut de Biologie Physicochimique, 13 rue Pierre et Marie Curie, 75005 Paris, France

Summary I. Introduction A. Study of Photosystem II B. Study of the Cytochrome Complex and Photosystem I II. Kinetic Analysis of the Fluorescence Yield A. Kinetic Analysis of the Fluorescence Yield Under Continuous Illumination B. Measurement of Fluorescence Yield Monitored by Modulated Light C. Measurement of Fluorescence Yield Monitored by Short Detecting Flashes III. Fluorescence Emission Spectra at Low Temperature IV. Delayed Fluorescence Measurements V. Oxygen Measurements VI. Absorption Spectroscopy A. Spectrophotometric Method using a Continuous Detecting Beam B. Spectrophotometers using Detecting Flashes VII. Photoacoustic Measurements A. Modulated Light Excitation B. Flash Excitation VIII. Conclusion and Perspectives Appendix A: Estimation of the Signal-to-Noise Ratio in Fluorescence Measurements Appendix B: Flash Spectrophotometer Acknowledgment References

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Summary Functional studies of living unicellular algae such as Chlamydomonas provide the simplest way to collect

information on the mechanisms of photosynthetic exciton and electron transfer. Physiological function and

regulation, which can be impaired upon isolation of subcellular compartments, are best studied in intact cells.

Functional analysis in vivo is particularly well suited for characterizing mutant strains obtained either by

random or by site-directed mutagenesis. At variance with isolated thylakoid membrane preparations, it should

be taken into account that in intact cells the thylakoid membranes are in permanent interaction with the other

compartments of the cell. The most important parameter that influences the properties of the photosynthetic

chain is the ATP/ADP ratio. Derived from this observation, optimal conditions for studying Photosystem II, Photosystem I and the cytochrome complex are discussed. The main biophysical techniques commonly applied to in vivo studies of exciton and electron transfer are described. Special emphasis is laid on the kinetic analysis of the fluorescence yield under continuous illumination, which is the most popular technique used for rapid characterization ofthe photosynthetic electron

J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 433–449. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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transfer chain. Other techniques particularly well adapted to in vivo studies, such as absorption spectrophotometry using detecting flashes, amperometric detection of oxygen evolution, and photoacoustic techniques, are also presented in detail. The optimal signal-to-noise ratio and time resolution that can be expected from these different techniques are discussed. A few experimental examples ofthe use of these techniques are presented.

I. Introduction Functional analysis of the photosynthetic apparatus is one ofthe main sources of information on the basic mechanisms of the photosynthetic process, from the molecular to the cellular level. The photosynthetic apparatus is well adapted to kinetic analysis, since the electron transfer reactions can be easily triggered by light in purified complexes, isolated membranes or living cells. Most of our knowledge ofthe primary photosynthetic processes has been obtained with thylakoid membranes isolated from plants, or solubilized membrane complexes. Nevertheless, the isolation process may impair function, and it suppresses most ofthe physiological regulation. Thus, in vitro studies have to be compared with functional analysis under physiological conditions. For in vivo studies, the functional analysis of unicellular algae in suspension is an easier task than that of plants. In addition, the large number of mutants of the photosynthetic apparatus of Chlamydomonas reinhardtii that have been obtained either by random mutagenesis or by genetic engineering makes this unicellular alga one of the best models for in vivo studies. Isolation of thylakoid membranes from C. reinhardtii cells affects the organization and activity of the photosynthetic components. Because of the procedure used to break the cells, such membranes are of lower functional quality than the thylakoids isolated from plant leaves. Consequently, functional studies on C. reinhardtii are generally performed either on whole cells, or on solubilized and purified protein complexes. Concerning the material used, one should be aware that Chlamydomonas is subject to frequent spontaneous mutations, which may induce low respiratory activity and a low growth rate under heterotrophic Abbreviations: AC – alternating current; DC – direct current; DCHC – dicyclohexyl-18-crown-6; DCMU – 3-(3,4-dichlorophenyl)-1, 1-dimethylurea; FCCP – carbonylcyanide-ptrifluoromethoxyphenylhydrazone; PAM – pulse-amplitudemodulation fluorometer; PS I –Photosystem I; PS II – Photosystem II; rms – root mean square; Nd:YAG – Neodymium:Yttrium Aluminum Garnet

conditions. Photosynthetic activity can be also impaired by lesions in some of the membrane complexes. The condition of the starting wild-type strain should be tested for growth in minimum medium under rate-limiting light intensity (around 1500 lux). When studying the photosynthetic process in vivo, one has to take into account that the thylakoid membrane is in close interaction with compartments such as the stroma, the cytosol and the mitochondria. The most important parameter that influences the kinetic and thermodynamic properties of the photosynthetic chain is the ATP/ADP ratio. In Chlamydomonas and Chlorella cells, the concentration of ATP and ADP equilibrates between the different cellular compartments. Thus the ATP/ADP ratio is determined by the overall equilibrium between ATP consumption and synthesis, irrespective of where they take place in the cell. Membrane photosynthetic reactions, respiration and fermentation participate in the synthesis of ATP, whereas all the energyconsuming reactions result in its consumption. The ATP/ADP ratio in the stroma controls directly or indirectly the kinetics of a large fraction of the electron transfer reactions, the redox state of the intersystem electron carriers, and the lateral organization of proteins within the thylakoid membrane. First of all, according to Mitchell (1966), the transmembrane electrochemical gradient of in volts), which is the sum of an electrical protons and of an osmotic component component is given by the equation

where n is the number of protons translocated through the membrane ATP synthase per ATP molecule synthesized, and K is the equilibrium constant for ATP hydrolysis. Assuming a constant phosphate the transmembrane electroconcentration chemical gradient of protons is linearly related to the logarithm ofthe ratio [ATP]/[ADP]. The electrostatic exerts a control on the equilibrium component

Chapter 22

In vivo Measurements

constant and the rate of the transmembrane electron transfer reactions. As the pH of the stromal compartment, which is an highly buffered medium, is fairly constant, the osmotic component modulates the internal pH of the lumen, and thus the equilibrium constant and the rate of the electron transfer reactions involving a release of protons in the lumen (Rumberg and Siggel, 1969). The electrochemical proton gradient and the concentration of nucleotides also control the activity of the membrane ATPase (Davenport and McCarty, 1981), and consequently the lifetime of the light-induced membrane potential. A decrease of the ATP/ADP ratio results in an enhancement of the glycolytic pathway (Pasteur effect), which in turn induces an increase of the stromal NADPH concentration, and consequently a reduction of the plastoquinone pool, presumably via a reversible NADPH-plastoquinone reductase (Rébeillé and Gans, 1988; Bulté et al., 1990; Gans and Rébeillé, 1990). Therefore, the ATP/ADP ratio exerts an indirect control on the lateral distribution of the light-harvesting complexes within the thylakoid membrane (state transitions, see Chapter 30, Keren and Ohad), via the modulation of the redox state of the plastoquinone pool (Gans and Rébeillé, 1990; Bulté et al., 1990). The ATP/ADP ratio also controls directly or indirectly the distribution of the complexes between the appressed cytochrome and the non-appressed regions of the membrane (Vallon et al., 1991).

A. Study of Photosystem II In Chlamydomonas cells in the presence of oxygen, mitochondrial ATP synthesis leads to a high ATP/ ADP ratio in all the cell compartments, generating a large permanent electrochemical proton gradient across the thylakoid membrane (Joliot and Joliot, 1980). As discussed above, the high ATP concentration shifts the plastoquinone pool towards the oxidized state (between 60 and 90% oxidized). The electrical component of the permanent electrochemical proton gradient has been estimated to be 40–80 mV in Chlorella under aerobic conditions, by measuring the absorption changes associated with non-linear electrochromic probes present in the thylakoid membrane (Joliot and Joliot, 1989). Similar values apply to Chlamydomonas. Addition of uncouplers such as FCCP or DCHC, or of specific inhibitors of the respiratory mitochondrial chain,

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induces a large decrease of the permanent electrochemical gradient of protons, but also leads to the reduction of the plastoquinone pool, and consequently to the inhibition of PS II activity (Bulté et al., 1990; Gans and Rébeillé, 1990). Thus in vivo analysis of Photosystem II activity and of oxygen evolution, which requires the presence of an oxidized pool of electron acceptors, is possible only under aerobic conditions and in the absence of uncoupler, i.e. in the presence of a large electrochemical proton gradient. We shall see that this point has important functional implications. Firstly, the electrochemical proton gradient modifies the rate constants of most electron transfer reactions. In isolated chloroplasts, the electrochemical proton gradient accelerates the back reactions between oxidized electron donors and reduced electron acceptors of PS II (Miles and Jagendorf, 1969; Barber and Kraan, 1970; Wraight and Crofts, ... are the successive oxidation 1971). If states of the water-splitting enzyme according to Kok et al. (1970), in DCMU-treated unicellular algae, and the the rate of the back reaction luminescence emission are decreased by a factor of about 5 to 10 upon addition of uncoupler, owing to the collapse of the electrochemical proton gradient (Joliot and Joliot, 1980; Bennoun, 1982). Moreover, the membrane potential decreases the concentration of active PS II centers (Diner and Joliot, 1976) by a mechanism not yet understood. The electrochemical proton gradient also accelerates the decay of the membrane potential, owing to the activation of the membrane ATPase. Thus the large electrochemical proton gradient which exists in the presence of oxygen imposes severe limitations to the study of the electron transfer chain. They can be overcome by pre-treating the cells with benzoquinone at approx. (Diner and Joliot, 1976). At this concentration, benzoquinone irreversibly inhibits the enzymatic processes involving nucleotides such as ATP and NADPH (Bulté and Wollman, 1990 ). After washing out the benzoquinone, the membrane ATPase and the reducing pathways such as NADPH-plastoquinone reductase remain blocked. Thus, under such conditions the plastoquinone pool stays oxidized, even in the absence of a permanent electrochemical proton gradient. Furthermore, when benzoquinone is added in the presence of oxygen, the Chlamydomonas cells are blocked in state 1 (Bulté and Wollman, 1990), where most of the mobile antenna is connected to PS II centers. Thus benzoquinone-

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436 treated cells provide a material equivalent to isolated chloroplasts, and well suited to the study of Photosystem II activity, oxygen evolution, and more generally open electron flow from Photosystem II to Photosystem I. It is noteworthy that benzoquinone treatment considerably slows down plastoquinol oxidation in the dark. As proposed by Bennoun (Chapter 35), this suggests that the oxidation of plastoquinol in vivo proceeds via a reversible NADPH plastoquinone reductase which is inhibited by the benzoquinone treatment, and not via an oxidase included in the thylakoid membrane (chlororespiration) (see also Peltier and Thibault, 1988).

B. Study of the Cytochrome Photosystem I

are anaerobiosis in the presence of an uncoupler at low concentration. In the following, we shall describe briefly the main biophysical techniques that can be used for the functional analysis of photosynthesis in unicellular algae in vivo. Most of these techniques have been developed in the laboratory, using a large variety of designs. We shall focus on the main constraints that should be taken into account to optimize in vivo analysis, and give some examples of the use of the different techniques. The reader is also referred to the review of Kramer and Crofts (1996) on the measurement of photosynthetic electron transport in intact plants.

Complex and II. Kinetic Analysis of the Fluorescence Yield

The cyclic electron flow involving Photosystem I complex is better studied in and the cytochrome anaerobic conditions, where most of the mobile antenna (Wollman and Delepelaire, 1984; Delepelaire and Wollman, 1985; Delosme et al., 1994, 1996) and complexes (Vallon et al., 1991) the cytochrome are localized in the non-appressed regions, in proximity to PS I centers (state 2). In these conditions, where the plastoquinone pool is fully reduced, electron complex is transfer within the cytochrome triggered by charge separation in PS I. It is noteworthy, however, that even under anaerobic conditions a permanent electrochemical proton gradient is observed (Joliot and Joliot, 1989). The value of the electrochemical gradient, about halved with respect to aerobic conditions, remains stable for hours. This implies that ATP is synthesized via a fermentation process (Joliot and Joliot, 1989). However this value, indicative of the ATP/ADP ratio in anaerobiosis, and thus of the fermentation efficiency, varies from one algal culture to another, and is higher in Chlorella than in Chlamydomonas. The permanent electrochemical proton gradient observed in anaerobiosis includes both an electrical and an osmotic component. Addition of uncoupler in anaerobic conditions increases by a factor of about 2–4 the rate of plastoquinol oxidation at the site of the cytochrome complex (Joliot and Joliot, 1994; Finazzi et al., 1997). As shown by the accelerating effect of nigericin (G. Finazzi, unpublished), this rate is mainly dependent upon the pH in the lumen, i.e. upon the osmotic component of the electrochemical proton gradient. Thus the best conditions to study the electron complex transfer reactions within the cytochrome

Fluorescence measurements represent the first optical method which has been applied to the analysis of electron and exciton transfer processes within the photosynthetic apparatus of living algae or leaves of higher plants (Kautsky and Hirsch, 1931; Dutton et al., 1943). It is still the most popular technique used for the rapid characterization of mutants of unicellular algae. Part of the light absorbed by chlorophyll and accessory pigments is re-emitted as fluorescence of chlorophyll, which peaks around 685 nm. The fluorescence yield varies between 2 and 6%, and is mainly dependent on the redox state of the primary of Photosystem II, but also on quinone acceptor various other parameters, such as the organization and number of pigments connected to PS II, the redox state ofthe PS II electron donors (S states), and the transmembrane electrochemical proton gradient. This variety of parameters often precludes an unambiguous interpretation of the fluorescence data.

A. Kinetic Analysis of the Fluorescence Yield Under Continuous Illumination Analysis of the light-induced changes of the fluorescence yield under continuous illumination is the basic technique that provides the richest source of information on the properties of the antennae and of the electron transfer chain. In this technique, darkadapted algae are illuminated by a continuous beam, which induces an electron flow involving both photosystems. We shall first discuss the relation between the time resolution and the signal-to-noise ratio. Suppose that we measure the integral of the

Chapter 22 In vivo Measurements number N of photoelectrons emitted by the photodetector during a time interval which is short compared with the overall kinetic changes as the time resolution of studied. We shall define the method. Integration of the number of photocan be easily achieved electrons emitted during using digital integration. The analog signal is sampled using an analog-to-digital converter with a period The time resolution can be shorter than adjusted by summing a series of n digital samples with As in all optical methods, the signalto-noise ratio depends upon the number N. If i is the photocathode current (number of photoelectrons per second), the quantum noise (rms) will be equal to and the signal-to-noise ratio is proportional to As an example, the noise level would be 1% (rms) when When studying the kinetics of fluorescence yield changes, the time resolution should be such that is small the fraction of PS II centers hit during enough to hardly induce any significant fluorescence yield increase (~1% of the PS II centers). Consequently, when varying the intensity I of the actinic light, the signal-to-noise ratio must be optimized together with the time resolution, bearing in mind that to fulfill the above prerequisites, the product (or I. ) should be kept constant. Therefore, i. should be inversely proporthe time resolution tional to the intensity of actinic light. We have computed (see Appendix A) the order of magnitude of the theoretical quantum noise. The result of this calculation shows that most generally, the quantum noise is not a limiting parameter. One can conclude that the poor signal-to-noise ratio of many fluorescence transients reported in the literature is due to a poor optical design of the instrument, to an erroneous adjustment of the time resolution, or to the intensity fluctuations of the light source. Various light sources can be used: tungsten-halogen sources, light-emitting diodes, continuous lasers, xenon arc lamps. Tungsten-halogen sources, lightemitting diodes, and lasers equipped with power regulation are among the light sources that do not degrade the signal-to-noise ratio by introducing intensity fluctuations. When using light-emitting diodes, the electrical power and the duration of illumination should be limited, to avoid drift in intensity due to heating of the diodes. Any heterogeneity of illumination will induce variations of the photochemical rate constants, depending upon the position of algae within the

437 cuvette. This distorts the observed fluorescence kinetics, and may lead to erroneous interpretations. Homogenization of light intensity in a plane perpendicular to the actinic beam can be achieved using a light pipe to illuminate the cuvette. However, a major cause of heterogeneity of illumination lies in light absorption by the biological sample. Using green light, which is weakly absorbed by chlorophyll, or illuminating two opposite sides of the cuvette when using an actinic light which is more strongly absorbed, can minimize the heterogeneity. The opening time of the actinic beam often limits the time resolution of fluorescence kinetic measurements. With conventional light sources, an opening time of about 1 ms can be obtained with electromagnetic shutters. An opening time shorter than can be provided by light-emitting diodes with an appropriate power supply, or lasers with an acoustooptic modulator. Convenient filters should block the fraction of actinic light that is transmitted or scattered toward the light detector. This requires that the wavelength of the actinic beam be shorter than that of fluorescence emission. The actinic light can be eliminated by using colored red filters with a cutoff around 660 to 690 nm. Upon illumination by actinic light, these colored filters generally emit a red fluorescence, which is superimposed on chlorophyll fluorescence. This leads to an overestimation of the chlorophyll fluorescence yield. The effect can be minimized by placing large band interference filters between the cuvette and the colored filter to decrease the excitation of the colored filter. Figures 1 and 2 show the fluorescence induction in dark-adapted cells of the C. reinhardtii mutant ccslcomplex, in the ac206 devoid of cytochrome presence or in the absence of DCMU. The time The rise-time of the light-emitting resolution is diodes is negligible compared to the time resolution. The small amplitude of the DCMU-induced increase of the initial fluorescence level, and the well-defined inflexion point, show that prior to addition ofDCMU, were in most of the secondary electron acceptors their oxidized form (Velthuys and Amesz, 1974). A cuvette of 10 mm thickness is used, illuminated on both sides by 594 nm light-emitting diodes. The More diluted chlorophyll concentration is about cell suspensions would be suitable as well. Conversely, we have tested that the well-defined shape of the induction curves remains unchanged up to chlorophyll concentrations as high as

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modulated technique is well adapted for the analysis of the state transitions induced by chromatic effects (Bonaventura and Myers, 1969).

C. Measurement of Fluorescence Yield Monitored by Short Detecting Flashes B. Measurement of Fluorescence Yield Monitored by Modulated Light The fluorescence yield can be monitored using a weak modulated beam of constant intensity, as in the PAM technique which can be applied by use of a commercially available instrument (Schreiber, 1986). Using an AC amplifier with a narrow bandwidth centered on the frequency of the modulated beam, or preferably a lock-in amplifier, it is possible to reject the DC components induced by continuous actinic light. The intensity of the modulated signal is proportional to the fluorescence yield. The main limitation of this method is that the intensity of the modulated beam should be sufficiently low so as to not induce any significant actinic effect. This leads to a low signal-to-noise ratio or poor time resolution, which do not permit detailed kinetic studies. Therefore, this type of device is convenient for measuring changes of the fluorescence yield under quasi-steady state conditions, upon illumination by continuous actinic light of various intensities and wavelengths. Particularly suited to study the effect of far-red actinic light on the fluorescence yield, the

In this technique, the fluorescence yield is sampled by short detecting flashes of low energy. Using a single beam device, the signal-to-noise ratio is limited by the energy fluctuations of the xenon flash (~2%) (Joliot and Joliot, 1977). A much better signal-tonoise ratio is achieved when the fluorescence signal is divided by a reference signal proportional to the energy of the detecting flash (Joliot et al., 1980; Kramer et al., 1990), using a double beam device similar to the flash spectrophotometer described in Appendix B. The field of application of the flash technique is similar to that of the modulated technique, but the signal to noise ratio is higher, and the actinic effect of the detecting flashes can be generally neglected. This technique is particularly well adapted to follow the kinetics of fluorescence yield changes following illumination by short actinic flashes (Joliot and Joliot, 1977). In the case ofxenon actinic flashes, is limited by the artifact the time resolution due to the luminescence emission induced by the strong actinic flash.

Chapter 22 In vivo Measurements III. Fluorescence Emission Spectra at Low Temperature Lowering the temperature greatly improves the resolution of the emission spectra, by sharpening the individual bands of the different fluorescent components. Moreover, the fluorescence yield of some long-wavelength emitting components is considerably increased. In C. reinhardtii, the broad fluorescence band from PS II observed at room temperature can be resolved into several components upon cooling the cells to liquid nitrogen temperature (77K) (Wollman and Bennoun, 1982; Tapie et al., 1984). Two emission peaks appear at 686 and 694 nm. A third peak near 715 nm originates from the PS I core, whereas at room temperature PS I is almost non-fluorescent. In mutants lacking the PS I core, a large emission band peaking around 707 nm dominates every other component at 77K. In the wild type, this emission arising from the PS I antenna is quenched by the PS I core. (PS II versus Changes in the ratio PS I fluorescence) have been extensively used as an indicator of state transitions. This ratio decreases upon transition from state 1 to state 2, as more excitation energy is directed towards the PS I centers at the expense of the PS II centers (Wollman and Delepelaire, 1984). Gans and Rébeillé (1990) observed that blocking mitochondrial ATP production ratio, leads to a decrease of the suggestive of a transition to state 2 via reduction of the plastoquinone pool. This interpretation was confirmed by Bulté et al. (1990), who demonstrated that the observed changes in the 77K-fluorescence spectrum indeed reflect reversible state transitions controlled by the intracellular demand for ATP.

IV. Delayed Fluorescence Measurements Delayed fluorescence is emitted in the same spectral band as fluorescence, and is associated with back reactions between PS II electron donors and acceptors (Lavorel, 1969; Barbieri et al., 1970). The luminescence intensity decays rapidly with time, according to a multi-exponential function. Detection of luminescence at times shorter than 1 ms after an actinic flash requires sophisticated devices that will not be described in this chapter. With the simplest device, in which the photoreceptor is not protected

439 from actinic flash excitation, luminescence kinetics can be analyzed from 10 ms to 10 s following the actinic flash. Luminescence measurements in vivo represent a powerful tool to qualitatively estimate the electrochemical proton gradient ( Joliot and Joliot, 1980), which modulates the rate of the back reaction between PS II electron donors and acceptors. In the presence of DCMU, the luminescence intensity depends mainly on the value of the membrane potential component, as the reduction of the state into hardly involves any proton uptake (Rappaport and Lavergne, 1991). In the absence of DCMU, the intensity of luminescence is much lower, owing to Under these conditions, the low concentration of the luminescence intensity depends both on the electrical and the osmotic component of the electrochemical proton gradient. Thermoluminescence is a light emission occurring when a material preilluminated at low temperature is warmed gradually in darkness. The study of themoluminescence provides useful information on photosynthetic transport, mostly in Photosystem II. On that subject, the reader is referred to reviews of Sane and Rutherford (1986) and Inoue (1996).

V. Oxygen Measurements Most of the methods used to measure photosynthetic oxygen emission in living algae are based on amperometric detection, using most generally a platinum electrode negatively polarized with respect to a reference electrode. In the Clark electrode (Clark, 1956), which is commercially available, a thin layer of Teflon permeable to oxygen protects the platinum electrode from any contaminant, inhibitor or redox component present in the suspension. The sensitivity of this type of electrode is not sufficient to measure oxygen evolution under weak light or flash excitation. The time resolution of a few seconds does not allow a detailed kinetic analysis of oxygen emission. Amperometric methods using a bare platinum electrode are much more sensitive. In one type of technique, the algal suspension is included in a circular closed cuvette (0.1 to 1 ml volume) equipped with a stirrer rotating at 3000 rpm (Joliot, 1956, 1965). The sensitivity is sufficient to measure the oxygen evolved following individual short flashes, with a signal-to-noise ratio better than 10. The noise level is proportional to the concentration of oxygen,

440 and for this reason is considerably decreased when working at low oxygen concentration (less than 10% of air saturation). The experiments are generally performed at a temperature close to 0° C, to slow down respiration and most biological processes. Under these conditions, the time resolution of 0.1 to 0.5 s allows a refined kinetic analysis of oxygen concentration changes. This technique allows measurement of oxygen evolution as well as that of oxygen consumption due to respiration or to the Mehler reaction (reduction ofoxygen via PS I electron acceptors (Mehler, 1951)). Measurement of oxygen evolution at the onset ofan illumination by saturating light reveals an ‘oxygen gush,’ with an amplitude proportional to the amount ofoxidized plastoquinone present prior to illumination (Joliot, 1965). This technique allows the measurement of absolute amounts of oxygen evolved, and therefore it can be used to determine the concentration of the PS II reaction centers and of the oxidized pool of plastoquinone on a chlorophyll basis. Blinks and Skow (1938) and Haxo and Blinks (1950) have introduced stationary electrodes, and were able to analyze oxygen emission under continuous light with high sensitivity. The complex time response of these methods does not allow quantitative analysis of the kinetics of oxygen emission. These methods have been adapted for measurements under modulated light or for short flash excitation (Joliot and Joliot, 1968). Chloroplasts or algae are sedimented on a horizontal platinum electrode (surface 0.2 to ). The high electrochemical capacitance requires the use of an amplifier with low input impedance, in order to limit the time constant to less than 1 ms. In the case of algae such as Chlamydomonas or Chlorella, optimal conditions are achieved when the cells form a monolayer at the surface of the electrode. When the photosynthetic material is subjected to modulated illumination (frequency <200 Hz), the amplitude of the modulated amperometric current is proportional to the average rate of oxygen emission. The modulated signal is detected using a lock-in amplifier, which rejects the components due to continuous illumination or to the oxygen uptake associated with respiration or the Mehler reaction. A time resolution of about 10 ms is obtained for a modulation frequency of 100 Hz. This method has the advantage of an excellent signal-to-noise ratio, and is especially well adapted to the measurement of action spectra under weak modulated light (1 photon per second per PS II

Pierre Joliot, Daniel Béal and René Delosme reaction center). The same technique can be used to measure the amount of oxygen evolved under excitation by a series of flashes. Provided the duration of the flash is sufficiently short (<1 ms), the amplitude of the signal is proportional to the amount of oxygen evolved. The amount evolved by a single flash is measured with a high signal-to-noise ratio

VI. Absorption Spectroscopy Redox change transients of a large fraction of the electron carriers involved in the photosynthetic electron transport chain can be measured in vivo by absorption spectroscopy. Electron transfer processes are induced by actinic illumination of the photosynthetic material (flashes or continuous light). The photoinduced absorption changes are measured by the variations of transmission of a weak monochromatic beam (detecting beam) passing through the biological sample. In the visible spectrum, the order of magnitude of the transmission changes induced by short saturating actinic flashes is Owing to the low amplitude of the biological signal, the noise level is a limiting parameter, which requires, in most of the cases, averaging of several experiments. The noise level decreases as the inverse square root of the number of experiments averaged. In a properly designed instrument, the noise level should be essentially limited by the quantum noise. If i is the photocathode the time resolution, the signal-tocurrent and noise ratio is, as in the fluorescence methods, where i is the intensity of the photocathode current (see Section II. A). As i is proportional to the intensity of the detecting beam, the signal-to-noise ratio is mainly limited by the integrated actinic effect of this beam during the period of measurement. This constraint is particularly severe in the case of Chlamydomonas cells or chloroplasts of higher plants, in which the antenna pigments increase the cross section of the reaction centers by a factor of about 200. The actinic effect of the detecting beam is inversely proportional to the surface of the cuvette. Thus increasing the surface of the cuvette, and therefore the amount of biological material studied, can minimize the actinic effect of the detecting beam. Kinetic analysis of electron transfer reactions in vivo is made more difficult by the strong light scattering of algal suspensions. Increasing the solid angle of collection of the measuring light can

Chapter 22

In vivo Measurements

minimize this apparent absorption. Large surface ) should be placed close photodetectors (about to the cuvettes. Using silicon photodiodes as photodetectors can minimize the quantum noise. The quantum yield of these photodetectors is close to 0.4 to 0.7 between 400 nm and 700 nm, versus about 0.06 for the S20 type photocathode of a photomultiplier in the green region of the spectrum.

A. Spectrophotometric Method using a Continuous Detecting Beam Most of the instruments devoted to analysis of photosynthetic electron transfer reactions are single beam spectrophotometers using continuous monochromatic light as the detecting beam (Witt, 1967). In the case of isolated chloroplast suspensions, which have better optical properties than algal suspensions, the highest signal-to-noise ratio reported in the for a time resolution literature is approx. of about 1 ms. On the basis of the published data, it is difficult to conclude whether the noise level is limited by the quantum noise, by the intensity of the detecting beam or by the fluctuations ofthe detecting beam. In order to minimize the integrated actinic effect of the detecting beam, the detecting beam should be opened only a short time before the onset ofthe actinic illumination, and the illumination should be as short as possible. In most cases, one must average the results from a large number of experiments to improve the signal-to-noise ratio. Few sets of data have been published reporting in vivo measurements with this type of conventional spectrophotometer.

B. Spectrophotometers using Detecting Flashes In this type of spectrophotometer, absorption is sampled by short monochromatic flashes of low energy produced by xenon tubes (Joliot et al., 1980; Joliot and Joliot, 1986; Kramer and Crofts, 1990). A detailed description of this type of instrument is presented in appendix B. In a new technique under development, monochromatic flashes are provided by a tunable optical parametric oscillator (OPO) pumped by a Nd:YAG laser (P. Joliot and D. Béal, unpublished). These instruments are double beam spectrophotometers, in which the monochromatic beam is divided in two beams illuminating a reference and a measuring cuvette. The time resolution is limited by the duration of the detecting flash, i.e.

441 in the case of xenon flashes, or 10 ns in the case of laser flashes. The kinetics of the absorption changes following a strong actinic flash are sampled by a series of appropriately spaced weak detecting flashes. In the case of multiphasic kinetics, the time interval between flashes should be adapted to the time constants of the different kinetic phases studied. About 20 detecting flashes are enough to characterize the time course of absorption change over a time to 10 s. In window which extends from spectrophotometers using xenon detecting flashes, technical reasons limit the shortest interval between two subsequent flashes to 5 ms. Therefore, kinetic ms requires analysis in the time interval several independent experiments. The signal-to-noise for photoelectrons emitted ratio (rms) is about by the photodetector per detecting flash, which corresponds roughly to the theoretical quantum noise. Suspensions of optical density up to 1 can be studied with this excellent signal-to-noise ratio, using xenon detecting flashes. The same signal-to-noise ratio is obtained with laser detecting flashes, but the higher available energy of the detecting flashes allows the analysis of algal suspensions of higher optical density (up to 3), yielding higher biological signals. In the case of the xenon flash device, when less than 20 detecting flashes are used, the integrated actinic effect ofthese flashes can be neglected. In the red region of the spectrum, the transmission signals are perturbed by chlorophyll fluorescence resulting from excitation either by the strong actinic flashes or by the detecting flashes. In this spectral range, accurate measurements require that no light-induced change occur in the fluorescence yield. These conditions are satisfied when the PS II centers are blocked in a closed state, or in mutants devoid of PS II centers. Therefore, experiments in the red region will address electron transfer reactions triggered by PS I, such as cyclic electron flow. When using red actinic flashes, the time resolution is limited to 1 ms, owing to the artifact associated with the large fluorescence emission excited by the actinic flash. In the UV spectral range, measurements are possible only at wavelengths higher than 290 nm, because of the large absorption band of the proteins. Flash spectrophotometric methods are well adapted to in vivo analysis, in view of their high signal-tonoise ratio. In several cases, the analysis of electron transfer reactions does not require the averaging of data from several experiments. This is particularly important when the experiment requires long dark

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adaptation. The signal-to-noise ratio can be considerably improved by using mutants with low antenna content, such as the BF4 strain of C. reinhardtii, particularly when working in the spectral range of the main absorption bands of chlorophyll. It is worth noting that in such mutants the absorption changes due to the formation of the membrane potential (electrochromic shift) are very small, as most of the electrochromic probes associated with the peripheral antenna are lacking (Lavergne et al., 1984). Figure 3 shows the absorption changes measured at 515 nm upon illumination by a saturating flash (curves A and B) or a weak flash (curve C), in a C. reinhardtii mutant devoid ofPS II, under anaerobic conditions. The absorption changes at 515 nm are mainly due to the electrochromic shift of membrane pigments (Junge and Witt, 1968), reflecting the formation of a delocalized membrane potential. The time course of the absorption change has been analyzed in the microsecond to second time range following the actinic flash. A fast absorption increase, is mainly due to the completed in less than

Pierre Joliot, Daniel Béal and René Delosme

formation ofthe membrane potential associated with the PS I charge separation (phase a). In addition, a fast absorption increase reflects the formation of a triplet state of carotenoid, which decays in the 0–50 time range. In the absence of oxygen, which is an efficient quencher of the triplet state, the lifetime of It is noteworthy the carotenoid triplet is about that in this particular mutant, the carotenoids that generate this signal are associated with PS I centers. time range, one observes a slow In the increase of the membrane potential (phase b) associated with electron and proton transfer within the cytochrome complex (Joliot and Delosme, 1974; Joliot and Joliot, 1994). Figure 4 shows the kinetics of phase b on a linear time scale, after normalization of the signal amplitudes measured at and deconvolution of the decay of the membrane potential. Addition of FCCP, which collapses the permanent electrochemical proton gradient, accelerates phase b by a factor of about 2 (Joliot and Joliot, 1994). Decreasing the energy of the flash further accelerates phase b (Joliot and Joliot, 1994). In the 20 ms to 5 s range, the absorption change at 515 nm reflects the decay ofthe membrane potential. In the absence of FCCP (curve A ofFig. 3), the decay is mainly due to dissipation of the proton gradient through the proton channel of the In the membrane ATP synthase

Chapter 22

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In vivo Measurements

presence of FCCP (curve B of Fig. 3), the ATP synthase is inactivated, and the decay ofthe membrane potential is mainly due to proton leaks induced by the uncoupler. The flash spectrophotometer can be used as a conventional spectrophotometer to analyze the absorption spectrum of algae in quasi-steady state conditions, as shown in Fig. 5. This spectrum (Joliot and Joliot, 1988) was obtained under anaerobic conditions in the mutant S52 of Chlorella sorokiniana, which lacks PS II and a large part of the pigment antenna. The spectrum shown in Fig. 5 is the difference between two spectra (not shown) drawn in different conditions. A first spectrum (1) was drawn after 1 hour of dark incubation. In these conditions, most of the cytochrome b in the cytochrome complex is in its reduced form. In order to compensate the drift due to settling ofthe algae, two spectra have been averaged: the first one was collected from short to long wavelengths, and the second one in the opposite direction. A second spectrum (2) was collected on algae dark-adapted for 30 s after illumination by a saturating flash. Under these conditions, heme is fully reduced, while most of stays in its oxidized form. The spectrum heme shown in Fig. 5 is the difference (1) – (2), which displays a pure reduced-minus-oxidized spectrum of heme

VII. Photoacoustic Measurements Upon excitation of an absorbing sample by light, volume changes occur in the sample and the surrounding medium, generating pressure waves. In dark-adapted photosynthetic material, two types of volume changes have to be considered. The first one is the expansion due to thermal conversion of part of the absorbed light energy (photothermal effect). The second one is due to conformational rearrangements associated with photoreactions. When all the reaction centers are closed (upon addition of inhibitors and saturating background light), absorbed light energy is completely dissipated The corresponding as heat, yielding reference signal is used for internal calibration ofthe amount of energy absorbed by the sample. The measures the energy stored difference in dark-adapted material, and, when divided by the efficiency of photochemical energy storage. Further division by (the wavelength of the radiation) gives, in relative units, the overall quantum yield of photochemical energy conversion. Note that the energy storage measurements are possible only can be neglected or subtracted from the when overall photoacoustic signal (see below). Different techniques have been developed to measure the photoinduced volume changes, using

444 either continuous modulated light or flash excitation.

A. Modulated Light Excitation Excitation by intensity-modulated light has been widely used in photoacoustic studies. Modulated heat is emitted by the illuminated sample at the same frequency as the excitation light. A gas-coupled microphone detects the thermally induced pressure wave. Any energy-storing product decaying with a time constant larger than the reciprocal of the angular modulation frequency is detected. Then, as the frequency increases, one can expect to sense more primary steps and measure more energy storage (Cahen et al., 1978). Note that the gas microphone in the measures primarily thermal expansion gas phase (owing to the large expansion coefficient of air), without any noticeable contribution of conformation changes (Lasser-Ross et al., 1980). In addition, Malkin and Cahen (1979) pointed out that gas exchanges could also give rise to modulated volume changes superposed on the photothermal changes, and Bults et al. (1982) showed that at low modulation frequency (approx. 100 Hz and below), a considerable fraction of the photoacoustic signal results from direct pressure modulation by modulated oxygen evolution (photobaric effect), whereas at high frequency (above 200 Hz), the main contribution is from conversion of modulated heat to modulated pressure. Modulated-light photoacoustic spectroscopy has been applied extensively to the evaluation of photosynthetic energy storage and oxygen evolution in vivo, especially at the Weizmann Institute in Rehovot. Canaani et al. (1989) studied energy distribution and state transitions in living cells of C. reinhardtii, by measuring photochemical energy storage and oxygen evolution, evaluated from the in-phase and quadrature (90° out of phase) vectorial components of the modulated photoacoustic signal, respectively. The same authors, followed by Ravenel et al. (1994), measured the energy storage induced by cyclic electron transfer around PS I.

B. Flash Excitation The use of photoacoustic spectroscopy to measure flash-induced volume changes in photosynthetic materials was introduced by Callis et al. (1972), and developed by Arata and Parson (1981). The volume and enthalpy changes accompanying photoreaction

Pierre Joliot, Daniel Béal and René Delosme in bacterial chromatophores (Callis et al., 1972) and reaction centers (Arata and Parson, 1981) were measured by a capacitor microphone in close contact with the sample suspension, on a time scale from to 1 s following the flash. Direct contact between the liquid phase and the diaphragm of the microphone revealed, in addition to the thermal expansion a contraction of approx. 33 Å per electron transferred, likely due to local electrostatic interactions between the photoinduced positive and negative charges and the surrounding medium (electrostriction) (Arata and Parson, 1981). These two components of the photoacoustic signal could be resolved by measuring the volume changes at two different temperatures, assuming that only is temperature dependent. An attractive feature of the flash detection used was the possibility of analyzing the complete relaxation kinetics of the light-induced volume changes. However, the response Another limitation ofthis time was limited to technique was the large volume of the cuvette (approx. 15 ml), which required the use of a large amount of biological material. A much higher time resolution (in the nanosecond to microsecond range) was introduced using laser optoacoustic spectroscopy, in which the heat released by a sample after absorption of a laser pulse is detected by a piezoelectric transducer (Patel and Tam, 1979). In the classical version, the acoustic wave is detected at right angles from the laser beam. The time resolution of heat detection is restricted by the duration of the laser pulse, the time response of the piezoelectric detector, and the transit time of the acoustic pulse across the diameter of the laser beam. per mm) is usually limiting. The latter (about Application of this technique to highly scattering materials such as intact plant tissues required a special optical arrangement to cancel the scattered-light induced signals. Following the first application in vivo by Jabben and Schaffner (1985) on intact leaves, a number of studies appeared in subsequent years, especially from S. E. Braslavsky and colleagues. More recently, Delosme et al. (1994) described a new high-sensitivity photoacoustic spectrometer but using operating in the same time window of a quite different geometry. The measuring pulsed light is distributed evenly (through a glass light guide) on the whole upper face (14 mm diameter) of a thin layer of photosynthetic material thickness). The total volume of the cuvette is only At the lower face of the measuring cell, the

Chapter 22

In vivo Measurements

445 presence of oxygen (see Section I. A.). In spectrum a, PS I and PS II reaction centers were in their photoactive state. The spectrum shows a small depression in the 650–680 nm range (compared to the 700 nm region), implying that a significant fraction of the energy absorbed by the peripheral antenna (LHC II) is not transferred to open reaction centers. In spectrum b, PS II activity was blocked by addition of DCMU and hydroxylamine, thus only PS I reaction centers were active. In this case, the spectral dependence of the quantum yield of PS I shows that most of the LHC II antenna does not transfer its excitation to PS I centers, whereas PS II (difference spectrum a – b) is substantially sensitized. An opposite situation is observed in state 2 (not shown), where most of the LHC II antenna is able to transfer its excitation to PS I centers (Delosme et al., 1994, 1996).

VIII. Conclusion and Perspectives fraction of incident light which has not been absorbed by the layer is reflected upwards by a glass-coated aluminum mirror. A piezoelectric ceramic (16 mm diameter) is glued to the back (lower) face of the mirror and detects the pressure waves propagating in the direction of the laser beam (note that another front-face illumination cell has been described by Melton et al. (1989)). In the technique used by Delosme et al., the theoretical response time corresponds to the transit time (about 30 ns) of the thickness of the sample. sound wave across the However, the authors adapted the instrument for detection in the range, using a ceramic of 1 MHz resonance frequency. The high signal-to-noise ratio of the method allows detection by very weak monochromatic flashes, which do not induce any significant actinic effect (about 1 photon per 400 reaction centers). This method has been used to analyze the spectral dependence of the quantum yield of the photosynthetic process in C. reinhardtii. The quantum yield spectra give information on the efficiency of energy transfer between the different pigments (chlorophyll and carotenoids) and PS I and PS II reaction centers, and are particularly useful for the quantitative study ofstate transitions. Figure 6 shows the spectral dependence of the quantum yield measured in cells of C. reinhardtii blocked in state 1 by benzoquinone treatment in the

In this chapter, we have shown that a refined functional analysis of the photosynthetic process can be performed on intact cells of Chlamydomonas, using biophysical techniques especially suited for this type of studies. Contrary to most biological processes, the photosynthetic electron transfer reactions can be easily triggered. The photons represent a ‘substrate’ that can be rapidly introduced within a living cell in a non destructive manner. The techniques described above allow one to measure the perturbations induced by light in a large variety of natural probes, including intact cells and purified photosynthetic complexes. Very likely, technical improvement of several other physical methods, such as magnetic spectrometry (EPR and NMR) will make them usable for in vivo studies (Amesz and Hoff, 1996). One can predict that, in a near future, there will be a growing interest for the analysis of the photosynthetic process in its normal physiological environment. The rapid progress of the reductionist approaches (mainly at the molecular level) will lead to a progressive shift of the interest of scientists toward the analysis of the structure-function relationship at higher levels of integration. In this domain, functional studies on living unicellular algae such as Chlamydomonas will provide an experimental basis for a better understanding of the mechanisms of photosynthesis, from the molecular to the cellular level.

446 Appendix A: Estimation of the Signal-toNoise Ratio in Fluorescence Measurements The quantum noise of a fluorescence technique can be estimated by computing the number of photoelectrons emitted by the photodetector during the time interval (see Section II.A). Let us suppose that a cuvette of volume V is filled with a suspension of Chlamydomonas cells, at the concentration of a liquid culture at the end of the exponential phase (about chlorophyll per ml). This corresponds to a concentration of approx. chlorophyll, or PS II centers. Assuming that during a time on average each PS II center absorbs 0.01 photon (see section IIA), the number of photons absorbed by the PS II centers in the cuvette is Assuming a minimum fluorescence quantum yield of approx. 2% ( level), the number of fluorescence photons emitted over a solid angle of Let us suppose that the fluorescence light is collected in a solid angle of 0.5 steradian, and that 25% of the collected light is transmitted by the red filter. Then 1 % of the total fluorescence emission reaches the photodetector. When using a photomultiplier equipped with a S20 type photocathode (quantum yield 0.05 in red light), the number of photoelectrons emitted during is Using a cuvette of 0.1 ml or the number N of photoelectrons available is and the signal-to-noise ratio is about 300. The same type of approach can be used to compute the quantum noise of a spectrophotometric method, taking into account the time resolution, and the acceptable actinic effect of the detecting light during the measurements.

Appendix B: Flash Spectrophotometer In the double beam spectrophotometer that has been developed by P. Joliot and D. Béal (Fig. 7A), short monochromatic flashes sample the absorption. A xenon flash (Hamamatsu) illuminates the entrance slit of a monochromator (Jobin et Yvon). This monochromator of large aperture (f/2) is equipped with a concave holographic grating (Jobin et Yvon, 1200 grooves/mm, 150 mm diameter), which covers a large spectral range (280 to 750 nm). From the output slit of the monochromator (a), the beam is divided in two parts (c), to illuminate a reference and

Pierre Joliot, Daniel Béal and René Delosme a measuring cuvette (e) filled with the suspension of algae. The light transmitted through the measuring and reference cuvettes is measured by two photodiodes PIN (Hamamatsu S3204-06) of large surface (18 × 18 mm) (h), allowing collection of scattered light over a wide angle. The photocurrent is integrated The integrated with a time constant of about output signals from the two preamplifiers enter a differential amplifier. The output signal from the differential amplifier is sampled after the onset of the flash. In the same way, the output signal from the preamplifier of the reference photodiode is after the flash. The time resolution of sampled i.e. the time interval between the the method is onset of the flash and the sampling. A 16 bit analogdigital converter digitizes the sampled signals from the reference preamplifier and the differential amplifier. The ratio

is computed for each detecting flash. Provided the number of photoelectrons available on both channels and the fluctuations of the reference is larger than and the measuring beam are well correlated (see is independent below), the ratio of the energy fluctuations of the flash, even when the reference and the measuring channel are not accurately balanced (ratiometer). Using a gain 10 differential amplifier and a 16 bit analog-digital converter, the ratio R is digitized with a precision of i.e. better than the noise level The computer generates the pulses which trigger the detecting flashes and the actinic light (flash or continuous illumination). In a flash spectrophotometer, the more critical parameter is the optical symmetry between the reference and the measuring beam. For instance, when using a conventional beam splitter (semitransparent mirror) to separate the reference and the measuring beam, the noise level at the output of the differential amplifier could be larger than irrespective of the number ofphotoelectrons available from the photodetector. In this case, the energy fluctuations on the reference and the measuring channel are mainly uncorrelated. A satisfying degree of symmetry is obtained only when the light emitted by the xenon flash bulb is mixed and randomized several times by a series of solid and optical fiber

Chapter 22

In vivo Measurements

light pipes. The xenon flashbulb is applied against the input of an optical-fiber light pipe, in which the fibers are randomly distributed between the input and the output face. The output of this optical-fiber light pipe is applied against a quartz light pipe with a frosted entrance face. The output of the quartz light pipe illuminates the entrance slit of the monochromator. The section of the quartz and the opticalfiber light pipes is equal to that of the slit of the monochromator (3 × 20 mm). A second quartz light pipe (3 × 20 × 30 mm) (b in Fig. 7A) is interposed between the output slit of the monochromator (a) and the common arm of an Y-shaped optical-fiber light pipe (c), which divides the detecting beam into a reference and a measuring beam. The fibers are randomly distributed between the two branches. These light pipes in series absorb a large fraction of the light emitted by the flash, which explains the need for a large aperture monochromator. In addition, the measuring and reference cuvette should be identical and filled with biological material at the same concentration. With well symmetrically arranged reference and measuring beams, and when the number N of photoelectrons available per detecting flash is

447

then the noise level in the ratio such as R is limited by the quantum noise. For lower values ofN, the noise of the preamplifiers is limiting, while for higher value of N the noise is limited by the remaining small optical asymmetry between the reference and the measuring beam. Cuvettes of various shapes can be placed at the output of the Y-shaped light guide, provided that the reference and measuring cuvettes are identical. In the instrument shown in Fig. 7B, the reservoir (b) and cuvette (e) are made of stainless steel. Actinic and detecting light enter the same optical face of the cuvette (5 mm × 5 mm × 15 mm), using a light pipe (d) in which the optical fibers ofthe detecting and the actinic beam are mixed. In this device, both cuvettes can be indifferently used for the reference or the measurement. The illuminated sample can be rapidly renewed with dark-adapted material, and can be incubated either aerobically or anaerobically in the reservoir. Owing to the small size of the cuvette, the absorption of the sample is stabilized in less than 4 s after renewing the sample. The reservoir could be adapted for redox and pH titration.

448 Acknowledgment The authors are pleased to thank F. Rappaport for critical reading of the manuscript.

References Amesz J and Hoff AJ (eds) (1996) Part two: Magnetic resonance. In: Biophysical Techniques in Photosynthesis, pp 209–313. Kluwer Academic Publishers, Dordrecht Arata H and Parson WW (1981) Enthalpy and volume changes accompanying electron transfer from P-870 to quinones in Rhodopseudomonas sphaeroides reaction centers. Biochim Biophys Acta 636: 70–81 Barber J and Kraan GPB (1970) Salt-induced light emission from chloroplasts. Biochim Biophys Acta 197: 49–59 Barbieri G, Delosme R and Joliot P (1970) Comparaison entre l’émission d’oxygène et l’émission de luminescence à la suite d’une série d’éclairs saturants. Photochem Photobiol 12: 197–206 Bennoun P (1982) Evidence for a respiratory chain in the chloroplast. Proc Natl Acad Sci USA 79: 4352–4356 Blinks LR and Skow RK (1938) The time course ofphotosynthesis as shown by a rapid electrode method for oxygen. Proc Natl Acad Sci USA 24: 420–427 Bonaventura C and Myers J (1969) Fluorescence and oxygen evolution from Chlorella pyrenoidosa. Biochim Biophys Acta 189: 366–383 Bulté L and Wollman FA (1990) Stabilization ofstates I and II by p-benzoquinone treatment of intact cells of Chlamydomonas reinhardtii. Biochim Biophys Acta 1016: 253–258 Bulté L, Gans P, Rébeillé F and Wollman FA (1990) ATP control on state transitions in vivo in Chlamydomonas reinhardtii. Biochim Biophys Acta 1020:72–80 Bults G, Horwitz B A, Malkin S and Cahen D (1982) Photoacoustic measurements of photosynthetic activities in whole leaves: Photochemistry and gas exchange. Biochim Biophys Acta 679: 452–465 Cahen D, Malkin S and Lerner El (1978) Photoacoustic spectroscopy of chloroplast membranes; listening to photosynthesis. FEBS letters 91: 339–342 Callis JB, Parson WW and Gouterman MM (1972) Fast changes of enthalpy and volume on flash excitation of Chromatium chromatophores. Biochim Biophys Acta 267: 348–362 Canaani O, Schuster G and Ohad I (1989) Photoinhibition in Chlamydomonas reinhardtii: Effect on state transition, intersystem energy distribution and Photosystem I cyclic electron flow. Photosynth Res 20: 129–146 Clark LC (1956) Monitor and control of blood and tissue oxygen tensions. Trans Am Soc Art Int Organs 2: 41–45 Davenport J W and McCarty RE (1981) Autocatalytic activation of thylakoid ATPasc. I n : Akoyunoglou G (ed) Photosynthesis, Vol II, pp 859–865. Balaban International Science Services, Philadelphia Delepelaire P and Wollman FA (1985) Correlations between fluorescence and phosphorylation changes in thylakoid membranes of Chlamydomonas reinhardtii in vivo: A kinetic analysis. Biochim Biophys Acta 809: 277–283

Pierre Joliot, Daniel Béal and René Delosme Delosme R, Béal D and Joliot P (1994) Photoacoustic detection of flash-induced charge separation in photosynthetic systems. Spectral dependence of the quantum yield. Biochim Biophys Acta 1185: 56–64 Delosme R, Olive J and Wollman FA (1996) Changes in light energy distribution upon state transitions: An in vivo photoacoustic study of the wild type and photosynthesis mutants from Chlamydomonas reinhardtii. Biochim Biophys Acta 1273: 150–158 Diner B and Joliot P (1976) Effect of the transmembrane electric field on the photochemical and quenching properties of Photosystem II in vivo. Biochim Biophys Acta 423: 479–498 Dutton HJ, Manning WM and Duggar BM (1943) Chlorophyll fluorescence and energy transfer in the diatom Nitzschia closterium. J Phys Chem 47: 308–313 Finazzi G, Büschlen S, de Vitry C, Rappaport F, Joliot P and Wollman FA (1997) Function-directed mutagenesis of the cytochrome complex in Chlamydomonas reinhardtii: Involvement of the cd loop of cytochrome in quinol binding to the site. Biochemistry 36: 2867–2874 Gans P and Rébeillé F (1990) Control in the dark of the plastoquinone redox state by mitochondrial activity in Chlamydomonas reinhardtii. Biochim Biophys Acta 1015: 150–155 Haxo FT and Blinks LR (1950) Photosynthetic action spectra of marine algae. J Gen Physiol 33: 389–422 Inouc Y (1996) Photosynthetic thermoluminescence as a simple probe of Photosystem II electron transport. In: Amesz J and Hoff AJ (eds) Biophysical Techniques in Photosynthesis, pp 93–107. Kluwer Academic Publishers, Dordrecht Jabben M. and Schaffner K (1985) Pulsed-laser induced optoacoustic spectroscopy of intact leaves. Biochim Biophys Acta 809: 445–451 Joliot P (1956) Dispositif ampérométrique de mesure de la photosynthèse. C R Acad Sci Paris 243: 677–680 Joliot P (1965) Cinétiques des réactions liées à 1’émission d’oxygène photosynthétique. Biochim Biophys Acta 102: 116– 134 Joliot P and Delosme R (1974) Flash-induced 519 nm absorption change in green algae. Biochim Biophys Acta 357: 267–284 Joliot P and Joliot A (1968) A polarographic method for detection of oxygen production and reduction of Hill reagent by isolated chloroplasts. Biochim Biophys Acta 153:625–634 Joliot P and Joliot A (1977) Evidence for a double hit process in Photosystem II based on fluorescence studies. Biochim Biophys Acta 462: 559–574 Joliot P and Joliot A (1980) Dependence ofdelayed luminescence upon adenosine triphosphatase activity in Chlorella. Plant Physiol 65: 691–696 Joliot P and Joliot A (1986) Proton pumping and electron transfer in the cytochrome b/f complex of algae. Biochim Biophys Acta 849: 211–222 Joliot P and Joliot A (1988) The low-potential electron-transfer chain in the cytochrome b/f complex. Biochim Biophys Acta 933: 319–333 Joliot P and Joliot A (1989) Characterization of linear and quadratic electrochromic probes in Chlorella sorokiniana and Chlamydomonas reinhardtii. Biochim Biophys Acta 975:355– 360 Joliot P and Joliot A (1994) Mechanism of electron transfer in the cytochrome b/f complex of algae: Evidence for a scmiquinonc

Chapter 22 In vivo Measurements cycle. Proc Natl Acad Sci USA 91: 1034–1038 Joliot P, Béal D and Frilley B (1980) Une nouvelle méthode spectrophotométrique destinée à 1’étude des réactions photosynthétiques. J Chim Phys 77: 209–216 Junge W and Witt HT (1968) On the ion transport system of photosynthesis. Investigations on a molecular level. Z Naturforsch 23b: 244–254 Kautsky H and Hirsch A ( 1 9 3 1 ) Chlorophyllfluoreszenz und Kohlensäureassimilation. Natnrwissenschaften 19: 964 Kok B, Forbush B and McGloin M (1970) Cooperation of charges in photosynthetic evolution – I. A linear four step mechanism. Photochem Photobiol 1 1 : 457–475 Kramer DM and Crofts AR (1990) Demonstration of a highlysensitive portable double-flash kinetic spectrophotometer for measurement of electron transfer reactions in intact plants. Photosynth Res 23: 231–240 Kramer DM and Crofts AR (1996) Control and measurement of photosynthetic electron transport in vivo. In: Baker NR (ed) Photosynthesis and the Environment, pp 25–66. K l u w e r Academic Publishers, Dordrecht Kramer DM, Robinson HR and Crofts AR (1990) A portable multi-flash kinetic fluorimeter for measurement of donor and acceptor reactions of Photosystem 2 in leaves of intact plants under field conditions. Photosynth Res 26: 181–193 Lasser-Ross N, M a l k i n S and Cahen D (1980) Photoacoustic detection of photosynthetic activities in isolated broken chloroplasts. Biochim Biophys Acta 593: 330–341 Lavergne J, Delosme R, Larsen U and Bennoun P (1984) Mutants of Chlorella sorokiniana: A new material for photosynthesis studies. II. Improved spectroscopic analysis ofelectron transfer in mutant strains. Photochem Photobiophys 8: 207–219 Lavorel J (1969) On a relation between fluorescence and luminescence in photosynthetic systems. I n : Metzner H (ed) Progress in Photosynthesis Research, Vol II, pp 883–898. H. Laupp Jr. Tübingen Lasser-Ross N, M a l k i n S and Cahen D (1980) Photoacoustic detection of photosynthetic activities in isolated broken chloroplasts. Biochim Biophys Acta 593: 330–341 M a l k i n S and Cahen D (1979) Photoacoustic spectroscopy and radiant energy conversion: Theory of the effect with special emphasis on photosynthesis. Photochem Photobiol 29: 803– 813 Mehler A ( 1 9 5 1 ) Studies on the reaction of i l l u m i n a t e d chloroplasts. Mechanism of the reduction of and other Hill reagents. Arch Biochem Biophys 33: 65–77 Melton LA, Ni T and Lu Q (1989) Photoacoustic calorimetry. A new cell design and improved analysis algorithms. Rev Sci Instrum. 60: 3217–3223 M i l e s CD and Jagendorf AT (1969) Ionic and pH transitions triggering chloroplast post-illumination luminescence. Arch Biochem Biophys 129: 7 1 1 – 7 1 9 Mitchell P (1966) Chemiosmotic coupling in oxidative and photosynthetic phosphorylation. Biol Rev 41: 445–502

449 Patel CKN and Tam AC (1979) Optoacoustic spectroscopy of liquids. Appl Phys Lett 34:467–470 Peltier G and Thibault P (1988) Oxygen-exchange studies in Chlamydomonas mutants deficient in photosynthetic electron transport: evidence for a Photosystem II-dependent oxygen uptake in vivo. Biochim Biophys Acta 936: 319–324 Rappaport F and Lavergne J (1991) Proton release during successive oxidation steps of the photosynthetic water oxidation process: stoichiometries and pH dependence. Biochemistry 30: 10005–10012 Ravenel J, Peltier G and Havaux M (1994) The cyclic electron pathways around Photosystem I in Chlamyclomonas reinhardtii as determined in vivo by photoacoustic measurements of energy storage. Planta 193: 251–259 Rébeillé F and Gans P (1988) Interaction between chloroplasts and mitochondria in microalgae: role of glycolysis. Plant Physiol 88: 973–975 Rumberg B a n d Siggel U (1969) pH changes in the inner phase of the thylakoids during photosynthesis. Naturwissenschaften 56: 130–132 Sane PV and Rutherford WA (1986) Thermoluminescence from photosynthetic membranes. In: Govindjee, Amesz J and Fork DC (eds) Light Emission by Plants and Bacteria, pp 329–360. Ac ademic Press, Orlando Schreiber U (1986) Detection of rapid induction kinetics with a new type of high-frequency modulated chlorophyll fluorometer. Photosynth Res 9: 261–272 Tapie P, Choquet Y, Breton J, Delepelaire P and Wollman FA (1984) Orientation of Photosystem-I pigments: Investigation by low-temperature linear dichroism and polarized fluorescence emission. Biochim Biophys Acta 767: 57–69 Vallon O, Bulté L, Dainese P, Olive J, Bassi R and Wollman FA (1991) Lateral redistribution of cytochrome complexes along thylakoid membranes upon state transitions. Proc Natl Acad Sci USA 88: 8262–8266 Velthuys BR and Amesz J ( 1 9 7 4 ) Charge accumulation at the reducing side of system 2 of photosynthesis. Biochim Biophys Acta 333: 85–94 Witt HT (1967) Direct measurements of reactions in the to second range by single and repetitive excitations with pulses of electromagnetic waves. In: Claesson S (ed) Fast Reactions and Primary Processes in Chemical Kinetics, Nobel Symposium 5, pp 8 1 – 9 7 . Interscience Publishers, New York Wollman FA and Bennoun P (1982) A new chlorophyll-protein complex related to Photosystem I in Chlamydomonas reinhardtii. Biochim Biophys Acta 680: 352–360 Wollman FA and Delepelaire P (1984) Correlation between changes in light energy distribution and changes in thylakoid membrane. Polypeptide phosphorylation in Chlamydomonas reinhardtii. J Cell Biol 98: 1–7 Wraight CA and Crofts AR ( 1 9 7 1 ) Delayed fluorescence and the high-energy state of chloroplasts. Eur J Biochem 19: 386–397

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Chapter 23

New Digital Imaging Instrument For Measuring Fluorescence and Delayed Luminescence Pierre Bennoun and Daniel Béal

Institut de Biologie Physico-Chimique, Centre National de la Recherche Scientifique,

UPR 9072, 13, rue Pierre & Marie Curie, 75005 Paris, France

Summary I. Introduction II. Setup for Fluorescence and Delayed Luminescence Video Imaging III. Digital Fluorescence Imaging Related to Photosynthetic Electron Transfer IV. Digital Fluorescence Imaging Related to the Permanent Thylakoid Electrochemical Gradient V. Digital Delayed Luminescence Imaging Related to Light-Induced and Permanent Thylakoid Electrochemical Gradient Acknowledgments References

451 452 452 452 453 455 457 457

Summary We describe a new digital imaging instrument designed to monitor fluorescence and delayed luminescence of photosynthetic systems (algal or bacterial colonies grown on Petri dishes, leaves). The setup includes lightemitting diodes, a cooled line transfer charge-coupled device (CCD) camera, a personal computer and appropriate software which allows capture of pictures and kinetics in real time during illumination. This instrument provides fluorescence images under actinic or non-actinic illumination and makes the fluorescence induction kinetics of individual colonies readily accessible with good time resolution. It brings great refinement to procedures for screening colonies that are altered in photosynthetic electron transfer owing to appropriately computed fluorescence images. This instrument allows, for the first time, to capture delayed luminescence images, thus opening the way to screening for mutants that have an altered permanent or light-induced electrochemical gradient across thylakoid membranes.

J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 451–458. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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I. Introduction The primary processes of photosynthesis include both redox reactions through an electron transfer chain and vectorial charge transfers which generate an electrochemical gradient across thylakoid membranes. The chlorophyll fluorescence emission provides a non-invasive method for studying photosynthetic electron transfer in thylakoid membranes in vivo. The fluorescence induction observed upon a dark-light transition provides direct information on the overall electron flow through the photosynthetic electron transport chain, inasmuch as fluorescence is inversely related to the photochemical activity of Photosystem II (PS II) reaction centers. This approximation is indeed valid at the beginning of the light phase (for details, see Chapter 22, Joliot et al.). Screening mutant colonies of algae that have impaired photosynthetic electron transfer according to their in vivo chlorophyll fluorescence emission has thus been exploited successfully for a long time (Bennoun and Levine, 1967; Garnier, 1967; Bennoun and Chua, 1976; Bennoun et al., 1978; Bennoun and Delepelaire, 1982). Recent improvements of this method were achieved by incorporating video imaging technology (Omassa et al., 1987; Daley et al., 1989; Fenton and Crofts, 1990; Goldman and Youvan, 1995; Kramer and Crofts, 1996). The delayed luminescence ofphotosynthetic systems results from a back reaction occurring between the primary photoproducts of PS II reaction centers, which is stimulated by the electrochemical gradient across thylakoid membranes (see Chapter 22, Joliot et al.). Delayed luminescence is thus a direct sensor of this gradient, but its use in the context of screening mutant colonies of algae has been limited by its very low intensity. The high sensitivity of our instrument allows, for the first time, to perform delayed luminescence digital imaging and therefore to detect algal colonies displaying alterations of their permanent or light-driven thylakoid electrochemical gradient. We report on a new fluorescence/ luminescence digital imaging instrument, which significantly improves the detection of algal colonies that are photosynthetically impaired (Bennoun and Béal, 1997). Abbreviations: CCD–charge coupled device; CR–dicyclohexyl18-crown-6; DCMU–3-(3,4-dichlorophenyl)-1,1-dimethylurea; LED – light emitting diode; PQ – plastoquinone; PS I – Photosystem I; PS II – Photosystem II; QA–primary quinone of PS II reaction centers; Ø – diameter

Pierre Bennoun and Daniel Béal II. Setup for Fluorescence and Delayed Luminescence Video Imaging The instrument is depicted in Scheme 1. The camera used allows the capture of a complete video-image in 40 ms, i.e. a frame made of two fields. The video signal is captured using a Matrox PIP 1024 B acquisition card with 1 Mb memory, which allows the acquisition of four pictures (512 × 512 × 8 bits). The acquisition times are distributed at will during an illumination. Any algebraic combination of pictures can be computed in order to highlight a given type of information. A Pentium P 120 personal computer is used. Our homemade software written in C controls the progress of the experiments, the treatment of the data and their visualization. Using the synchronization signals of the video camera, the printer port controls the LED current, the electromechanical shutter and the integration mode of the camera. The display is achieved using false colors, providing visual amplification of the gradation intensity in the image; black and white display is also available for video or laser printing. For delayed luminescence imaging, the red filter is omitted and an electromechanical shutter is used in order to protect the CCD during actinic flashes (the closing and opening cycle lasts 10 ms). The camera is used in the integration mode while illumination is given by a series of flashes of 400 duration fired 80 ms apart. Delayed luminescence is thus detected for 70 ms at each cycle and integrated over the number of cycles determined by the number of flashes. III. Digital Fluorescence Imaging Related to Photosynthetic Electron Transfer In order to analyze leaves, or algal colonies grown in a Petri dish, it is important to illuminate a large area in an homogenous way. The combination of lightemitting diodes and a light pipe meets this requirement. A good homogeneity of illumination throughout the photosynthetic material is obtained by using wavelengths of light which are poorly absorbed by this material (594 nm in the present case). A correction is made for the difference in distances of the various points of the picture from the camera lens and for the defects in homogeneity of illumination. This can be achieved by dividing the fluorescence picture by the picture of a plain translucent surface at the same location. The

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acquisition of four pictures is possible at any time of the fluorescence rise during a continuous illumination (acquisition time is 40 ms). One such image, acquired at t = 2000 ms, is shown in Fig. 1A. In practice, screening algal mutant colonies on a Petri dish is best achieved on computed images resulting from algebraic combinations of these pictures. The computed images are designed so as to optimize the detection of a given class of mutants, as well as to correct for the various amount ofchlorophyll analyzed by each pixel (due to variable thickness of colonies, for instance). These qualities are demonstrated in Figs. 1A, C and D and Fig. 2B. The possibility of selecting individual colonies on a plate is given by positioning the cursor with a mouse and recording their fluorescence kinetics in one sweep with a good signal-to-noise ratio and a good time resolution (Figs. 1B and 2B). Up to six zones of a plate can be selected on a picture and the fluorescence of each zone computed and normalized to the area of this zone (the area of a zone can either fit automatically the area of a colony, or be restricted to a constant pixel number). The software allows significant measurements to be made down to colony diameters of 0.5 mm (4 × 4 pixels). Each point of the fluorescence rise corresponds to an integration of the signal for 40 ms. In addition to the detection of defects in the photosynthetic electron transfer, it is possible to evaluate the pool size of electron acceptors to PS II. As shown in Fig. 2, the decrease of this pool size induced by addition of DCMU is readily detected (DCMU isolates the primary quinone QA from the plastoquinone pool). IV. Digital Fluorescence Imaging Related to the Permanent Thylakoid Electrochemical Gradient The thylakoid electrochemical gradient consists of a permanent component observed in dark-adapted algae and a light-driven component. The permanent component exists in dark-adapted algae even in the synthase (Bennoun, absence of chloroplast 1982) and is thought to result from the hydrolysis of mitochondrial ATP by a thylakoid-bound vacuolartype of ATPase (Bennoun, 1994). Since the primary photoproducts of PS II reaction centers are localized on either side of the thylakoid membranes, their recombination rate is enhanced by the electrochemical gradient across these membranes (Kraan et al., 1970;

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Joliot and Joliot, 1974; Lavergne and Etienne, 1980). The back reaction is accompanied by a decrease in fluorescence due to the reoxidation of the primary semiquinone and by a delayed luminescence emission due to charge recombination (Arthur and Strehler, 1957; Lavorel, 1968; Bennoun, 1971). In the presence of DCMU, the reoxidation of proceeds entirely through charge recombination. The permanent thylakoid gradient can then be estimated through by measuring the reoxidation of fluorescence detection. Fluorescence imaging of this back reaction is achieved here by taking fluorescence pictures with detecting flashes of light which do not

Pierre Bennoun and Daniel Béal

This is in appreciably shift the redox state of contrast to fluorescence imaging of photosynthetic systems usually achieved by using a strong continuous illumination that is actinic and detecting at the same time. Figure 3 shows an image computed from several fluorescence pictures captured in the course of the back reaction following a single subsaturating actinic flash (see figure legend). The four colonies that have been treated with the ionophore CR are readily distinguishable: in the presence of CR, the permanent gradient is abolished and the rate ofthe back reaction reaches its lowest value (Bennoun, 1994). As shown below, a detection of this gradient can also be

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achieved, even in the absence of DCMU, through delayed luminescence imaging.

V. Digital Delayed Luminescence Imaging Related to Light-Induced and Permanent Thylakoid Electrochemical Gradient Light-induced delayed luminescence is an increasing function of the following parameters: i) The number of active PS II centers. ii) The rate of the backreaction, which is enhanced by the electrochemical gradient built up across thylakoid membranes. iii) The chlorophyll fluorescence yield which is mainly and which controlled by the redox state of determines the probability of an exciton to yield a photon. The three parameters should be taken into account when luminescence is to be used as a sensor of the thylakoid electrochemical gradient. In the absence of an active chloroplast synthase, the light-driven electrochemical gradient coupled to photosynthetic electron transfer cannot be dissipated to form ATP. Under high light, it will reach abnormally high values and delayed luminescence will be stimulated. Mutant strains deficient in chloroplast synthase have been characterized previously in this way (Bennoun and Chua, 1976; Bennoun et al., 1980; Bennoun and Delepelaire, 1982a; Woessner et al., 1984). Because of its very low intensity, the delayed luminescence light emitted

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by individual colonies could not be recorded either on an infrared film or using a simple CCD video camera. This difficulty is now overcome with the help of a cooled CCD camera. The low temperature allows the accumulation of photoelectrons on the CCD for a long time with a very low background noise. Since each reading introduces a significant noise to the signal, it is highly advantageous to sum up several experiments in a single reading. Delayed luminescence imaging to reveal the lightdriven electrochemical gradient of algal colonies is achieved by a series of short illuminating flashes fired 80 ms apart. For wild-type colonies, this interval is long enough for the dissipation of the electrochemical gradient built up by one flash through ATP synthesis before the next flash of the series is fired. However, in the case of mutant colonies lacking synthase, an abnormally high active gradient is formed during the series of flashes and luminescence is stimulated. The luminescence picture shown in Fig. 4B allows a clear distinction between the two kinds of colonies. This distinction no longer exists when one uses a series of flashes adequately spaced out (1s or more), to allow relaxation of the light-driven electrochemical gradient between two ATP synthase is flashes, whether active present or not (Fig. 4A). The ratio of these two pictures is insensitive to possible variations in the amount of chlorophyll or of PS II centers in each colony, as well as to optical defects. It truly reflects the stability of the light-induced electrochemical gradient (Fig. 4C). The mutant of Chlorella synthase, sorokiniana, deficient in shown in Fig. 4 (SL8.1) was identified by application of this method of delayed luminescence imaging. Previous attempts at screening such mutants of C. sorokiniana had failed, owing to lack of an appropriate procedure. Delayed luminescence imaging to reveal the permanent thylakoid electrochemical gradient is achieved using a series of non saturating flashes sufficiently spaced out (Fig. 4D): colonies treated with CR, in which this gradient is abolished, are readily distinguishable by their very low luminescence yield. The isolation of mutants with alterations in the permanent gradient should allow progress towards an understanding of its origin and function. One should note that the intensity of delayed luminescence light detected in the experiments depicted in Fig. 4 is about a thousand times weaker than the intensity of fluorescence light detected in the experiments

457 depicted in Fig. 1. A linear gray scale was used in all figures of this chapter for the convenience of the editor, although false color display is ordinarily used and allows a much better discrimination between colonies with different brilliance. A color version of Fig. 4 is shown as Color Plate 4 together with the scale of false color used.

Acknowledgments The authors are pleased to acknowledge R. Delosme and O. Vallon for critical reading of the manuscript. This work was supported by UPR 9072 of the Centre National de la Recherche Scientifique.

References Arthur WE and Strehler BL (1957) Studies on the primary process in photosynthesis. I: Photosynthetic luminescence multiple reactants. Arch Biochem Biophys 70: 507–526 Bennoun P (1971) Étude de la luminescence des chloroplastes d’épinards en présence de DCMU. C R Acad Sci Paris 273: 2654–2657 Bennoun P (1982) Evidence for a respiratory chain in the chloroplast. Proc Natl Acad Sci USA 79: 4352–4356 Bennoun P (1994) Chlororespiration revisited: mitochondrialplastid interactions in Chlamydomonas. Biochim Biophys Acta 1186: 59–66 Bennoun P and Béal D (1997) Screening algal mutant colonies with altered thylakoid electrochemical gradient through fluorescence and delayed luminescence digital imaging. Photosynth Res 51: 161–165 Bennoun P and Chua NH (1976) Methods for the detection and characterization of photosynthesis mutants in Chlamydomonas reinhardtii. In: Bücher TH, Neupert W, Sebald Wand Werner S, (eds), Genetics and Biogenesis of Chloroplasts and Mitochondria, pp 33–39. Elsevier Publishers, Amsterdam Bennoun P and Delepelaire P. (1982) Isolation of photosynthesis mutants in Chlamydomonas. In: Edelman M, Hallick RB and Chua NH, (eds.) Methods in Chloroplast Molecular Biology, pp 25–38. Elsevier Publishers, Amsterdam Bennoun P and Levine RP (1967) Detecting mutants that have impaired photosynthesis by their increased level of fluorescence. Plant Physiology 42: 1284–1287 Bennoun P, Masson A, Piccioni R and Chua NH (1978) Uniparental mutants of Chlamydomonas reinhardtii defective in photosynthesis. I n : Akoyunoglou and ArgiroudiAkoyounoglou JH, (eds) Chloroplast Development, pp 721– 726. Elsevier Publishers, Amsterdam Bennoun P, Masson A and Delosme M (1980) A method for complementation analysis of nuclear and chloroplast mutants of photosynthesis in Chlamydomonas. Genetics 95: 39–47 Daley PF, Raschke K, Ball TJ and Berry JA (1989) Topography of photosynthetic activity of leaves obtained from video images of chlorophyll fluorescence. Plant Physiol 90: 1233–1238

458 Fenton JM and Crofts AR (1990) Computer aided fluorescence imaging of photosynthetic systems. Photosynth Res 26: 59–66 Gamier J (1967) Une méthode de détection par photographic de souches d’algues vertes émettant une fluorescence anormale. CR Acad Sci Paris 265D: 874–877 Goldman ER and Youvan DC (1995) Imaging spectroscopy and combinatorial mutagenesis of the reaction center and light harvesting II antenna. In: Blankenship RE, Madigan MT and Bauer CE (eds) Anoxygenic Photosynthetic Bacteria, pp 1257– 1268. Kluwer Academic Publishers, Dordrecht Joliot P and Joliot A (1974) Comparative study of the 520 nm absorption change and delayed luminescence in algae. In: Avron M (ed.) Proceedings ofthe Third International Congress on Photosynthesis pp 25–39. Elsevier Publishers, Amsterdam Kraan GPB, Amesz J, Velthuis BR and Steemers RG (1970) Studies on the mechanism of delayed and stimulated delayed fluorescence of chloroplasts. Biochim Biophys Acta 223: 129– 145 Kramer DM and Crofts AR (1996) Control and measurement of

Pierre Bennoun and Daniel Béal photosynthetic electron transport in vivo. In: Baker NR (ed) Photosynthesis and the Environment, pp 25–66. Kluwer Academic Publishers, Dordrecht Lavergne J and Etienne AL (1980) Radiative and non radiative recombination in Photosystem II following flash illumination. In: Akoyunoglou G (ed.) Proceedings of the Fifth International Congress on Photosynthesis, Vol III, pp 939–948. Balaban Intl Sci Publishers, Philadelphia Lavorel J (1968) Sur une relation entre fluorescence et luminescence dans les systémes photosynthétiques. Biochim Biophys Acta 153: 727–730 Omassa K, Shimazaki K, Aiga I, Larcher W and Onoe M (1987) Image analysis of chlorophyll fluorescence for diagnosing the photosynthetic system of attached leaves. Plant Physiol 84: 748–752 Woessner JP, Masson A, Harris E, Bennoun P, Gillham NW and Boynton JE (1984) Molecular and Genetic Analysis of the Chloroplast ATPase of Chlamydomonas. Plant Mol Biol 3: 177–190

Chapter 24 The Structure, Function and Biogenesis Of Cytochrome Complexes Francis-André Wollman

Service de photosynthèse, UPR/CNRS 9072, Institut de Biologie Physico-Chimique,

13 rue Pierre et Marie Curie, 75005 Paris, France

Summary I. General Traits II. Biochemical and Structural Studies A. Biochemical Composition B. Topology C. State of Oligomerization III. Functional Studies A. Functional Properties of the Isolated Complex B. Electron Transfer in vivo C. Other Functions IV. The pet Genes A. Gene Localization and Sequence B. Transcription C. Translation D. Nuclear Control on Chloroplast pet Gene Expression V. Biogenesis and Assembly A. Biogenesis of Individual Subunits 1. Biogenesis of Cytochrome f 2. Biogenesis of Cytochrome

B. Membrane Insertion of Cytochrome Subunits C. Assembly of Cytochrome Complexes 1. The Two-Step Model 2. An Assembly-Mediated Concerted Accumulation 3. Epistatic Regulation of Protein Synthesis and Post-Translational Degradation 4. A Molecular Mechanism for the Control by Epistatic Synthesis 5. Proposal for an Assembly Pathway D. Degradation of Assembled Cytochrome b6f Complexes. VI. Concluding Remarks Acknowledgments References

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Summary In most photosynthetic prokaryotes, cytochrome-containing complexes—cytochrome and cytochrome are also part of respiratory chains, whereas the photochemical reaction centers and their light-harvesting antennae function only in photosynthesis. Thus, genetic approaches, which are intimately associated with the possibility to inactivate and restore a function, have met with limited success in the case of cytochromecontaining complexes because their inactivation is usually lethal. This is in great contrast with the situation in Chlamydomonas reinhardtii which shows dispensable photosynthesis and can be grown at the expense of coupledexogeneous reduced carbon sources, utilizing a mitochondrial-based and cytochrome electron flow. Thus, Chlamydomonas is the only photosynthetic organism which has been extensively used in studies. Owing to the genetics of Chlamydomonas, a number of cytochrome mutants were cytochrome isolated both by classical and transformation mutagenesis. These strains offered a unique opportunity to in a photosynthetic advance the understanding of the structure, biogenesis and function of cytochrome eukaryote. This chapter reviews studies from the past decade, which combine mutational approaches with recent progress in membrane protein biochemistry and time-resolved flash spectroscopy. These studies have in photosynthesis and its value as a model system for brought to light both the pivotal role of cytochrome general issues in the field of organellar gene expression and protein assembly.

I. General Traits The major features of the cytochrome complex from Chlamydomonas reinhardtii are identical to those observed in vascular plant chloroplasts or cyanobacteria. These protein complexes share a number of structural and functional similarities with bacterial and mitochondrial cytochrome bc complexes, but they are of simpler biochemical composition (for reviews, see Trumpower and Gennis, 1994; Kallas, 1994; Cramer et al., 1996). The oligomeric proteins have two quinone-binding sites and contain four redox cofactors: two b-type hemes, one c-type heme and one iron-sulfur center. These cofactors carry out the redox reactions within the protein complex, which result in plastoquinol oxidation and reduction of plastocyanin. Electron complex is transfer through the cytochrome accompanied by a net translocation of protons across the thylakoid membranes. When Chlamydomonas is placed in copper-depleted medium, the electron acceptor plastocyanin is replaced by a soluble cytochrome, Cyt (see Chapter 31, Merchant). Abbreviations: DBMIB – 2,5-dibromo-3-methyl-6-isopropyll,4-benzoquinone; DNP-INT–2-iodo-2´,4´,4´-trinitro-3-methyl6-isopropyl diphenyl ether; CES – controlled by epistatic synthesis; – midpoint potential; FCCP – carbonyl cyanide hydrazone; OEC – oxygen evolving complex; PC – plastocyanin; PQ–plastoquinone; –plastoquinol; PS–Photosystem; SD – Shine-Dalgarno; TM – transmembrane; UTR – untranslated region; – proton electrochemical gradient

Thus cytochrome complexes participate in oxygenic photosynthesis by transferring electrons from PS II to PS I and contributing to the required for ATP synthesis. There is a three fold rationale for studying the complex from Chlamydomonas cytochrome reinhardtii rather than from other sources. For the biochemist and the crystallographer, Chlamydomonas cultures are an ideal source of starting material. This organism is easily grown in the laboratory and yields large amounts of thylakoid membranes with reproducible properties. For the biophysicist, the unicellular nature and small size of Chlamydomonas are well-suited for spectrophotometric studies in vivo. Since the cells can be grown in heterotrophic conditions, strains which display severely altered electron transfer properties can be compared with the wild type. This comparative approach is now boosted by site-directed mutagenesis techniques which offer the possibility to use a potentially unlimited set of strains with altered cytochrome complexes. Last, but not least, Chlamydomonas should be a particularly appropriate organism for the study of the electron transfer route from the cytochrome bf complex to PS I as the atomic structures of the participating proteins—cytochrome f (Berry EA., Huang LS, Chi Y, Zhang Z, Malkin R and Fernandez-Velasco JG, unpublished), plastocyanin (Redinbo et al., 1993) and cytochrome (Kerfeld et al., 1995)—have been solved in recent years. For the geneticist and the cell biologist, the genetics and

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gene transformation techniques available with this alga provide unique tools for the study of the regulation of pet gene expression in eukaryotes and for the dissection of the biogenesis pathway of the protein complex. Most of the studies that I will describe below relate to one of these opportunities offered by Chlamydomonas. Additional information or comments on some aspects of cytochrome complexes and their pet genes will be found elsewhere in this volume, in particular in Chapters 22, Joliot et al.; 31, Merchant; 10, Stern and Drager; 13, Perret et al.; 14, Olive and Wollman.

II. Biochemical and Structural Studies

A. Biochemical Composition Early work on the subunit composition of the complex in C. reinhardtii started cytochrome with the observation that there were similar pleiotropic deficiencies among the thylakoid membrane polypeptides in various mutants lacking cytochrome activity (Lemaire et al., 1986). These mutants were identified by their altered fluorescence properties (Bennoun and Delepelaire, 1982; see Chapter 35, Bennoun). The major subunits of the protein complex were identified by a comparison of the set of polypeptide deficient in the mutant membranes with the polypeptides copurifying in cytochrome fractions. This approach led to the identification of three chloroplast- encoded subunits, cytochrome f, cytochrome and subunit IV, and one nucleus-encoded subunit, the Rieske protein. A fifth putative subunit, which is nucleus-encoded, suV, was found to be missing from mutants deficient in complexes but was present at early cytochrome steps of its purification procedure (Lemaire et al., 1986). It was lost in the more purified, and still active, preparations of cytochrome complexes (Wynn et al., 1988; Pierre et al, 1995).This 19.5 kDa polypeptide is therefore not required for the enzyme to be active in electron transfer in vitro. Its relation to cytochrome complexes under physiological conditions—whether it participates in its biogenesis or in functional interactions with other photosynthetic proteins—remains to be investigated. A breakthrough in the biochemical characterization complex arose with the use of of the cytochrome a new class of neutral detergents. In the present case, Chlamydomonas thylakoid membranes were solu-

461 bilized in Hecameg, rather than in the more widely used octyl-glucopyranoside (Pierre et al., 1995). A complex, which showed purified cytochrome high rates of electron transfer from decylplastoquinol to plastocyanin, was recovered in two steps, involving sucrose gradient fractionation and hydroxylapatite chromatography. The purified protein contained the four above-mentioned major subunits in 1:1 stoichiometric ratios (Pierre et al., 1995). Three additional small subunits in the 4 kDa range were discovered subsequently. Two of them were identified as the petG and petL chloroplast gene products, the third one being encoded by the nuclear gene PetM (Pierre and Popot, 1993; Schmidt and Malkin, 1993; Ketchner and Malkin, 1996; de Vitry et al., 1996; Takahashi et al., 1996). Besides the hemes and iron-sulfur center, a chlorophyll molecule has been recognized only recently as a genuine fifth cofactor of the protein complex. It has been studied in some detail in Chlamydomonas where it was reproducibly found in preparations, in a 1:1 purified cytochrome stoichiometry with cytochrome f (Pierre et al., 1997). Two lines of evidence further support the identification of this chlorophyll as a cofactor of the complex: it is only slowly exchanged cytochrome chlorophyll and its with externally added signature by resonance Raman spectroscopy is indicative of a well defined binding environment (Pierre et al., 1997). Flash spectroscopy studies have independently demonstrated the existence of a local electrochromic shift, attributed to a particular chlorophyll molecule, upon electron transfer at Qo, one of the two quinone binding sites of the protein complex (A. Joliot and Joliot, 1995).

B. Topology The transverse organization of the cytochrome and complexes in the membranes has been addressed in several organisms but not in C. rein­ hardtii. However, the well-conserved primary sequences of the transmembrane subunits allows one to extrapolate the major conclusions drawn from these studies (Fig. 1). Cytochrome and subunit IV have four and three transmembrane (TM) helices respectively, now termed helices A,B,C,D and helices E,F,G (Widger et al., 1984), with stroma-located Ntermini. The primary sequence and biochemical behavior of the three 4 kDa subunits are consistent with each having one TM helix, with their N-termini

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facing the lumen (de Vitry et al., 1996). Mature cytochrome f, from residue 32 to 317 of the coding region (Buschlen et al., 1991), is also attached to the membrane by only one TM helix, from residue 283 to 302, located towards its C-terminal end (Kuras et al., 1995b). The polypeptide ends in the stroma by a short C-terminal extension of 15 residues, but the bulk of the protein (residue 32-282) resides in the lumen compartment where the c-heme is also located. In Fig. 1, it is represented by the crystal structure of the homologous soluble domain of cytochrome f from turnip (Martinez et al., 1994). Consistent with this transverse organization of cytochrome f, the introduction of a stop codon at position 283 by sitedirected mutagenesis, just upstream of the Cterminal TM helix, yielded a transformant strain expressing a truncated form of cytochrome f released in a soluble form in the thylakoid lumen (Kuras et al., 1995b). The interaction of the Rieske protein with the membrane has long been a controversial issue. In Fig. 1, one part of the Rieske protein is represented by the crystal structure of a fragment of the homologous protein from beef-heart mitochondria (Iwata et al., 1996). The most recent structural data arising from crystallization of the entire cytochrome bc complexes from beef-heart mitochondria, support the view that the Rieske protein is anchored in the membrane by a transmembrane segment at its Nterminus (Xia et al., 1997). However, the Rieske complexes can be protein from cytochrome

Francis-André Wollman

released from Chlamydomonas membranes without the use of detergents, which is a major criterion for the identification of a peripheral membrane protein (Breyton et al., 1994). In its absence, the other subunits still assemble and accumcytochrome ulate (see Section V.C. 1; Lemaire et al., 1986), which is reminiscent of the accumulation of the transmembrane PS II core in the absence ofthe peripheral OEC subunits (de Vitry et al., 1989). Although the Rieske protein has no cleavable lumen-targeting sequence, the N-terminal domain of the mature protein, which could also be considered as a regular TM helix, has features consistent with an uncleaved targeting sequence (de Vitry, 1994). Beef-heart cytochrome bc complexes have a number of distinctive features as compared to those of Chlamydomonas, which include the presence of additional subunits which are peripheral to the membrane, the core proteins and may stabilize the transmembrane orientation of the N-terminal hydrophobic domain ofthe Rieske protein. Therefore the possibility that the Rieske protein from Chlamydomonas does not have a transmembrane orientation should not yet be disregarded.

C. State of Oligomerization The protein complex is isolated as a dimer with an apparent molecular mass of about 310 kDa (Breyton et al., 1997). It can be converted to a monomer of

Chapter 24 Cytochrome

complexes

app. molecular mass 128 kDa, when the preparation is further delipidated by increasing the detergent concentration. The Rieske protein is usually lost during the conversion. However, upon mild dissociation with detergents, cytochrome complexes lose the Rieske protein while remaining dimers (Breyton et al., 1997). Furthermore, dimeric complexes lacking the Rieske protein cytochrome can be recovered from several mutant strains of Chlamydomonas (Takahashi et al., 1996; Finazzi et al., 1997; C. de Vitry, unpublished). These observations exclude the idea that the Rieske protein is itself required for dimerization. The PetL subunit has been suggested to play a role in stabilizing the dimeric state of the protein complex (Breyton et al., 1997) Two dimensional crystallization of the purified complex has been achieved recently cytochrome (Mosser et al., unpublished). A projection map of the negatively stained crystal was calculated at 8Å resolution. The protein complex crystallizes as a dimer with a long axis of 88Å, an external diameter of 53 Å per monomer and an internal groove of about 14Å. At present, there is no evidence for a dimer-based complexes, neither in function of cytochrome vitro nor in vivo. Thus, the isolation of a purified complex in a dimeric form raises the cytochrome question of its native state of oligomerization in the thylakoid membranes. The freeze-fracturing technique has been used to monitor the size of a cytochrome complex both in situ, by estimating the size of particles missing from cytochrome mutant membranes (Olive et al., 1986) and after reconstitution of the purified protein as a dimer or a monomer into liposomes (Breyton et al., 1997). The reconstitution experiments suggested apparent diameters of about 110Å and 80Å for the dimer and monomer respectively. The comparative measurements of freeze-fracture particles in the wild type mutants led to the and several cytochrome conclusion that two size classes of particles, 80100Å and 110–130Å in diameter, were mainly affected. In contrast with PS I and PS II which tend to segregate in distinct membrane domains, complexes have multiple locations cytochrome in the thylakoid membranes and are present in both stacked and unstacked membrane domains (Olive et al., 1986; Vallon et al., 1991). According to the diameters of the particles missing in Chlamydomonas complexes, those mutants lacking cytochrome retained on the exoplasmic face of the stacked

463 domains are about 80Å in diameter (Olive et al., 1986; Olive et al., 1992), which is consistent with a monomeric state, whereas those on the unstacked periplasmic faces are in the 110Å size range, which is consistent with a dimeric state (Olive et al., 1986). Upon transition from state I to state II, which is best suited for cyclic electron flow (Bulte et al., 1990), there is an enrichment of the cytochrome complexes in the unstacked membrane regions (Vallon et al., 1991). It is then tempting to suggest that the various locations would correspond to functional heterogeneity, with monomeric complexes being involved in linear cytochrome electron flow in the stacked membrane regions, complexes would whereas dimeric cytochrome perform cyclic electron flow in the unstacked membrane regions. However, this proposal conflicts with the fully complexes dimeric nature of the cytochrome extracted from the thylakoid membranes (Breyton et al., 1997). The two views can be reconciled only if one assumes that the detergent has little access to the stacked membrane domains, in which monomeric complexes would be trapped. That cytochrome PS II and LHCII protein complexes, which are similarly trapped in the stacked domains, are not extracted by Hecameg solubilization (Pierre et al., 1995), gives some credit to this assumption. On the other hand, it has been suggested that cytochrome complexes may also associate with other photosynthetic protein complexes (Wollman and Bulte, 1989). These interactions would result in the formation of particles of various sizes upon freezefracture, as would also the monomer/dimer equilibrium and could account for the freeze-fracture mutants (Olive et al., 1986). data in cytochrome Thus the proposal that both the dimer and monomer form of the protein may exist in vivo needs to be assessed critically in further studies.

III. Functional Studies

A. Functional Properties of the Isolated Complex Electron transfer from durohydroquinone to plastocyanin was shown to be DBMIB and DNPINT sensitive in an early preparation of cytochrome from C. reinhardtii (Wynn et al., 1988). In the most recent preparations (Pierre et al., 1995), which show a stigmatellin-sensitive improved activity of

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270 ± 60 electrons per Cyt f, the absorbance maxima of the and bands for cytochrome f are at 554 and 523 nm and are at 564 and 534 nm for cytochrome In these preparations, cytochrome f exhibited an Em of +330mV and cytochrome two Ems of –84 and –158 mV.

B. Electron Transfer in vivo Cytochrome complexes undergo a series of electron transfer reactions, driven by PS I photooxidation (Fig. 2). There is a branched pathway from the Qo site leading to electron transfer along a high potential chain—via the Rieske protein, cytochrome f and PC—and a low potential chain—via the and hemes and the Qi site. Electron transfer along both chains depend on an oxidant-induced concerted reduction of cytochrome f and cytochrome by a reduced plastoquinol at the Qo site, next to the lumenal face. An oxidized quinone, bound to the Qi site of the protein, on the opposite side of the membrane next to the stromal face, can then be reduced at the expense of the two b hemes, whose orientation and position within the protein complex allow the transfer ofan electron across the membrane (for a recent review see Cramer et al., 1996). These reactions can be followed in vivo by time-resolved flash spectroscopy (see Chapter 22, Joliot et al.). Both the oxidation and the reduction of cytochrome f are extremely sensitive to the resting proton electrochemical gradient. In addition, the oxidation kinetics for cytochrome f in vivo also depend on the extracellular osmotic pressure (R. Delosme and F.A. Wollman, unpublished). Thus, the half-time for cytochrome f oxidation in uncoupled conditions, usually achieved by an FCCP treatment, varies from at 5mM osmolarity to about at 50 about mM osmolarity (Delosme, 1991; Kuras et al., 1995; Soriano et al., 1996; Zhou et al., 1996; Finazzi et al., 1997). In the absence of FCCP, the oxidation kinetics are locked in a slow state, with half-times of about (Finazzi et al., 1997; Delosme, 1991). These variations could reflect the dependence of the interaction between cytochrome f and its natural oxidant, plastocyanin, on the lumenal ionic strength (Qin and Kostic, 1992). This hypothesis has been tested by site-directed mutagenesis: the positivelycharges patch at the surface of cytochrome f, which comprises five lysines thought to be involved in the electrostatic docking of plastocyanin through negative patches, was modified by replacing either two, three

or all five lysines with neutral residues (Soriano et al., 1996). Strangely enough, the rate ofcytochrome f oxidation was only marginally affected (by less than 25%) in the transformed strains, whereas its rate of reduction was decreased at most by a factor of two. On the other hand, the three-fold increase in concentration of cytochrome f in the lumen, which is observed upon replacement of the membrane-bound form by a truncated and soluble form, does not accelerate significantly its rate of oxidation (Kuras et al., 1995b). Moreover, the reaction retains its uncoupler sensitivity. The seemingly preserved interaction of plastocyanin with various forms of cytochrome f suggests that oxidation rates do not measure a collision-limited process. Either some step other than the oxidation of cytochrome f by plastocyanin, such as the release of oxidized plastocyanin from PS I, is rate-limiting, or there is restricted diffusion of plastocyanin in the lumenal space which prevents it from reacting with cytochrome f by a diffusion-limited, collisional process. The latter view is supported by the fact that all the above-described experiments were performed in the presence of FCCP which promotes transition to state II in vivo (see Chapter 30, Keren and Ohad). This treatment may induce the formation of a supercomplex through interactions between PS I, plastocyanin and the lumenal part of cytochrome f

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(Delosme, 1991; Finazzi et al., 1997). According to this view, intra-supercomplex electron transfer would be responsible for the fast oxidation kinetics observed for cytochrome f in green algae. In contrast, slow oxidation kinetics, observed in the absence of FCCP, would be produced upon dissociation of this supercomplex and would reflect a collisional process. In the latter conditions, one may expect a further decrease in the rates of cytochrome f oxidation in the lysine mutants. This prediction is consistent with the oxidation kinetics recorded in more oxidizing conditions with another set of lysine mutants, which included substitutions with negatively charged residues: cytochrome f oxidation was five to ten times slower in some of these strains (J. G. FernandezVelasco, J. Zhou and R. Malkin, unpublished). Clearly, an analysis of the kinetics of cytochrome f oxidation in the absence of FCCP in the lysine mutants of Soriano et al. (1996) would improve our understanding of the electron transfer mechanism on the donor side of PS I. Subsequent reduction of oxidized cytochrome f by plastoquinol bound at the Qo site occurs with half-times ranging from about 7 ms in coupled conditions, to about 2.5 ms when uncoupled with FCCP (Finazzi et al., 1997) or by some other means (Zhou et al., 1996). An intriguing observation is the higher uncoupler sensitivity of the rate of cytochrome f reduction in two widely different cytomutants, one lacking the PetL subunit chrome (Takahashi et al., 1996), the other having an altered Qo site (Finazzi et al., 1997): in both cases, the mutation caused a larger functional alteration in the absence than in the presence of FCCP. This could not be attributed to a larger resting proton electrochemical potential as the rates of cytochrome f oxidation, which are also dependent, were unchanged. Interestingly, the two strains showed a loosened biochemical interaction of the Rieske protein with complex. It suggests the rest of the cytochrome interaction between that there may be a plastoquinol and the Rieske protein, which is the first oxidant in the reaction leading to cytochrome f reduction at the Qo site. The folding ofthe Qo site in the mutants, which is, in part, borne by the Rieske protein (Link and Iwata, 1996, and references therein), may be more perturbed in the presence of a than in its absence. Mapping of this quinone binding site by mutagenesis has been undertaken in bacterial and mitochondrial cytochrome bc complexes (Brasseur et al., 1996) but not yet in Chlamydomonas. In one

465 instance, a deep alteration of the Qo site was observed, due to a 12 amino acid duplication of the lumenal cd loop connecting helices C and D in cytochrome (Finazzi et al., 1997). As a result, the affinity of the Qo site for plastoquinols, which was estimated to be in the wild type, decreased by two orders of magnitude. The Qo site was also altered in a series of site-directed mutants, in which the glutamic residue from the conserved PEWY sequence of suIV, was replaced by bulky residues or a glutamine or an asparagine (F. Zito, G. Finazzi, P. Joliot and F.-A. Wollman, unpublished). The former substitutions caused a decrease in the rate of the concerted reduction at the Qo site with no change in its affinity for plastoquinols. The latter substitutions markedly increased the stoichiometry of charges translocated across the membrane per electron transferred to cytochrome f. Surprisingly, the reduction signals of the b-hemes remained very low in the mutants. This observations suggest that a pathway for charge translocation through a proton pump or a semiquinone cycle may be favored when a glutamine or an asparagine residue replaces the glutamate in the PEWY sequence. Because the free N-terminus of mature cytochrome f serves as an axial ligand for the c-heme iron (Martinez et al., 1994), changes in the rates of cytochrome f reduction were assessed by site-directed mutagenesis of one of the first three N-terminal residues, Y-P-V (Zhou et al., 1996). In particular, the possibility that the aromatic ring of the well conserved terminal tyrosine, which is oriented parallel to the heme plane (Martinez et al., 1994), plays a critical role in electron transfer from the Rieske protein to the c-heme, was investigated. Zhou et al. (1996) found no evidence for such a role: substitution of by aromatic or non aromatic residues yielded rates of cytochrome f reduction comparable to those in the wild type. In contrast, substitution of by a valine decreased the reduction rates more than ten-fold, which could indicate that the proline turn may be required for the proper orientation of the liganding N-terminal residue.

C. Other Functions Cytochrome complexes from the green alga Chlorella sorokiniana were shown, by differential spectrophotometry, to interact with a soluble electron carrier, a high spin hemoprotein which could be a cytochrome c´ (P. Joliot and Joliot, 1988). The same

Francis-André Wollman

466 species has been detected in Chlamydomonas (R. Delosme and P. Joliot unpublished). Its possible correspondence with the 19.5 kDa polypeptide, originally considered as a fifth subunit of the Cyt complex (Lemaire et al., 1986) requires further investigation. Chlamydomonas has also been the first photosynthetic organism in which the implication complexes in the reversible protein of cytochrome phosphorylation process, underlying the regulation mechanism of state transitions, has been recognized (Lemaire et al., 1987; Wollman and Lemaire, 1988). It has been shown subsequently that cytochrome complexes undergo lateral migration from stacked to unstacked membrane domains upon transition to state II (Vallon et al., 1991) (for details, see Chapter 30, Keren and Ohad).

IV. The pet Genes

A. Gene Localization and Sequence The five chloroplast pet genes have been localized and sequenced on the circular chloroplast chromosome of C. reinhardtii. They are all uninterrupted continuous and they encode proteins whose sequences show high conservation with those found in complexes from vascular plants, cytochrome ranging from about 60% to 90% identical residues (Bertsch and Malkin, 1991; Buschlen et al., 1991; Matsumoto et al., 1991; Fong and Surzycki, 1992; Huang and Liu, 1992; Takahashi et al., 1996). The pet genes are scattered on the chloroplast genome, with, according to the nomenclature in Harris (1989), petA, and petD positioned on restriction fragments R1, petB on R9, petG on R2 and petL on R6. The petA gene and the downstream petD gene are separated by an intergenic region, of about 2 kb, which comprises ORF 112 for which there is no evidence of expression (Sturm et al., 1994). Their relative organization is conserved in C. moewusii (Boudreau et al., 1994). The petB sequence is approx. 0.8 kb upstream of the protease gene clpP (Huang et al., 1994), petG is localized about 0.5 kb downstream of the PS II gene psbL (Fong and Surzycki, 1992) and petL is localized approximately 0.6 kb downstream of the PS II gene psbC (Takahashi et al., 1996). The nuclear genes PetC and PetM are interrupted by four and two introns respectively (de Vitry, 1994; Ketchner and Malkin, 1996). These genes have not been mapped with respect to the 18 nuclear linkage groups of Chlamydomonas.

B. Transcription The transcripts for the nuclear genes PetC and PetM are respectively 1.5 kb and 0.6 kb long and most likely represent intron-free mRNAs (de Vitry, 1994; de Vitry et al., 1996; Ketchner and Malkin, 1996). Like most of the other chloroplast mRNAs from C. reinhardtii, the petA petB and petD mRNAs accumulate mainly as monocistronic transcripts of 1.3–1.4 kb, 0.8–0.9 kb and 0.9 kb respectively (Matsumoto et al., 1991; Chen et al., 1993; Kuras and Wollman, 1994; Zito et al., 1997). Evidence for the presence of active promoters immediately upstream of petD and petA has been provided through uidA gene fusion (Sakamoto et al., 1993) or DNA deletion analyses (Sturm et al., 1994; Chapter 10, Stern and Drager). A TATAAT motif, for a promoter upstream of petG has been proposed by Fong and Surzycki (1992) but, the presence of this particular motif may be fortuitous because the chloroplast genome is AT rich. Despite the accumulation of monocistronic transcripts, one cannot draw the conclusion that the pet genes are only transcribed as monocistronic units. Besides the monocistronic petA and petD transcripts, a 4.4 kb petA/petD cotranscript accumulates in strains carrying a deletion in the promoter region of the petD gene (Sturm et al., 1994). This observation argues for redundant petD transcription from distal and proximal promoters, with 5´ processing events generating the mature monocistronic petD transcript with a 362 nt long 5´UTR (Sakamoto et al., 1994b). Other petA transcripts with sizes larger than the most abundant 1.3 kb monocistronic transcript are detected by mRNA blot hybridization (Matsumoto et al., 1991; Sturm et al., 1994). These transcripts, about 3.4 and 2.3 kb long, were first suggested to originate from alternative transcription start sites upstream of petA (Matsumoto et al., 1991). However, 5´ mapping of petA transcripts by primer extension analysis resolved three distinct 5´ ends, all of which are clustered within a 20 base region, about 270 bases upstream of the translation initiation codon (Matsumoto et al., 1991). Because of the cotranscription of petA with the downstream petD gene, it is more likely that the 3.4 and 2.3 kb transcripts differ at their 3´ ends and correspond to processing intermediates between the 4.4 kb polycistronic and the 1.3 kb monocistronic petA transcript. This hypothesis is strengthened by the size changes of these transcripts, or even their disappearance, in various chloroplast transformants with deletions or substitutions in the intergenic petA/

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petD region (R. Kuras, Y. Choquet, unpublished).

C. Translation Whether there are potential ribosome-binding sites in the 10–30 nt upstream of the initiation codons of the pet genes has not yet been established. A ShineDalgarno (SD) element was not detected upstream of petG (Fong and Surzycki, 1992). An AAGA sequence, 12 nt upstream of the petA initiation codon, was considered in one work (Matsumoto et al., 1991) but not in another (Sakamoto et al., 1994a), as a potential SD element. In the latter work, a potential SD element, GGA, 10 nt upstream of the petD initiation codon, was converted to a TTA motif, (Sakamoto et al., 1994a). This experiment led to question the requirement for Shine-Dalgarno motifs for chloroplast translation as translation of the petD transcripts was unaffected in the TTA mutants. However, other 5´UTR sequences, about 30–50 bases and 160–260 bases upstream of the AUG codon appear critical for translation (Sakamoto et al., 1994a). Nucleotides immediately upstream of the initiation codon, in positions –1 to –3, are also critically involved in translation initiation (Chen et al., 1995;Y. Choquet and D. Drapier unpublished). AUG initiation codons for either petD or petA were replaced by cognate codons, which can substitute for AUG in E coli (Chen et al., 1993; Chen et al., 1995) In most cases, translation still occurred, albeit at much lower efficiency. Moreover, translation was still initiated at the same site, with no alternative upstream or downstream codon substituting for the mutated ATG codon. Translation initiation was inhibited by substitutions at the first position of the ATG codon and was much less sensitive to substitution at the third than at the second position. Thus petD translation can be initiated only at one site and is under the control of several elements far upstream in the 5´UTR.

D. Nuclear Control on Chloroplast pet Gene Expression A number of nuclear mutants lacking cytochrome activity and showing specific alterations in the expression of only one of the chloroplast pet genes were recovered either after conventional mutagenesis (Howe and Merchant, 1992; Girard-Bascou et al., 1995) or by random integration of transforming DNA (Gumpel et al., 1995). Expression of the pet genes was then characterized at the transcriptional,

467 translational and post-translational levels. The loci mutated in these strains identified two sets ofnuclear products, but the number ofnuclear loci participating in the specific expression of the pet genes is certainly by far greater than what is known presently and shown in Table 1. A first set of factors is involved in cofactor binding to the pet gene products. It corresponds, at least, to four nuclear loci, CCS1,2,3,4 involved in c-heme attachment to cytochrome f and four nuclear loci, CCB1,2,3,4, involved in b-heme binding to cytochrome (see, for details, Section V and Chapter 30, Merchant). Another set of nuclear products acts on the chloroplast pet genes at the posttranscriptional or translational levels. It involves, at least, one nuclear locus in the maturation/stabilization of each of the chloroplast transcripts, petA, petB, petD and petG, thereby named MCA1, MCB1, MCD1 and MCG1, respectively, and one locus acting on petA translation, TCA1, (Girard-Bascou et al., 1995). Recently, the action of one of these nuclear factors, the MCD1 gene product, has been characterized in more depth. Based on the use of chimeric genes, it was shown to act on the 5´ UTR of petD mRNA, presumably by preventing its 5´-3´ exonucleolytic degradation (Drager et al., 1998).

V. Biogenesis and Assembly

A. Biogenesis of Individual Subunits 1. Biogenesis of Cytochrome f The petA gene sequence shows the presence of a cleavable lumen-targeting leader peptide of 31 residues, ending by an AQA motif, a typical target site for a lumenal processing peptidase (Buschlen et al., 1991a; Kuras et al., 1995a). Consistent with these assignments, replacement of the AQA site by an LQL sequence allowed the detection, in 5 min pulses, of an uncleaved, membrane bound, precursor form. Nevertheless, the LQL-containing precytochrome f was still processed—but more slowly than the wild type precytochrome f to some other forms of lower apparent molecular mass (Kuras et al., 1995a). The presequence contains a hydrophobic stretch long of nine residues, ending with a glycine residue and bordered on its N-terminal side by a positively charged arginine. The role of this protein motif in translocation has been demonstrated by the isolation of translocation-incompetent strains obtained by replacing one of the hydrophobic residues by a charged residue

468

(Smith and Kohorn, 1994). A stroma-located precursor form was detected in one of the mutants. This approach led to the identification of a translocation core of 7 aa, from T13 to G19. Substitutions outside this core had no effect on translocation. Interestingly, Baillet and Kohorn (1996) recovered intragenic suppressor mutations, which further altered the presequence of cytochrome f but restored translocation and processing to the mature form. The new substitutions occurred at the position of the above-mentioned arginine, which demonstrated that this residue was dispensable for translocation. Furthermore these mutants had a hydrophobic translocation core, now 10–11 residue long but shifted towards the N-terminus by at least three residues (for a more detailed discussion, see Chapter 13, Perret et al). The translocation machinery itself has not been characterized. It may be used jointly by other transmembrane proteins as cytochrome f translocation mutants were reported to have some decrease in D1 and LHCII membrane insertion (Smith and Kohorn, 1994). The identification of three nuclear loci, the mutant alleles ofwhich restore translocation of a translocation-incompetent variant of precytochrome f, opens the way to a molecular characterization of the translocator (Smith and Kohorn, 1994).

Francis-André Wollman

A major step in the biogenesis of cytochrome f is the conversion from the apo to holoform upon covalent binding of the c-heme. This process has been studied in great detail in Chlamydomonas (Howe and Merchant, 1994b). It affects in parallel the biogenesis of the other lumenal c-type cytochrome, cytochrome (Howe and Merchant, 1992). A detailed presentation of the heme-binding mechanism is presented in Chapter 30 (Merchant). Briefly, c-heme binding occurs through the formation of thioether bonds between the vinyl side chains of the heme and two cysteinyl residues, which are part of a conserved heme binding motif CxxCH, with the conserved histidine serving as one axial ligand for the heme iron. This process is catalyzed by an enzyme complex whose active site is on the lumen side ofthe thylakoid membranes (Howe and Merchant, 1994a). A genetic mutants which also analysis of the cytochrome display a minus phenotype, led to the identification of at least four nuclear genes and one chloroplast gene controlling c-heme attachment, whose products may be part of a cytochrome maturation complex (Howe and Merchant, 1992; Girard-Bascou et al., 1995; Howe et al., 1995; Xie and Merchant, 1996; Xie et al., 1998). Conversion of apo- to holocytochrome f is too rapid to be detected by pulse-labeling studies. When the apo- to holo- conversion is blocked, due to the

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Cytochrome

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replacement of the cysteinyl ligands by some unrelated residue, the apocytochrome is degraded rapidly, with a half-life of about 30 min (Kuras et al., 1995a). The crystal structure of a C-terminal truncated form of turnip cytochrome f has revealed that one of the axial ligands of the c-heme iron is the N-terminal amino group of Y1, which is produced upon maturation of the precursor form (Martinez et al., 1994). This unique feature among c-type cytochromes strongly suggested that the heme-binding and processing steps were intimately coordinated during cytochrome f biogenesis. It was reasonable to predict that processing of the precursor form would be a prerequisite for heme binding, or that covalent heme attachment might catalyze a conformation of the preprotein suitable for its processing. Contrarily to either of these expectations, the two steps can occur independently. Pre-apocytochrome f is processed to its mature form in heme-binding mutants (Howe and Merchant, 1992; Howe et al., 1995; Kuras et al., 1995a). Conversely, the precursor form of cytochrome f was shown to bind heme in a mutant having an altered processing site (Kuras et al., 1995a). It should be emphasized that none of these observations precludes the two processes from being kinetically coupled in the wild type. They are both catalyzed by membrane-bound enzymes, active on the lumen side of the thylakoids, which could well be part of a cytochrome maturation complex, active on both cytochrome and cytochrome f. The soluble Cterminal truncated form of cytochrome f obtained by introducing a stop codon at position 283 of the petA gene, accumulates the holoform with no evidence for the parallel production of the apoform, which should have been detected in pulse-labeling experiments (Kuras et al., 1995a; Kuras et al., 1995b). This observation supports the view that the heme binds to the cytochrome when it is still membraneassociated which means binding to the precursor form in the case of the C-terminal truncated cytochrome f. Thus, heme ligation should precede protein processing, a proposal consistent with heme binding to the precursor form which was detected in the processing mutant (Kuras et al., 1995a).

2. Biogenesis of Cytochrome The two hemes, and associated with cytochrome use two pairs of histidines as their axial ligands. The histidines are located towards opposite sides of the membrane in TM helices B and D (Kuras et al.,

469 1997). Replacement of either of the four histidines by some unrelated residue yields protease-sensitive variants, which can be easily cytochrome distinguished from the wild type form upon electrophoresis (Zito et al., 1997). In addition, substitution of or coordinating histidines leads to distinct electrophoretic forms. Unexpectedly, a set of nuclear mutants showed similar cytochrome alterations (Kuras et al., 1997). These comparative observations led to the conclusion that at least four nuclear loci, CCB1-CCB4, contribute specifically to the association of the b hemes with cytochrome shows unusual The hemoform of cytochrome resistance to denaturing treatments like solubilization in boiling SDS or acetone extraction (Kuras et al., 1997). The stability of the interaction between the b hemes and the protein may originate from some posttranslational modification of cytochrome catalyzed by the CCB factors, possibly involving covalent heme binding by some linkage other than that which occurs with the c-type hemes.

B. Membrane Insertion of Cytochrome Subunits Cytochrome f is translated from thylakoid-bound polysomes both in pea (Gray et al., 1984) and in Chlamydomonas (D. Drapier, unpublished). Therefore it is most likely co-translationally inserted in the thylakoid membranes. It is translocated through the Sec pathway (Rothstein et al., 1985; Mould et al., 1995). The N-terminus of the mature polypeptide is located in the lumen because of the removal of the lumen-targeting presequence. However translation and insertion are not obligatorily coupled, as a stromal-located precursor form of cytochrome f was detected in translocation mutants (Smith and Kohorn, 1994). The Rieske protein has been proposed to be translocated in the lumen via a non cleavable targeting sequence (de Vitry, 1994). The rationale for this proposal came from the physico-chemical properties of the Rieske protein which behaves as a peripheral membrane protein located on the lumen surface of the thylakoid membranes (Breyton et al., 1994). If the N-terminal hydrophobic segment of the Rieske protein were a genuine TM helix, then the insertion issue would not be different from that of most of the bi- or polytopic thylakoid polypeptides which retain their N-terminus on the side of membrane insertion: the insertion process involves docking of the protein

Francis-André Wollman

470 by its N-terminal region at the stromal surface of the thylakoid membranes and subsequent translocation and partitioning of the rest of the protein between the lumen and membrane compartments, according to the hydrophobicity of the various protein domains. Among the exceptions to this rule are two cytochrome subunits, the PetG and PetM products, whose Ntermini are located in the lumen although they have no lumenal targeting presequence (de Vitry et al., 1996). As discussed by the authors, this unusual orientation could result from, or be achieved through, the absence of positively charged residues on the very short N-terminal segment, only four residues long, which protrudes in the lumen. Following the same rationale, the sequence data on the third small subunit, PetL(Takahashi et al., 1996), cytochrome predicts that it should also have its N-terminus facing the lumen.

C. Assembly of Cytochrome

Complexes

1. The Two-Step Model Early studies with photosynthesis mutants from C. reinhardtii have pointed to concerted accumulation complex of the various subunits of the cytochrome (Bendall et al., 1986; Lemaire et al., 1986). A number of independent mutants, whether of chloroplast or nuclear origins, displayed the same pleiotropic loss in all the constitutive subunits ofthe protein complex. In contrast to this behavior, one nuclear mutant, ac21, was shown to accumulate more than 50% of the wild type levels of cytochrome and cytochrome f in the absence of detectable amounts of the Rieske protein (Bendall et al., 1986; Lemaire et al., 1986). It was thus proposed that cytochrome assembled in two steps. A first step comprised the concerted accumulation and assembly of a cytochrome moiety, made of the major transmembrane subunits encoded in the chloroplast: cytochrome f, cytochrome and suIV. In a second step, the nucleus-encoded Rieske protein assembled with the cytochrome moiety (Lemaire et al., 1986). It has been confirmed recently, with two nuclear mutants altered in the PetC gene, that the presence ofthe Rieske protein is not required for the concerted accumulation of the other subunits complex (C. de Vitry and Y. of the cytochrome Choquet, unpublished). The association ofthe Rieske protein with the cytochrome moiety of the protein complex is very sensitive to its structural integrity. It dissociates more easily from purified cytochrome

complexes which either lack the PetL subunit (Takahashi et al., 1996) or display structural modifications in the cd loop of the cytochrome subunit (Finazzi et al., 1997).

2. An Assembly-Mediated Concerted Accumulation Gene deletion studies provided a more detailed insight into the concerted accumulation of cytochrome subunits. Deletion of the petA, petB or petD chloroplast genes (Kuras and Wollman, 1994) and, to a somewhat more limited extent, deletion of the petG chloroplast gene (Berthold et al., 1995), prevented the accumulation of the entire set of cytochrome subunits. In marked contrast, exponentially growing cells bearing a deletion of the petL chloroplast gene still accumulated, up to 50% of the wild type levels, of a fully functional cytochrome complex (Takahashi et al., 1996). Changes in the primary sequence of one subunit only, presumably by altering its folding, similarly prevented accumulation of the whole cytochrome complex. For instance, substitution of a proline for a leucine residue in the sequence, next to which cytochrome participates in coordination of led to a drop in all cytochrome, subunits (Zito et al., 1997). Preventing heme-binding to either cytochrome f (Howe and Merchant, 1992; Kuras et al., 1995a) or cytochrome (Kuras et al., 1997) had the same pleiotropic effect as that of a deletion of the petA or petB genes. This was most likely caused by the rapid proteolytic disposal of the apocytochrome end product: for instance, apocytochrome f was barely detectable when accumulation was assessed in a mutant blocked in the conversion to the holoform by substitution of the cysteinyl heme ligands (Kuras et al., 1995a). Thus, there is overwhelming evidence that the concerted subunits is an accumulation of the cytochrome assembly-mediated process.

3. Epistatic Regulation of Protein Synthesis and Post-Translational Degradation Since the pet genes are not organized in a polycistronic transcriptional unit on the chloroplast genome (Buschlen et al., 1991), there is little chance that their synthesis is coordinated at the transcriptional level. Thus the concerted accumulation of the subunits, revealed by their pleiotropic cytochrome mutants, was likely loss in numerous cytochrome

Chapter 24

Cytochrome

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to originate from some (post)-translational regulation. Studies with deletion mutants showed that there was a dual regulation process operating both at the posttranslational and translational levels (Kuras and Wollman, 1994; Wollman et al., 1995). The rates of synthesis and half-lives of the major chloroplast complex were subunits of the cytochrome compared in petA, petB and petD deletion mutants (Kuras and Wollman, 1994). The rates of synthesis of and suIV were independent of the cytochrome expression of their assembly partners. In contrast, their half-lives were considerably shortened in the deletions strains. Whereas these subunits were not degraded over a three hour time period in the wild type strain, cytochrome and suIV had half-lives of about 45 min only in the absence of cytochrome f. SuIV had an even shorter half-life, about 15 min, in a subunit which is the absence of cytochrome probably required for its proper folding (Kuras and Wollman, 1994). The behavior of cytochrome f was widely different. It showed about the same resistance to proteolytic degradation in the membranes, whether its assembly partners were present or absent. Nevertheless, it was present at only 10% of the wild type level in the absence of cytochrome or su IV This could be explained only by a reduced rate of synthesis. Indeed 5 min pulse-labeling experiments showed that, in the absence of its assembly partners, cytochrome f was labeled at only 10% of its rate in the wild type strain (Kuras and Wollman, 1994). This observation defines cytochrome f as a CES protein, Controlled by Epistatic Synthesis: its synthesis at a wild type rate requires the presence of some other subunit of the same protein complex (Wollman et al., 1995). In contrast with the ten-fold drop in cytochrome f synthesis in the petB and petD deletion strains, a three-fold increased rate was observed in strains lacking the C-terminal anchor of cytochrome f (Kuras et al., 1995b) or in strains unable to undergo apo to holo conversion of cytochrome f (Kuras et al., 1995a). Based on these contrasting effects, it was suggested that the concentration of the unassembled C-terminal domain of cytochrome f controlled, by some negative feedback mechanism, the rate of cytochrome f translation (Kuras et al., 1995;Wollman et al., 1995): C-terminal truncated cytochrome f, as well as the rapidly degraded apocytochrome f, would show increased rates of synthesis because they do not accumulate a C-terminal-located regulatory motif in the thylakoid membranes. In contrast the extensive

471 accumulation of the unassembled C-terminal domain in assembly mutants, amounting to 10% of the total concentration of cytochrome f present in the wild type, would repress petA translation. In the wild type, the regulatory motif is shielded by its assembly complexes. Therefore, the within cytochrome amount of unassembled cytochrome f, exposing a translational down-regulation motif, would depend directly on the rate at which the other subunits are delivered for assembly in the thylakoid membranes. According to this view, the epistatic regulation of cytochrome f synthesis corresponds to fine tuning of the rate of cytochrome f synthesis by its rate of complexes. assembly in cytochrome

4. A Molecular Mechanism for the Control by Epistatic Synthesis (CES) Some molecular insight into the CES mechanism came with the use of chimeric petA genes (Choquet et al., 1998). The resident petA gene was replaced by a chimeric gene, in which the endogenous 5´UTR was replaced by that of the atpA gene. The resulting transformants were phototrophic and could be crossed subsequently with a nuclear mutant in which suIV is not expressed. The double mutants, with the chimeric petA gene expressed in the absence of suIV, showed extensive accumulation of unassembled cytochrome f Accordingly, the rate of synthesis of cytochrome f now made from this chimeric gene, became independent of the presence or absence of suIV: the epistatic control of cytochrome f synthesis was lost with the replacement of the 5´petA UTR by an unrelated 5´UTR sequence. Moreover, the 5´petA UTR fused to the reporter aadA gene was sufficient to confer the epistatic regulation to the level of antibiotic resistance brought about by aadA: strains were poorly resistant in the absence of suIV but highly resistant when the apo to holo-cytochrome f conversion was prevented. Thus, the 5´petA UTR is the target for an interaction of the carboxy terminal domain of cytochrome f with the petA message, to down regulate further initiation of translation of cytochrome f. This interaction develops only when the carboxy terminal domain is not shielded by complex. Whether assembly in the cytochrome the interaction involves direct molecular contacts between the stromal C-terminal segment of cytochrome f and the 5´UTR of petA transcripts is not yet known. However, the regulation could well occur through competition of binding of a trans-

472 lational activator to the C-terminal domain and the 5´UTR of the petA transcript. Mutations at the TCA1 locus specifically prevent synthesis of cytochrome f (Girard-Bascou et al., 1995). The Tcal factor is thus a good candidate for a putative translational activator involved in the epistatic regulation of cytochrome f synthesis.

5. Proposal for an Assembly Pathway The wealth of data pertaining to biogenesis issues for complex provides a reasonable the cytochrome frame for a typical assembly pathway of an oligomeric protein in the thylakoid membranes (Fig. 3). Chloroplast transcripts, associated with specific nuclear factors, become translatable on thylakoidbound ribosomes to provide co-translational insertion of transmembrane proteins. Polypeptide modifications, i.e. processing or cofactor addition, occur early in the biogenesis of individual subunits, possibly through specialized protein complexes of the CcsA, Ccs1-4 type (see Fig. 3). Accumulation ofthe various subunits is driven by their assembly, which protects most of them against post-translational degradation and controls the rate ofsynthesis ofsome subunits by a CES mechanism, possibly involving nuclear factors of the Tcal type. The assembly pathway itself is a step by step process in which the folding of subcomplexes serves as a template for the subsequent association of additional subunits.

D. Degradation of Assembled Cytochrome Complexes. There are several situations in which assembled cytochrome complexes were shown to undergo specific degradation. Towards the end of their exponential phase of growth, cells from mutants lacking the PetL subunit (Takahashi et al., 1996) or having an altered cd loop on the lumen side of (Finazzi et al., 1997), lose their cytochrome complexes. In both cases, the authors cytochrome presented biochemical evidence for weakened interactions between the Rieske protein and the rest of the protein complex. Since cytochrome complexes are also prone to proteolytic degradation in aging cultures of mutants specifically lacking the Rieske protein (de Vitry and Wollman, unpublished), proteolytic attack of the protein complex may occur through some site exposed on the lumenal face ofthe

Francis-André Wollman membranes, which is usually masked by its interaction with the Rieske protein. A proteolytic attack of cytochrome f from the lumen side of the membranes has also been considered by Kuras et al. (1995a), based on the much larger sensitivity of apocytochrome f to degradation when membrane-bound than when released in a soluble form in the lumen, due to the truncation of its C-terminus. complexes undergo specific Cytochrome degradation when wild type cells of Chlamydomonas are grown in nitrogen-free medium to induce gametogenesis (Bulte and Wollman, 1992). Although little is known on the degradation process, it seems to be triggered by the energetic status of the starved cells: when mitochondrial electron transport is inhibited during nitrogen starvation, cytochrome complexes are no longer degraded. As a result, nitrogen-starved cells tend to keep coupled electron transfer activity in only one ATP-generating organelle. complexes may then be degraded by Cytochrome an ATP-dependent protease which, although it is active in the chloroplast compartment, would also sense the rate of mitochondrial ATP production.

VI. Concluding Remarks The data selected from this chapter provide only a limited view of the recent progress made in the field studies. Structural studies using of cytochrome preparations from higher plants or cytochrome cyanobacteria are currently progressing in parallel with those from Chlamydomonas and have already provided valuable information, particularly in the case of the Rieske protein (Zhang et al., 1996) and cytochrome f (Martinez et al., 1994). In addition, several maize mutants altered in some aspects of pet gene expression have been described recently (Barkan et al., 1994). These should allow us to assess whether biogenesis in key features of cytochrome Chlamydomonas are conserved in multicellular photosynthetic eukaryotes. Studies of the cytochrome complex from Chlamydomonas reinhardtii will certainly expand further in the future as, today, Chlamydomonas is still the one cytochrome eukaryote which can most easily accommodate mutations inactivating this protein complex. The tools for modifying cis-acting sequences of the pet genes and protein motifs from the constitutive subunits,

Chapter 24

Cytochrome

complexes

including the Rieske protein, are now available. Functional and biogenesis studies should further benefit from this molecular approach. Among the challenging issues are the elucidation of the role of the chlorophyll cofactor in the cytochrome complex, the possible regulatory function of the monomer to dimer conversion in photosynthetic electron flow and a better understanding of the electrogenic mechanism which accompanies electron transfer through the cytochrome complex. This controversial issue may come closer to an end when mutations altering or inactivating the Qi site will be convincingly demonstrated. It should be apparent from this review article that studies of cytochrome biogenesis have progressed

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tremendously in the past decade. Translocation mutants for cytochrome f should provide the opportunity to tackle the organization of the translocation process for a membrane protein and to understand whether the translocator is or is not a single machinery branched downstream to three or four routes with separate regulatory controls (Chapter 13, Perret et al.). The study of the assembly of the whole protein complex should benefit greatly from the recent technical advances and should help us to understand whether the involvement of some nuclear factor in the translation of cytochrome f represents an actual nuclear control of cytochrome assembly through a CES mechanism.

474 Acknowledgments Many thanks to René Delosme, Giovani Finazzi and Catherine de Vitry for critical reading of the manuscript, to Cecile Breyton, Yves Choquet and Giovanni Finazi for their help in drawing Figs. 1 and 2 and to Javier Fernando-Velasco, Jean-Luc Popot, David Stern and Catherine deVitry for communication of preprints or unpublished data.

References Baillet B and Kohorn BD (1996) Hydrophobia core but not amino-terminal charged residues are required for translocation of an integral thylakoid membrane protein in vivo. J Biol Chem 271: 18375–18378 Barkan A, Walker M, Nolasco M and Johnson D (1994) A nuclear mutation in maize blocks the processing and translation of several chloroplast mRNAs and provides evidence for the differential translation of alternative mRNA forms. EMBO J 13: 3170–3181 Bendall DS, Sanguansermsri M, Girard-Bascou J and Bennoun P (1986) Mutations in Chlamydomonas reinhardtii affecting the cytochrome bf complex. FEBS Lett 203: 31–35 Bennoun P, Delepelaire P (1982) Isolation of photosynthesis mutants in Chlamydomonas. In: Edelman M, Hallick RB and Chua N-H (eds) Methods in Chloroplast Molecular Biology, pp 25–38. Elsevier Biomedical Press, Amsterdam Berthold DA, Schmidt CL and Malkin R (1995) The deletion of petG in Chlamydomonas reinhardtii disrupts the cytochrome bf complex. J Biol Chem 270, 49: 29293–29298 Bertsch J and Malkin R (1991) Nucleotide sequence of the petA (cytochrome f ) gene from the green alga, Chlamydomonas reinhardtii. Plant Mol Biol 17: 131–133 Boudreau E, Otis C and Turmel M (1994) Conserved gene clusters in the highly rearranged chloroplast genomes of Chlamydomonas moewusii and Chlamydomonas reinhardtii. Plant Mol Biol 24: 585–602 Brasseur G, Sangas AS and Daldal F (1996) A compilation of mutations located in the cytochrome b subunit of the bacterial and mitochondrial bc(1) complex. Biochim Biophys Acta 1275: 61–69 Breyton C, de Vitry C and Popot J-L (1994) Membrane association of cytochrome subunits: The Rieske Protein of Chlamydomonas reinhardtii is an extrinsic protein. J Biol Chem 269: 7597–7602 Breyton C, Tribet C, Olive J, Dubacq JP and Popot J-L (1997) Dimer to monomer conversion of the cytochrome complex: Causes and consequences. J Biol Chem 272: 21892–21900 Bulté L and Wollman F-A (1992) Evidence for a selective destabilization of an integral membrane protein, the cytochrome complex, during gametogenesis in Chlamydomonas reinhardtii. Eur J Biochem 204: 327–336 Bulté L, Gans P, Rebèillè F and Wollman F-A (1990) ATP control on state transitions in vivo in Chlamydomonas reinhardtii. Biochim Biophys Acta 1020: 72–80 Buschlen S, Choquet Y, Kuras R and Wollman F-A (1991)

Francis-André Wollman Nucleotide sequences of the continuous and separated petA, petB and petD chloroplast genes in Chlamydomonas reinhardtii. FEBS Lett 284: 257–262 Chen X, Kindle K and Stern D (1993) Initiation codon mutations in the Chlamydomonas chloroplast petD gene result in temperature-sensitive photosynthetic growth. EMBO J 12: 3627–3635 Chen X, Kindle KL and Stern DB (1995) The initiation codon determines the efficiency but not the site of translation initiation in Chlamydomonas chloroplasts. Plant Cell 7: 1295–1305 Choquet Y, Stern DB, Wostrikoff K, Girard-Gascou J and Wollman FA (1998) Translation of cytochrome f is autoregulated through the 5´ untranslated region of petA mRNA in Chlamydomonas chloroplasts. Proc Natl Acad Sci USA, in press Cramer WA, Soriano GM, Ponomarev M, Huang D, Zhang H, Martinez SE and Smith JL (1996) Some new structural aspects complex and old controversies concerning the cytochrome of oxygenic photosynthesis. Annu Rev Plant Physiol Plant Mol Biol 47: 477—508 Drager R, Girard-Bascou J, Choquet Y, Kindle K and Stern D (1998) In vivo 5´-3´ exonuclease degradation of an unstable chloroplast mRNA. Plant J 13: 85–96 de Vitry C (1994) Characterization ofthe gene ofthe chloroplast Rieske iron-sulfur protein in Chlamydomonas reinhardtii: Indications for an uncleaved lumen targeting sequence. J Biol Chem 269: 7603–7609 de Vitry C, Olive J, Drapier D, Recouvreur M and Wollman F-A (1989) Posttranslational events leading to the assembly of Photosystem II protein complex: A study using photosynthesis mutants from Chlamydomonas reinhardtii. J Cell Biol 109: 991–1006 de Vitry C, Breyton C, Pierre Y and Popot J-L (1996) The 4 kDa nuclear-encoded PetM polypeptide of the chloroplast cytochrome complex. J Biol Chem 271: 10667–10671 Delosme R (1991) Electron transfer from cytochrome f to Photosystem I in green algae. Photosynth Res 29: 45–54 Finazzi G, Buschlen S, de Vitry C, Rappaport F, Joliot P and Wollman F-A (1997) Function-directed mutagenesis of the complex in Chlamydomonas reinhardtii: cytochrome inquinol binding Involvement of the cd loop of cytochrome to the Qo site. Biochemistry 39: 2867–2874 Fong SE and Surzycki SJ (1992) Organization and structure of plastome psbF, psbL, petG and ORF712 genes in Chlamy­ domonas reinhardtii. Curr Genet 21: 527–530 Girard-Bascou J, Choquet Y, Gumpel NJ, Culler D, Purton S, Merchant S, Laquerriere F, Wollman F-A (1995) Nuclear control of the expression of the chloroplast pet genes in Chlamydomonas reinhardtii. In: Mathis P(ed) Photosynthesis: From Light to Biosphere, pp 683–686. Kluwer Academic publishers, Dordrecht Gray JC, Phillips AL, Smith A (1984) Protein synthesis in chloroplasts. In: Ellis R (ed) Chloroplast Biogenesis, pp 137– 163. Cambridge University Press, Cambridge Gumpel NJ, Ralley L, Girard-Bascou J, Wollman FA, Nugent JH and Purton S (1995) Nuclear mutants of Chlamydomonas reinhardtii defective in the biogenesis of the cytochrome complex. Plant Mol Biol 29: 921–932 Harris E (1989) The Chlamydomonas Sourcebook. Academic Press, San Diego, Howe G and Merchant S (1992) The biosynthesis of membrane

Chapter 24 Cytochrome

complexes

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475 reinhardtii. Biochim Biophys Acta 851: 229–238 Lemaire C, Girard-Bascou J, Wollman F-A (1987) Characcomplex subunits and studies on the LHCterization of the kinase in Chlamydomonas reinhardtii using mutant strains altered in the complex, pp 655–658. I n : Biggins J (ed) Progress in Photosynthesis Research. Martinus Nijhoff Publishers, Dordrecht Link TA and Iwata S (1996) Functional implications of the structure of the ‘Rieske’ iron-sulfur protein of bovine heart mitochondrial cytochrome bc(1) complex. Biochim Biophys Acta 1275: 54–60 Martinez SE, Huang D, Szczepaniak A, Cramer WA and Smith JL (1994) Crystal Structure of Chloroplast Cytochrome f reveals a novel cytochrome fold and unexpected heme ligation. Structure 2: 95–105 Matsumoto T, Matsuo M and Matsuda Y (1991) Structural analysis and expression during dark-light transitions of a gene for cytochrome f in Chlamydomonas reinhardtii. Plant Cell Physiol 32: 863–872 Mould RM, Knight JS, Bogsch E and Gray JC (1995) Translocation of cytochrome f across the chloroplast thylakoid membrane. In: Mathis P (ed) Photosynthesis: From Light to Biosphere, pp 747–750. Kluwer Academic Publishers, Dordrecht Olive J, Vallon O, Wollman F-A, Recouvreur M and Bennoun P (1986) Studies on the cytochrome complex. II Localization of the complex in the thylakoid membranes from spinach and Chlamydomonas reinhardtii by immunocytochemistry and freeze-fracture analysis of mutants. Biochim Biophys Acta 851: 239–248 Olive J, Recouvreur M, Girard-Bascou J and Wollman FA (1992) Further identification of the exoplasmic face particles on the freeze-fractured thylakoid membranes: A study using double and triple mutants from Chlamydomonas reinhardtii lacking various Photosystem II subunits and the cytochrome complex. Eur J Cell Biol 59: 176–186 Pierre Y and Popot J-L (1993) Identification of two 4 kDa complex from Chlamy­ miniproteins in the cytochrome domonas reinhardtii. C R Acad Sci Paris 316: 1404–1409 Pierre Y, Breyton C, Kramer D and Popot J-L (1995) Purification complex of and characterization of the cytochrome Chlamydomonas reinhardtii. J Biol Chem 49: 29342–29349 Pierre Y, Breyton C, Lemoine Y, Robert B, Vernotte C and Popot J-L (1997) On the presence, role and evolutionary origin of a complex. J molecule of chlorophyll a in the cytochrome Biol Chem 272: 21901–21908 Popot J-L, Pierre Y, Breyton C, Lemoine Y, Takahashi Y, Rochaix J-D (1995) Purification and composition of the complex from Chlamydomonas reinhardtii. cytochrome In: Mathis P (cd) Photosynthesis: From Light to Biosphere, pp 507–512. Kluwer Academic Publishers, Dordrecht Qin L and Kostic NM (1992) Electron-transfer reactions of cytochrome f with flavin semiquinones and with plastocyanin. Importance of protein-protein electrostatic interactions and of donor-acceptor coupling. Biochemistry 31:5145–5150 Redinbo MR, Cascio D, Choukair MK, Rice D, Merchant S and Yeates TO (1993) The 1.5 Å crystal structure of plastocyanin from the green alga Chlamydomonas reinhardtii. Biochemistry 32:10560–10567 Sakamoto W, Kindle KL and Stern DB (1993) In vivo analysis of Chlamydomonas chloroplast petD gene expression using stable

476 transformation ofbeta-glucuronidase translational fusions. Proc Natl Acad Sci USA 90: 497–501 Sakamoto W, Chen X, Kindle KL and Stern DB (1994a) Function of the Chlamydomonas reinhardtii petD 5´ untranslated region in regulating the accumulation of subunit IV ofthe cytochrome complex. Plant J 6: 503–512 Sakamoto W, Sturm NR, Kindle KL and Stern DB (1994b) petD mRNA maturation in Chlamydomonas reinhardtii chloroplasts: Role of 5´ endonucleolytic processing. Mol Cell Biol 14: 6180–6186 Schmidt CL and Malkin R (1993) Low molecular weight subunits associated with the cytochrome complexes from spinach and Chlamydomonas reinhardtii. Photosynth Res 38: 73–81 Smith TA and Kohorn BD (1994) Mutations in a signal sequence for the thylakoid membrane identify multiple protein transport pathways and nuclear suppressors. J Cell Biol 126: 365–374 Soriano GM, Ponomarev MV, Tae GS and Cramer WA (1996) Effect of the interdomain basic region of cytochrome f on its redox reactions in vivo. Biochemistry 35: 14590–14598 Sturm NR, Kuras R, Buschlen S, Sakamoto W, Kindle KL, Stern DB and Wollman FA (1994) The petD gene is transcribed by functionally redundant promoters in Chlamydomonas reinhardtii chloroplasts. Mol Cell Biol 14: 6171–6179 Takahashi Y, Rahire M, Breyton C, Popot J-L, Joliot P and Rochaix J-D (1996) The chloroplast ycf7 (petL) open reading frame of Chlamydomonas reinhardtii encodes a small functionally important subunit of the cytochrome complex. EMBO J 15: 3498–3506 Trumpower BL and Gennis RB (1994) Energy transduction by cytochrome complexes in mitochondrial and bacterial respiration: The enzymology of coupling electron transfer reactions to transmembrane proton translocation. Annu Rev Biochem 63:675–716 Vallon O, Bulte L, Dainese P, Olive J, Bassi R and Wollman Fcomplexes A (1991) Lateral redistribution of cytochrome along thylakoid membranes upon state transitions. Proc Natl Acad Sci USA 88: 8262–8266 Widger WR, Cramer WA, Herrmann RG and Trebst A (1984) Sequence homology and structural similarity between

Francis-André Wollman cytochrome b ofmitochondrial complex III and the chloroplast complex: Position of the cytochrome b hemes in the membrane. Proc Natl Acad Sci USA 81: 674–678 Wollman F-A, Bulte L (1989) Towards an understanding of the physiological role of state transitions. In: Hall DO and Grassi G (eds) Photoconversion Processes for Energy and Chemicals, pp 198–207. Elsevier, Amsterdam Wollman F-A and Lemaire C (1988) Studies on kinase-controlled mutants from state transitions in Photosystem II and Chlamydomonas reinhardtii which lack quinone-binding proteins. Biochim Biophys Acta 85: 85–94 Wollman F-A, Kuras R, Choquet Y (1995) Epistatic effects in thylakoid protein synthesis: The example of cytochrome f. In: Mathis P (ed) Photosynthesis: From Light to Biosphere, pp 737–742. Kluwer academic publishers, Dordrecht. Wynn RM, Bertsch J, Bruce BB and Malkin R (1988) Green algal cytochrome complexes: Isolation and characterization from Dunaliella salina, Chlamydomonas reinhardtii and Scenedesmus obliquus. Biochim Biophys Acta 935: 115—122 Xia D, Yu C-A, Kim H, Xia J-Z, Kachurin A, Zhang L, Yu L and Deisenhofer J (1997) Crystal structure of the cytochrome complex from bovine heart mitochondria. Science 277: 60–66 Xie Z and Merchant S (1996) The plastid-encoded ccsA gene is required for heme attachment to chloroplast c-type cytochromes. J Biol Chem 271:4632–4639 Xie Z, Culler D, Dreyfuss B, Kuras R, Wollman F-A, GirardBascon J and Merchant S (1998) Genetic analysis of chlorplast c-type cytochrome assembly. Genetics 148: 681–692 Zhang H, Carrell CJ, Huang D, Sled V, Ohnishi T, Smith JL and Cramer WA (1996) Characterization and crystallization of the lumen side domain of the chloroplast Rieske iron-sulfur protein. J Biol Chem 271: 31360–31366 Zhou J, Fernandez-Velasco JG and Malkin R (1996) N-terminal mutants of chloroplast cytochrome f. J Biol Chem 271: 1–8 Zito F, Kuras R, Choquet Y, Kossel H and Wollman F-A (1997) in Chlamydomonas reinhardtii Mutations of cytochrome disclose the functional significance for a proline to leucine conversion by petB editing in maize and tobacco. Plant Mol Biol 33: 79–86

Chapter 25 Assembly and Function of the Chloroplast ATP Synthase Heinrich Strotmann

Institut für Biochemie der Pflanzen, Heinrich Heine Universität Düsseldorf,

D-40225 Düsseldorf, Germany

Noun Shavit* and Stefan Leu

The Doris and Bertie Black Center for Bioenergetics in Life Sciences,

Ben Gurion University of the Negev, Beer Sheva 84105, Israel

Summary I. Introduction II. Structure of A. Isolation and Subunit Composition B. Three-Dimensional Structure III. Molecular Genetics of A. Genes, Gene Arrangement, Gene Distribution and Evolution of B. Expression of atp Genes Complex C. Biogenesis and Assembly of the D. ATP Synthase Mutants and Site-Directed Mutagenesis of C. reinhardtii ATP Synthase IV. Mechanism of A. Kinetic Properties of and B. Catalytic Mechanism C. Coupling Mechanism V. Regulation of

A. ATP Hydrolyzing Activity of and B. Activation and Thiol Modulation of by Nucleotides C. Regulation of VI. Conclusions References

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Summary The type ATPase (‘ATP synthase’) is a central enzyme of bacterial, mitochondrial and is the chloroplast energy transducing membranes. The function of the chloroplast enzyme complex formation of ATP from ADP and phosphate at the expense of energy derived from a transmembrane electrochemical proton potential difference which is built up by photosynthetic electron transport (photophosphorylation). Recent results on the molecular genetics, structure and mechanism of the chloroplast enzyme, supplemented with data from its bacterial and mitochondrial counterparts, convey a more complete which requires the interplay between and precise image of this enzyme complex. The biosynthesis of the nuclear and plastid genome, depends on a variety of regulatory factors. The nuclear genes are controlled by *Deceased on 19th of June 1997

J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondriain Chlamydomonas, pp. 477–500. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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Heinrich Strotmann, Noun Shavit and Stefan Leu

light and by plastid derived factors, the expression of the chloroplast-synthesized subunits is regulated at the supplied important transcriptional and the translational level. The crystal structure of the mitochondrial structural information concerning the catalytic sites of the enzyme, and mutagenesis and crosslinking studies with the E. coli enzyme helped to localize important domains of interaction of the part with It is now wellestablished that catalysis and energy coupling involve movements of the ATPase structure, and regulation of also involves conformational changes. The structural basis and the molecular mechanisms of those motions are subjects of intensive research. C. reinhardtii has become a model organism for the investigation of mechanisms and biogenesis, due to its easy cultivation, the advances in molecular characterization of the ATP synthase genes, the possibilities for site-directed mutagenesis in the plastid as well as in the nuclear genome and improved methods for biochemical analysis. These studies will be especially helpful for the understanding of mechanistic and regulatory aspects that are specific for chloroplast ATP synthesis. This chapter reviews the state of knowledge about photosynthetic ATP synthases with special reference to work performed with C. reinhardtii.

I. Introduction The formation of ATP from ADP and phosphate in chloroplasts or cyanobacteria (‘photophosphorylation’) is energetically coupled to photosynthetic electron transport via a transmembrane proton gradient. While electrons flow through the electron transfer chain, protons are translocated across the thylakoid membrane into the thylakoid lumen. A transmembrane proton gradient is thus created and maintained due to the low proton permeability of the membrane. The electrochemical energy stored in the gradient is the driving force for ATP formation. ATP is formed when the protons flow back along the gradient through the membrane-bound reversible proton translocating ATPase (ATP synthase). Most recent results indicate that the translocation of four protons is required for synthesis of one molecule of ATP (Berry and Rumberg, 1996; van Walraven et al., 1996). The structure and biogenesis of the ATP synthase, the catalytic and coupling mechanism and their regulation are the subjects of this article. While most results on structure and mechanism have so far been obtained on ATP synthases from vascular plants, more data concerning biogenesis and assembly of Abbreviations: AMPPNP – – membrane sector of the chloroplast proton translocating ATPase (ATP synthase); -peripheral catalytic sector of the chloroplast proton translocating ATPase (ATP synthase); – chloroplast proton translocating ATPase (ATP synthase); DCCD – N,N´-dicylohexylcarbodiimide; – transmembrane electrochemical potential difference of the proton; – transmembrane pH difference; – membrane sector of the proton translocating ATPase (ATP synthase); – peripheral catalytic sector of the proton translocating ATPase (ATP synthase); – proton translocating ATPase (ATP synthase)

this enzyme have been contributed from research with Chlamydomonas reinhardtii. This organism is now also amenable to the application of molecular genetic methods for investigations of the mechanism of ATP synthesis. Photophosphorylation resembles oxidative phosphorylation which occurs in mitochondria and bacteria. The compositions and structures of the ‘oxidative’ and ‘photosynthetic’ ATP synthases are very similar and the catalytic mechanisms are essentially the same. Hence many results obtained with the mitochondrial or bacterial enzyme may be applied to the photosynthetic ATP synthases and vice versa.

II. Structure of

A. Isolation and Subunit Composition The ATP synthase complex is composed of two sectors, the thylakoid membrane-embedded responsible for transmembrane proton translocation (see also, Frasch, and the extrinsic catalytic 1994; Gromet-Elhanan, 1996; McCarty, 1996; Mills, 1996; Richter and Mills, 1996). The molecular weight of the holocomplex is about 540 kD. The sector of C. reinhardtii accounts for 420 kD as determined by gel exclusion chromatography and 390 kD as calculated from its sedimentation coefficient (Merchant et al., 1983). Thylakoid membranes from spinach chloroplasts contain about 1 nmol per mg chlorophyll (Strotmann et al., 1973). is located in the stroma-exposed, non-appressed areas of the thylakoid system; the sector is exposed towards the stroma. C. reinhardtii

Chapter 25 Assembly and Function of the Chloroplast Synthase

thylakoid membranes contain less per chlorophyll than spinach thylakoid membranes and, as in land plants, the ratio may vary depending on complex consists of growth conditions. The nine different subunits. The primary structures of all nine subunits are known from the sequences of the corresponding genes from several plant, cyanobacterial and algal species. can be solubilized by the nonionic detergent octylglucoside and subsequently isolated by sucrose gradient centrifugation (Pick and Racker, 1979). The isolated complex reconstituted into liposomes is capable of synthesizing ATP when energized by an artificial gradient generated by an acid/base transition (Possmayer and Gräber, 1994). When it is coreconstituted into liposomes with a light-dependent proton pump like bacteriorhodopsin, this system can produce ATP continuously in the light (Pick and from C. reinhardtii was Racker, 1979). isolated by similar techniques (Lemaire and Wollman, 1989a; Fiedler et al., 1995). The N-terminal sequences of all C. reinhardtii subunits have been determined (Fiedler et al., 1995) and the complete sequences of subunits are known seven C. reinhardtii from the analysis of the corresponding genes. Treatment of isolated thylakoid membranes with a from the dilute EDTA solution releases membranes (Avron, 1963) and this partial complex exhibits ATPase activity after it is suitably ‘activated’. membranes are unable to maintain The a proton gradient in the light since the uncovered is an open pore, but photophosphorylation can be restored to such vesicles by reconstitution with Thylakoid membranes, extensively isolated washed with pyrophosphate, release almost pure when they are resuspended in a sucrose-containing solution (Strotmannetal., 1973). Further purification to homogeneity may be achieved by subsequent anion exchange chromatography (Binder et al., 1978). consists of five different subunits, Thus isolated named by Greek letters, in the stoichiometry Another method of removal involves treatment of thylakoid membranes with chloroform (Younis et al., 1977). is released into the aqueous phase while the membranes gather in the chloroformwater interface. The extracted by chloroform, however, lacks the subunit and is not active in reconstitution of phosphorylation. Both isolation protocols, which were developed can be employed for originally for higher plant (Selman-Reimer the isolation of C. reinhardtii

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et al., 1981; Piccioni et al., 1981). The subunit stoichiometry of the four subunit C. reinhardtii preparation as determined from distribution of label was into uniformly (Merchant et al., 1983). A model of is shown in Fig. 1a and some properties of the subunits are listed inTable 1. Information concerning the function of the individual subunits has been obtained by various labeling techniques, by dissociation-reconstitution experiments and by analysis of the properties of the isolated subunits. The three and three subunits of spinach contain altogether six non-catalytic and catalytic nucleotide binding sites (Girault et al., 1988) and subunit contains a phosphate binding site (Pougeois etal., 1983). The isolated and subunits are able to bind nucleotides (Issartel and Vignais, 1984; Rao et al., 1988; Mills and Richter, 1991; Bar-Zvi et al., complex extracted with LiCl from 1992). The thylakoid membranes (Avital and Gromet-Elhanan, 1991) exists as a mixture of various stoichiometric assemblies and displays very low ATPase activity, but the preparation is stabilized into a Mg-ATPase complex by the fungal toxin tentoxin active (Gromet-Elhanan and Avital, 1992). Isolated subunit stabilizes the catalytic core (Gao et al., 1995; Miwa and Yoshida, 1989), and this is probably one function of in the natural complex. Moreover, subunit plays a role in energy transduction, and is involved in redox regulation of of chloroplasts from spinach (Schumann et al., 1985) and C. rein­ hardtii (Ross et al., 1995, 1996; Fiedler et al., 1997) to (see Section VB). Subunit seems to link (Beckers et al., 1992). The subunit was proposed also to control proton flux through (Engelbrecht and Junge, 1990). The subunit was shown to inhibit ATPase activity when added to depleted of subunit (Richter et al., 1984), and conformational changes of play a role in the (Richter and McCarty, induced activation of 1987; Komatsu-Takaki, 1992). Site-directed mutagenesis of E. coli suggests that this subunit may be involved in coupling as well (Zhang et al., 1994). consists of four different subunits, each of which has at least one hydrophobic trans-membrane helix. For the chloroplast ATP synthase, the subunits are designated with the Roman numerals I to IV and They are their stoichiometry is: homologous to the cyanobacterial constituents b, b´, c and a. The bacterial has only three different subunits but the mitochondrial enzyme

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contains additional polypeptides. Feng and McCarty (1990) reported the isolation of spinach from by zwittergent 3-12 treatment followed by DEAE-Trisacryl anion exchange chromatography. After incorporation into asolectin liposomes, and this resulted in could be rebound to reconstitution of energy transducing activities. Subunits III (‘proteolipid’) and subunit IV are involved in the transport of protons through Subunit I which can be crosslinked with the subunit (Beckers et al., 1992), seems to form a link between and Studies with zero length

Heinrich Strotmann, Noun Shavit and Stefan Leu

crosslinkers on the E. coli enzyme suggest that both the and subunits are involved in connections between the polar loop of the proteolipid molecule and the and subunits (Haughton and Capaldi, 1995; Watts et al., 1995; Zhang and Fillingame, 1995). On the other hand, Wetzel and McCarty complex and (1993 a) isolated a incorporated the partial complex into liposomes. the vesicle membranes could After removal of bind either or (Wetzel and McCarty, 1993b) suggesting that the subunit was not essential for binding.

Chapter 25 Assembly and Function of the Chloroplast Synthase

B. Three-Dimensional Structure Because of the similarities in the primary structures of the subunits, we may expect that results concerning the three-dimensional structure of the enzyme from higher plant chloroplasts may be applicable to the C. reinhardtii enzyme as well. High resolution electron microscopy of the detergent solubilized (Fig. 1b) after negative staining with spinach uranyl acetate shows a tripartite organization consisting of a head of 110 Å diameter and 83 Å length, a base part of 62 Å diameter and 83 Å length and a connecting stalk of 27 Å diameter and 37 Å length (Gogol et al., 1987; Boekema et al., 1988; Gräber et al., 1990). The head consists mainly of the three and three subunits whose alternate arrangement was demonstrated by decoration with subunit-specific antibodies (Tiedge et al., 1985). The base is formed by the membrane-domains of the subunits. The stalk is thought to comprise the extramembrane pieces of subunits I and II, part of and the subunits and (A recent report by Lill et al. (1997), however, suggests that subunit is attached rather than being located in to the top region of the stalk.) Subunits I and II have one trans-membrane helix at their N-terminal ends. The rest of these two polypeptides is hydrophilic and extends towards The hairpin-shaped proteolipid (subunit III) molecule consists of two membrane-spanning antiparallel connected by a polar loop that is exposed to (Girvin and Fillingame, 1995). Subunit IV is highly hydrophobic, has at least five membranespanning helices, and almost no extramembrane domains. The 10 to 12 copies of the proteolipid probably assemble to form a ring. It is not yet clear whether the membrane helices of subunits I, II and IV are located within or outside of this ring. An asymmetric model with subunit IV being located outside of the proteolipid cylinder is preferred by several workers in the field (Schneider and Altendorf, 1987; Vik and Antonio, 1994; Cross and Duncan, 1996). The sector of the beef heart mitochondrial ATP synthase (Fig. 1c) has been crystallized and its structure was resolved at 2.8 Å resolution by X-ray analysis (Abrahams et al., 1994). The structure confirms the alternating arrangement of the and subunits. The two polypeptides, which share about 20% sequence identity, are almost identical in their

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three dimensional structures and are arranged like the slices of an orange. The top part of the hexagonal structure is formed by a ‘crown’ consisting of the Nterminal domains of the and subunits that are arranged in six sheets each. The nucleotide binding sites are located in the central domain. In agreement with previous assumptions, the three catalytic sites reside mainly on the subunits at the interfaces with the subunits. The three non-catalytic nucleotide binding sites whose functional significance is unclear, are located mainly on the subunits at the interfaces with the subunits. The C-terminal domains of the and subunits consist of mainly helical structural elements. In the subunit, this domain can assume different positions depending on whether the catalytic site is occupied by a nucleotide or is empty. The C-terminal and the N-terminal domains of the subunit, which form reach into the central ring (Fig. 1c). More than 50% of cavity of the the subunit protrudes out of the basal part of and is assumed to participate in the formation of the stalk. This part ofthe subunit as well as subunits and are invisible in the crystal structure, probably owing to their high conformational flexibility. The invisible portion of the subunit contains the sequence responsible for redox regulation (Section V.B). Recently the three-dimensional structure of isolated subunit from E. coli was resolved by NMR methods. This polypeptide is L-shaped and consists of an Nterminal 10-stranded structure and a C-terminal part consisting of two antiparallel helices (Wilkens et al., 1995). The C-terminal part forms cross-links with a cluster of acid amino acids (‘DELSEED region’) in the C-terminal third of subunit (Dallmann et al., 1992), whereas the N-terminal part of seems to be in contact with the subunit (Wilkens et al., 1995; Watts et al., 1996) and with (Zhang et al., 1994; Zhang and Fillingame, 1995). in the complex is not The structure of necessarily identical to the structure of isolated mitochondrial Actually, electron cryomicroscopic analysis of two-dimensional crystals indicated that is rather different from that of the top view of the latter being highly asymmetric (Böttcher et al., 1995a). Unclear also is the dimension and structure of the stalk. Finally, certain features of the structure are at variance with results from biochemical and biophysical studies. For example the so-called ‘dark site cysteine’ of spinach subunit is the second amino acid from the C-

482 terminus and, according to the three-dimensional should be located in structure of mitochondrial the top part of —hidden in the central cavity of the hexagon. This cysteine could, however, be crosslinked with C89 of the subunit with the bifunctional maleimide N,N´-o-phenylenebismaleimide (Weiss and McCarty, 1977; Fritsche and Junge, 1996) although according to the three-dimensional structure it should be located at the opposite pole of i.e. more than 70 Å apart. On the basis of energy transfer experiments with covalently attached fluorescent and is no probes, the distance between more than 30 Å (Cerione et al., 1983). These examples of discrepancies show that, at this state of knowledge, has to the presentation of structural details of be considered preliminary.

III. Molecular Genetics of

A. Genes, Gene Arrangement, Gene Distribution and Evolution of Structural homologies indicate an evolutionary archaerelationship between vacuolar bacterial and type (Nelson, 1992). The common origin might have been the proton-translocating ATPase of a progenote that was the ancestor of archaebacteria, eukaryotes, and eubacteria including cyanobacteria. It seems that the type has evolved when the eubacteria came into being; since then the principal features of the

Heinrich Strotmann, Noun Shavit and Stefan Leu enzyme have remained remarkably unchanged. It is proposed that eukaryotic cells subsequently acquired the enzyme by endosymbiosis of cyanobacteria-like and purple bacteria-like prokaryotes that are presumed to be the ancestors of the chloroplasts and mitochondria, respectively (Gray, 1989). In E. coli and other bacteria, the three genes and the five genes are encoded in a single operon, called the atp or unc operon (Fig. 2). In Rhodo­ genes are located in two spirillum rubrum, the separate gene clusters, one comprising the genes and the other comprising the genes. This is supposed to be the primordial arrangement, and it has assembled from a membrane suggests that ion pore and a soluble ATPase (Falk and Walker, 1988). The same gene arrangement as in E. coli is also found in cyanobacteria, but there the operon is split into two different clusters, the atpA cluster harboring all genes and the genes atpD and atpA, and the atpB cluster containing atpB and atpE. Depending on specific species of cyanobacteria, the gene for subunit (atpC) is either separately encoded or attached to the atpA cluster. The gene cluster of Rs. rubrum and the atpA cluster of cyanobacteria, gene, atpG, respectively, contains an additional coding for subunit b´. Apparently atpG and atpF which show some sequence homology, originated from the same ancestral gene by duplication. The and have about 20% sequences of subunits identical amino acids suggesting that the two genes atpA and atpB also evolved from a common ancestral gene by duplication. The arrangement of the genes in

Chapter 25

Assembly and Function of the Chloroplast Synthase

the atpA operon is I-H-G-F-D-A(-C) and in the atpB operon B-E. During the evolution of chloroplasts from the cyanobacteria-like prokaryotic ancestors the gene clusters and the principal arrangement of the genes in the operons remained unchanged (Fig. 2). An exception is Chlamydomonas spp. where the ATPase genes are spread over the entire chloroplast genome due to extensive species-specific rearrangements in the chloroplast genome (Woessner et al., 1987; Boudreau et al., 1994). In contrast, the atpB­ atpE operon structure is maintained in the chloroplast genome of the green alga Chlorella (Yoshinaga et al., 1988). In the course of the evolution of eukaryotic plant cells up to three ATP synthase genes have been successively transferred from the chloroplast genome subunit to the nucleus. Except for AtpC, all genes are still in the chloroplast genome in the chromophyte Odontella sinensis (Pancic et al., 1992), the rhodophyte Porphyra purpurea (Reith and Munholland, 1995) and the glaucophyte Cyanophora paradoxa (Stirewalt et al., 1995). In higher plants gene AtpG are nuclear genes AtpC, AtpD and the (Herrmann et al., 1983) and it seems that the same gene distribution is true for C. reinhardtii and other green algae. Hence AtpC was among the first of the genes that were transferred to the nuclear genome in the common eukaryotic ancestor of all plants, and AtpD and AtpG were transferred later at the beginning of the evolution of the chlorophyll a+b-containing plants. As is the case for other protein complexes of the photosynthetic membrane, the genes for the most conserved and functionally most important subunits, i.e. atpA, atpB and atpH, were maintained in the chloroplast genome (Table 1). In contrast, only two or three subunit genes of the mitochondrial ATP synthase remained in the mitochondrial genomes of animal cells and Saccharomyces cerevisiae. Most mitochondrial subunits are nucleus-encoded. As yet, C. reinhardtii is the only organism where no ATP synthase genes at all were found in the mitochondrial genome (Michaelis et al., 1990). The mitochondrial ATP synthase of C. reinhardtii seems also to be exceptional in other respects. The enzyme which has recently been isolated and characterized, appears to consist of 14 polypeptides (Nurani and Franzén, 1996). However, only the and subunits could be clearly identified. The genes Atp1 (Nurani and Franzén, 1996) and Atp2 (Franzén and Falk, 1992) have been identified and sequenced. In contrast

483

to the corresponding subunits from all other organisms studied so far, the subunit contains a Cterminal extension of 66 amino acids, resulting in a subunit with a molecular weight of 63 kDa. The mature subunit has an N-terminal extension of 15– 18 residues. It will be interesting to study the consequences of these alterations on the properties of the C. reinhardtii mitochondrial enzyme.

B. Expression of atp Genes Since is composed of subunits encoded in the nucleus and in the chloroplast genome, multiple regulatory mechanisms are required to coordinate the expression of the corresponding genes. In land plant cells the atpA operon is transcribed as one long transcript and is subsequently processed to yield a complex pattern of smaller mRNA molecules (Herrmann et al., 1983, 1985). Expression of the genes in the required stoichiometry may be achieved by additional promoter elements found within this operon or by the complex mRNA processing mechanism of the initial transcript. In the atpB operon of spinach, the last four bases of the atpB reading frame are the first four bases of the atpE gene. The transcript of this operon appears not to be processed and expression of the and subunit in a 3:1 ratio is probably controlled at the translational level (Hennig and Herrmann, 1986). In C. reinhardtii, the ATP synthase genes located in the chloroplast genome are either monocistronically expressed, like atpB (Woessner et al., 1986), or arranged in new multicistronic operons, like atpA (Levy et al., 1997), atpE (Robertson et al., 1990) and atpI (C. Hauser, personal communication). The promoter regions and the 5´ and 3´ untranslated sequences of these genes do not share any obvious similarities, and thus the mechanisms coordinating their expression are not immediately obvious (Fiedler et al., 1997). In spinach, the nuclear genes AtpC, AtpD, and AtpG are expressed synchronously in response to light, organ-specific factors and plastid-derived signals (Bolle et al., 1996). The nuclear genes contain promoter elements that are regulated by signals from the plastids. This was demonstrated for the spinach AtpC and AtpD genes where expression is decreased in the tissues that contain photobleached plastids or where carotene biosynthesis is inhibited (Bolle et al., 1994). In spite of the obvious similarities in expression of these genes, the promoters and regulatory elements

484 of the AtpC and AtpD genes are surprisingly different (Bolle et al., 1996). On the other hand, biosynthesis of the chloroplastmade subunits may be regulated at the translational level by nucleus-derived factors that consequently have to be expressed in coordination with the nucleusencoded subunits. Expression of the C. reinhardtii atpA and atpB genes (as well as of many other C. reinhardtii chloroplast genes) was shown to be regulated by various, nucleus-encoded factors affecting specifically transcription, stability or translation of specific mRNAs (Drapier at al., 1992). These factors may correspond to a set of recently identified proteins which bind specifically to the 5´ untranslated region of chloroplast mRNAs, depending on their rate of translation (Hauser et al., 1996). In addition to the requirements for stoichiometric subunit expression, the expression of the ATP synthase genes is affected by diurnal cycle and fluctuations in environmental conditions (e.g. light and temperature). Various systems like greening of etiolated or embryonic cells of vascular plants (Gruissem, 1989), cells synchronized in the diurnal light-dark cycle (Leu et al., 1990b, Salvador et al., 1993) or greening of yellow mutant cells in C. reinhardtii (Wallach et al., 1972; Malnoe et al., 1988) have been employed to study the regulatory mechanisms involved in the biogenesis of chloroplast proteins. Regulation of expression in response to light has been studied by analysis of the ATP synthase activity in thylakoid membranes during greening of the C. reinhardtii mutant y-1 (Wallach et al., 1972). It was shown that the rise ofATP synthase activity precedes the accumulation of chlorophyll and the appearance of photosystems I and II. Pulselabeling of synchronized C. reinhardtii cells indicated are expressed early that the and subunits of in the cell cycle, specifically, in the first half of the 12 h light period, while chlorophyll, PS I and PS II are synthesized in the middle or the second half of the light period (Herrin and Michaels, 1984). Quantification of transcription and transcript levels for atpA and atpB demonstrated that the expression of these genes is controlled by increased transcriptional activity at the beginning of the light period (Leu et al., 1990b). Peaks in mRNA abundance were observed at 4 h in the light part of the cycle when translation was also found to be maximal (Leu et al., 1989). In contrast to other chloroplast proteins, however, which are only translated in the light, atpA and atpB mRNAs can be translated in the dark from

Heinrich Strotmann, Noun Shavit and Stefan Leu the small pool of available mRNA if the cells are supplied with acetate (Michaels and Herrin, 1990). The and subunits can also be synthesized by C. reinhardtii y-1 cells lacking chlorophyll (Malnoe et al., 1988).

C. Biogenesis and Assembly of the Complex The nucleus-encoded subunits are translated as soluble precursor proteins that are imported posttranslationally into the chloroplast. Their transit sequences are proteolytically cleaved before they assemble into the enzyme complex. The transit of C. reinhardtii is 35 (Yu and sequence of is 33 amino acids Selman, 1988) and that of long (Manthey et al., 1997) and they show similar into features. The import of C. reinhardtii isolated chloroplasts and its processing to mature has been demonstrated (Yu et al., 1988). In land plants, the chloroplast-located genes atpI and atpF contain short N-terminal sequences that are processed in the course of their integration into the thylakoid membranes (Hennig and Herrmann, 1986), and it is assumed that these N-terminal extensions act as signal sequences. The same seems to apply to C. reinhardtii (Fiedler et al., 1997). In C. reinhardtii, at least four different mechanisms contribute to determine the N-terminus of the mature chloroplastsynthesized proteins. The atpA and atpE genes are translated from the first AUG codon, but this methionine is removed from the polypeptide. The atpB gene is translated from the second methionine of the reading frame and the methionine is removed, or alternatively, if it is translated from the first methionine of the reading frame it is processed at the twelfth amino acid residue. The chloroplast atpI gene is synthesized as a precursor protein with a leader peptide of 8 amino acids that is subsequently processed (C. Hauser, personal communication). Finally, as in other organisms, the first methionine of the translation product of the atpH gene was found to be formylated. The chloroplast encoded subunits and were shown to be translated by thylakoid associated polysomes in land plants (Bhaya and Jagendorf, 1985) and C. reinhardtii (Herrin and Michaels, 1985). It has to be questioned whether this is indicative of a compartmentation of translation within the chloroplast, since soluble proteins are also translated on membrane-associated polysomes (Hattori and

Chapter 25 Assembly and Function of the Chloroplast Synthase

Margulies, 1986; Breidenbach et al., 1990). The and subunits synthesized by run off translation of thylakoid-bound C. reinhardtii chloroplast polysomes (Herrin and Michaels, 1985), or by in vitro translation of atpB mRNA in the reticulocyte lysate (Leu et al., 1990a), or in vivo in the presence of cycloheximide which inhibits synthesis of the subunit (Merchant and Selman, 1984), were assembled into the complex. It was therefore concluded that the thylakoid membrane harbors small pools of unassembled subunits that allow complex assembly when additional subunits are synthesized (Herrin and Michaels, 1985). An alternative explanation could be that the existing complex can partly dissociate under the experimental conditions employed and reassemble with in vitro translated subunits (Merchant and Selman, 1984). The pathway and mechanism of assembly of chloroplast ATP synthase is unclear. E. coli mutants lacking either or accumulate ATPase-active or functional proton channels, respectively (Aris et al., 1985; Klionsky and Simoni, 1985). In C. reinhardtii, mutants that are unable to synthesize one of the ATP synthase subunits owing to deletion of the gene or a defect in its expression (Table 2), do not accumulate unassembled subunits in the chloroplast even though all the remaining subunits are synthesized (Piccioni et al., 1981; Lemaire and Wollmann, 1989b, Robertson et al., 1990; Drapier et al., 1992; Fiedler et al., 1997). This indicates that all the unassembled subunits are rapidly degraded. In one very interesting C. reinhardtii assembly mutant, caused by two missense mutations in the atpA gene, the and subunits associate and form inclusion bodies in the chloroplast stroma, but they bind very poorly to the thylakoid membrane (Ketchner et al., 1995). This might indicate that the first step in ATP complex, synthase assembly is formation ofthe and subunits can readily bind as to which the shown by in vitro reconstitution studies (Richter et might then bind al., 1984; Gao et al., 1995). The which is already assembled in the thylakoid to membrane. An alternative pathway might be binding of the smaller subunits and to the complex as demonstrated in vitro by Engelbrecht and Junge hexamer (1992), and subsequent binding of the to this preassembled stalk. and from tobacco overexpressed Subunits in E. coli could be reconstituted to a fully active catalytic core complex with the help of a mixture of purified spinach chloroplast chaperones. Crucial components were Cpn60 and Cpn24, but other

485

components were also necessary (Chen and Jagendorf, 1994). The subunit prepared in soluble is able to reconstitute into form from spinach Rs. rubrum chromatophores (Richter et al., 1986). To what extent the chaperonin-like function of the subunit observed in vitro (Avni et al., 1991) is relevant to complex assembly in vivo remains to be clarified. Numerous missense mutations that prevent assembly of the ATP synthase complex have been identified, mostly in E. coli, but also in yeast and C. reinhardtii. These include many mutations in the Nand subunit, which terminal domains of the confirms the importance of this domain for stability of the ATP synthase complex as suggested by the partial three-dimensional structure of mitochondrial (Abrahams et al., 1994). Interestingly, in some of these mutants the binding of the portion to is specifically disturbed (Senior, 1990), which suggests that this (most membrane distant) domain of the complex nevertheless is indirectly involved in binding of to the portion. A number of heterologous complementation and in vitro reconstitution experiments demonstrated that chloroplast and bacterial subunits can substitute functionally for each other (Richter et al., 1986; Galmiche et al., 1994; Chen et al., 1995), and that can be reconstituted with bacterial and vice versa (Lill et al., 1993). The latter observation is surprising considering the very low sequence similarity between the subunits that are assumed to mainly I, be located at the interface between to II, and It appears that the subunit-subunit interactions are primarily determined by conserved secondary structure elements.

D. ATP Synthase Mutants and Site-Directed Mutagenesis of C. reinhardtii ATP Synthase A large number of non-photosynthetic strains with mutations in the chloroplast ATP synthase were identified and isolated in the course of producing nonphotosynthetic C. reinhardtii mutants, and these map to both the chloroplast and nuclear genomes (Harris, 1989). Several of these mutants have been analyzed at the molecular level; the best characterized strains are listed in Table 2. Plastid-inherited mutations could be assigned by complementation and recombination analysis to five groups, possibly representing five different chloroplast-encoded ATP synthase genes (Woessner et al., 1984). An important

486

finding of these studies was that nuclear mutations in other than the structural ATP synthase genes can affect specifically the synthesis of ATP synthase subunits by affecting transcript stability or translation. depends on This indicates that synthesis of specific regulatory factors (Section III.B). Techniques for manipulating the plastidic and nuclear genes of C. reinhardtii allow one to analyze chloroplast proteins by site directed mutagenesis.

Heinrich Strotmann, Noun Shavit and Stefan Leu

Strains FUD50 and ac-u-c-1-20 (= cc373) (Table 2) which carry deletions in the atpB gene were logical target recipient cells for introduction of mutated atpB genes. First complementation experiments with the atpB gene employed the subcloned Bam10 fragment or subclones thereof (Boynton et al., 1988; Avni et al., 1992; Hu et al., 1997). To facilitate the mutagenesis of atpB and to make the atpA gene accessible to mutational analysis as well, suitable

Chapter 25 Assembly and Function of the Chloroplast Synthase

recipient strains and complementing plasmids were designed. Deletion mutants were produced in the cell wall-deficient strain cw15 of C. reinhardtii by replacing the atpA and atpB reading frames with the aadA cassette (Leu et al., 1995). Likewise, suitable mutagenesis/complementation vectors for both genes were produced which facilitates the production of mutated genes and allows insertion of marker cassettes in the vicinities of these mutated genes (Hu et al., 1997). To make the mutant ATP synthase complex accessible to thorough functional analysis, protocols to isolate well-coupled thylakoid membranes from these strains were developed (Fiedler et al., 1997), and the catalytic properties of the C. reinhardtii were reassessed (Hu et al., 1997). A large number of strains with mutated atpA and atpB genes has been generated of which only a few have been characterized so far (Sections IV and V). Besides many mutations that had no effect on catalysis and a few that prevented the assembly of an ATP synthase complex, a third unexpected category was revealed. This third category appeared lethal to the cell. Specifically, when the mutated genes, linked to the aadA cassette, were introduced via transformation, spectinomycin resistant colonies appeared at normal frequency. However, the colonies could not be maintained and died after two to three weeks. A few (out of hundreds of transformants carrying various different mutations of the third category) that did survive, displayed abnormal integration of the mutated atpA/atpB DNA. This interesting phenomenon (S. L. and N. S. unpublished) is probably unique to the chloroplast systems and requires further study. A system that allows the introduction and analysis of site directed mutations in the nuclear gene AtpC was also developed. The AtpC was ‘knocked out’ by transformation of the strain nit1-305 (cw15) which is null for nitrate reductase (Smart and Selman, 1991), to generate a strain without a wild type copy of AtpC. The resulting strain is a suitable recipient for mutated versions of AtpC which are introduced by cotransformation with the cloned nitrate reductase gene (Ross et al., 1995, 1996). The results of mutageneses of AtpC will be discussed in Section V.B.

IV. Mechanism of

A. Kinetic Properties of

and

Dependent on the electrochemical potential of the

487

thylakoid membrane, the complex catalyzes either the formation or the hydrolysis of ATP. This reaction is coupled with the vectorial translocation of protons across the membrane. Isolated should, in principle, be able to catalyze ATP hydrolysis. However, in contrast to bacterial and mitochondrial the ATPase activity of land plant chloroplast is latent and has to be elicited. Activation procedures like heat or proteolytic treatment which induce Caof vascular plants, dependent ATPase activity in were not effective for the from C. reinhardtii. However, 50 mM DTT stimulated a Mg-dependent ATPase about twofold relative to the activity of the latent enzyme, and alcohols or other solvents stimulated a Mg-dependent ATPase up to 25 fold. Ethanol also de-stabilized the enzyme by rendering it more susceptible to heat denaturation (SelmanReimer et al., 1981; Kneusel et al., 1982). The response of the Mg-ATPase activity of C. reinhardtii to octylglucoside and tentoxin was similar to preparations from tentoxin insensitive that of vascular plants (Pick et al., 1982). Initially C. was found to exhibit a rather high reinhardtii ‘pre-activated’ ATPase activity of at 37 °C (Selman-Reimer et al., 1981; Kneusel et al., 1982), but recent results with purified in the presence of protease inhibitors indicated that the non-elicited activity was less than (Hu et al., 1997). Determination of the ATPase activities of this preparation yielded stimulation of MgATPase activity by ethanol, methanol and octylglucoside by a factor of the of about 40. The catalytic core complex C. reinhardtii enzyme has recently been prepared according to the procedure of Richter et al. (1984). The basal Mg-ATPase activity of the complex was very similar to that of and ethanol had the same stimulatory effect as in the complete complex (Richter et al., (Hu et al., 1997). As for spinach 1994), removal of did not elicit Mg-ATPase activity of the C. reinhardtii enzyme, but in contrast to of vascular plants, no significant Ca-dependent ATPase activity was induced by removal of the subunit (D. Hu, unpublished). forms bidentate nucleotide-metal ion complexes with the phosphate chains of ADP and ATP, respectively, and these Mg-nucleotide complexes are the actual substrates for (Zhou and Boyer, 1992). ATP hydrolyzing activity of heat or trypsin however, is most efficient in the activated because free presence of rather than

488 is less inhibitory than free (Hochman and behaves differently Carmeli, 1981). C. reinhardtii from the higher plant in this respect: CaATP was or a poor substrate but MgATP MnATP were good substrates of C. reinhardtii although the free metal ions were inhibitory (SelmanReimer et al., 1981; Kneusel et al., 1982). An enzyme carrying two mutations displays MgATPase activity similar to that of the wildtype enzyme and also displays significant CaATPase activity in the complex (D. Hu, unpublished). This suggests that the CaATPase can be induced by minor changes of activity of the primary structure of the subunit. The kinetic properties of the activity seem to be complicated and difficult to determine, as free free ATP and product ADP act as inhibitors. In the for Mg-ATP of about 0.2 absence of ethanol, a mM was obtained (Selman-Reimer et al., 1981) while in the presence of ethanol non-linear double ratio reciprocal plots were obtained. If the is kept constant at 2/3, the enzyme displays Michaelisof about 1 mM for ATP in Menten kinetics with a the presence of ethanol (Hu et al., 1997). At substrate concentrations in the physiological range, the kinetics of photophosphorylation can be described by the Michaelis-Menten equation. Enzyme (Strotmann et al., kinetic analysis at clamped 1990) using spinach thylakoids indicated that there is no particular compulsory order for substrate binding: rather the two substrates display random binding (Kothen et al., 1995). A comparison of some kinetic parameters for spinach and C. reinhardtii is given in Table 3. While the for phosphate is the same, the affinity for ADP and the maximal velocity appear to be considerably lower for photophosphorylation by C. reinhardtii thylakoids than for the higher plant thylakoids. At saturating the rate of phosphorylation is probably determined by the amount of the ATP synthase molecules per surface area. The fact that C. reinhardtii thylakoids exhibit only 40% of the phosphorylation rates obtained with spinach thylakoids, may be well explained by the density in the former. lower

B. Catalytic Mechanism The formation of ATP includes the elimination of a molecule of water from ADP and phosphate. phosphate and Experiments employing ADP, respectively, indicated that the

Heinrich Strotmann, Noun Shavit and Stefan Leu

bridge oxygen atom between the and the phosphate of ATP comes from the ADP molecule (Avron and Sharon, 1960). Conversely, during ATP hydrolysis the water-oxygen appears in the inorganic bridge oxygen remains with phosphate and the the ADP. ATP hydrolysis catalyzed by mitochondrial (Webb et al., 1980) or bacterial (Senter et al., 1983) was conducted by employing the analog of ATP containing and as further substituents in the group, and the Substituent inversion reaction was run in was observed in the released phosphate, suggesting mechanism with a pentacoordinated an bipyramidal intermediate. Since all biochemical onestep phosphoryl transfer reactions proceed with substituent inversion, it can be concluded that the process of oxidative phosphorylation and certainly photophosphorylation, too, do not involve a phosphorylated intermediate. Investigations with various natural nucleotides and nucleotide analogs have shown that the adenine moiety and the diphosphate chain are responsible for ADP recognition and orientation in the catalytic niche (Strotmann and Bickel-Sandkötter, 1984). The protein domains involved in catalysis and substrate binding have been identified by mutagenesis and by affinity labeling (Futai et al., 1989; Boyer, 1993) and, for crystallized beef heart mitochondrial by Xray analysis (Abrahams et al., 1994). Several amino acid residues, previously recognized by other methods, were found to be located in the catalytic and noncatalytic nucleotide binding sites (Fig. 3). The adenine binding domain is close to corresponds (numbering of C. reinhardtii); to a conserved tyrosine which was shown previously in mitochondrial to be photolabeled by 2-azidoADP (Garin et al., 1986). The phosphate chain binding domain, the so-called ‘P-loop’, comprises the amino acids to and this domain also contains a A bound water molecule is hydrogen-bonded to the carboxylic group of (C. reinhardtii

Chapter 25

Assembly and Function of the Chloroplast Synthase

numbering) and is appropriately positioned to promote in-line nucleophilic attack on the of ATP in ATP hydrolysis. Exchange of this glutamic acid for glutamine by site-directed mutagenesis in C. reinhardtii resulted in loss of photophosphorylation and loss of photoautotrophic growth (Hu et al., 1996). A water molecule is not present at the equivalent position of the non-catalytic nucleotide binding site in the subunit which is occupied by a glutamine Mutation of to E in C. reinhardtii had no noticeable consequences on the enzyme activity (S. Leu, unpublished). The threefold symmetry of the subunit couples implies that three identical catalytic sites are present

489

Catalysis may proceed with only one site per (‘uni-site catalysis’) when the substrate concentration is lower than the enzyme concentration. Under these conditions the turnover is very slow (Gräber and Labahn, 1992). It is, however, greatly accelerated by ADP and phosphate indicating that at least one additional catalytic site must be involved in normal catalysis (‘multisite catalysis’). Since the sites are in different functional states and are assumed to change their states alternatingly, the cooperation is explained by means of ‘alternating site models.’ In the ‘three site binding change’ hypothesis (Boyer, 1993), all three catalytic sites are assumed to pass sequentially through three distinct states, such that at any given

490 time each site is in a different state from that of the other two. In the three-dimensional structure of mitochondrial (Abrahams et al., 1994) the three catalytic sites are in different conformations due to differential substrate loading. One site contains ADP, the second the nonhydrolyzable ATP analogue AMPPNP, and the third is empty. The three different subunits form different ‘catch’ points with the two central helices of the subunit. During catalysis, the three subunits are thought to change their positions relative to and thereby pass through the different conformations. This is supported by recent experimental results with indicating that subunit rotates within isolated hexagon during ATP hydrolysis (Duncan et the al., 1995; Sabbert et al., 1996; Zhou et al., 1996; Noji et al., 1997). Thus functional rotation ofthe catalytic sites may be related to the physical rotation of the subunit. However, it should be mentioned that the participation of all three catalytic sites is still a matter of debate. There are experimental results indicating that only two catalytic sites are involved in steady state catalysis and the third one is in a nonfunctional state (Shapiro and McCarty, 1990; Berden et al., 1991). The natural tetrapeptide tentoxin is a powerful of some plants, but is ineffective for inhibitor of others (Steele et al., 1978). Tentoxin sensitivity or is determined by tolerance of higher plant subunit. the amino acid at position 83 of the Tentoxin sensitive enzymes have an aspartate at this position, but tolerant enzymes contain glutamate. C. reinhardtii belongs to the latter category (Avni et al., 1992), the corresponding amino acid is Replacement of the peptide domain of the C. reinhardtii subunit around by the corresponding sequence ofthe tobacco subunit (exchange of five amino acid residues) indeed yielded a tentoxin sensitive C. reinhardtii ATP synthase. The alone also converted C. reinhardtii exchange to a susceptible enzyme (Hu et al., 1997). however, did not induce The exchange tentoxin sensitivity. Position 72 (or 83 in spinach) is located in the crown domain of at the interface between and subunit. Seemingly, the more bulky glutamate and lysine residues prevent access of the tentoxin molecule to the binding site which itself is probably hydrophobic and involves other amino acids. Investigations of the mode of action of this inhibitor showed that binding of 1 mol of tentoxin per mol of specifically inhibits multisite catalysis (Hu et al.,

Heinrich Strotmann, Noun Shavit and Stefan Leu 1993). The results suggest that the crown domain— although far away from the catalytic sites— participates in the transmission of conformational signals between the catalytic sites. The wedge-shaped tentoxin which fits into the interfacial domain, might be visualized to inhibit an essential movement of and subunits relative to each other.

C. Coupling Mechanism In an aqueous environment, the equilibrium constant for the ATPase reaction favors greatly the formation of hydrolysis products. A shift of the reaction equilibrium towards ATP synthesis could be achieved in a catalytic niche of low water activity. At a water activity of instead of 55 M for example, the free energy change for the reaction would be instead of at 25 °C. Experiments conducted under uni-site conditions (Gräber and Labahn, 1992) indeed showed that the ratio of the complex over the concentration of concentration of enzyme-ATP complex is about 0.4 irrespective of thylakoid energization. On the other hand, the dissociation constants for ADP and phosphate were decreased and the dissociation constant for ATP was increased by membrane energization by several orders of magnitude. These results suggest strongly that the energy of the proton gradient must be invested into the binding of ADP and and the release of the ATP which might be achieved by active closure or opening of the catalytic sites, as proposed by Boyer and his colleagues. The ‘energy-linked binding change mechanism’ which is extensively discussed in a review by Boyer (1993) is widely accepted. Since the electrochemical gradient acts indirectly by changing the conformation of the enzyme, such a mechanism may be designated an ‘indirect’ or ‘conformational coupling mechanism.’ The crucial question is where and how the cyclic conformational changes are induced while the protons are translocated across the complex. membrane through the The actual energy transducing domain of the complex may be visualized as a screened domain containing pairs of acidic and basic amino acid residues which form salt bridges. Protonation of conjugate bases from the positively charged side of the membrane may break the salt bridges, and thereby induce a structural change which is somehow transmitted to the catalytic site. The site is opened, ATP is released and ADP and phosphate are bound to

Chapter 25 Assembly and Function of the Chloroplast Synthase

the active site. Substrate binding may induce closure of the catalytic site and simultaneous opening of the protonation domain to the negatively charged membrane side. The release of protons into this phase may restore the initial state. This general mechanism would also be applicable to the synthase of the bacterium Propionigenium modestum (Dimroth, 1994). The ratio may be determined by the number of protonatable groups involved. Under the influence of and the proton gradient, the interaction between is weakened owing to protonation of amino acid residues at the interface between the two partial complexes, and this effect is reversed by which suggests a relationship to energy transduction in phosphorylation (Fiedler et al., 1994). Accordingly the coupling between the proton gradient and ATP interface. synthesis may take place at the Dicyclohexyl carbodiimide (DCCD) is a classical DCCD ‘energy transfer inhibitor’ which acts on modifies the carboxyl group of the glutamic acid E61 (in E. coli: D61) of subunit III (=c) (Fillingame, 1992; Zhang and Fillingame, 1994), and thereby E/ blocks the translocation of protons through D61 is located almost in the middle of the C-terminal and is the only charged amino transmembrane acid within the hydrophobic membrane-spanning domain. Other essential amino acids in subunit III/c were identified by mutagenesis of E. coli including R41 and Q42 located in the polar loop which connects the two helices and faces the sector (Hatch et al., 1993; Fraga et al., 1994). In addition to the subunit III molecules, subunit IV (=a) has been proposed to be involved in the process of proton translocation. The subunits III and IV were proposed to form a water-filled transmembrane channel which is permeable only to specific cations (Junge et al., 1992). Kinetic measurements have proton channel of pea chloroplasts shown that the has a high conductivity, is very specific for (Lill et al., 1986) and is voltage-gated (Wagner et al., 1989). Subunit IV/a has at least five trans-membrane helices. Helix 4 is amphiphilic and has a face with partly conserved polar and charged amino acids. It was proposed that these amino acids together with E/D61 of the subunit III/c form a proton pore by interconnecting hydrogen bonds (Cox et al., 1986). A hypothesis for the mechanism of the energy transducing ATP synthase states that the primary energy transducing process is proton translocationdriven rotation induced in (Cox et al., 1984, 1986;

491

Cross, 1992; Vik and Antonio, 1994), which is propagated through the stalk via the subunit to the active sites. With every 120° turn the structural changes necessary for operation of the catalytic sites occur (Abrahams et al., 1994; Duncan et al., 1995; Cross and Duncan, 1996; Sabbert et al., 1996; Zhou et al., 1996). Mutagenesis studies have shown that in E. coli the amino acids R210, E219 and H245 of subunit a are critical for translocation (Howitt et al., 1988). For the E. coli enzyme, Vik and Antonio (1994) suggested that a ring consisting of 10 subunit c molecules functions as a rotor attached to subunit a which functions as a stator element. Residue a-R210 forms a salt bridge with a c-D61 residue ofone ofthe subunits of the subunit c cylinder. Protonation by a proton from the positively charged membrane side may dissociate the salt bridge, and cause a ratchettype motion of the rotor so that the unprotonated cD61 residue of the adjacent subunit c will associate with the a-R210. The amino acids a-E219 and aH245 are required to accept and guide the proton to the negative membrane side. The hypothesis could explain why binding of one molecule of DCCD per is sufficient to block continuous translocation (Sigrist-Nelson et al., 1978). The rotational movement of the cylinder of subunits c, might be transmitted to via the subunit. A connection between subunit c and appears possible on the basis of the recently complex of determined position of in the E. coli (Watts et al., 1995; Zhang and Fillingame, 1995). Rotation of the two helices of the subunit within the hexagon would then provide a mechanism for coupling the driving force to ATP synthesis. If so, a structure would be required which fixes the core to the membrane against the rotating axial subunits. Such an element has not been identified as yet. Lill et al. (1997) propose that subunit I subunit could connected to an represent such a device. is The mechanism of energy transduction in far from being solved and it should be emphasized that the rotational coupling mechanism discussed in this chapter remains speculative at present. In contrast to isolated where the catalytic activity matches the rotational speed, rotation of subunit in the complete complex at the speed which would be required for phosphorylation, has not been demonstrated. Attempts to measure rotations by phosphorescence emission anisotropy of the probe erythrosin isothiocyanate introduced into the subunit of have failed (Musier-Forsyth and Hammes, 1990).

Heinrich Strotmann, Noun Shavit and Stefan Leu

492 V. Regulation of

A. ATP Hydrolyzing Activity of

and

Plants or algae in their natural habitat experience diurnal cycles of light and darkness. Also, during the day, changes in light intensity are experienced as the illumination varies between bright sunshine and shade. Accordingly, the proton motive force produced in the chloroplasts is variable, too. At low proton motive force, the ATP synthase could work in the opposite direction, i.e. to hydrolyze ATP. Therefore, appears essential (Strotmann regulation of and Bickel-Sandkötter, 1984; Ort and Oxborough, 1992). Indeed, dark adapted chloroplasts show virtually no ATP hydrolyzing activity. In vitro ATP hydrolyzing activity of thylakoid membranes is obtained only after preillumination in the presence of a thiol compound like dithiothreitol (‘lighttriggered ATPase’). The isolated and are latent ATPases, too, and need activation by proteases, heat, alcohols, detergents and other means (Section IV.A). This suggests that the enzyme is indeed highly regulated.

B. Activation and Thiol Modulation of Photophosphorylation by thylakoid membranes from spinach chloroplasts shows a sigmoidal dependence on the proton gradient, the half maximal rate being of 3.4. But when the thylakoid obtained at a membranes are preilluminated in the presence of is shifted to 2.8 dithiothreitol, the midpoint (Junesch and Gräber, 1987). A similar shift is observed with C. reinhardtii thylakoid membranes (Fiedler et is al., 1997). This regulatory mechanism of called ‘thiol modulation’. Subunit of land plant or C. reinhardtii contains two cysteines separated by five amino acids between them. In spinach these two positions are and in C. reinhardtii and (Yu and Selman, 1988). The two cysteines can be oxidized to form a disulfide bridge. The reversible reduction of this disulfide bridge is the basis of thiol modulation (Nalin and McCarty, 1984). The regulatory segment is missing in other subunits including those from cyanobacterial (Cozens and Walker, 1987; Werner et al., 1990) or from the diatom Odontella sinensis (Pancic and Strotmann, 1993). In the subunit of the cyanobacterium

Synechocystis sp. 6803, a plant chloroplast-like regulatory segment was inserted by mutagenesis (Werner-Grüne et al., 1994), and this resulted in acquisition of thiol-modulation without change of the phenotype (Werner-Grüne et al., 1994; Krenn et al., 1995, 1997). Inversely, one or both cysteines were replaced by serines in the regulatory sequence of the C. reinhardtii subunit, and this resulted in the loss of thiol modulation in the strain carrying the mutation (Ross et al., 1995). The mutant strains, however, were able to grow photoautotrophically like the wild type strain. The spacer region between the two cysteines also seems to be important. For and instances, mutations prevented redox regulation (Ross et al., 1996). The natural reductant for the subunit of is a chloroplast thioredoxin (Mills et al., 1980). Recently it has been demonstrated with spinach thylakoid membranes in vitro that thioredoxin-f is more efficient than thioredoxin-m (Schwarz et al., complex 1997). The formation of a has been observed by fluorescence energy transfer between introduced fluorochromes (Dann and McCarty, 1992). Complex formation seems to be rate-limiting whereas the subsequent reduction step is fast (Schwarz et al., 1997). As thioredoxin is reduced by the photosynthetic electron transport chain via ferredoxin and ferredoxin-thioredoxin reductase (see Chapter 26, Jacquot et al.), the reduction of in vivo is dependent on light. Its reoxidation in the dark, on the other hand, most probably involves oxidized thioredoxin (Dann and McCarty, 1992). Efficient reduction of the disulfide bond requires a transmembrane proton gradient. The membrane energization causes a series of conformational changes of e.g. a change in the conformation and location of subunit within the complex (Richter and McCarty, 1987; Komatsu-Takaki, 1992) and a change of the structure of the catalytic sites which effects the release of a tightly bound nucleotide (Schumann and Strotmann, 1981; Bar-Zvi and Shavit, 1982). These reactions are related with the activation of the arrested enzyme, it thus seems that the proton gradient in addition to being the driving force for ATP synthesis, is also a kinetic factor. Membrane energization causes the hidden disulfide bridge of subunit to become accessible to the reductant (Schumann et al., 1985). Hence, in the intact is controlled by light in two chloroplast,

Chapter 25 Assembly and Function of the Chloroplast Synthase

different ways. Proton gradient dependent activation is the prerequisite for the reduction of the disulfide bridge by the photosynthetically reduced thioredoxin. Possibly the structural changes involved in activation are not principally different from those which drive catalysis of ATP formation (Section IV.C). Both, the process of activation and the catalytic process include changes ofaffinity for nucleotides of the catalytic sites, and the ‘activating’ and the ‘driving’ protons can not be distinguished kinetically (Groth and Junge, 1995). Hence activation might be the step connected with protonation which is repeated in every catalytic cycle. The enzyme is deactivated upon light-dark oxidized transition. The ATPase activity of in the subunit decays immediately with relaxation of the proton gradient, but if the subunit is reduced, the deactivation is much slower and activity lasts for some minutes. This is the reason why only are capable chloroplasts with thiol-modulated of hydrolyzing added ATP after transition from light to dark. In C. reinhardtii, however, the light-triggered ATP hydrolyzing activity was found to be very low suggesting a more rapid deactivation of the thiol modulated enzyme at the transition from light to dark (Fiedler et al., 1997). is thiol modulated, enzyme activity When as a function of the proton gradient has a midpoint of 2.2 (Junesch and Gräber, 1987). Under the same conditions, the midpoint of phosphorylation is 2.8. From this it was concluded that phosphorylation is energetically for reduced controlled and that the profile of phosphorylation represents its thermodynamic dependency. On the other hand, for the oxidized enzyme, the midpoint of phosphorylation is 3.4 and the profile is concluded to indicate the activation equilibrium of (Junesch and Gräber, 1987). the oxidized

C. Regulation of

by Nucleotides

The obvious advantage of thiol modulation is the more efficient utilization of light energy at small proton gradients (Mills and Mitchell, 1984; Junesch and Gräber, 1987; Ort and Oxborough, 1992). On the other hand, the thiol modulated enzyme seems to be less flexible to react fast to changes of the light regime due to the slow inactivation at the light/dark transition. Actually, some ATP is hydrolyzed in the dark in suspensions of thylakoid membranes

493

(Strotmann et al., 1987) as well as in intact chloroplasts (Altvater-Mackensen and Strotmann, 1988) until a critical ADP/ATP ratio is reached. The mechanism which provides control of under these circumstances is the deactivation by ADP. ADP deactivates by tight binding to the enzyme (Schumann and Strotmann, 1981; Bar-Zvi and Shavit, 1982), and it has been shown that the respective binding site is one ofthe three catalytic sites (Zhou et al., 1988). The tight binding ofADP and the velocity of inactivation of the ATPase activity is retarded by inorganic phosphate (Dunham and Selman, 1981; Schumann and Strotmann, 1981). The essential features of regulation are summarized in Fig. 4. Isolated also contains tightly bound nucleotides which may be the cause for the latency of ATPase activity. New studies indicate that purified complex both conC. reinhardtii and the tain 1 mol per mol enzyme of ADP tightly bound to one of the catalytic sites (D. Hu, unpublished). Treatment with ethanol (Kneusel et al., 1982) or octylglucoside (Pick and Bassilian, 1983) leads to acceleration of the release of tightly bound ADP. Acceleration of release of tightly bound nucleotides from membrane bound ATP synthase by methanol was also observed (Anthon and Jagendorf, 1984). This may thus be the reason for activation of the ATPase activity by these compounds. In fact, the C. reinhardtii enzyme is no longer inhibited by free ADP in the presence of 20% ethanol (Kneusel et al., 1982). The interaction of nucleotides and phosphate also comaffects the proton conductivity of the plex. The proton conductivity measured in the absence of ADP and phosphate is not exclusively due to the nonspecific leakiness of the thylakoid membrane, but to a significant extent due to leakiness of the complex (‘proton slip’). The pro ton leakiness is abolished, or at least reduced, by tight nucleotide binding and is converted to a productive proton flow upon binding ofboth ADP and phosphate (Groth and Junge, 1993). On the other hand, at high phosphorylation potentials and high proton gradients, ATP binding triggers the opening of the proton channel (Strotmann et al., 1986) and this reaction may be conceived as a valve which protects the thylakoid lumen from over-acidification under conditions where protons are effectively pumped into the thylakoid lumen but phosphorylation is excluded.

494

VI. Conclusions A great deal of recent progress in ATP synthase research is based on molecular genetic and structural work, in particular the elucidation of a partial structure of the mitochondrial We have good hopes that the molecular mechanism of ATP hydrolysis will be unraveled in near future. Much less understood and more difficult to investigate is the mechanism of coupling and energy transduction that enables the complex to form ATP. The investigation of the interface seems to be structure of and of the an important prerequisite, but as protein movements and conformational changes are involved, new molecular biophysical techniques may also be required. All basic mechanistic principles can be studied in ATP synthases of either bacteria, mitochondria or chloroplasts. However, the photosynthetic ATP synthases have specific regulatory properties which must be investigated in detail. Chlamydomonas may be a useful model organism for the study of those aspects of the chloroplast enzyme. Moreover, the expression of the ATP synthase genes and its control, the interplay between the plastid genome and nucleus genome and the assembly of the ATP synthase subunits are insufficiently investigated. The advantages of the Chlamydomonas system over land plants could be exploited for such studies as well. References Abrahams JP, Leslie AGW, Lutter R and Walker JE (1994) Structure at 2.8 Å resolution of from bovine heart

Heinrich Strotmann, Noun Shavit and Stefan Leu

mitochondria. Nature 370: 621–628 Altvater-Mackensen R and Strotmann H (1988) Contents of endogenous adenine nucleotides and ATPase activity of isolated whole chloroplasts. Biochim Biophys Acta 934: 213–219 Anthon GE and Jagendorf AT (1984) Effect of methanol on spinach thylakoid ATPase. Biochim Biophys Acta 723: 358– 365 Aris JP, Klionsky DJ and Simoni RD (1985) The subunits of the Escherichia coli synthase are sufficient to form a functional proton pore. J Biol Chem 260: 11207–11215 Avital S and Gromet-Elhanan Z (1991) Extraction and purification of the subunit and an active complex from the spinach chloroplast Synthase. J Biol Chem 266: 7067–7072 Avni A, Avital S and Gromet-Elhanan Z (1991) Reactivation of beta subunit by trace amounts of the chloroplast the alpha subunit suggests a chaperonin-like activity for alpha. J Biol Chem 266: 7317–7320 Avni A, Anderson JD, Holland N, Rochaix JD, Gromet-Elhanan Z and Edelman M (1992) Tentoxin sensitivity of chloroplasts determined by codon 83 of subunit of proton-ATPase. Science 257: 1245–1247 Avron M (1963) A coupling factor in phototphosphorylation. Biochim Biophys Acta 77: 699–702 Avron M and Sharon N (1960) A study of photophosphorylation with oxygen-18. Biochem Biophys. Res Commun 2: 336–339 Bar-Zvi D and Shavit N (1982) Modulation of the chloroplast ATPase by tight binding of nucleotides. Biochim Biophys Acta 681: 451–458 Bar-Zvi D, Bar I, Yoshida M and Shavit N (1992) Covalent binding of 3´-O-(4-benzoyl)benzoyl adenosine-5´-triphosphate (BzATP) to the isolated and subunits and the core complex of binding of BzATP prevents association of the and subunits and induces dissociation of the core complex. J Biol Chem 267: 11029–11033 Beckers G, Berzborn RJ and Strotmann H (1992) Zero-length crosslinking between subunits and I of the ATPase of chloroplasts. Biochim Biophys Acta 1101: 97–104 Berden JA, Hartog AF and Edel CM (1991) Hydrolysis of ATP by can be described only on the basis of a dual-site mechanism. Biochim Biophys Acta 1057: 151–156 Berry S and Rumberg B (1996) ration at the unmodulated

Chapter 25 Assembly and Function of the Chloroplast Synthase

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Chapter 26

Molecular Aspects of Components of the Ferredoxin/Thioredoxin Systems Jean-Pierre Jacquot, Mariana Stein, Stéphane Lemaire,

Paulette Decottignies, Pierre Le Maréchal

Institut de Biotechnologie des Plantes, ERS 569 CNRS, Université de Paris-Sud,

Bâtiment 630, F-91405 Orsay Cedex, France

Jean-Marc Lancelin

RMN Biomoléculaire, ESA CNRS 5078, Université Claude Bernard - Lyon I,

Bat. 308, CPE-Lyon, F-69622 Villeurbanne, France

Summary I. Introduction II. Ferredoxin Dependent Systems A. Molecular Studies of Ferredoxin 1. Isolation andSequence Determination 2. cDNA and Gene Isolation and Characterization 3. Expression in Escherichia coli and Mutagenesis B. Molecular Studies of Ferredoxin-NADP Oxidoreductase 1. Aminoacid and cDNA Sequences 2. Methylation Properties III. Thioredoxin Dependent Systems A. NADPH and Ferredoxin Dependent Thioredoxin Reductases B. Thioredoxins 1. Isolation and Sequence Analysis 2. Comparative Gene Structure 3. Deduced Cleavage Sites of the Preproteins 4. NMR Deduced Three-dimensional Structure 5. Biochemical Properties and Mutagenesis IV. Conclusion Acknowledgment References

501 502 505 505 505 505 506 507 507 507 508 508 508 508 509 510 510 510 511 512 512

Summary In the chloroplasts of eukaryotic photosynthetic cells, ferredoxin and thioredoxin are linked in a redox cascade which activates several catalysts of the Calvin cycle by molecular reduction of selected disulfide bridges. These two redox constituents are also present in Chlamydomonas reinhardtii and their properties are described. Ferredoxin is a nucleus-encoded protein synthesized as a 126 amino-acid precursor which is processed to a 94 amino-acid mature protein. In contrast with the intronless genes of ferredoxin in land plants, the C. reinhardtii ferredoxin gene (Frx1) contains a single intron and is present in very few copies. In chloroplasts, ferredoxin is located at a metabolic branchpoint. Ferredoxin distributes electrons to many biochemical pathways involving J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 501–514. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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reductive chemistry; e.g. reduced ferredoxin is a substrate of the enzymes ferredoxin oxidoreductase (FNR) and ferredoxin-thioredoxin reductase (FTR). Several characteristics of electron transfer from ferredoxin to FNR and FTR will be described. Evidence will also be presented that FNR is subject to post-translational modification (methylation) in C. reinhardtii. As in land plants, C. reinhardtii cells contain several types of thioredoxin, with different subcellular localization. Thioredoxin h is nucleus-encoded, located in the cytosol and does not require a transit sequence. On the other hand, thioredoxin m (106 amino-acids in its mature form) is also nucleus-encoded, synthesized as a precursor of 140 amino-acids and located in the chloroplast. The genes coding for the two thioredoxins (Trx1 and Trx2) have been isolated and sequenced. In addition, the molecular 3D-structures of these two proteins will be presented as well as some of their biochemical characteristics.

I. Introduction Chloroplast ferredoxin is a small soluble protein (molecular weight ca 10 kDa), which transfers electrons via a 2Fe-2S center by shifting from the to the state. In the chloroplast, this protein is reduced by light and the photosynthetic electron transfer chain (Arnon, 1988). Ferredoxin is located at a metabolic branchpoint. Reduced ferredoxin distributes electrons to many biochemical pathways, thus playing a key role in NADP photoreduction, sulfur and nitrogen assimilation as well as in redox-linked photoregulation of enzymes. A summary of the sites of interaction for ferredoxin is shown in Fig. 1. At the molecular level, [2Fe-2S] ferredoxin is constituted of a polypeptide of ca 96 amino-acids, to which an iron-sulfur center is attached through four cysteine ligands. The [2Fe-2S] ferredoxins show a very high degree of identity (typically ca 70%) in all photosynthetic organisms studied, ranging from cyanobacteria to land plants (Matsubara and Hase, 1983). Cyanobacteria and land plants do, however, also contain related [2Fe2S] ferredoxin sequences with lower similarities (for example, five different sequences are present in the Synechocystis sp. 6803 genome) (Kaneko et al., 1996). Due to its ability to interact with numerous proteins, ferredoxin has also been subjected to site directed mutagenesis. Most of these studies have been performed either with the protein from Anabaena sp. or with the spinach protein. Depending on the protein studied, the results differ a little bit, but it is generally assumed that the site of interaction of ferredoxin with the other molecules involves Abbreviations: DTNB–5-5´ dithiobis nitrobenzoic acid; FNR – ferredoxin NADP+ oxidoreductase; FAD – flavin adenine dinucleotide; FTR – ferredoxin-thioredoxin reductase; NADPMDH – NADP-malate dehydrogenase; NTR – NADPH dependent thioredoxin reductase; TRX – thioredoxin

electrostatic interactions with essential negative charges on the ferredoxin molecule and positive charges on its protein partners. A patch of negative charges located near the C-terminus ofthe ferredoxin molecule has been proposed to play a major role in these interactions (de Pascalis et al., 1993; Hurley et al., l993; Piubelli et al., 1996). Thioredoxins are small molecular weight (ca 12 kDa), generally extremely thermostable proteins which possess disulfide oxidoreductase activity. These properties are due to the existence of a very reactive disulfide bridge located in a protrusion of the molecule (Holmgren, 1989). The thioredoxin active site is overwhelmingly conserved and features the following sequence: -Trp-Cys-Gly-Pro-Cys-(Lys/Arg)While bacteria, anaerobic photosynthetic prokaryotes and animal cells rely exclusively on NADPH and a flavoprotein called NADPH-thioredoxin reductase to reduce thioredoxin, all aerobic photosynthetic organisms do contain an alternate reducing system constituted by the photosynthetic electron transfer chain, ferredoxin and the protein ferredoxin-thioredoxin reductase (Buchanan, 1991). The two systems coexist in eukaryotic photosynthetic cells. A schematic representation of the two reducing systems in plants is shown in Fig. 2. Besides containing two thioredoxin reducing systems, eukaryotic photosynthetic cells also have the peculiarity of containing multiple isoforms of thioredoxin. These thioredoxin isoforms can be differentiated both by sequence analysis and by their kinetic properties. Sequence analysis indicates that three different thioredoxin groups are present in eukaryotic photosynthetic cells (Jacquot et al., 1997a). The first contains proteins similar in structure to the prokaryotic thioredoxins (model, Escherichia coli). These proteins are nucleus-encoded, chloroplast-

Chapter 26

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localized and reduced by ferredoxin and the enzyme ferredoxin-thioredoxin reductase. In vitro, these proteins catalyze efficiently the reduction/ activation of the target enzyme NADP-malate dehydrogenase and accordingly they have been called thioredoxin m. A second group features nucleus-encoded chloroplastic proteins which are extremely specialized in activating a catalyst of the Calvin cycle, fructose1,6-bisphosphatase. These proteins called thioredoxin f differ much in their primary structure from other known thioredoxins, a feature that is related to their catalytic properties. As thioredoxin m, thioredoxin f is reduced by the ferredoxin/ ferredoxinthioredoxin reductase system. The third type of

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protein is also nucleus-encoded, but located outside the chloroplasts, presumably in the cytosol of the cells. As it is also present in non photosynthetic cells, it has been called thioredoxin h (for heterotrophic type). Multiple genes have been isolated for thioredoxin h from Arabidopsis thaliana and the enzyme targets of these proteins are not well known. Thioredoxin h is reduced by NADPH and the flavoprotein NADPH-thioredoxin reductase (Buchanan et al., 1994). Sequence analysis indicates that thioredoxin h shares a rather high degree of identity with mammalian thioredoxins (Rivera-Madrid et al., 1995; Sahrawy et al., 1996). It is also interesting to mention that thioredoxin has been reported to occur

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in plant and animal mitochondria (Bodenstein-Lang et al., 1989) Research in the thioredoxin area is currently extremely active for several reasons. One of these is that it was discovered that thioredoxin can modulate the activity of transcription factors (such as NFKB, fos and jun) and receptors (including the glucocorticoid receptor in animal cells and the S-locus receptor kinase in plants) by redox regulation in vitro (Babiychuk et al., 1994; Schenk et al., 1994; Bower et al., 1996). In C. reinhardtii, it has been proposed that it could regulate translation by modulating the binding ofactivator proteins to chloroplast messenger RNA (Danon and Mayfield, 1994). As a consequence, thioredoxin is likely to be involved in signaling pathways and developmental processes. Second, thioredoxin is an excellent model for structural studies

(involving NMR and crystallography) because of its high stability and relatively simple architecture. Third, thioredoxin is becoming ever more used in biotechnological applications, including the production of recombinant proteins which are difficult to obtain in a soluble form. This problem has often been solved by expressing them as a fusion product with Escherichia coli thioredoxin. The present article will focus on the ferredoxinand thioredoxin-dependent systems in C. reinhardtii. It will describe how the ferredoxin and thioredoxin proteins were isolated, their cDNAs and genes cloned, and expression systems devised. The results of site directed mutagenesis studies, which provide an understanding of the interactions of ferredoxin and thioredoxins with their protein partners will be presented. The present state of knowledge concerning

Chapter 26 Ferredoxin and Thioredoxin Systems the pathways of reduction of thioredoxins in C. reinhardtii will be described, as well as the structures of the two C. reinhardtii thioredoxins m and h.

II. Ferredoxin Dependent Systems

A. Molecular Studies of Ferredoxin

1. Isolation and Sequence Determination In order to isolate the nucleic acid sequences coding for C. reinhardtii ferredoxin (and thioredoxins), the proteins were isolated and purified from large batch cultures. Since ferredoxin (like thioredoxins) has a small molecular mass (being composed of a polypeptide ofca 100 amino-acids), it was quite easy to establish the primary structure of this protein by direct amino-acid sequencing. In the case of ferredoxin, the protein is very abundant in algal cells, which further facilitated the study. C. reinhardtii ferredoxin was found to be a single polypeptide of 94 amino-acids (Schmitter et al., 1988). A comparison of the amino-acid sequences of several land plant and cyanobacterial ferredoxins with the protein of C. reinhardtii is shown in Fig. 3. As expected, the amino-acid identity is extremely high (ca 70%), and the 4 cysteines involved in the binding of the [2Fe2S] iron-sulfur center are strictly conserved.

2. cDNA and Gene Isolation and Characterization Taking advantage of the coding bias of C. reinhardtii, oligonucleotides were synthesized in order to clone

505 the cDNA fragment coding for the mature part of the protein by application of PCR (Rogers et al., 1992). The sequence predicted by the cDNA was identical to the protein sequence except that Thr 7 was replaced by Ser. Interestingly, the codon preference of the ferredoxin sequence matched extremely well the coding bias reported for the nuclear genes of C. reinhardtii (Campbell and Gowri, 1990), except for Ala residues for which the GCC and GCT codons were found to be equally used. The cDNA fragment has been used in order to isolate cDNA and genomic sequences coding for ferredoxin by classical hybridization techniques. A full length cDNA sequence has been obtained and since this protein is nucleus-encoded but chloroplast located, it features a transit peptide (Stein et al., 1993). The preferredoxin protein contains 126 amino acids (molecular mass 13250 Da), which is cleaved into a mature 94 amino-acid protein (9908 Da). As a consequence the transit peptide contains 32 aminoacids (3342 Da) with the following sequence: MAMAMRSTFAARVGAKPAVRGARPASRMSCMA

The C. reinhardtii ferredoxin transit peptide is composed of only 11 different amino-acid residues, and eight amino-acids, most notably the aromatics (except one Phe) and the acidic ones, are absent. The absence ofnegative charges is a feature characteristic of transit peptides (see Chapter 13, Ferret et al.). On the other hand, this sequence is extremely rich in Met, Ala and Arg, and hence extremely positively charged. This property is also overwhelmingly found in all chloroplastic transit sequences. One problem with transit sequences however is that they exhibit very little homology to one another when the primary

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structure is considered and it is thus difficult to recognize cleavage sites for example. It has thus been postulated that the recognition motifs for the chloroplastic envelope lie at a higher structural degree (e.g. secondary or tertiary structure). In order to test this hypothesis, we have used NMR to determine the structure of the ferredoxin transit sequence. While the chemically synthesized peptide is essentially unstructured in aqueous solution, the addition of trifluoroethanol, a compound known to mimic amphiphilic media, promoted the formation of an helix between residues 3 and 13 (Lancelin et al., 1994). It is thus possible that, at the contact of the hydrophobic chloroplast envelope, the N-terminus of the preferredoxin polypeptide is organized into an helix able to recognize the outer membrane and which facilitates the subsequent transport and integration of the protein inside the chloroplast stroma. The gene coding for ferredoxin (Frx1) has been isolated after screening ofa genomic library (Stein et al., 1995). Southern blot analyses indicate that it is likely to be present at a very low copy number (possibly only one or two copies per genome). The C. reinhardtii gene is split into two exons of 59 and 67 amino-acids by an intron situated between residues E59 and E60 (precursor numbering). The first exon codes for the transit peptide (32 amino acids) and the first 27 amino acid residues of the mature protein. It is remarkable that the C. reinhardtii ferredoxin gene is the only genomic sequence isolated so far, coding for a [2Fe-2S] ferredoxin which contains an intron. All ferredoxin genes isolated from land plants so far are intronless (Elliot et al., 1989; Vorst et al., 1990; Bringloe et al., 1995). This property indeed suggests that C. reinhardtii is not situated in the same evolutionary branch as land plants. The structure of the ferredoxin gene is compared on a schematic drawing to those of thioredoxins m and h (see below Fig. 6).

3. Expression in Escherichia coli and Mutagenesis The cDNA coding for C. reinhardtii ferredoxin has been introduced by PCR into the expression plasmid pET-3d (Studier et al., 1986). When using this construction, ferredoxin is produced as a recombinant protein by Escherichia coli cells and can be purified as a holoprotein with its associated iron-sulfur center. Due to the nature of the construction (fusion), the

recombinant protein was found to start with the sequence: AYKVT–Other than this additional N-terminus Ala residue, the recombinant ferredoxin was indistinguishable from the algal ferredoxin, whether considering its spectroscopic properties (EPR and UV-visible absorption) or its kinetic behavior (Rogers et al., 1992). It thus constitutes an excellent model for site directed mutagenesis studies. Due to its numerous protein-protein interactions (see Fig. 1), ferredoxin is certainly a fascinating protein for such studies. Especially interesting, is the property that the interaction sites of ferredoxin with ferredoxin NADP oxidoreductase (FNR) and ferredoxin-thioredoxin reductase (FTR) are not equivalent. Several lines of evidence indicate that C-terminal negative charges play an important role for the interaction with these two proteins, and in addition, other negative charges closer to the center of the polypeptide are important for the interaction with FNR (de Pascalis et al., 1993; Hurley et al., 1993; Piubelli et al., 1996). We have brought our own contribution to this problem by preparing by PCR several mutants of ferredoxin and testing them with FNR (cytochrome c reduction assay) and FTR (NADP-malate dehydrogenase activation assay) (Table 1). It was extremely exciting to observe that the loss of a single negative charge (on Glu 91, recombinant protein numbering, and not of its neighbor Glu 92) completely suppressed reactivity with FTR but not with FNR (Jacquot et al.,

Chapter 26 Ferredoxin and Thioredoxin Systems 1997b). In addition, the triple mutant D25A, E28Q, E29Q was also inactive with FTR but not with FNR. The availability of these specific mutants suggests that they could be used as tools to evaluate the in vivo effect of the ferredoxin thioredoxin system. At this point, homologous recombination is not very easy for nuclear genes in C. reinhardtii. Should it become feasible, it would be extremely interesting to introduce these mutations back into the algal cells.

507 In addition, the combination of these two papers gives indications on at least one transit sequence of FNR which is as follows: MASLRKPSNHADRACSRRLRVATRVAGRRMCRPVA

B. Molecular Studies of Ferredoxin-NADP Oxidoreductase (FNR)

This sequence contains 35 amino acids and is composed of 15 different amino-acids with a very high predominance of R+A (40% of the sequence). As for ferredoxin, acidic and aromatic amino acids are rare or absent. As for ferredoxin, –1 of the cleavage site lies an Ala residue.

1. Aminoacid and cDNA Sequences

2. Methylation Properties

Since ferredoxin establishes strong electrostatic interactions with its protein partners, ferredoxinSepharose columns can be used to isolate FNR and FTR from algal extracts. This procedure was used in combination with 2´5´ ADP chromatography to purify FNR from C. reinhardtii cells to homogeneity. The FNR protein was found to be also very abundant and monomeric with a molecular mass of ca 35 kDa and one FAD per subunit. Despite its relatively high molecular mass, the complete amino-acid sequence of the protein was then determined by direct sequencing (Decottignies et al., 1995). The FNR polypeptide was found to be constituted by a sequence of 320 amino-acids with a calculated molecular mass of 35685 Da (36470 Da with the FAD moiety). The C. reinhardtii FNR sequence showed ca 35% identity when compared to other plant or algal FNR’s, with some stretches of amino-acids extremely conserved. A cDNA sequence was also determined for ca 80% of the mature protein, following the same strategy as for ferredoxin (i.e. PCR cloning using biased oligonucleotides). The sequence deduced from the cDNA was exactly identical to the sequence determined by direct amino-acid sequencing (Decottignies et al., 1995). A complete cDNA sequence of C. reinhardtii FNR, including the deduced structure of the transit sequence (as ferredoxin, FNR is nuclear encoded and imported into the chloroplast) has also been determined by Kitayama et al. (1994). In general this sequence is identical to the one published in Decottignies et al. (1995) except that the sequence 90-EVPHGTRLYS-99 was replaced by EVPTARLYS. In addition, two differences (F109G and A212G) were observed. The analysis of these sequences may therefore indicate that there are at

least two isogenes coding for FNR in C. reinhardtii.

Besides indicating the exact cleavage site ofthe FNR preprotein, the direct amino-acid sequencing of C. reinhardtii FNR also gave another important piece of information: the protein is post-translationally modified. It was indeed found that two amino acids (K83 and K89) are trimethylated and one residue (K135) dimethylated (see Fig. 4). (Decottignies et al., 1995). This conclusion is based on three observations: (i) the HPLC elution profile of the corresponding derivatized amino-acids after sequencing was consistent with trimethylated and dimethylated lysines; (ii) the molecular mass of the corresponding peptides (estimated by laser desorption mass spectrometry) was also consistent with the proposed modification; (iii) cDNA analysis indicated codons for lysine residues at the corresponding positions. Of these three methylated residues, one (K89) was proposed to be essential for the interaction with ferredoxin in spinach (Aliverti et al., 1994). Apparently these three lysine residues are not methylated in other photosynthetic organisms and it is not yet known why this modification occurs in C. reinhardtii. The catalytic and binding properties of the methylated and unmethylated proteins are

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being thoroughly investigated as well as the protein methylation system. It is noteworthy that this type of post-translational modification could only be detected by direct amino-acid sequencing. This indicates that although the technique is more tedious and time consuming than DNA sequencing, it can also bring additional information that could not have been uncovered by studying genes exclusively.

plants (Jacquot et al., 1994), this protein has a subunit molecular weight ca 36 kDa and contains a bound FAD molecule. It was found to cross react with antisera to spinach leaf NTR and was most active with C. reinhardtii thioredoxin h in the DTNB reduction reaction. There is no protein sequence or DNA sequence for NTR yet.

B. Thioredoxins Ill.Thioredoxin Dependent Systems

A. NADPH and Ferredoxin Dependent Thioredoxin Reductases There are two different thioredoxin reduction systems in C. reinhardtii as depicted in Fig. 2. The first one is dependent on photoreduced ferredoxin, and the thioredoxin reduction is catalyzed by ferredoxinthioredoxin reductase (FTR). This enzyme has been purified to homogeneity and shown to be a heterodimer containing an iron-sulfur center. The subunits had molecular masses ofca 10 and 13 kDa (Huppe et al., 1990). Thus, in many respects the algal protein resembles the higher plant FTR (subunit composition, presence of a [4Fe-4S] iron-sulfur center). Furthermore, C. reinhardtii FTR had excellent catalytic activity with its own ferredoxin as a substrate, but could also use spinach ferredoxin, albeit not as efficiently. No protein sequence or DNA sequence is available for the C. reinhardtii protein yet. There is also evidence that NADPH-thioredoxin reductase (NTR) is present in C. reinhardtii cells (Huppe et al., 1991). Like the enzyme from land

1. Isolation and Sequence Analysis As discussed in the introductory part, thioredoxins are low molecular weight heat stable proteins. These two properties were used to isolate two ofthe proteins (namely thioredoxin m and thioredoxin h) from algal extracts and purify them to homogeneity (Decottignies et al., 1990, 1991). As for ferredoxin and FNR, the amino-acid sequence of thioredoxins m and h was determined by direct sequencing of the native polypeptide and of purified peptide fragments. These sequences are shown in Fig. 5 and compared with those of Escherichia coli, spinach thioredoxin m, A. thaliana thioredoxin h, and human lymphocyte thioredoxin, proteins with which they share extensive sequence similarity. All thioredoxins clearly show strong identities at the active site and extensive homologies in the C-terminal part following the active site sequence. On the other hand, the similarities are lower in the N-terminal part (as observed with many other proteins). C. reinhardtii thioredoxin m is more similar to prokaryotic type thioredoxin (i.e. Escherichia coli and spinach thioredoxin m), and thioredoxin h has more homology with A. thaliana

Chapter 26 Ferredoxin and Thioredoxin Systems

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thioredoxin h or with human thioredoxin. cDNA and genomic sequences have in turn been isolated from cDNA and genomic libraries by PCR cloning followed by classical hybridization techniques (Jacquot et al., 1992; Stein et al., 1995). Both proteins are nucleusencoded and the deduced sequences are identical to those found by direct amino-acid sequencing, except for the following changes (E7D, E8D, E53C and D85C for thioredoxin m and an additional A113 in thioredoxin h). These differences could reflect the existence of several genes for thioredoxin m; for thioredoxin h it could result from an error in the sequencing of the protein. Nevertheless, Southern blot analyses suggest that there are very few genes for both thioredoxin m and h in C. reinhardtii (unpublished).

2. Comparative Gene Structure Analysis of the genes coding for thioredoxin m and h indicates that the coding sequences are split by introns. The thioredoxin m (Trx2) gene structure is simple with a single intron. The first exon codes for the transit sequence (34 amino-acids) plus the 77 Nterminus sequence ofthe mature protein. The second exon codes for the last C-terminal 29 amino-acids. Overall the open reading frame for thioredoxin m codes for a preprotein of 140 amino-acids with a molecular mass of 15101 Da. The gene for thioredoxin h (Trx1) is more complex being split in 4 exons. The genomic sequence indicates that thioredoxin h is a protein of 113 amino- acids with a molecular mass of 11817 Da. As for the ferredoxin gene, the thioredoxin h genomic sequence seems to be richer in introns than other plant thioredoxin h genes which lack the equivalent of the second intron (Stein et al., 1995, Sahrawy et al., 1996). In fact differences in the number and positions of introns within C. reinhardtii genes compared to other plant genes are frequent and were for example reported for the RbcS sequences (Goldschmidt-Clermont and Rahire, 1986). All these results confirm that C. reinhardtii has diverged early in the evolution from the branch of vascular plants. A comparative structure of the three genes (ferredoxin, thioredoxin m and thioredoxin h) isolated and sequenced is shown in Fig. 6. All genes have consensus splice sites and introns of the 0 type (Breathnach and Chambon, 1981). In addition, all coding sequences show the reported codon bias documented for other C. reinhardtii genes. Table 2 shows the comparative

frequency of A+T bases in the 5' UTR, the exon coding sequence, the introns and 3' part of the genes. In all three sequences, this frequency is lowest in the translated regions and slightly higher in all other parts of the genes, as in other C. reinhardtii sequences like the cytochrome gene (Hill et al., 1991). Still, the A+T frequency is generally rather low even in the other parts ofthe genes, compared to other organisms

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that use them preferentially to the G+C combination (type Dictyostelium discoideum) (Wetterauer et al., 1988).

3. Deduced Cleavage Sites of the Preproteins The sequence of the thioredoxin m gene has also provided the structure of the transit peptide of this nuclear encoded chloroplastic protein. It contains 34 ammo-acids and has the following sequence:

proteins in Escherichia coli cells (Stein et al., 1995). In addition, the cultures can be grown in the presence of stable isotopes such as either or which facilitates the subsequent NMR analysis of the two proteins. After assignment of the resonances, the secondary structures of thioredoxin m and h have been determined (Lancelin et al., 1993; Mittard et al., 1995). Both thioredoxins were found to have a similar organization with a central sheet surrounded by helices. In both proteins, the secondary structural units were found to be organized as follows:

MALVARRAAVPSARSSARPAFARAAPRRSVVVRA

It consists of only eight different amino-acids with a very high predominance of A+R (56%). Once again, acidic amino acids are absent and aromatics very scarce. Overall, these studies have established the sequence ofthe transit peptides for three stromal, nucleus-encoded proteins, since the N-termini were established for each of the mature polypeptides. When comparing these three transit sequences, no identities appear. Several papers have dealt with the comparison of the transit peptides in plants and also in C. reinhardtii (Franzen et al., 1990). The chloroplastic transit sequences listed in these compilations are not always as firmly established (some are based on sequence comparisons, and Nterminal sequences of the mature proteins were not always determined). It remains however that (i) there is no clear similarity between the transit sequences, (ii) there is no apparent conserved cleavage signal when looking at the primary structures. The only characteristic which seems frequent is an Ala residue at position –1 of the cleavage site. Combining this observation with the existence ofan helix in the Nterminus of the ferredoxin transit sequence (and the prediction that similar structures are bound to be formed in other transit sequences, Franzen et al., 1990), we propose that the common determinant for cleavage of the chloroplastic transit sequences may be the presence ofan Ala residue at a given structural distance ofthe helix which seems to be characteristic of the chloroplastic sequences but different in mitochondrial sequences (Lancelin et al., 1996).

4. NMR Deduced Three-dimensional Structure The cDNA sequences coding for thioredoxin h and m have been introduced by PCR into the expression plasmid pET-3d, and the resulting constructions allow extremely efficient production of the corresponding

The 3-dimensional solution structures of both C. reinhardtii thioredoxin h and m have been solved recently (Mittard et al., 1997; J.-M. Lancelin, L. Guilhaudis, V. Mittard, M. J. Blackledge, D. Marion, M. Stein and J.-P. Jacquot, unpublished). These two structures show a high degree of similarity and are shown in Fig. 7. There is however one major difference: the first helix of thioredoxin h is one turn longer than in thioredoxin m (this is also apparent in the amino acid sequence comparison of Fig. 5, where 3 amino-acids are lacking in the m sequence). Several structures of thioredoxin are now available in the literature. The human lymphocyte and Escherichia coli proteins have been thoroughly described (Eklund et al., 1984; Qin et al., 1994, Weichsel et al., 1996). The algal thioredoxin m has a structure which is similar to the prokaryotic type thioredoxin (i.e. E. coli) and the thioredoxin h is more similar to the human lymphocyte type. Besides the two structures described here, few structures of plant thioredoxins have been solved and only another one published to date (Gleason, 1994, Saarinen et al., 1995, Jacquot et al., 1997).

5. Biochemical Properties and Mutagenesis It was observed early that thioredoxin h and m do not display the same reactivity either with the reducing systems or with the target enzymes. The presumed cytosolic thioredoxin h is preferentially reduced by NADPH and NTR and its reactivity with chloroplast target enzymes is lower than thioredoxin m. On the other hand, the chloroplastic thioredoxin m is preferentially reduced by the light dependent system (ferredoxin+FTR) and activates chloroplast targets such as NADP-malate dehydrogenase more efficiently (Decottignies et al., 1990, 1991; Huppe et al., 1990,

Chapter 26

Ferredoxin and Thioredoxin Systems

1991). These properties were fully confirmed when using the corresponding recombinant proteins obtained by genetic engineering from E. coli cells (Stein et al., 1995). This suggests that surface located determinants are responsible for these specificities. In addition, the recombinant technology has allowed us to obtain large quantities of the two thioredoxins. We could thus determine that despite their similar global folding, the two thioredoxin proteins have very different thermal stabilities. While thioredoxin m is extremely thermostable (much as the E. coli protein), thioredoxin h is much more labile, being irreversibly denatured after 3 min incubation at ca 70°C in its oxidized form. Unusual spectral characteristics were also found for thioredoxin h, possibly linked to the presence of a specific tryptophan helix situated in the Nresidue (W13) in the terminal part (Stein et al., 1995). Site-directed mutants of both thioredoxin m and h have now been generated in order to uncover (i) the molecular characteristics responsible for the differential thermal properties of the two thioredoxin molecules, (ii) the amino-acids

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responsible for the spectral characteristics of thioredoxin h. While we have not yet found out the structural basis of the high stability of thioredoxin m, the mutations W13F (thioredoxin h) and F10W (thioredoxin m) have shown that residue W13 is indeed responsible for these characteristics in thioredoxin h (S. Lemaire and J.-P. Jacquot, unpublished). Other mutations, including active site mutations (C34S in thioredoxin m and W35A and C39S in thioredoxin h) have also been generated in order to unravel the physiological role of the two thioredoxin molecules in algal cells. The proteins carrying active site mutations are apparently redox inactive, at least in the NADP-malate dehydrogenase activation test (J.-P. Jacquot, unpublished).

IV. Conclusion This report summarizes the properties of ferredoxin and thioredoxins of C. reinhardtii. Ferredoxin, FNR, thioredoxin m and h have been purified to homo-

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geneity from algal cells and their primary sequence and biochemical properties determined. This has in turn led to the isolation of the cDNAs and eventually of the genes coding for these proteins. It has been shown that C. reinhardtii cells contain the full set of thioredoxin systems (i.e., a chloroplastic system reduced by ferredoxin and FTR, and a cytosolic system reduced by NADPH and NTR). Several questions, however, deserve to be further studied: (i) although FTR and NTR, the two reducing enzymes, have been isolated and characterized, there is still no structural information on these important enzymes; (ii) the targets of the chloroplast and cytosolic thioredoxins are still largely unknown and this is certainly an exciting area of investigation. In this respect, the recent determination of an NADPmalate dehydrogenase sequence is extremely interesting (O. Ocheretina, H. Tellioglu and R. Scheibe, unpublished, accession # Genbank X98357). There is also a report about the sequence of sedoheptulose-l,7-bisphosphatase, one of the Calvin cycle enzymes regulated by the thioredoxin system in plants (Hahn and Kück, 1994). A sequence of glutamine synthetase, an enzyme reportedly activated by thioredoxin in C. reinhardtii (Florencio et al., 1993; see Chapter 33, Fernandez et al.) has also been published very recently (Chen and Silflow, genbank accession number # U46208). Another potential chloroplast target of thioredoxin m is the coupling factor for which a mutant lacking the nuclear encoded subunit has been isolated in C. reinhardtii (Smart and Selman, 1991). Mutated versions of the subunit have in turn been reintroduced, confirming the regulatory role of a nine amino acid insertion containing two cysteine residues (Smart and Selman, 1993; Ross et al., 1995, 1996; Chapter 25, Strotmann). (iii) Other molecules similar to thioredoxins are bound to be present in C. reinhardtii. This is the case of thioredoxin f which has been characterized but neither purified to homogeneity nor sequenced or expressed. Another type of thioredoxin-like molecule was also reported recently as part of the dynein complex of flagella (Patel-King et al., 1996). This finding is physiologically very significant with respect to the earlier observation that flagellar

movement is redox-controlled (Kooijman, 1988; Beck and Haring, 1996). (iv) Thioredoxins were recently found to be secreted molecules in animal and plant cells (it is a major component of phloem sap in rice) (Wollman et al., 1988; Ishiwatari et al., 1995). Furthermore, it has been shown that it can interact with receptors in higher plants (Bower et al., 1996). These properties suggest that thioredoxins could act in signal transduction pathways. An interesting question in C. reinhardtii is whether such a function is indeed also possible and whether thioredoxins can also be secreted and serve as extracellular messenger between the individual algal cells; (v) It will also be of interest to study the system responsible for the methylation of FNR in C. reinhardtii. Likewise, the reactivity and affinity of the ferredoxin mutants with the PsaC subunit is of high interest (Farah et al., 1995).

Acknowledgment The authors would like to thank Myroslawa MiginiacMaslow for critical reading of the manuscript.

References Aliverti A, Ermanno-Corrado M and Zanetti G (1994) Involvement of lysine-88 of spinach ferredoxin-NADP+ reductase in the interaction with ferredoxin. FEBS Lett 343: 247–250 Arnon DI (1988) The discovery of ferredoxin: The photosynthetic path. Trends Biochem Sci 13: 30–33 Babiychuk E, Kushnir S, Van Montagu M, and Inze D (1994) The Arabidopsis thaliana apurinic endonuclease Arp reduces human transcription factors Fos and Jun. Proc Natl Acad Sci USA 91: 3299–303 Beck CF and Haring MA (1996) Gametic differentiation of Chlamydomonas. Intern Rev Cytol 168: 259–302 Bodenstein-Lang J, Buch A and Follmann H (1989) Animal and plant mitochondria contain specific thioredoxins. FEBS Lett 258: 22–26 Bower MS, Matias DM, Femandes-Carvalho E, Mazzurco M, Gu T, Rothstein SJ and Goring DR (1996) Two Members of the thioredoxin-h family interact with the kinase domain of a Brassica S-locus receptor kinase. Plant Cell 8: 1641–1650 Breathnach R and Chambon P (1981) Organization and expression of eukaryotic split genes coding for proteins. Annu Rev

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Ferredoxin and Thioredoxin Systems

Biochem 50: 349–383 assimilation in Buchanan BB (1991) Regulation of the oxygenic photosynthesis: The ferredoxin/thioredoxin system. Arch Biochem Biophys 288: 1–9 Bringloe DH, Dyer TA and Gray JC (1995) Developmental, circadian and light regulation of wheat ferredoxin gene expression. Plant Mol Biol 27: 293–306 Buchanan BB, Schurmann P and Jacquot J-P (1994) Thioredoxin and metabolic regulation. Seminars in Cell Biology 5: 285– 293 Campbell WH and Gowri G (1990) Codon usage in higher plants, green algae and cyanobacteria. Plant Physiol 92: 1–11 Danon A and Mayfield SP (1994) Light-regulated translation of chloroplast messenger RN As through redox potential. Science 266: 1717–1719 Decottignies P, Schmitter JM, Jacquot J-P, Dutka S, Picaud A and Gadal P (1990) Purification, characterization and complete amino-acid sequence of a thioredoxin from a green alga Chlamydomonas reinhardtii. Arch Biochem Biophys 280: 112–121 Decottignies P, Schmitter JM, Dutka S, Jacquot J-P and MiginiacMaslow M (1991) Characterization and primary structure of a second thioredoxin from the green alga Chlamydomonas reinhardtii. Eur J Biochem 198: 505–512 Decottignies P, Le Maréchal P, Jacquot J-P, Schmitter JM and Gadal P (1995) Primary structure and post-translational modification of ferredoxin-NADP-reductase from Chlamy­ domonas reinhardtii. Arch Biochem Biophys 316: 249–259 de Pascalis AR, Schürmann P. and Bosshard HR (1994) Comparison of the binding sites of plant ferredoxin for two ferredoxin-dependent enzymes. FEBS Lett 337: 217–220 Eklund H, Cambillau C, Sjöberg BM, Holmgren A, Jörnvall H, Hoög JO and Bränden CI (1984) Conformational and functional similarities between glutaredoxin and thioredoxins. EMBO J 3: 1443–1449 Elliott RC, Pedersen TJ, Fristensky B, White MJ, Dickey LF and Thompson WF (1989) Characterization of a single copy gene encoding ferredoxin I from pea Plant Cell 1: 681–690 Farah J, Frank G, Zuber H and Rochaix JD (1995) Cloning and sequencing of a cDN A clone encoding the photosystem I PsaD subunit from Chlamydomonas reinhardtii. Plant Physiol 107: 1485–1486 Florencio FJ, Gadal P, and Buchanan BB (1993) Thioredoxinlinked activation of the chloroplast and cytosolic forms of Chlamydomonas reinhardtii glutamine synthetase. Plant Physiol Biochem 31: 649–655 Franzeii LG, Rochaix JD and von Heijne G (1990) Chloroplast transit peptides from the green alga Chlamydomonas reinhardtii share features with both mitochondrial and higher plant presequences. FEBS Lett 260: 165–168 Gleason FK (1994) Thioredoxins in cyanobacteria: structure and redox regulation of enzyme activity. In: Bryant DA (ed) The Molecular Biology of Cyanobacteria, pp 715–729. Kluwer Academic Publishers, Dordrecht Goldschmidt-Clermont M and Rahire M (1986) Sequence, evolution and differential expression of the two genes encoding variant small subunits of ribulose bisphosphate carboxylase/ oxygenase in Chlamydomonas reinhardtii. J Mol Biol 191: 421–432 Halm D and Kück U (1994) Nucleotide sequence of a cDNA

513 encoding the chloroplast sedoheptulose-l,7-bisphosphatase from Chlamydomonas reinhardtii. Plant Physiol 104: 1101– 1102 Hill KL, Li HH, Singer J and Merchant S (1991) Isolation and structural characterization of the Chlamydomonas reinhardtii J Biol Chem 266: 15060–15067 gene for cytochrome Huppe HC, de Lamotte-Guéry F, Jacquot J-P and Gadal P (1990) The ferredoxin-thioredoxin system of a green alga, Chlamy­ domonas reinhardtii. Planta 180: 341–351 Huppe HC, Picaud A, Buchanan BB and Miginiac-Maslow M (1991) Identification of an NADP/ thioredoxin system in Chlamydomonas reinhardtii. Planta 186: 115–121 Ishiwatari Y, Honda C, Kawashima I, Nakamura S, Hirano H, Mori S, Fujiwara T, Hayashi H and Chino M (1995) Thioredoxin h is one of the major proteins in rice phloem sap. Planta 195: 456–463 Jacquot J-P, Stein M, Hodges M, and Miginiac-Maslow M (1992) PCR cloning of a nucleotidic sequence coding for the mature part of Chlamydomonas reinhardtii thioredoxin. Nucl Ac Res 20: 617 Jacquot J-P, Rivera-Madrid R, Marinho P, Kollarova M, Le Maréchal P, Miginiac-Maslow M and Meyer Y (1994) Arabidopsis thaliana NADPH thioredoxin reductase. J Mol Biol 235: 1357–1363 Jacquot J-P, Lancelin J-M and Meyer Y (1997a) Thioredoxins: Structures and functions in plant cells. New Phytologist 136: 543–570 Jacquot J-P, Stein M, Suzuki A, Liottet S, Sandoz G and MiginiacMaslow M (1997b) Residue Glu 91 of Chlamydomonas reinhardtii ferredoxin is essential for electron transfer to ferredoxin-thioredoxin reductase. FEBS Lett 400: 293–296 Kaneko T, Sato S, Kotani H, Tanaka A, Asamizu E, Nakamura Y, Miyajima N, Hirosawa M, Sugiura M, Sasamoto S, Kimura T, Hosouchi T, Matsuno A, Muraki A, Nakazaki N, Naruo K, Okumura S, Shimpo S, Takeuchi C, Wada T, Watanabe A, Yamada M, Yasuda M and Tabata S (1996) Sequence analysis of the genome of the unicellular Cyanobacteriuin Synechocystis sp. strain PCC6803. II. Sequence determination of the entire genome and assignment of potential protein-coding regions. DNA Res 3: 109–136 Kitayama M, Kitayama K and Togasaki RK (1994) A cDNA clone encoding a ferredoxin-NADP+ reductase from Chlamydomonas reinhardtii. Plant Physiol. 106: 1715–1716 Kooijman R (1988) Signal transduction and the regulation by light of sexual agglutinability in the green alga Chlamydomonas eugametos. Thesis of Doctorate, University of Amsterdam Koradi R, Billeter M and Wüthrich K (1996) MOLMOL: A program for display and analysis of macromolecular structures. J Mol Graphics 14: 51–55 Lancelin J-M, Stein M and Jacquot J-P (1993) Secondary structure and protein folding of recombinant chloroplastic thioredoxin Ch2 from the green alga Chlamydomonas reinhardtii as determined by J Biochem 114: 421–431 Lancelin J-M, Bally I, Arlaud GJ, Blackledge M, Cans P, Stein M and Jacquot J-P (1994) NMR structures of ferredoxin chloroplastic transit peptide from Chlamydomonas reinhardtii promoted by trifluoroethanol in aqueous solution. FEBS Lett 343: 261–266 Lancelin J-M, Gans P, Bouchayer E, Bally I, Arlaud GJ and Jacquot J-P (1996) NMR structures of a mitochondrial transit

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peptide from the green alga Chlamydomonas reinhardtii. FEBS Lett 391: 203–208 Matsubara H and Hase T (1983) Phylogenetic considerations of ferredoxin sequences in plants particularly algae. In: Jensen U and Fairbrother DE (eds) Proteins and Nucleic Acids in Plants, pp 168–181. Springer-Verlag, Berlin Mittard V, Morelle N, Brutscher B, Simorre JP, Marion D, Stein M, Jacquot J-P, Lirsac PN and Lancelin J-M (1995) 1H, 13C, 15N-NMR resonance assignments of oxidized thioredoxin h from the eukaryotic green alga Chlamydomonas reinhardtii using new methods based on two-dimensional triple-resonance NMR spectroscopy and computer-assisted backbone assignment. Eur J Biochem 229: 473–485 Mittard V, Blackledge MJ, Stein M, Jacquot J-P, Marion D and Lancelin J-M (1997) NMR structure of oxidised thioredoxin h from the eukaryotic green alga Chlamydomonas reinhardtii. Eur J Biochem 243: 374–383 Patel-King RS, Benashaki SE, Harrison A, King SM. 1996. Two functional thioredoxins containing vicinal dithiols from the Chlamydomonas reinhardtii outer dynein arm. J Biol Chem 271: 6283–6291 Piubelli L, Aliverti A, Bellintani F and Zanetti G (1996) Mutations of Glu92 in ferredoxin I from spinach leaves produce proteins fully functional in electron transfer but less efficient in supporting NADP+ photoreduction. Eur J Biochem 236: 465– 469 Qin J, Clore CM and Gronenborn AM (1994) The high-resolution three-dimensional solution structures of the oxidized and reduced states of human thioredoxin. Structure 2: 503–522 Rivera-Madrid R, Mestres D, Marinho P, Jacquot J-P, Decottignies P, Miginiac-Maslow M and Meyer Y (1995) Evidence for five divergent thioredoxin h sequences in Arabidopsis thaliana. Proc Natl Acad Sci USA 92: 5620–5624 Rogers WJ, Hodges M, Decottignies P, Schmitter JM, Gadal P and Jacquot J-P (1992) Isolation of a cDNA fragment coding for Chlamydomonas reinhardtii ferredoxin and expression of the recombinant protein in Escherichia coli. FEBS Lett 310: 240–245 Ross SA, Zhang MX and Selman BR (1995) Role of the Chlamydomonas reinhardtii coupling factor 1 gamma-subunit cysteine bridge in the regulation ofATP synthase. J Biol Chem 270: 9813–9318 Ross SA, Zhang MX and Selman BR (1996) A role for the disulfide bond spacer region ofthe Chlamydomonas reinhardtii coupling factor 1 gamma-subunit in redox regulation of ATP synthase. J Bioenerg Biomembr 28: 49–57 Saarinen M, Gleason FK and Eklund H. (1995) Crystal structure of thioredoxin-2 from Anabaena. Structure 3: 1097–1108

Sahrawy M, Hecht V, Lopez-Jaramillo J, Chueca A, Chattier Y and Meyer Y (1996) Intron position as an evolutionary marker of thioredoxins and thioredoxin domains. J Mol Evol 42: 422– 431 Schenk H, Klein M, Erdbrugger W, Droge W, and SchulzeOsthoff K (1994) Distinct effects of thioredoxin and antioxidants on the activation of transcription factors NFkappa B and AP-1. Proc Natl Acad Sci USA 91: 1672–1676 Smart EJ and Selman BR (1991) Isolation and characterization of a Chlamydomonas reinhardtii mutant lacking the gammasubunit of chloroplast coupling factor 1 (CF1). Mol Cell Biol 11: 5053–5058 Smart EJ and Selman BR (1993) Complementation of a Chlamydomonas reinhardtii mutant defective in the nuclear gene encoding the chloroplast coupling factor 1 gammasubunit (atpC). J Bioenerg Biomembr 25: 275–284 Stein M, Jacquot J-P and Miginiac-Maslow M (1993) A cDNA clone encoding Chlamydomonas reinhardtii preferredoxin. Plant Physiol 102: 1349–1350 Stein M, Chedozeau B and Jacquot J-P (1995) Cloning and sequencing of a ferredoxin gene (Genbank U29516) from Chlamydomonas reinhardtii. Plant Physiol 109: 721 Stein M, Jacquot J-P, Jeannette E, Decottignies P, Hodges M, Lancelin J-M, Mittard V, Schmitter JM and Miginiac-Maslow M (1995) Chlamydomonas reinhardtii thioredoxins; structure of the genes coding for the chloroplastic m and cytosolic h isoforms: Expression in Escherichia coli of the recombinant proteins, purification and biochemical properties. Plant Mol Biol 28: 487–503 Studier FW and Moffat BA (1986) Use of bacteriophage T7 RNA polymerase to direct selective high level expression of cloned genes. J Mol Biol 189: 113–130 Vorst O, van Dam F, Oosterhoff-Teertstra R, Smeekens S and Weisbeek P (1990) Tissue-specific expression directed by an Arabidopsis thaliana pre-ferredoxin promoter in transgenic tobacco plants. Plant Mol Biol 14: 491–499 Weichsel A, Gadaska JR, Powis G and Montfort W (1996) Crystal structures of reduced oxidized and mutated human thioredoxins: Evidence for a regulatory homodimer. Structure 4: 735–751 Wetterauer B, Jacquot J-P and Véron M (1992) Thioredoxins from Dictyostelium discoideum are a developmentally regulated multigene family. J Biol Chem 267: 9895–9904 Wollman EE, d’Auriol L, Rimsky L, Shaw A, Jacquot J-P, Wingfield P, Graber P, Dessarps F, Robin P, Galibert F, Bertoglio J and Fradelizi D (1988) Cloning and expression of a cDNA for human thioredoxin. J Biol Chem 263: 15506– 15512

Chapter 27 Genetic Engineering of Rubisco Robert J. Spreitzer Department of Biochemistry, University of Nebraska, Lincoln, NE 68588-0664, U.S.A.

Summary I. Introduction A. Genetic Engineering B. Basic Rubiscology C. Chlamydomonas Rubisco II. Chloroplast Genetic Screening and Selection A. Recovery of Chlamydomonas rbcL mutants B. Complementing Substitutions III. Directed Mutagenesis and Chloroplast Transformation A. Effects on Catalysis B. Interactions With Rubisco Activase IV. Rubisco Nuclear Mutants A. Screening and Selection B. Insertional Mutagenesis and Deletion of the RbcS Genes V. Conclusion and Perspective Acknowledgments References

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Summary Because of its pivotal role in photosynthetic fixation, ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) has become a focus for genetic engineering. An increase in carboxylase activity or a decrease in the inhibition of carboxylase activity would directly improve plant productivity. Much is known about the Rubisco catalytic mechanisms, a number ofX-ray crystal structures have been solved, and directed mutagenesis of prokaryotic enzymes has probed the importance of individual amino-acid residues. However, as in most cases of enzyme engineering, it is difficult to deduce which changes are needed to make Rubisco better. The eukaryotic green alga Chlamydomonas reinhardtii offers other approaches to this problem. Classical genetic methods, which are difficult to apply in prokaryotes or higher plants, have been used to recover mutations at random, identify interactions between amino-acid residues, and discover other genes that influence the ultimate expression of Rubisco function. Several regions outside the active site have now been identified that control catalytic efficiency. Molecular genetic methods have also become well established for Chlamydomonas. Directed mutagenesis and gene transformation can be used to investigate the nucleus- and chloroplast-encoded Rubisco subunits, which likely serve as the best models for ultimately engineering the Rubisco of crop plants.

J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 515–527. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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I. Introduction Ribulose-1,5-bisphosphate (RuBP) carboxylase/ oxygenase (Rubisco) may be the most important enzyme on earth because it enables atmospheric carbon to be captured by life on the planet (for recent reviews see Schneider et al., 1992; Spreitzer, 1993; Hartman and Harpel, 1994). Essentially all of the carbon in a human body (as well as much of the carbon in the clothes upon it) was at one time or another fixed from the atmosphere by Rubisco. Rubisco serves our needs for food and fiber by and RuBP to catalyzing the reaction between form two molecules of phosphoglycerate within the fixation is chloroplasts of plants. However, net considerably reduced under normal atmospheric is mutually conditions due to the fact that competitive with at the same active site. Furthermore, oxygenation of RuBP generates one molecule of phosphoglycerate and one molecule of phosphoglycolate, and this latter product is the first intermediate in a pathway that ultimately leads to the If carboxylation could be increased release of or oxygenation decreased, an increase in net fixation and plant productivity would be realized.

A. Genetic Engineering There has been much interest in genetically engineering Rubisco to increase its carboxylation efficiency (Schneider et al., 1992; Spreitzer, 1993; Hartman and Harpel, 1994; Gutteridge and Gatenby, 1995). Because Rubisco is essential for the survival of crop plants, strategies for genetic manipulation have been limited. Furthermore, the active-site large subunit is coded by a polyploid gene (rbcL) within the chloroplast, and the small subunit is coded by a family of RbcS genes within the nucleus (reviewed by Dean et al., 1989). It is quite difficult to change all of these rbcL or RbcS gene copies to produce a change in the holoenzyme. Several rbcL mutations have been identified in higher plants (Winter and Herrmann, 1988; Avni et al., 1989; Shikanai et al., 1996), but these mutants must be maintained in tissue culture or as chimeras that contain both wildtype and mutant rbcL alleles. It has also been possible Abbreviations: CABP – 2-carboxyarabinitol 1,5-bisphosphate; for for Rubisco – ribulose- 1,5bisphosphate carboxylase/oxygenase; RuBP – ribulose 1,5bisphosphate; for carboxylation; for oxygenation; specificity factor

Robert J. Spreitzer to delete the rbcL gene in tissue culture and rescue such plants by introducing rbcL into the nucleus (Kanevski and Maliga, 1994). However, this system presently depends on a cotransformation strategy that does not permit selection for a functional, or improved, Rubisco. A eukaryotic holoenzyme has not yet been expressed in Escherichia coli (e.g., Gatenby et al., 1987;Cloney et al., 1993), precluding the use of directed mutagenesis for examining plant Rubisco in vitro. Because of these limitations, many scientists have used directed mutagenesis of prokaryotic Rubisco enzymes to investigate the structural basis ofcatalysis (reviewed by Spreitzer, 1993; Hartman and Harpel, 1994). For the most part, these studies rely on the homodimeric large-subunit enzyme of the bacterium Rhodospirillum rubrum or the plant-type enzyme of the cyanobacterium Synechococcus, both of which can be expressed in and isolated from E. coli (reviewed by Hartman and Harpel, 1994; Tabita, 1994). Although substitutions at the active-site residues generally eliminate or greatly reduce carboxylase activity, several have been particularly useful for defining large-subunit interactions (e.g., Larimer et al., 1987), catalytic residues (e.g., Lorimer et al., 1993; Harpel and Hartman, 1996), and reaction intermediates (e.g., Harpel et al., 1995), Much has been learned at a rapid pace about the enzyme active site from these studies (reviewed by Hartman and Harpel, 1994), but it remains difficult to deduce which changes need to be made to improve the enzyme. Because the active-site residues are essential for carboxylation, they may not be the best targets for engineering a better Rubisco. It has apparently been difficult to use other genetic approaches for investigating prokaryotic Rubisco. Although rbcL gene elimination or replacement has been achieved for some prokaryotes (Pierce et al., 1989; Falcone and Tabita, 1991; Amichay et al., 1993; Wang et al., 1993), it is not known whether lethal directed mutations can be identified efficiently by screening transformed prokaryotic cells. A prokaryotic rbcL mutant has not yet been recovered by genetic screening or selection, and prokaryotic directedmutant Rubisco has not yet been expressed in vivo. All of the problems associated with the genetic manipulation of Rubisco in crop plants and prokaryotes have been circumvented by using classical genetic methods and newly-established molecular methods in Chlamydomonas reinhardtii. Chlamydomonas is clearly not of immediate

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Engineering Rubisco

agronomic importance, but one can anticipate that the information gained with this eukaryote could ultimately be applied to the improvement of land plants (Kanevski and Maliga, 1994). At the same time, the type of information being accumulated may prove useful for understanding the structurefunction relationships of a truly fascinating plant enzyme.

B. Basic Rubiscology The Rubisco holoenzyme in green plants and algae has a molecular mass of about 560 kD and is comprised of two different subunits, each present in eight copies. The small subunit is coded by a family of RbcS nuclear genes, synthesized as a 20-kD precursor outside the chloroplast, and processed to 15 kD during transport into the chloroplast (reviewed by Schmidt and Mishkind, 1986; Dean et al., 1989). In most cases, a single rbcL chloroplast gene encodes the 55-kD large subunit, which assembles with small subunits via the action of molecular chaperones (reviewed by Gutteridge and Gatenby, 1995). X-ray crystallography of the bacterial and plant enzymes has revealed that the C-terminal domain of each active site, and has large subunit forms an confirmed earlier studies (Larimer et al., 1987) indicating that residues in the N-terminal domain of an adjacent large subunit contribute to active-site structure (Knight et al., 1990; Curmi et al., 1992; Newman and Gutteridge, 1993; Andersson, 1996). Loops between the alternating strands and helixes of the barrel contain residues that interact with the transition-state analog 2-carboxyarabinitol 1,5bisphosphate(CABP) (Knight et al., 1990; Schreuder et al, 1993). Small subunits are required for maximal carboxylation (Andrews, 1988), but the details of their contribution to Rubisco structure and function are not yet understood (reviewed by Spreitzer, 1993; Tabita, 1994). Rubisco must also be activated via the carbamylation of large-subunit Lys-201 and (Lorimer et al., subsequent coordination with 1976; Lorimer, 1981). This process is facilitated by the nucleus-encoded Rubisco-activase enzyme (reviewed by Portis, 1992, 1995; Salvucci and Ogren, 1996). The ratio of carboxylation to oxygenation at any specified concentrations of and is defined by the Rubisco specificity factor, where V is the of carboxylation or oxygenation, and K is the for

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or respectively (Laing et al., 1974). A proton is abstracted from RuBP to form the 2,3-enediolate of RuBP, which is common to both carboxylation and then compete and oxygenation. Unbound is for this substrate (Pierce et al., 1986), and ultimately determined by the differential stabilization ofthe carboxylation and oxygenation transition states (Chen and Spreitzer, 1991, 1992). The value of has increased during evolution, with photosynthetic bacteria generally having the lowest values and land plants having the highest values (Jordan and Ogren, 1981). Several eukaryotic, nongreen algae have values that are even higher (Read and Tabita, 1992, 1994). These observations indicate that further beneficial specificity may be possible. improvements in However, is only one measure ofcatalytic efficiency. Increases in are not beneficial if they occur at the expense of a significant decrease in or (Laing et al., 1974). In other words, one needs to consider carboxylation rate as well as in the design of a better enzyme (reviewed by Spreitzer, 1993).

C. Chlamydomonas Rubisco Much is known about the molecular biology of Chlamydomonas Rubisco. In fact, transit peptides were first discovered by investigating the translocation of the Chlamydomonas small subunit into the chloroplast (Dobberstein et al., 1977; Schmidt et al., 1979). The rbcL gene has been cloned and sequenced from chloroplast DNA (Dron et al., 1982). This gene has served as a model for studying chloroplast gene promoters and patterns ofexpression in general (e.g., Salvador et al, 1993; Klein et al., 1994; Hwang et al., 1996). The two members of the RbcS gene family have been sequenced (Goldschmidt-Clermont and Rahire, 1986) and studied with regard to their differential expression in light and darkness (Goldschmidt-Clermont, 1986; GoldschmidtClermont and Rahire, 1986). Transient decreases in concentration may also inhibit small-subunit accumulation at the level of translation (Winder et al., 1992). The single nuclear gene for Rubisco activase has been isolated from Chlamydomonas (Roesler and Ogren, 1990). Several chaperonin genes have also been identified in Chlamydomonas that likely play a role in Rubisco assembly (Thompson et al., 1995). In addition to these achievements, it is also important to point out, relative to the present discussion, that the first physically-defined chloro-

518 plast mutation was recovered in Chlamydomonas, and this mutation was found within the rbcL gene (Spreitzer and Mets, 1980; Dron et al., 1983). was Furthermore, the first change in Rubisco found via the analysis of a Chlamydomonas rbcL mutant (Chen et al., 1988).

II. Chloroplast Genetic Screening and Selection Chlamydomonas remains the only organism in which photosynthesis-deficient mutants resulting from mutations in either the nuclear or chloroplast genome can be routinely recovered. Such mutants can be maintained with acetate medium in darkness, a condition under which Chlamydomonas synthesizes a complete photosynthetic apparatus (Spreitzer and Mets, 1981; Spreitzer et al., 1988a). Levine and coworkers demonstrated the utility of nuclear-gene mutants for dissecting photosynthetic electron transport (reviewed by Levine, 1969). Wang and coworkers have used pigment mutants to investigate chlorophyll synthesis (e.g., Ford and Wang, 1980; Huang and Wang, 1986). The discovery that 5fluorodeoxyuridine can reduce the ploidy of the chloroplast genome enabled the successful screening for acetate-requiring chloroplast mutations (Wurtz et al., 1979). The discovery that most photosynthetic mutants are also sensitive to light (they grow better in the dark with acetate than in the light with acetate) (Spreitzer and Mets, 1981, 1982; Spreitzer and Ogren, 1983a) led to an increase in the spectrum of mutant types (Spreitzer and Mets, 1980, 1981; Spreitzer and Ogren, 1983b). These were important discoveries, which now allow the detailed genetic dissection of any chloroplast-encoded photosynthetic component (Spreitzer et al., 1992).

A. Recovery of Chlamydomonas rbcL mutants As shown in Table 1, ten rbcL mutations have been recovered by screening acetate-requiring Chlamydomonas mutants for defective Rubisco (Spreitzer and Mets, 1980; Spreitzer et al., 1988a) or by screening for alleles of physically-defined rbcL mutants (Spreitzer and Ogren, 1983b; Spreitzer et al., 1992). One of the mutants, 68-4PP, was recovered by screening temperature-conditional, acetaterequiring mutants (Spreitzer et al., 1988a; Chen et al., 1988). All of these Rubisco-deficient strains are

Robert J. Spreitzer quite sensitive to light, and must be maintained with acetate in darkness to prevent selection for revertants, suppressors, or suppressors of the associated light sensitivity (Spreitzer et al., 1992). Because the lightsensitivity suppressors can cause secondary defects in Photosystem II activity or pigmentation (Spreitzer and Ogren, 1983a), their presence could confuse the biochemical analysis of the primary mutant defect. Mutant screening differs fromdirected mutagenesis in an important way. Although the mutants are recovered by random screening, they are not distributed randomly. They define only the essential structural and functional regions of the enzyme. We now know that Rubisco carboxylation must be reduced to less than about 15% of the wild-type rate for a mutant cell to express an acetate-requiring phenotype (Spreitzer et al., 1984; Spreitzer and Chastain, 1987). Furthermore, based on observed rbcL mutation frequencies, it is also known that more than 80% ofall rbcL missense mutations fail to reduce carboxylation below the 15% threshold (Spreitzer, 1993). The rbcL missense mutations cause amino-acid substitutions in several discrete parts of the Rubisco large subunit. Most of these regions were previously shown by chemical modification to be important for catalysis (reviewed by Hartman and Harpel, 1994). The most interesting observation from these aminoacid substitutions is that all but one (T173I; Spreitzer et al., 1988b) affect residues that do not directly coordinate with the CABP transition-state analog. Although they do affect residues that are generally conserved in eukaryotic Rubisco enzymes, their effects on Rubisco activity or assembly could not be predicted by simply looking at the Rubisco crystal structures (Knight et al., 1990; Curmi et al., 1992; Newman and Gutteridge, 1993; Andersson, 1996). Furthermore, L290F and V331A were the first aminoacid substitutions found to influence the value of Rubisco (Chen et al., 1988; Chen and Spreitzer, 1989). The Rubisco rbcL mutants have also been useful for mapping chloroplast genes (Mets and Geist, 1983), establishing chloroplast gene transformation (Boynton et al., 1988), defining chloroplast heteroplasinicity relative to chloroplast tRNA suppressors (Spreitzer et al., 1984; Spreitzer and Chastain, 1987; Zhang and Spreitzer, 1990; Yu and Spreitzer, 1992), demonstrating the essentiality of Rubisco holoenzyme for pyrenoid assembly (Rawat et al., 1996), and examining the potential supra-

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Engineering Rubisco

molecular organization of Calvin cycle enzymes (Belknap and Togasaki, 1982). The rbcL nonsense mutants also confirmed earlier studies with proteinsynthesis inhibitors (Schmidt and Mishkind, 1983) which indicated that small subunits are synthesized, transported to the chloroplast, but then degraded in the absence of large subunits (Spreitzer and Ogren, 1983b; Spreitzer et al, 1985a). However, the rbcL mutants have been most useful for allowing the application of additional genetic methods aimed at understanding the structural basis ofRubisco catalytic efficiency.

B. Complementing Substitutions Because the rbcL mutations exist in vivo, and the mutant strains can be kept alive on acetate in darkness, genetic selection can be exploited for recovering additional mutations as intragenic or intergenic suppressors. These second-site mutations are not recovered randomly. They are selected as functional changes that complement a primary defect, and thereby define structural interactions within the enzyme. In practice, photosynthesis-competent revertants are selected that must have an improved Rubisco. Only true revertants have been found for the 106C(G171D),69-12Q(T173I), and 67-2AA(G237S) missense mutants (Spreitzer et al., 1982, 1985b; R.J. Spreitzer, unpublished), indicating the importance of these residues or regions for Rubisco catalysis or stability. Informational suppression accounts for reversion of the nonsense mutants (Spreitzer et al., 1984; Spreitzer and Chastain, 1987; Zhang and Spreitzer, 1990; Yu and Spreitzer, 1992), as well as

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for two revertants of missense mutant 28-7J (Thow et al., 1994). However, detailed analysis of the photosynthesis-competent revertants of the 28-7J (R217S), 31-4E (G54D), 45-3B (V331A), and 684PP (L290F) missense mutants has revealed additional amino-acid substitutions in the large subunit. Pseudorevertants of mutant 31-4E have shown that G54A and G54V substitutions in the N-terminal domain can restore holoenzyme stability, and that the G54V substitution causes a decrease in (Spreitzer et al., 1995) (Table 2). Within the Cdomain, intragenic suppression terminal, of the 28-7J (R217S) mutation restored holoenzyme stability and produced a double-mutant enzyme (Thow et al., (R217S/A242V) with a reduced 1994). These two substitutions occur at residues that are close to the activator Lys-201 residue, and likely influence by affecting the placement of this residue in the active site (Chen and Spreitzer, 1992; Thow et al., 1994). Intragenic suppression of the 68-4PP (L290F) and 45-3B (V331A) mutations arose from of the amino-acid substitutions that increased mutant enzymes (Chen and Spreitzer, original 1989; Chen et al., 1991; Hong and Spreitzer, 1997) (Table 2). As of today, these are the only amino-acid substitutions that increase both and to produce ‘better’ enzymes (Spreitzer, 1993), but these enzymes are not better than the wild-type enzyme (Chen et al., 1991; Hong and Spreitzer, 1997). The L290F/A222T and L290F/V262L substitutions appear to act at the interface between Rubisco large and small subunits (Fig. 1) (Hong and Spreitzer, 1997), indicating that quite distant substitutions can affect active-site catalytic efficiency. The V331A/T342I and V331 A/

520

G344S substitutions complement for size within the loop 6 (Fig. 2) (Chen hydrophobic core of and Spreitzer, 1989; Chen et al, 1991). This loop contains a catalytic lysyl residue (Lorimer et al., 1993), and is now thought to be a flexible flap that closes over substrate in the active site (Knight et al., 1990; Schreuder et al., 1993; Larson et al., 1995). Inspection of the kinetic constants for the mutant

Robert J. Spreitzer

and revertant enzymes reveals that is primarily responsible for the growth phenotype (Table 2). For example, the of R217S/A242V Rubisco is lower than that ofV331A Rubisco, but the R217S/A242V strain can grow photosynthetically presumably due to a higher (Chen and Spreitzer, 1989; Thow et al., usually taken 1994). Substantial decreases in as a measure of catalytic efficiency, are of little

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importance as long as remains at (R217S/A242V) or above (V331A/G344S) 10% of the wild-type value (Table 2). Because Chlamydomonas, like other single-celled phototrophs, contains a mechanism for internally (reviewed by Spalding concentrating et al., 1985, see also chapter in this volume; Badger and Price, 1994), the value of Rubisco appears to have little impact on net fixation. In other words, Chlamydomonas Rubisco (due to its higher would be more efficient than higher-plant enzymes if both could be compared in Chlamydomonas, even though the higher-plant enzymes have higher values (Jordan and Ogren, 1981). The complementing substitutions identified by screening and selection (Table 2) are relatively far from those active-site residues that coordinate with the CABP transition-state analog (Knight et al., 1990; Andersson, 1996). Nonetheless, they define regions and interactions that can influence and catalytic efficiency. Perhaps such regions would be better targets for attempting to engineer an improved Rubisco.

III. Directed Mutagenesis and Chloroplast Transformation In contrast to chloroplast transformation technology in vascular plant systems, it is now a routine matter to transform genes in the chloroplast of Chlamydomonas

521

(reviewed by Boynton and Gillham, 1993). The biolistic approach proved useful for replacing mutant rbcL (Boynton et al., 1988), as well as for insertionally inactivating wild-type rbcL (Newman et al., 1991).

A. Effects on Catalysis The first Chlamydomonas rbcL mutants recovered by directed mutagenesis and chloroplast transformation (N123G and S379A photosynthesisdeficient mutants) were created to test whether aminoacid substitutions in prokaryotic enzymes produced the same biochemical effects when introduced into eukaryotic Rubisco (Chène et al., 1992; Harpel and Hartman, 1992; Lee and McFadden, 1992; Soper et al., 1992; Zhu and Spreitzer, 1994). For example, whereas an S379 A substitution in Rs. rubrum Rubisco increased at the expense of a decrease in (Harpel and Hartman, 1992), the same substitution in the Chlamydomonas enzyme decreased both and (Table 3), and produced a photosynthesis-deficient phenotype (Zhu and Spreitzer, 1994). It is thus apparent, and perhaps not surprising, that there are subtle differences in the structure-function relationships of prokaryotic and eukaryotic Rubisco enzymes. It is also reasonable to assume that these subtle differences account for the differences in catalytic efficiency between the prokaryotic and eukaryotic enzymes, indicating that Chlamydomonas Rubisco may be a better model ultimately for deducing

522 approaches to engineer crop-plant enzymes. Because the primary structure of the Chlamydomonas large subunit is more than 85% identical with the primary structures of higher-plant large subunits, it should be feasible to determine which residue differences account for the differences in catalysis. Once identified, such residues or regions might serve as suitable targets for attempting to improve crop-plant enzymes. Random genetic screening and in vivo genetic selection in Chlamydomonas have already identified a number of largesubunit regions that are essential for catalysis and that play a role in determining (Table 2) (Chen et al., 1988, 1991; Chen and Spreitzer, 1989; Thow et al., 1994; Spreitzer et al., 1995; Hong and Spreitzer, 1997). Directed mutagenesis and chloroplast transformation have been used to ‘correct’ the primary structure of the loop-6 region of Chlamydomonas Rubisco so that it is nearly identical to the loop 6 of higher-plant Rubisco (Zhu and Spreitzer, 1996). Individual substitutions (L326I, V341I, and M349L) had little effect on Rubisco catalysis (Table 3), but the L326I substitution caused a decrease in the amount of holoenzyme in vivo. The purified L326I mutant Rubisco also has a decreased thermostability in vitro. These effects can be partially alleviated by the M349L substitution (Zhu and Spreitzer, 1996), indicating that van der Waals interactions between residues 326 and 349 are important for Rubisco stability in vivo and in vitro (Fig. 2) (Knight et al., 1990; Andersson, 1996). However, the photosynthesis-competent L326I/M349L double-mutant enzyme has a decrease

Robert J. Spreitzer in and (Zhu and Spreitzer, 1996). It is thus apparent that holoenzyme stability and catalytic efficiency can be interconnected, and that there must be other higher-plant Rubisco residues that compensate for the presence of this pair of residues. Alternatively, it is interesting to consider whether an engineered I326L/L349M double substitution in higher-plant Rubisco might yield an improved enzyme.

B. Interactions With Rubisco Activase Directed mutagenesis and Chlamydomonas chloroplast transformation have also been exploited to probe the interaction between the Rubisco large subunit and Rubisco activase. Spinach and Chlamydomonas Rubisco cannot be activated by tobacco activase, and tobacco Rubisco cannot be activated by spinach or Chlamydomonas activase (Wang et al., 1992). Because the large-subunit primary structure of tobacco Rubisco differs from those of spinach and Chlamydomonas Rubisco at a surface location (Portis, 1995), residues at this region were examined by changing Chlamydomonas residues to those found in tobacco (Larson et al., 1997). A K356Q substitution in the large subunit of Chlamydomonas Rubisco had no effect on activation, but a P89R substitution produced a dramatic effect. The P89R enzyme can no longer be activated by spinach activase but can now be activated by tobacco activase (Larson et al., 1997). In other words, the large-subunit site that includes residue 89 interacts with Rubisco activase, and also determines the specificity of the interaction.

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Engineering Rubisco

IV. Rubisco Nuclear Mutants The classical genetic approach to study Rubisco biochemistry has another advantage. Genes that control the ultimate expression ofthe genes of interest (i. e., RbcS and rbcL genes) can also be identified either as mutants that have biochemical phenotypes similar to those of structural-gene mutants or as intergenic suppressors of structural-gene mutations.

A. Screening and Selection Several mutants resulting from nuclear gene mutations that affect specifically Rubisco stability, activity, or expression have been recovered. Most of these mutant strains were isolated by screening temperature-conditional, acetate-requiring Chlamydomonas mutants for defective Rubisco (Spreitzer et al., 1988a, 1992). One was recovered as a suppressor of the temperature-conditional 68-4PP rbcL mutant (Chen et al., 1990). Genetic analysis has shown that none of these mutations resides within either of the two RbcS genes (Chen et al., 1990; Spreitzer et al., 1992; Gotor et al., 1994; R. J. Spreitzer, unpublished). Instead, they all act posttranslationally, causing decreased (or, for the 68-4PP suppressor, increased) Rubisco thermostability in vivo and in vitro (Chen et al., 1990, 1993; Gotor et al., 1994). These nuclear mutations also cause alterations in the kinetic constants of purified Rubisco (Spreitzer et al., 1992), and, in two cases, they alter (Chen et al., 1990; Gotor et al., 1994). The molecular basis underlying the phenotype of the nuclear Rubisco mutants is not yet known. Nonetheless, these genes might also serve as future targets for approaches to improving Rubisco. One other nonconditional, acetate-requiring, nuclear-gene mutant appears to be blocked specifically in transcription ofthe chloroplast rbcL gene (Hong and Spreitzer, 1994). Perhaps further analysis of this mutant will increase our understanding of rbcL gene transcription, and, in so doing, will provide clues for increasing the expression of rbcL or other chloroplast genes.

B. Insertional Mutagenesis and Deletion of the RbcS Genes One can assume that it would be difficult to recover RbcS mutants by genetic screening in Chlamydomonas because there are two RbcS genes (Goldschmidt-Clermont and Rahire, 1986; Chen et

523 al., 1990). If either gene is sufficient for growth, the probability of knocking out both by point mutations is the square of the probability of knocking out either. It has been extremely difficult to ask questions about the role of the eukaryotic small subunit in Rubisco function because plants and green algae have multiple RbcS genes (reviewed by Dean et al., 1989). Without being able to eliminate all of the resident RbcS gene copies, it is difficult to introduce foreign or directed mutant RbcS genes into any plant (reviewed by Spreitzer, 1993). Because nuclear transformation in Chlamydomonas occurs via nonhomologous recombination, it has become possible to use random insertional mutagenesis to tag genes of interest (Adam et al., 1993; Tarn and Lefebvre, 1993). While performing random insertional mutagenesis of Chlamydomonas, Khrebtukova and Spreitzer (1996) recently recovered a mutant strain that lacks both members of the RbcS gene family. This was possible because the two RbcS genes reside at a single ~8-kb locus (GoldschmidtClermont and Rahire, 1986), and insertional mutagenesis in Chlamydomonas is often associated with deletions as large as 23 kb (Tam and Lefebvre, 1993). The RbcS deletion mutant lacks both Rubisco subunits because large-subunit synthesis is blocked at the level of translation in this strain (Khrebtukova and Spreitzer, 1996). Nonetheless, transformation of the mutant with either RbcS gene restores photosynthesis, and the Rubisco holoenzyme accumulates to substantial levels in both light- and dark-grown cells (Khrebtukova and Spreitzer, 1996). The multiple RbcS genes of plants and green algae are often differentially regulated and encode slightly different proteins (e.g., Goldschmidt-Clermont and Rahire, 1986; Meagher et al., 1989; Dedonder et al., 1993; Meier et al., 1995). Whereas the two RbcS genes of Chlamydomonas encode mature proteins that differ at four residues, a chimeric gene, encoding two variant residues from each small subunit, also produced photosynthesis-competent transformants of the RbcS deletion mutant (Khrebtukova and Spreitzer, 1996). Thus, sequence differences between members of this RbcS gene family are also not essential for the formation of a functional Rubisco. It seems possible that multiple RbcS genes are required only for the production of more Rubisco. Because either of the two Chlamydomonas RbcS genes is sufficient for the production of a functional holoenzyme, it will now be possible to employ directed mutagenesis and molecular-genetic methods

524 for addressing questions of eukaryotic small-subunit function in Chlamydomonas. Small subunits are more divergent than large subunits. This raises the possibility that small subunits may account for the and observed species variation in Rubisco carboxylation efficiency (Jordan and Ogren, 1981). Previous studies support this possibility (Schneider et al., 1990; Read and Tabita, 1992).

V. Conclusion and Perspective One may be skeptical or enthusiastic about the prospects for engineering an improved Rubisco, but the fact of the matter is that we still do not know enough about Rubisco to design a better enzyme rationally. With the versatility of Chlamydomonas chloroplast and nuclear genetics, ease of creating directed mutants of the prokaryotic enzymes (reviewed by Hartman and Harpel, 1994; Tabita, 1994), and potential for engineering chloroplast genes in crop plants (Kanevski and Maliga, 1994), the tools are now available for making rapid progress.

Acknowledgments I thank Carolyn M. O’Brien for reviewing the manuscript, and John C. Szot and Yvonne J. O. Arendsen-Wilms for reviewing the manuscript and creating the figures. Financial support from the United States Department of Agriculture (National Research Initiative) and Department of Energy is gratefully acknowledged.

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Robert J. Spreitzer unassembled ribulose-1,5-bisphosphate carboxylase small subunits in chloroplasts. Proc Natl Acad Sci USA 80: 2632– 2636 Schmidt GW and Mishkind ML (1986) The transport of proteins into chloroplasts. Annu Rev Biochem 55: 879–912 Schmidt GW, Devillers-Thiery A, Desruisseaux H, Blobel G and amino acid sequences of Chua NH (1979) precursor and mature forms of the ribulose-1,5-bisphosphate carboxylase small subunit from Chlamydomonas reinhardtii. J Cell Biol 83: 615–622 Schneider G, Knight S, Andersson I, Brändén CI, Lindqvist Y and Lundqvist T (1990) Comparison of the crystal structures Rubisco suggests a functional role for the small of and subunit. EMBO J 9: 2045–2050 Schneider G, Lindqvist Y and Brändén CI (1992) Rubisco: Structure and mechanism. Annu Rev Biophys Biomol Struct 21: 119–143 Schreuder HA, Knight S, Curmi MG, Andersson I, Cascio D, Brändén CI and Eisenberg D (1993) Formation of the active site of ribulose-1,5-bisphosphate carboxylase/oxygenase by a disorder-order transition from the unactivated to the activated form. Proc Natl Acad Sci USA 90: 9968–9972 Shikanai T, Foyer CH, Dulieu H, Parry MAJ and Yokota A (1996) A point mutation in the gene encoding the Rubisco large subunit interferes with holoenzyme assembly. Plant Mol Biol 31: 399–403 Soper TS, Larimer FW, Mural RJ, Lee EH and Hartman FC (1992) Role of asparagine-111 at the active site of ribulose1,5-bisphosphate carboxylase/oxygenase from Rhodospirillum rubrum as explored by site-directed mutagenesis. J Biol Chem 267: 8452–8457 Spalding MH, Spreitzer RJ and Ogren WL (1985) Use ofmutants in the analysis of the pathway of Chlamydomonas reinhardtii. In: Lucas WJ and Berry JA (eds) Inorganic Carbon Uptake by Aquatic Photosynthetic Organisms, pp 361–375. American Society of Plant Physiologists, Rockville Spreitzer RJ (1993) Genetic dissection of Rubisco structure and function. Annu Rev Plant Physiol Plant Mol Biol 44: 411–434 Spreitzer RJ and Chastain CJ (1987) Heteroplasmic suppression of an amber mutation in the Chlamydomonas chloroplast gene that encodes the large subunit of ribulosebisphosphate carboxylase/oxygenase. Curr Genet 11: 611–616 Spreitzer RJ and Mets L (1980) Non-mendelian mutation affecting ribulose-1,5-bisphosphate carboxylase structure and activity. Nature 285: 114–115 Spreitzer RJ and Mets L (1981) Photosynthesis-deficient mutants of Chlamydomonas with associated light-sensitive phenotypes. Plant Physiol 67: 565–569 Spreitzer RJ and Mets L (1982) An assessment of arsenate selection as a method for obtaining nonphotosynthetic mutants of Chlamydomonas. Genetics 100: 417–425 Spreitzer RJ and Ogren WL (1983a) Nuclear suppressors of the photosensitivity associated with defective photosynthesis in Chlamydomonas reinhardtii. Plant Physiol 71: 35–39 Spreitzer RJ and Ogren WL (1983b) Rapid recovery of chloroplast mutations affecting ribulosebisphosphate carboxylase/ oxygenase in Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 80: 6293–6297 Spreitzer RJ, Jordan DB and Ogren WL (1982) Biochemical and genetic analysis of an RuBP carboxylase/oxygenase mutant

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and revertants of Chlamydomonas reinhardtii. FEBS Lett 148: 117–121 Spreitzer RJ, Chastain CJ and Ogren WL (1984) Chloroplast gene suppression of defective ribulosebisphosphate carboxylase/oxygenase in Chlamydomonas reinhardtii: Evidence for stable heteroplasmic genes. Curr Genet 9: 83–89 Spreitzer RJ, Goldschmidt-Clermont M, Rahire M and Rochaix JD (1985a) Nonsense mutations in the Chlamydomonas chloroplast gene that codes for the large subunit of ribulosebisphosphate carboxylase/oxygenase. Proc Natl Acad Sci USA 82: 5460–5464 Spreitzer RJ, Rahire M and Rochaix JD (1985b) True reversion of a mutation in the chloroplast gene encoding the large subunit of ribulosebisphosphate carboxylase/oxygenase in Chlamydomonas. Curr Genet 9: 229–231 Spreitzer RJ, Al-Abed SR and Huether MJ (1988a) Temperaturesensitive photosynthesis-deficient mutants of Chlamydomonas reinhardtii. Plant Physiol 86: 773–777 Spreitzer RJ, Brown T, Chen Z, Zhang D and Al-Abed SR (1988b) Missense mutation in the Chlamydomonas chloroplast gene that encodes the Rubisco large subunit. Plant Physiol 86: 987–989 Spreitzer RJ, Thow G, Zhu G, Chen Z, Gotor C, Zhang D and Hong S (1992) Chloroplast and nuclear mutations that affect Rubisco structure and function in Chlamydomonas reinhardtii. In: Murata N (ed) Research in Photosynthesis, Vol 3, pp 593– 367. Kluwer Academic Publishers, Dordrecht Spreitzer RJ, Thow G and Zhu G (1995) Pseudoreversion substitution at large-subunit residue 54 influences the specificity of chloroplast ribulose-bisphosphate carboxylase/ oxygenase. Plant Physiol 109: 681–686 Tabita FR (1994) The biochemistry and molecular regulation of carbon dioxide metabolism in cyanobacteria. In: Bryant DA (ed) The Molecular Biology of Cyanobacteria, pp 437–467. Kluwer Academic Publishers, Dordrecht Tam LW and Lefebvre PA (1993) Cloning of flagellar genes in Chlamydomonas reinhardtii by DNA insertional mutagenesis. Genetics 135: 375–384 Thompson MD, Paavola CD, Lenvik TR and Gantt JS (1995) Chlamydomonas transcripts encoding three divergent plastid chaperonins are heat-inducible. Plant Mol Biol 27: 1031–1035 Thow G, Zhu G and Spreitzer RJ (1994) Complementing

527 substitutions within loop regions 2 and 3 of the

active site influence the

specificity of chloroplast ribulose-1,5-bisphosphate carboxylase/oxygenase. Biochemistry 33: 5109–5114 Wang X, Modak HV and Tabita FR (1993) Photolithoautotrophic fixation in Rhodobacter sphaeroides growth and control of and Rhodospirillum rubrum in the absence of ribulose bisphosphate carboxylase-oxygenase. J Bacteriol 175: 7109– 7114 Wang ZY, Snyder GW, Esau BD, Portis AR Jr and Ogren WL (1992) Species-dependent variation in the interaction of substrate-bound ribulose-1,5-bisphosphate carboxylase/ oxygenase (Rubisco) and Rubisco activase. Plant Physiol 100: 1858–1862 Winder TL, Anderson JC and Spalding MH (1992)Translational regulation of the large and small subunits of ribulose bisphosphate carboxylase/oxygenase during induction of the mechanism in Chlamydomonas reinhardtii. Plant Physiol 98: 1409–1414 Winter P and Herrmann RG (1988) A five-base-pair-deletion in the gene for the large subunit causes the lesion in the ribulose bisphosphate carboxylase/oxygenase-deficient plastome mutant sigma of Oenothera hookeri. Bot Acta 101: 68–75 Wurtz EA, Sears BB, Rabert DK, Shepherd HS, Gillham NW and Boynton JE (1979) A specific increase in chloroplast gene mutations following growth of Chlamydomonas in 5fluorodeoxyuridine. Mol Gen Genet 170: 235–242 Yu W and Spreitzer RJ (1992) Chloroplast heteroplasmicity is stabilized by an amber-suppressor tryptophan Proc Natl Acad Sci USA 89: 3904–3907 Zhang D and Spreitzer RJ (1990) Evidence for informational suppression within the chloroplast of Chlamydomonas reinhardtii. Curr Genet 17: 49–53 Zhu G and Spreitzer RJ (1994) Directed mutagenesis of chloroplast ribulose-bisphosphate carboxylase/oxygenase: Substitutions at large subunit asparagine 123 and serine 379 decrease specificity. J Biol Chem 269: 3952–3956 Zhu G and Spreitzer RJ (1996) Directed mutagenesis ofchloroplast ribulose-1,5-bisphosphate carboxylase/oxygenase: Loop-6 substitutions complement for structural stability but decrease catalytic efficiency. J Biol Chem 271: 18494–18498

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Chapter 28 Acquisition. Acclimation to Changing Carbon Availability Martin H. Spalding Department of Botany, Iowa State University, Ames, Iowa 50011-1020, U.S.A.

Summary I. Introduction II. Photosynthetic Carbon Assimilation A. Different Physiological States B.

-Concentrating Mechanism 1. Inorganic Carbon Transport 2. Site of

Elevation 3. Role of Carbonic Anhydrases 4. Role of Pyrenoid 5. Role of Thylakoid Lumen Carbonic Anhydrase 6. Model of CCM 7. Energy Supply C. Interaction With Photorespiration III. Induction of the CCM and Related Adaptations to Limiting

A. Gene Expression and Proteins Changes Regulated Changes in Cell Organization B. C. Cell Division Cycle Concentration Signal D. The E. Non-adapting Mutants Acknowledgments References

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Summary Aquatic organisms, including those such as Chlamydomoncas reinhardtii that inhabit the soil water solution, for photosynthetic carbon assimilation. Accordingly, face a variable supply of dissolved inorganic carbon C. reinhardtii has the ability to acclimate to the changing supply through a variety of responses, which mechanism (CCM). The CCM uses active include development of a transport, probably both at the plasmalemma and the chloroplast envelope, to accumulate a high concentration of bicarbonate in the chloroplast stroma. The initial enzyme of photosynthetic carbon assimilation, Rubisco, is located in a stromal structure called the pyrenoid, and one hypothesis suggests that dehydration of the the substrate for Rubisco, occurs only in the pyrenoid. However, an accumulated bicarbonate to supply carbonic anhydrase responsible for dehydration of the stromal bicarbonate pool apparently is located in the thylakoid lumen, so it may be restricted to thylakoid membranes encompassed by the pyrenoid. The high concentration generated at the site of Rubisco both increases the photosynthetic rate and suppresses oxygenation of RuBP, a wasteful side pathway. In addition to the changes demonstrably related to the function such as induction of of the CCM, C. reinhardtii exhibits several other changes upon acclimation to limiting J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 529–547. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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three additional carbonic anhydrases (one periplasmic and two mitochondrial), changes in the intracellular location of mitochondria, up-regulation of photorespiratory enzymes and up-regulation of several other genes of unknown function. Important transient changes in gene regulation also occur during the acclimation period before the CCM becomes functional, including an arrest of the cell division cycle and a decline in Rubisco synthesis. One of the key areas of interest currently under investigation is how the C. reinhardtii cells recognize or concentration and transduce that signal into gene expression changes needed for the change in induction of a functional CCM, as well as all the other changes seen upon acclimation to limiting

I. Introduction Aquatic and soil-borne photosynthetic organisms such as C. reinhardtii can be exposed to rapid and dramatic changes in the supply of dissolved inorganic for photosynthesis on a seasonal, daily or carbon even an hourly basis, largely as a consequence of sediment or soil respiration and the very slow (and in water, relative to air. diffusion of Because of this, C. reinhardtii and other aquatic photosynthetic organisms possess genetic programs that allow them to acclimate rapidly and specifically to changes in supply for photosynthesis. One of the key adaptations of most aquatic photosynthetic organisms is some mechanism to concentrate or when its supply is limited. The most scavenge extensively studied of these mechanisms has been mechanism (CCM) found in the microalgae and cyanobacteria. There has been interest in the algal CCM over about the past 20 years because it represents a fairly simple system capable of suppressing photorespiration and improving the assimilation under ambient efficiency of concentrations. More recent interest in the mechanisms by which aquatic photosynthetic organisms concentrations stems in acclimate to changing part from their potential as models for the mechanisms by which terrestrial plants may respond to the increasing atmospheric concentration. Because of its exceptional genetic characteristics, C. reinhardtii has become one of the primary model organisms, along with certain cyanobacteria, in which acclimation of aquatic photosynthetic organisms to changing concentrations are investigated. Thus much of the research on the microalgal CCM and acclimation of eukaryotic microalgae to limiting Abbreviations: CA – carbonic anhydrase; CCM – mechanism; carbon; concentration at which the velocity of the reaction is at half of its maximum; Rubisco – ribulose-l,5-bisphosphate carboxylase/ oxygenase

concentrations concentrations at or below the equilibrium with air) has been and continues to be carried out with this microalga.

II. Photosynthetic Carbon Assimilation

A. Different Physiological States An abundance of research reviewed by Briggs and Whittingham (1952), suggested that green microalgae could exist in two distinctly different physiological concentration (0.03% states depending on the vs. 1–5%) during growth. The physiological ‘efficiency’ of use in photosynthesis differed markedly in the two states, with cells exposed to high concentrations apparently being less efficient. Cells were found to be induced to change from one physiological state to the other by exposure to the concentration, with induction taking appropriate one to several hours depending on the direction of transfer and the algal strain. These changes of physiological state subsequently were confirmed for the green microalgae C. reinhardtii (Nelson et al., 1969; Graham et al. 1971), Scenedesmus obliquus (Findenegg, 1974), and Chlorella vulgaris (Hogetsu and Miyachi, 1977). The change of physiological state in algae associated with different concentrations during growth was for many years an unexplained curiosity, until Berry et al. (1976) demonstrated that concenC. reinhardtii cells grown at elevated trations (1–5% in air; cells) had photosynthetic characteristics very similar to terrestrial angiosperms with the C3 pathway of photosynthesis. However, when grown with air levels (air-adapted cells) the cells were much more of exhibiting efficient in their assimilation of photosynthetic characteristics similar to those observed in plants using the C4 pathway of photosynthesis. Relative to cells grown in elevated concentrations, air-adapted C. reinhardtii cells

Chapter 28

Acquisition

were found to have a much higher apparent affinity in photosynthesis. Whereas C. reinhardtii for enrichment exhibited a cells grown with of value for photosynthesis similar to the ribulose-1.5-bisphosphate carboxylase/oxygenase (Rubisco), the initial enzyme responsible for assimilation, air-adapted cells exhibited a value 10–100-fold lower than either that ofthe cells or the of Rubisco (Badger and Price, 1992). The air-adapted cells also exhibited compensation concentrations and much lower little or no oxygen inhibition of photosynthesis. Since via the C4 C. reinhardtii does not assimilate pathway ofphotosynthesis (Spalding, 1989), clearly there had to be some other explanation to account for these C4-like photosynthetic characteristics.

B.

Mechanism (CCM)

The high apparent affinity for and apparent inhibition of photosynthesis in airabsence of adapted C. reinhardtii were explained by the action inside the of a mechanism for concentrating cells, as demonstrated by Badger et al. (1980). In this landmark paper, Badger et al. (1980) demonstrated that active accumulation of was correlated with the acquisition of C4-like photosynthetic characteristics in air-adapted C. reinhardtii cells. They demonstrated that air-adapted but not C. reinhardtii cells exhibited an energy-dependent internal accumulation of to a level several-fold higher than that outside the cell. Thus the photosynthetic characteristics of C. reinhardtii were very C4-like, because, as in the C4 carbon assimilation pathway, C. reinhardtii was able to concentrate at the site of Rubisco. However, rather than a

531 metabolic carboxylation/decarboxylation sequence to pump into the Rubisco-containing compartment, C. reinhardtii and other microalgae use what appears to be a simpler system involving active into a Rubiscotransport and accumulation of containing internal compartment of the cell.

1. Inorganic Carbon Transport The requirement for active transport in the CCM was most clearly confirmed by isolation and characterization of a C. reinhardtii mutant (pmp1-1; see Table 1) that apparently lacks any capacity for transport and accumulation (Spalding et al., 1983b). Although the pmp1-1 mutant established the requirement for transport in the acclimation of C. concentrations at or reinhardtii to limiting below equilibrium with air), and several research groups have directed research at understanding transport, our understanding of transport in microalgae remains somewhat unclear nonetheless, both in terms of location and substrate specificity. The use of carbon isotope disequilibrium studies to identify the species taken up by C. reinhardtii has established that the major flux of into the cell across the occurs through direct uptake of plasmalemma via an active process (Marcus et al., 1984; Sültemeyer et al., 1989; Badger et al., 1994; Palmqvist et al., 1994b), although the data cannot transport at the distinguish between active plasmalemma and active transport at the chloroplast envelope following diffusion into the cell. From these and similar studies, there also is good evidence for direct bicarbonate transport across the plasmalemma, but generally at a lower rate than influx (Sültemeyer et al., 1989; Thielmann et al.,

532 1990; Badger et al., 1994; Palmqvist et al., 1994b). Therefore, it appears that both ofthe predominant species present in the aquatic environment can be used by C. reinhardtii cells, although they apparently prefer transport has been Chloroplast envelope demonstrated in intact, isolated chloroplasts from both C. reinhardtii and Dunaliella tertiolecta, but difficulties with obtaining high yields of intact, active chloroplasts apparently have precluded extensive investigation of the species transported (Moroney et al., 1987a; Goyal and Tolbert, 1989a; Sültemeyer et al., 1988). The light-dependent accumulation activity was demonstrated to be restricted to chloroplasts from air-adapted cells, since chloroplasts from or acetate-grown cells did not exhibit accumulating activity. and/or bicarbonate in Therefore, transport of microalgae may occur at the plasmalemma and/or the inner chloroplast envelope, but the mechanism of transport is not understood for uptake of either bicarbonate or across either membrane. There is evidence for the inhibition of uptake by the inhibitor vanadate both in whole cells and chloroplasts, suggesting that activity of a vanadatesensitive is essential for transport across both the plasmalemma and the chloroplast inner envelope (Palmqvist et al., 1988; Goyal and Tolbert, 1989a;Thielmann et al., 1990;Karlsson et al., 1994). The characteristics of the pmp1-1 mutant, which transport, have raised the completely lacks intriguing question as to how a single mutation apparently can eliminate virtually all accumulation in C. reinhardtii, even though there is good evidence demonstrating transport across both the plasmalemma and the chloroplast envelope. The answer to this intriguing question may be provided when the gene affected by the pmp1-1 mutation is cloned. It is reasonable to assume that direct bicarbonate uptake across the plasmalemma would require a transport, if it protein carrier, as would active occurs. However, if enters the cells by diffusion across the plasmalemma in response to active transport from the cytosol into the chloroplast, a plasmalemma protein carrier would not be needed Thus C. reinhardtii probably has at least one for transporters or transport and possibly two complexes in its plasmalemma. However, no plasmalemma transport proteins have been identified, nor have any proteins been identified as potential candidates for this function.

Martin H. Spalding The substrate specificity of chloroplast uptake has not been established, so it is difficult to speculate as to the number and type of carriers to expect in the chloroplast envelope. However, cDNA clones of two genes (Ccp1 and Ccp2; see Table 2) encoding inducible polypeptides closely-related, showing sequence similarity with the mitochondrial carrier protein superfamily have recently been identified (Chen et al., 1997). This superfamily of carrier proteins displays a fairly wide variety of transport substrate specificities and contains plastid and peroxisomal as well as mitochondrial carrier proteins (Chen et al., 1997). It is tempting to speculate that the Ccp proteins might be involved in transport across the chloroplast inner envelope, but much more information is needed before the function of these proteins is known. Even though neither the substrate specificity nor the location of transport have been clearly established by biochemical studies, evidence from mutational analyses do provide some insight into the final steps of the process of active accumulation. Mutants with lesions at the CA1 locus, long presumed to be defective in an internal carbonic anhydrase in (CA), over-accumulate but still are photosynthesis (Spalding et al., 1983a; Moroney et al., 1986a, 1987b; Suzuki and Spalding, 1989), a situation that can be mimicked or phenocopied by treatment with the membrane-permeant CA inhibitor ethoxyzolamide (Spalding et al., 1983a; Moroney et al., 1987b). The internal CA defective in the ca1-1 mutant recently was cloned by complementation (Funke et al., 1997) and found to be identical to a purified and cloned (Cah3 gene; plastid Genbank accession No. U40871) independently by Karlsson et al. (1995). Since the over-accumulated is unavailable to Rubisco, which uses as its substrate, mutants with lesions at the CA1 locus in the form of bicarbonate. must accumulate These results suggest that in wild-type C. reinhardtii bicarbonate is actively accumulated in the chloroplast and that the Cah3 gene product (ctCA1; see Table 1) is required for rapid dehydration of this accumulated at a bicarbonate to supply Rubisco with physiological rate. The apparent plastid location of ctCAl suggests that active accumulation of bicarbonate occurs in the chloroplast, but it is not yet clear whether the accumulation occurs throughout the chloroplast or is restricted to some compartment within the plastid. Since isolated chloroplasts from air-adapted

Chapter 28

Acquisition

C. reinhardtii exhibit active accumulation of it is reasonable to assume that the transported enters and is accumulated in the stroma as bicarbonate, regardless of which species of serves in the cytosol as transport substrate.

2. Site of

Elevation

It is clear that the site of elevation in C. reinhardtii must at least overlap with the site where bicarbonate is actively accumulated and must include both the is located and the site where site where the Rubisco is located. Active bicarbonate accumulation most likely occurs in the chloroplast as the result of active transport across the plastid inner envelope, and Rubisco appears to be exclusively located within the chloroplast pyrenoid. In a preliminary report, Karlsson et al. (1997a,b) have indicated that ctCAl appears to be localized in the thylakoid lumen. Depending on whether ctCA1 is located in the lumen of thylakoid membranes throughout the chloroplast or, as suggested below, restricted to those encomelevation passed by the pyrenoid, the site where

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is focused might be restricted to the pyrenoid or include the whole chloroplast. Further work on specific localization of ctCA1 should provide more insight on this question. It is as yet unclear how restricted the area of elevation is, in part because of the uncertainty of the ctCAl location and in part because it is not clear whether there are any barriers to restrict diffusion of away from the region in which it is produced. the It had previously been suggested that a plastid CA might be restricted to the pyrenoid in order to supply from dehydration of Rubisco effectively with accumulated bicarbonate without unnecessarily by dehydration of bicarbonate in the releasing stroma (Badger and Price, 1992). As part of this hypothesis, Badger and Price also suggested that the pyrenoid starch sheath might act as a diffusion barrier diffusion out of the pyrenoid. As to restrict discussed below, a thylakoid lumen location for ctCA1 would not necessarily preclude a pyrenoid-specific localization, since thylakoid membranes traverse the pyrenoid, but recent evidence appears to eliminate the pyrenoid starch sheath as playing any essential

534 role in the CCM (Plumed et al., 1996; Villarejo et al., 1996a). Since the chloroplast envelopes probably do not represent a significant diffusion barrier to the existence of any barrier to restrict diffusion of the away from its source, whether that be elevated the pyrenoid or the whole chloroplast, is still in question.

3. Role of Carbonic Anhydrases Even before the CCM was discovered, CA activity was implicated in the physiological state changes of concenmicroalgae associated with shifts in tration (Graham et al., 1971; Findenegg, 1976; Reed and Graham, 1977; Hogetsu and Miyachi, 1979). The search for CA and investigation of its role in microalgal carbon assimilation has continued ever since. In recent years it has become clear that three apparently independent evolutionary lines of CA can be found in various organisms (Hewett-Emmett and Tashian, 1996), and at least two CAs, have been found in of these lines, the C. reinhardtii (Fujiwara et al., 1990; Fukuzawa et al, 1990; Eriksson et al. 1996, Funke et al. 1997). The first CA identified in C. reinhardtii was the abundant periplasmic CA induced by limiting concentrations (Coleman and Grossman, 1984; Coleman et al., 1984), which has been studied rather extensively since its initial discovery. There are two periplasmic CAs in C. reinhardtii, represented by the Cah1 and Cah2 genes (Fujiwara et al., 1990; Rawat and Moroney, 1991; see Table 2). The products of these two genes, pCAl and pCA2, respectively, are and exhibit a very high degree of both sequence similarity, but the expression levels and expression patterns of the two are very different. The Cah2 gene is expressed predominantly in cells grown concentrations but, even under those in elevated conditions it is expressed at very low levels. The major periplasmic CA, pCA1, of C. reinhardtii, encoded by the Cah1 gene, is not expressed in elevated concentrations, but is expressed to very high conditions. levels under limiting The Cah1 gene product, pCA1, is the best known and most extensively studied putative component of the CCM, but it still is rather unclear whether it plays any essential role in the C. reinhardtii CCM. Using ostensibly membrane-impermeant CA inhibitors, Moroney et al. (1985) demonstrated that inhibition of pCA1 decreased the photosynthetic rate of C. reinhardtii cells significantly at limiting

Martin H. Spalding concentrations, but only at alkaline pH. These results suggest that the primary role of pCA1 is to dehydrate for active uptake, since bicarbonate to supply is the preferred species for uptake by by C. reinhardtii. If the active uptake of C. reinhardtii represents diffusion into the cell followed by active transport into the chloroplast (see above), rapid bicarbonate dehydration at the cell surface might be especially important to maximize concentration gradient across the plasmathe lemma, since it is this gradient that determines the rate of diffusion into the cell. On the other hand, in spite of the strong correlation between its activity and activity ofthe CCM, Williams and Turpin (1987) and Sültemeyer et al. (1990) have called the role of pCA1 into question. Although Williams and Turpin also observed decreased photosynthetic rates at alkaline pH using CA inhibitors, wall-less cells washed free of pCA1 activity, as judged by very sensitive mass spectrometric techniques, showed no such inhibition. Williams and Turpin concluded that the ostensibly impermeable CA inhibitors did penetrate the cells and inhibit internal CA isozymes. However, the CA inhibitors used by Moroney et al. (1985) included a dextran-bound sulfonamide far too large to cross membranes intact. Furthermore, Moroney et al. (1985) did not observe any inhibition of photosynthesis under acidic or neutral pH, conditions under which membrane permeable CA inhibitors certainly do inhibit photosynthesis (Spalding et al., 1983; Moroney et al., 1985). Thus the question of whether pCA1 plays any substantial role in the function of the CCM has not been clearly resolved and probably will not be resolved until a Cah1 mutant is identified and characterized. Inasmuch as the normal environment of C. reinhardtii is the soil (Harris, 1989), it is possible that a major role of pCA1 is facilitation of the dissolution of into the water layers surrounding soil particles, a function that also would be needed more under supply. conditions of limiting Intracellular CA isozymes have long been implicated in function of the C. reinhardtii CCM, and various internal CA activities have been reported over the past several years (Badger and Price, 1994). However, only very recently has significant progress been made in unambiguously identifying any internal CA. So far, at least one chloroplast-localized CA has been explicitly identified, the thylakoid lumen (ctCA1, encoded by the Cah3 gene) defective in mutants with lesions at the CA1 locus. Based on

Chapter 28

Acquisition

analyses of the several ca-1 alleles, it seems clear that, as discussed above, ctCA1 plays a role in the to Rubisco through dehyrapid provision of dration of the actively accumulated bicarbonate. In cell fractionation studies, ctCA1 apparently behaves as an insoluble CA, fractionating with the chloroplast membranes (Karlsson et al., 1995), but both soluble and insoluble (membrane associated) chloroplast CA activities have been reported for C. reinhardtii (Sültemeyer et al., 1990, 1995; Amoroso et al., 1996). It is possible that the soluble chloroplast CA might represent unsedimented ctCA1 (Funke et al., 1997), although the two forms are reported to have differential sensitivity to effectors and inhibitors (Amoroso et al., 1996). Also difficult to explain is the residual insoluble chloroplast CA activity reported by Sültemeyer et al. (1995) in the ca1-1 mutant, which should contain no ctCA1. A preliminary report of a purified chloroplast CA different from the ctCA1, a large CA associated with the membrane fraction of C. reinhardtii chloroplasts (Carlson, 1995), may represent the residual insoluble chloroplast CA reported in the ca1-1 mutant by Sültemeyer et al. (1995). Regardless of how many are eventually identified, it is not clear what role(s) additional chloroplast CA(s) might play in the CCM or in the general carbon assimilation of C. reinhardtii. In addition to the chloroplast CAs discussed above, there also is evidence for internal CA activity not localized in the plastid (Sültemeyer et al., 1990, 1995; Amoroso et al., 1996). It is possible that this activity corresponds to the two mitochon(mtCA1 and mtCA2, encoded by the drial Mca1 and Mca2 genes, respectively; see Table 2) recently identified (Eriksson et al., 1996), since both the CA activity reported by Amoroso et al. (1996) and the expression of Mca1 and Mca2 (Eriksson et al., 1996) are up-regulated by growth in limiting The Mca1 and Mca2 genes were identified as cDNA clones for an unidentified, 22 kD, limitinginduced mitochondrial protein (Eriksson et al., 1996), so the characterization of these genes as encoding is derived from their sequence The sequence similarity similarity to other between Mca1 and Mca2 is very high, and the deduced amino acid sequences of the mature proteins (mtCA1 and mtCA2) are identical. It is clear that mtCA1 and mtCA2 are identical to the 21 kD induced polypeptide (LIP-21; see Table 2) described by Geraghty et al. (1996), since the N-terminal amino acid sequences are identical

535

(A. Geraghty and M. Spalding, unpublished). It has been suggested that mtCA1 and mtCA2 might function in pH regulation (via hydration of photorespiratory to prevent alkalization of the mitochondrial matrix during rapid production of photorespiratory ammonia from the glycine decarboxylase complex and to facilitate diffusion of from the same source out of the mitochondrial matrix under high photorespiratory conditions (Eriksson et al., 1996). The roles suggested by Eriksson et al. assume that mtCA1 and mtCA2 are located in the matrix, but the sub-organellar location of these proteins has not been established. Furthermore, it is not entirely clear how hydration of photorespiratory to bicarbonate inside the mitochondria would facilitate its diffusion out of the mitochondria. In fact, it could be argued that, since the glycine which diffuses decarboxylase complex produces readily through membranes, rather than bicarbonate, which does not, the presence of CA in the mitochondrial matrix might be more likely to trap photorespiratory in the mitochondrion rather than to facilitate its diffusion out into the cytosol. On the other hand, if the CA activity was located in the mitochondrial intermembrane space it might be expected to function in preventing loss of the from the cell by trapping it as photorespiratory bicarbonate, which could freely diffuse through the mitochondrial outer membrane into the cytosol to be available for transport into the chloroplast. It will be interesting to learn more about the location of mtCA1 and mtCA2. If these suggested roles are authentic, one would expect to find CA activity in plant mitochondria as well, since photorespiratory fluxes in plants are at least as high ammonia and as those in C. reinhardtii under air-adapted conditions. It has been proposed that there might be a cytosolic CA in C. reinhardtii, although the solid evidence for one may have disappeared with the demonstration of the mtCAs. If cytosolic CA activity does exist in C. reinhardtii, it could play an important role in providing the appropriate substrate for the chloroplast-envelope transporter. It might also play an important role in trapping, as bicarbonate, any leaking from the chloroplast as well as that produced by the mitochondrion in photorespiration and TCA cycle activity. Interestingly, all these roles might also be played by a mitochondrial CA, if it were localized in the intermembrane space rather than the matrix, especially since the mitochondria in air-adapted C. reinhardtii are located in abundance at the cell

Martin H. Spalding

536 periphery between the chloroplast envelope and the plasmalemma (Geraghty and Spalding, 1996).

4. Role of Pyrenoid As the location of most, if not all, of the Rubisco and Rubisco activase in C. reinhardtii (Lacoste-Royal and Gibbs, 1987;Kuchitsu et al., 1988b, 1991; McKay and Gibbs, 1989, 1991; McKay et al., 1991)., the pyrenoid obviously plays some role in the carbon acquisition system of these algae. It is not clear how great that role is, and it is similarly unclear whether the pyrenoid is in any way essential to the operation of the CCM, other than as the site of Rubisco localization. It is intriguing to note that at least one alga that lacks a pyrenoid has also been found to lack a functional CCM (Palmqvist et al., 1994a). It has been suggested that the pyrenoid of C. reinhardtii might play a role in the CCM similar to the role apparently played by the carboxysome in cyanobacteria (Badger and Price, 1992, 1994). In cyanobacteria, Rubisco and CA are co-localized in the carboxysome of the cell, resulting in an elevated concentration only in this compartment (Price and Badger, 1989a). Rubisco is localized in the pyrenoid of C. reinhardtii, and some evidence has pointed to a co-localization of CA in the pyrenoid as well (Kuchitsu, 1991). However, recent preliminary evidence indicating a thylakoid lumen location for ctCA1 calls that into question (Karlsson et al., 1997a,b). Correlations between changes in the pyrenoid starch sheath and changes in concentration in C. reinhardtii and other microalgae have lent support to a potential role for the pyrenoid in the microalgal CCM (Kuchitsu et al., 1988a, 1991; Ramazanov et al., 1994; Geraghty and Spalding, 1996). In keeping release from with the hypothesis that the site of actively accumulated bicarbonate might be restricted to the pyrenoid, it has been suggested that the starch sheath might serve as a diffusion barrier to minimize from the pyrenoid and thus from the site loss of of Rubisco activity (Badger and Price, 1992, 1994). However, recent work with mutants of both C. reinhardtii and Chlorella pyrenoidosa lacking pyrenoid starch sheaths showed that the affinity of in photosynthesis was not the mutants for significantly different from that of wild-type cells (Plumed et al., 1996; Villarejo et al., 1996a). Thus any role as a diffusion barrier played by the pyrenoid starch sheath must be minimal at best.

5. Role of Thylakoid Lumen Carbonic Anhydrase It has long been suggested that, in microalgae, the acidic thylakoid lumen might participate in net dehydration of bicarbonate to supply to Rubisco at concentrations higher than the equilibrium in the (presumably) alkaline concentration of pyrenoid (Pronina and Borodin, 1993; Raven, 1997). The requirements for effective participation of such a system are that the thylakoid membrane transport (or be permeable to) bicarbonate and that CA activity be present in the thylakoid lumen but not in the compartment containing Rubisco. Raven (1997) has suggested such a system as a possible explanation for photosynthetic organisms which appear physiologically to have a CCM but which cannot be However, the demonstrated to accumulate preliminary report that ctCA1 is located in the thylakoid lumen in C. reinhardtii (Karlsson et al., 1997a,b) raises the possibility that such a system also might operate as part of a CCM. As noted by Raven (1997), bicarbonate uptake in the thylakoid lumen could and dehydration to uncouple photophosphorylation. However, the flux needed for bicarbonate dehydration to supply assimilation needs would require consumption only influx. of about 1/12 of the total photosynthetic This could, of course occur throughout the thylakoid influx lumen, consuming a small part of the uniformly, or it could be localized to a small part of the thylakoid lumen where a large proportion of the local influx would be consumed. Since Rubisco is localized to the pyrenoid in C. reinhardtii, it is tempting to speculate that ctCA1 is restricted to those thylakoids that traverse the pyrenoid and that the high bicarbonate concentration in the stroma has access to this same pyrenoidal thylakoid lumen, thus release into the pyrenoid Rubisco allowing for pool at a concentration even higher than that expected at equilibrium in the alkaline stroma/pyrenoid.

6. Model of CCM A recent model for the CCM in C. reinhardtii (Badger and Price, 1992, 1994), based in part on the model of the cyanobacterial CCM (Badger and Price, 1992), proposed that specific co-localization of Rubisco occur only in and CA and localized elevation of the pyrenoid of the chloroplast. As mentioned above, this model predicted that bicarbonate accumulated in the stroma and that specific co-localization of CA

Chapter 28

Acquisition

and Rubisco in pyrenoids allowed rapid dehydration only in the pyrenoid, where the of bicarbonate to could be utilized rapidly by Rubisco. This model further suggested that the starch sheath surrounding the pyrenoid might act as a diffusion barrier to the released in the pyrenoid. It seems loss of reasonably clear at this point that the pyrenoid starch sheath does not function in the CCM in any essential manner (Plumed et al., 1996;Villarejo et al., 1996a), but the other key components of this model, specific accumulation of bicarbonate in the stroma and exclusive co-localization of Rubisco and CA in the pyrenoid, are still viable, with some modification. As discussed above, the evidence for specific accumulation of bicarbonate in the chloroplast stroma is as strong as ever, based on the evidence that chloroplasts and that mutational loss or actively accumulate inhibition of chloroplast CA results in overaccumulation of which is unavailable to Rubisco. The idea of strict co-localization of Rubisco and CA in the pyrenoid has been dealt a setback by the preliminary report (Karlsson et al., 1997a,b) that ctCA1, which is defective in mutants of the CA1 locus, is localized in the thylakoid lumen. However,

537 such a localization would not preclude ctCA1 from being localized exclusively in the thylakoids encompassed by the pyrenoid. Thus effective colocalization of Rubisco and CA in the pyrenoid is still a possibility. Putting all the available information together, it is feasible to construct a new working model of the CCM in C. reinhardtii, as illustrated in Fig. 1. The salient features of the proposed model are that enters the cell either by carrier as bicarbonate or by that is transported into the diffusion as chloroplast, accumulating in the stroma as bicarbonate regardless of the form transported, and that the accumulated bicarbonate enters the lumen of the pyrenoidal thylakoids where it is dehydrated by ctCA1 to Rubisco in the pyrenoid. to provide substrate This model preserves the Rubisco/CA colocalization from the model of Badger and Price (1992, 1994), with some modification to account for the probable thylakoid lumen location of ctCA1. In doing so, it also incorporates some ofthe elements of the Pronina and Borodin (1993) thylakoid lumen CA model. Unlike the starch sheath diffusion barrier suggested by Badger and Price, this model does not

538 incorporate any physical barrier to diffusion of the released out of the pyrenoid. It is possible that the very high concentration ofRubisco in the pyrenoid may be sufficient in itself to consume most or all of before it has an opportunity to the released escape the pyrenoid. It also should be recognized that this model side-steps the question of which species serves as substrate for transport both across the plasmalemma and into the chloroplast. As indicated above, bicarbonate transport across the plasmalemma almost certainly occurs, but the transport is more evidence for plasmalemma equivocal. Since certainly can enter the cells by diffusion, that route of entry also is indicated. Even less is known about the species transported across the chloroplast inner envelope, but the critical aspect of this transport appears to be that bicarbonate arrives and accumulates in the chloroplast stroma rather than which species leaves the cytosol. However, the model appears to be consistent with most, if not all, of the currently available information about the function of the C. reinhardtii CCM.

7. Energy Supply It is clear that the CCM requires an additional input of energy above that required for operation of the Calvin cycle, and it is clear from many early experiments that photosynthesis is the source of the additional energy (Badger and Price, 1992). It is not clear how or in what form the additional energy is provided by photosynthesis, but it is reasonable to assume that a substantial part of the additional energy is required as ATP to drive active transport of However, considering the essential role played by ctCA1, its probable location in the thylakoid lumen and its suggested function in the CCM (Fig. 1), it seems apparent that at least some energy associated might be used with generation of the thylakoid to enhance the dehydration of bicarbonate in the thylakoid lumen. C. reinhardtii to Acclimation of limiting results in changes in the photochemical properties of the cells (Spalding et al., 1984; Sundblad et al., 1986; Palmqvist et al., 1990), indicating that the cells must adjust to the different energy demands of operating the CCM. Short term changes in the photochemical properties probably reflect the availability prior to transient stress of decreased development of CCM activity, and long term photochemical changes may reflect an increased ratio

Martin H. Spalding of PSI to PS II to provide extra ATP to the CCM (Palmqvist et al., 1990). Based on the apparent lack of light saturation for accumulation in the ca1-1 mutant, it appears that energy demands of the CCM might be substantial (Spalding, 1990). However, the details ofboth energy demands and energy supply for the CCM are yet to be fully established.

C. Interaction With Photorespiration The photorespiratory carbon oxidation cycle of C. reinhardtii is different from the classical plant photorespiratory carbon oxidation cycle, because these microalgae have no peroxisomes (Spalding, 1989). Therefore, instead of oxidizing glycolate to glyoxylate via glycolate oxidase in the peroxisome, C. reinhardtii and many other microalgae have been thought to oxidize glycolate to glyoxylate via a glycolate dehydrogenase in the mitochondrion, with other steps in the pathway that occur in the plant peroxisome thought also to be displaced to the mitochondrion (Fig. 2A; Spalding, 1989). The glycolate dehydrogenase has only been identified as an enzymatic activity in crude extracts, although cytochemical techniques have been used to localize it to the mitochondrion (Beezley et al., 1976), and much of the rest of the pathway has little or no evidence supporting its intracellular location. An alternative to the mitochondrial location of glycolate oxidation recently has been suggested by Goyal and Tolbert (1996), who observed that rapid oxidation of glycolate to glyoxylate can be demonstrated in C. reinhardtii and Dunaliella tertiolecta chloroplasts. They further reported that this glycolate oxidizing activity was mediated by a quinone and was associated with photosynthetic electron transport between Photosystem II and the cytochrome b/f complex. Based on this work, and making some assumptions about other parts of the photorespiratory pathway, an alternative photorespiratory pathway can be hypothesized for C. reinhardtii (Fig. 2B). Regardless of the path it takes to get there, glycine generated in the photorespiratory pathway of C. reinhardtii still is expected to undergo an oxidative decarboxylation in the mitochondrion to yield (from (Fig. 2). Thus under 2 glycines) serine, and normal conditions photorespiratory should be released, at least internally, if there is photorespiratory pathway flux in C. reinhardtii.

Chapter 28

Acquisition

Under high photorespiratory conditions, such as when C. reinhardtii cells are exposed concentrations, the cells simply to limiting excrete the glycolate produced rather than metabolizing it further through the downstream portion (past glycolate) of the photorespiratory pathway. One reason for this apparent waste of carbon is that C. reinhardtii may lack sufficient downstream photorespiratory pathway capacity to handle such a large flux of glycolate. Induction of the CCM should suppress most of the photorespiratory flux within a few hours, leaving only a short transition period of high photorespiratory flux. It may not be cost effective for the cells to have more downstream capacity than needed to handle the low flux found in elevated During acclimation of C. reinhardtii to limiting some photorespiratory enzymes also increase in activity, apparently via increased gene expression (Marek and Spalding, 1991; see Table 2). Thus the combination of decreased photorespiratory glycolate production and increased downstream photorespiratory pathway capacity appears to be sufficient to prevent glycolate excretion in fully air-adapted cells. The C4-like photosynthetic characteristics of airadapted C. reinhardtii provide ample evidence that photorespiratory activity is greatly suppressed by the CCM, but there also is abundant evidence that the photorespiratory flux is not completely suppressed. Estimates of photorespiratory pathway flux in airadapted C. reinhardtii, although lower than in cells lacking CCM activity, were found to be rather high

539

(Moroney et al., 1986b). Also, the increased activity of key photorespiratory enzymes suggests increased photorespiratory pathway flux in air adapted cells (Nelson and Tolbert, 1969; Marek and Spalding, 1991; Ramazanov and Cardenas, 1992, 1994). A C. reinhardtii mutant, pgp1-1, with an apparent deficiency in the photorespiratory enzyme phosphoglycolate phosphatase (Suzuki et al., 1990; see Table 1) has helped clarify the relationship between the CCM and photorespiration (Suzuki et al., 1990; Marek and Spalding, 1991). The very fact that this photorespiratory mutant requires elevated for rapid photoautotrophic growth demonstrates that the CCM does not completely suppress the oxygenase activity of Rubisco and photorespiration.

III. Induction of the CCM and Related Adaptations to Limiting

A. Gene Expression and Proteins Changes Relative to other adaptations, the CCM has received the most attention because of its effect, via increased intracellular concentration, on photosynthetic and photorespiratory characteristics (Spalding, 1989). However, C. reinhardtii exhibits a whole suite of adaptive responses to limiting including, as discussed below, induction (or derepression) of the CCM, transient down-regulation of Rubisco synthesis, dramatic changes in sub-cellular organization, and perturbation of the cell division

540 cycle. Coinciding with these functional changes, major changes in gene expression associated with acclimation to limiting occur, including both up-regulation and down-regulation of specific genes (Table 2). Along with induction of a functional CCM and other changes mentioned above, acclimation to also results in the appearance or limiting increased abundance of specific proteins, resulting from changes in expression of the corresponding genes. The induced proteins include the major periplasmic CA (pCA1), two membrane-associated proteins of 36 kD (Ccp1 and Ccp2) and 21 kD (mtCA1 and mtCA2) and two soluble polypeptides of 45–50 kD (Coleman and Grossman, 1984; Bailly and Coleman, 1988; Manuel and Moroney, 1988; Spalding and Jeffrey, 1989; Geraghty et al., 1990). The first identified of the induced proteins was a 37 kD polypeptide identified as a periplasmic CA (Coleman and Grossman, 1984). This periplasmic CA (pCA1) subsequently was found to be composed of two subunits, one of approximately 37 kD and another of 4 kD (Kamo et al., 1990; Ishida et al., 1993). Both subunits are encoded by the Cah1 gene (Fujiwara et al., 1990; Fukuzawa et al., 1990; Ishida et al., 1993; Roberts and Spalding, 1995), which is conditions. expressed only in limiting The 21 kD and 36 kD induced polypeptides also have been the subject of additional investigations (Geraghty et al., 1990; Mason et al., 1990; Ramazanov et al, 1993; Geraghty and Spalding, 1996), including cloning of the corresponding genes and identification of possible functions (Eriksson et al., 1996; Chen et al., 1997). The induced 21 kD polypeptide is the product of two genes (Mca1 and Mca2), which encode (mtCA1 and mtCA2) with two mitochondrial identical amino acid sequences (Eriksson et al., 1996). The mtCAs were identified by sequence analysis as and localized to mitochondria both by cell fractionation (Eriksson et al., 1996) and by immunocytochemistry (Geraghty and Spalding, 1996). The induced 36 kD polypeptide has been localized by cell fractionation to the chloroplast envelope (Ramazanov et al., 1993), and it also is encoded by two very similar genes (Ccp1 and Ccp2), which exhibit strong sequence similarity to the mitochondrial carrier protein superfamily (Chen et al., 1997). This carrier protein superfamily includes carriers from other organelles, including plastids (Chen et al., 1997). Expression of Cah1, Mca1, Mca2, Ccp1 and Ccp2

Martin H. Spalding all appear to be closely correlated during acclimation of C. reinhardtii to limiting with mRNA for Cah1, Ccp1 and Ccp2 all appearing within 1 h and protein within 2 h (Bailly and Coleman, 1988; Dionisio-Sese et al., 1990; Fujiwara et al., 1990; Geraghty et al., 1990; Spalding et al., 1991; Chen et al., 1997). The mRNA from these three genes also disappears quickly, with Cah1 mRNA and Ccp1 and Ccp2 translatable message both undetectable 1 h after exposure of air-adapted cells to elevated (Bailly and Coleman, 1988; Fujiwara et al., 1990; Spalding et al., 1991). Although a time course of Mca1 and Mca2 mRNA abundance has not been reported, the induction time course for the mtCAs is similar to the other three (Geraghty and Spalding, 1996), suggesting their mRNA abundance also will follow a time course similar to those of Cah1, Ccp1 and Ccp2. Since expression of the Cah1, Mca1, Mca2, Ccp1 and Ccp2 genes all are regulated at the level of mRNA abundance (Fujiwara et al., 1990; Fukuzawa et al., 1990; Eriksson et al., 1996; Chen et al., 1997) and their mRNA and protein levels appear to be closely correlated, it is possible that their expression may be coordinately regulated by a single mechanism, e.g., a single trans-acting factor, either an inducer or a represser. However, it should be noted that, although they all are controlled at the level of mRNA abundance, there is no published evidence demonstrating whether the regulation is at the transcriptional or post-transcriptional level for any of these five genes. Several other inducible genes have been identified by Burow et al. (1995) through differential screening of a cDNA library. They inducible clones, reported six different, one of which corresponded to the Cah1 gene. The sequence of another of the cDNA clones, (Lci1), was reported in the paper, but could not be identified based on homology to any known genes. The deduced amino acid sequence of Lci1 did appear to have four segments predicted to form transmembrane spanning helices, suggesting it might encode a transmembrane protein. Another of the cDNA clones (Att1) was subsequently identified as encoding alanine: aminotransferase, but it is not clear what role this aminotransferase might play in (Chen acclimation of C. reinhardtii to limiting et al., 1996). Along with the genes induced (or derepressed) in Table 2 also indicates some genes that limiting exhibit decreased expression in limiting

Chapter 28

Acquisition

including stable down-regulation of the minor periplasmic CA (Cah2; Fujiwara et al., 1990) and transient decrease in the synthesis of both subunits of Rubisco (the RbcS1, RbcS2 and rbcL gene products; Coleman and Grossman, 1984; Winder et al., 1992). Interestingly, although Cah2 expression also is controlled at the level ofmRNA abundance, it appears to be regulated in a manner inverse to the other five genes discussed above (Fujiwara et al., 1990). Both the Cah2 mRNA and gene product (pCA2) are more than in air-adapted C. abundant in reinhardtii cells (Fujiwara et al., 1990; Rawat and Moroney, 1991). In contrast to all the gene expression changes so far discussed, the transient decrease in biosynthesis of both Rubisco subunits during is controlled at the acclimation to limiting translational level (Winder et al., 1992). Other changes associated with acclimation of C. reinhardtii to limiting probably also reflect changes in gene expression. Besides the up-regulation of two soluble proteins of 45–50 kD mentioned earlier, substantial increases occur in the activity of photorespiratory enzymes, including a transient increase in phosphoglycolate phosphatase and more long-term increases in glycolate dehydrogenase and glutamine synthetase (Nelson and Tolbert, 1969; Marek and Spalding, 1991; Ramazanov and Cardenas, 1992, 1994). In these cases only enzymatic activity has been shown to increase, but evidence indicates that the increases are unlikely to have resulted from changes in effector concentrations, and so probably reflect changes in protein abundance resulting from changes in gene expression (Marek and Spalding, 1991).

B.

Changes in Cell Organization

Accompanying changes occurring at a molecular also involves level, acclimation to limiting substantial structural changes in C. reinhardtii cells. One well characterized change is an increased development of the starch sheath that surrounds the chloroplast pyrenoid (Kuchitsu et al., 1988a; Kuchitsu et al., 1991; Ramazanov et al., 1994; Geraghty and Spalding, 1996). Along with changes in pyrenoid starch, abundance of stromal starch also has been reported to decrease in air-adapted microalgae (Miyachi et al., 1986; Kuchitsu et al., 1988a), but recent work indicates that this decrease in stromal starch occurs only transiently and probably results from the depletion of starch reserves in

541 cells during the transition period before the CCM becomes functional (Geraghty and Spalding, 1996). Mitochondrial distribution also changes dramatically during acclimation of C. reinhardtii to limiting with mitochondria moving from a central position, within the cup of the chloroplast, to a peripheral position, between the chloroplast envelope and the plasmalemma (Geraghty and Spalding, 1996). Although the mechanism controlling this migration is unknown, it must involve changes in cytoskeleton organization, and the mitochondria probably pass through ‘gaps’ observed in the cup-shaped chloroplast. This mitochondrial migration is quite intriguing, especially since the induced mtCAs are localized specifically to the peripheral mitochondria in air-adapted cells (Geraghty and Spalding, 1996), and Eriksson et al. (1996) have reported two other induced polypeptides in C. rein­ hardtii mitochondria. The mitochondrial relocation and the induction of specific mitochondrial proteins argue that the mitochondria may play an important role in acclimation of C. reinhardtii to limiting but any discussion of possible function(s) for the mitochondria specifically in air-adapted C. rein­ hardtii, including those discussed above for the mtCAs, is highly speculative. Nonetheless, Geraghty and Spalding (1996) suggested that, because of their move to a peripheral location in air-adapted cells and the increased glycolate pathway flux in air-adapted cells (see above), the mitochondria might be involved in scavenging glycine or glycolate produced by the chloroplasts (see Fig. 2). Another suggestion was that the mitochondria might be involved in transport, so energization of plasmalemma relocation would place them in closer proximity to any plasmalemma transporters. The cytochrome oxidase-dependent respiratory pathway could be involved directly in ATP synthesis, or the alternative pathway could be involved indirectly to decrease excess reductant resulting from the export of ATP from the chloroplast via a PGA-triose phosphate shuttle. In support of a possible role for the alternative pathway, Goyal and Tolbert (1989b, 1990) have reported SHAM inhibition of accumulation and a higher level of alternative oxidase activity in airadapted C. reinhardtii.

C. Cell Division Cycle There may be some interaction between the cell

Martin H. Spalding

542 division cycle and the acclimation of C. reinhardtii availability. Several years ago, to changes in Marcus et al. (1986) demonstrated that the activity of the CCM, as well as the activity of pCA1 changed rhythmically in synchronously grown cultures of C. reinhardtii. Recently, those observations have been confirmed, at least for pCA1, by demonstration of a circadian rhythm in the expression of Cah1 (Rawat and Moroney, 1995; Fujiwara et al., 1996). A preliminary report also indicates that the cell division cycle of C. reinhardtii is transiently arrested during acclimation ofthe cells to limiting (Dillard and Spalding, 1997), which could have important implications for research on acclimation to changes concentration and induction of the CCM. in Because much of the research in these areas uses non-synchronous C. reinhardtii cultures and relatively short acclimation times (24 h or less), it is important to recognize that most of the cells in the culture may accumulate in G1 phase behind a cell division cycle blocks-point, thus at least temporarily becoming functionally synchronous and erasing the averaging effect of the non-synchronous culture with respect to, e.g., gene expression that follows a cell-cycle or circadian rhythm.

D. The

Concentration Signal

The signal regulating acclimation of C. reinhardtii to concentration is unknown (Coleman, changes in 1991). While acclimation is known to be under environmental control, it is not understood how the concentration. cells sense a change in the external Clearly, they must sense the change either directly as concentration or by an indirect effect on a cellular process, such as carbohydrate metabolism. Although investigations of the response of C. rein­ hardtii to changes in availability have focused on acclimation to limiting acclimation works in both directions, i.e., C. reinhardtii also acclimates to supply by changes in cell organization increased and repression of the CCM activity and of induced genes. The response of C. reinhardtii to changes in availability typically has been studied using expression of Cah1 as a reporter by monitoring either protein (pCA1) or mRNA abundance. Several studies have implicated a requirement for photosynthetic activity for induction (or derepression) of concentrations, since periplasmic CA by low light apparently was required for induction (Spalding

and Ogren, 1982; Spencer et al., 1983; Dionisio et al., 1989a,b; Dionisio-Sese et al., 1990), DCMU blocked induction in the light (Spencer et al. 1983; Dionisio et al. 1989a; Dionisio-Sese et al., 1990), and induction did not occur in non-photosynthetic mutants (Spalding and Ogren, 1982; Spencer et al., 1983, Villarejo et al., 1996b). Because, in addition to the apparent requirement for photosynthesis, induction of periplasmic CA was increased by increasing tension and was prevented by inhibitors of the photorespiratory pathway, it has been suggested deficiency might be signaled by a that a photorespiratory metabolite (Spalding and Ogren, 1982; Ramazanov and Cardenas, 1992; Villarejo et al., 1996b). However, this rather simplistic model for an induction signal fails to take into account reports indicating a rather more complicated signaling system. These include reports of Cah1 or CCM activity induction (or derepression) by low concentrations in the dark (Bailey and Coleman, 1988; Fett and Coleman, 1994; Rawat and Moroney, 1995; Villarejo, 1996b), enhanced Cah1 or CCM activity induction by non-photosynthetic blue light (Dionisio et al., 1989a,b; Dionisio-Sese et al., 1990; Borodin et al., 1994) and partial repression (or decreased induction) of Cah1 or CCM activity by mixotrophic growth with acetate as a carbon source (Spalding and Ogren, 1982; Moroney et al., 1987a; Coleman et al., 1991; Fett and Coleman, 1994; Ramazanov et al. 1994). Clearly the acclimation of C. reinhardtii to limiting involves a signaling system more complex than a change in the concentration of a photorespiratory metabolite, but, at present, it is difficult to incorporate all the available information on Cah1 and CCM induction into a cogent hypothesis for the signaling mechanism. Part of the difficulty in trying to understand the signal or signals involved in induction or derepression of the acclimation is that conflicting information has been reported from different research groups. For example, there are three reports of the concentrations in the induction Cah1 by low dark (Bailey and Coleman, 1988; Rawat and Moroney, 1995; Villarejo, 1996b), but there also are seven reports indicating the lack of Cah1 induction by low concentrations in the dark (Spalding and Ogren, 1982; Spencer et al., 1983; Dionisio et al., 1989a,b; Dionisio-Sese et al., 1990; Fujiwara et al., 1990; Fukuzawa et al., 1990). In these various reports, authors have used either pCA1 activity or protein abundance, or Cah1 mRNA abundance to indicate

Chapter 28

Acquisition

Cah1 induction, but there is no discernible relationship between the method of detection used and whether Cah1 induction was observed. Further, Villarejo et al. (1996b), who reported dark induction of Cah1 (as pCA1 protein abundance), also reported the lack of Mca1, Mca1, Ccp1 and Ccp2 induction (protein abundance) in the dark. Rawat and Moroney (1995), who reported dark induction of periplasmic CA in synchronously-grown C. reinhardtii, suggested that use of synchronous cultures might be important to see the response in the dark. Cah1 mRNA abundance oscillates in synchronous cultures, being abundant in the light and disappearing in the dark, but reappearing before the end of the dark period (Rawat and Moroney, 1995; Fujiwara et al., 1996). However, since the message evidently does not appear late in the dark period but does appear at when cells are grown in high at the that point if the cells are switched to low beginning of the dark period (Rawat and Moroney, 1995), the circadian rhythm or cell-cycle control of Cah1 transcription cannot explain how the cells concentration oftheir surroundings in detect the the darkphase. Furthermore, both Bailly and Coleman (1988) and Villarejo et al. (1996b) reported Cah1 induction in non-synchronous cultures in the dark. signal Not only is the nature of the unclear, it is not even clear that there is a limiting signal. The true signal may occur, not in limiting repressing the limitingbut in elevated acclimation. One might expect the situation in levels in the natural environment normally to be equilibrium with the air, but in the natural, soil concentration environment of C. reinhardtii, the generally is fairly elevated (Keeney et al., 1985; Fernandez et al., 1993). The possibility ofa repression is appealing, since signal generated in elevated it is somewhat analogous to other systems for metabolic regulation of gene expression, such as sugar regulation of photosynthesis-related genes in plants (Sheen, 1990, 1994) and glucose repression of alternative catabolic pathways in yeast (Gancedo, 1992). In fact, it may be useful to think of of gene expression in C. reinhardtii as analogous to the glucose repression system in Saccharomyces cerevisiae. In that system, the presence of a preferred carbon source, glucose, represses the transcription of genes for enzymes that catabolize less-desirable, alternate carbon sources such as sucrose or galactose (Gancedo, 1992). By analogy then, in C. reinhardtii, the presence of a rich

543 or acetate, may repress carbon source, either high the expression of genes necessary for growth in limiting concentrations. However, a repression is difficult to reconcile with signal in elevated the apparent photosynthesis requirement for full unless both an induction acclimation to limiting signal and lack of a repression signal are required for adaptations, as is the expression of the case in S. cerevisiae for expression of galactose catabolism genes (Gancedo, 1992). It also is difficult to incorporate the apparent blue light enhancement of Cah1 induction into a global, single-signal hypothesis for either induction or repression. If a blue-light receptor is somehow involved, it may be on a parallel or interacting signal pathway.

E. Non-adapting Mutants An important advance in understanding gene regulation mediated in C. reinhardtii by changes in concentration was made with the identification and characterization of a mutant, cia5, which apparently does not acclimate to limiting (Moroney et al., 1989; see Table 1). The cia5 mutant lacks induction of transport, induction of Cah1, Mca1 and Mca2, Ccp1 and Ccp2, Lci1, or any of the induced polypeptides, upunidentified regulation of phosphoglycolate phosphatase and glycolate dehydrogenase, and down-regulation of Rubisco biosynthesis (Moroney et al., 1989; Spalding et al., 1991; Marek and Spalding, 1991;Burow et al., 1996). The lack of any of these responses in cia5 argues that one signal transduction pathway regulates a very diverse set of adaptations, including transcriptional (pre-translational) up-regulation of Lci1, Cah1, Mca1, Mca2, Ccp1 and Ccp2, transcriptional (pre-translational) down-regulation of Cah2, transient, translational down-regulation of rbcL (chloroplast), RbcS1 (nuclear) and RbcS2 (nuclear) expression, long term up-regulation of glycolate dehydrogenase and transient up-regulation of phosphoglycolate phosphatase. Since cia5 appears to lack any response to limiting this mutant probably represents a defect in a component of the signal transduction pathway that is required either for induction or for derepression of all of the observed adaptations. By analogy with the limiting S. cerevisiae glucose repression system, cia5 would represent a derepression mutant, since, in the absence or acetate), the cia5 of sufficient carbon (either mutant lacks derepression of the genes necessary for

544 growth in limiting The cia5 mutant serves as a prototype for identification of other mutants with defects in the signal transduction pathway for acclimation of C. reinhardtii to changes in concentration. Based on the appearance of preliminary reports from three research groups describing these types of mutants in C. reinhardtii (Fukuzawa et al., 1997; Van and Spalding, 1997; Villand et al., 1997), it appears that this may be a fertile area to pursue. Acknowledgments The author wishes to thank Drs. Mats Eriksson, James Moroney and Gören Samuelsson for their cooperation in renaming some genes and gene products to conform with guidelines of the Plant Genome Nomenclature Commission. Thanks is also extended to the USDA-NRICGP for support of the author’s research in this area. References Amoroso G, Weber C, Sültemeyer DF and Fock H (1996) Intracellular carbonic anhydrase activities in Dunaliella tertiolecta (Butcher) and Chlamydomonas reinhardtii (Dangeard) in relation to inorganic carbon concentration during growth: Further evidence for the existence of two distinct carbonic anhydrases associated with the chloroplasts. Planta 199: 177–184 concentrating Badger MR and Price GD (1992) The mechanism in cyanobacteria and microalgae. Physiol Plant 84: 606–615 Badger MR and Price GD (1994) The role of carbonic anhydrase in photosynthesis. Ann Rev Plant Physiol Plant Mol Biol 45: 369–392 Badger MR, Kaplan A and Berry JA (1980) Internal inorganic carbon pool of Chlamydomonas reinhardtii. Evidence for a concentrating mechanism. Plant Physiol 66: 407–413 Badger MR, Palmqvist K and Yu J-W (1994) Measurement of fluxes in cyanobacteria and microalgae during steady-state photosynthesis. Physiol Plant 90: 529–536 concentration on Bailly J and Coleman JR (1988) Effect of protein biosynthesis and carbonic anhydrase expression in Chlamydomonas reinhardtii. Plant Physiol 87: 833–840 Beezley BB, Gruber PJ and Frederick SE (1976) Cytochemical localization of glycolate dehydrogenase in mitochondria of Chlamydomonas. Plant Physiol 58: 315–319 Berry JA, Boynton J, Kaplan A and Badger MR (1976) Growth and photosynthesis of Chlamydomonas reinhardtii as a function of concentration. Carnegie Inst Wash Yrbk 75: 423–432 Borodin V, Garderström P and Samuelsson G (1994) The effect of light quality on the induction of efficient photosynthesis

Martin H. Spalding under low conditions in Chlamydomonas reinhardtii and Chlorella pyrenoidosa. Physiol Plant 92: 254–260 Briggs GE and Whittingham CP (1952) Factors affecting the rate of photosynthesis of Chlorella at low concentrations of carbon dioxide and in high illumination. New Phytol 51: 236–249 Burow MD, Chen Z-Y, Mouton, TM and Moroney JV (1996) Isolation of cDNA clones induced upon transfer of Plant Mol Biol Chlamydomonas reinhardtii cells to low 31: 443–148 Carlson S (1995) Purification and characterization of the chloroplastic carbonic anhydrase from Chlamydomonas reinhardtii. PhD Thesis. Indiana University, Bloomington Chen Q and Silflow CD (1996) Isolation and characterization of glutamine synthetase genes in Chlamydomonas reinhardtii. Plant Physiol 112: 987–996 Chen Z-Y, Burow MD, Mason CB and Moroney JV (1996) A gene encoding an alanine: aminotransferase in Chlamydomonas reinhardtii. Plant Physiol 112: 677–684 Chen Z-Y, Lavigne LL, Mason CB and Moroney JV (1997) Cloning and overexpression of two cDNAs encoding the lowchloroplast envelope protein LIP-36 from Chlamydomonas reinhardtii. Plant Physiol 114: 265–273 Coleman JR (1991) The molecular and biochemical analyses of concentrating mechanisms in cyanobacteria and microalgae. Plant Cell Environ 14: 861–867 Coleman JR and Grossman AR (1984) Biosynthesis of carbonic anhydrase in Chlamydomonas reinhardtii during adaptation to Proc Natl Acad Sci USA 81: 6049–6053 low Coleman JR, Berry JA, Togasaki RK and Grossman AR (1984) Identification of extracellular carbonic anhydrase of Chlamydomonas reinhardtii. Plant Physiol 76: 472–477 Coleman JR, Luinenburg I, Majeau N and Provart N (1991) Sequence analysis and regulation of expression of a gene coding for carbonic anhydrase in Chlamydomonas reinhardtii. Can J Bot 69: 1097–1102 Dillard S and Spalding MH (1997) Transient arrest at the G1/S transition of the Chlamydomonas cell division cycle during adaptation to limiting Plant Physiol 114: 110 (supplement) Dionisio ML, Tsuzuki M and Miyachi S (1989a) Light requirement for carbonic anhydrase induction in Chlamydomonas reinhardtii. Plant Cell Physiol 30: 207–213 Dionisio ML, Tsuzuki M and Miyachi S (1989b) Blue light induction of carbonic anhydrase activity in Chlamydomonas reinhardtii. Plant Cell Physiol 30: 215–219 Dionisio-Sese ML, Fukuzawa H and Miyachi S (1990) Lightinduced carbonic anhydrase expression in Chlamydomonas reinhardtii. Plant Physiol 94: 1103–1110 Eriksson M, Karlsson J, Ramazanov Z, Garderström P and Samuelsson G (1996) Discovery of an algal mitochondrial carbonic anhydrase: Molecular cloning and characterization of a polypeptide in Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 93: 12031–12034 Fernandez, IJ, Son Y, Kraske CR, Rustad LE, David MB (1983) Soil carbon dioxide characteristics under different forest types and after harvest. Soil Sci Soc Am J 57: 1115–1121 Fett JP and Coleman JR (1994) Regulation of periplasmic carbonic anhydrase expression in Chlamydomonas reinhardtii by acetate and pH. Plant Physiol 106: 103–108 Findenegg GR (1974) Beziehungen zwischen Carboanhy-

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draseaktivität and Aufnahme von and bei der Photosynthese von Scenedesmus obliquus. Planta 116: 123– 131 Findenegg GR (1976) Correlations between accessibility of carbonic anhydrase for external substrate and regulation of by Scenedesmus obliquus. and photosynthetic use of Z Pflanzenphysiol 79: 428–437 Fujiwara S, Fukuzawa H, Tachiki A and Miyachi S (1990) Structure and differential expression of two genes encoding carbonic anhydrase in Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 87: 9779–9783 Fujiwara S, Ishida N and Tsuzuki M( 1996) Circadian expression of the carbonic anhydrase gene, Cah1, in Chlamydomonas reinhardtii. Plant Mol Biol 32: 745–749 Fukuzawa H, Fujiwara S, Yamamoto Y, Dionisio-Sese ML and Miyachi S (1990) cDN A cloning, sequence, and expression of carbonic anhydrase in Chlamydomonas reinhardtii: Regulation by environmental concentration. Proc Natl Acad Sci USA 87: 4383–4387 Fukuzawa H, Ishizaki K, Matsueda S, Miura K, Inoue T and requiring mutants Ohyama K (1997) Isolation of from Chlamydomonas reinhardtii by gene tagging. Plant Physiol 114: 214 (supplement) Funke RP, Kovar JL and Weeks DP (1997) Intracellular carbonic anhydrase is essential to photosynthesis in Chlamydomonas reinhardtii at atmospheric levels of Plant Physiol 114: 237–244 Gancedo JM (1992) Carbon catabolite repression in yeast. Eur J Biochem 206: 297–313 Geraghty AM and Spalding MH (1996) Molecular and structural A possible changes in Chlamydomonas under limiting mitochondrial role in adaptation. Plant Physiol 111: 1339– 1347 Geraghty AM, Anderson JC and Spalding MH (1990) A 36 induced polypeptide of Chlamy­ kilodalton domonas is distinct from the 37 kilodalton periplasmic carbonic anhydrase. Plant Physiol 93: 116–121 Goldschmidt-Clermont M and Rahire M (1986) Sequence, evolution and differential expression ofthe two genes encoding variant small subunits of ribulose bisphosphate carboxylase/ oxygenase in Chlamydomonas reinhardtii. J Mol Biol 191: 421–432 Goyal A and Tolbert NE (1989a) Uptake of inorganic carbon by isolated chloroplasts from air-adapted Dunaliella. Plant Physiol 89: 1264–1269 Goyal A and Tolbert NE (1989b) Variations in the alternative Plant oxidase in Chlamydomonas grown in air or high Physiol 89: 958–962 Goyal A and Tolbert NE (1990) Salicylhydroxamic acid (SHAM) inhibition of the dissolved inorganic carbon concentrating process in unicellular algae. Plant Physiol 92: 630–636 Graham D, Atkins CA, Reed ML, Patterson BD and Smillie RM (1971) Carbonic anhydrase, photosynthesis, and light-induced pH changes. In: Hatch MD, Osmond CB and Slatyer O (eds) Photosynthesis and Photorespiration, pp 267–274. Wiley Interscience, New York Harris EH (1989) The Chlamydomonas Sourcebook. A Comprehensive Guide to Biology and Laboratory Use. Academic Press, Inc, San Diego Hewett-Emmett D and Tashian RE (1996) Functional diversity,

545 conservation, and convergence in the evolution of the anhydrase gene families. Mol Phylogenet Evol 5: 50–77 Hogetsu D and Miyachi S (1977) Effects of concentration fixation in during growth on subsequent photosynthetic Chlorella. Plant Cell Physiol 18: 347–352 Hogetsu D and Miyachi S (1979) Role of carbonic anhydrase in fixation in Chlorella. Plant Cell Physiol photosynthetic 20: 747–756 Ishida S, Muto S and Miyachi S (1993) Structural analysis of periplasmic carbonic anhydrase 1 of Chlamydomonas reinhardtii. Eur J Biochem 214: 9–16 Kamo T, Shimogawara K, Fukuzawa H, Muto S and Miyachi S (1990) Subunit constitution of carbonic anhydrase from Chlamydomonas reinhardtii. Eur J Biochem 192: 557–562 Karlsson J, Ramazanov Z, Hiltonen T, Garderström P and Samuelsson G (1994) Effect of vanadate on photosynthesis Chlamydomonas and the ATP/ADP ratio in reinhardtii cells. Planta 192: 46–51 Karlsson J, Hiltonen T, Husic HD, Ramazanov Z and Samuelsson G (1995) Intracellular carbonic anhydrase of Chlamydomonas reinhardtii. Plant Physiol 109: 533–539 Karlsson J, Clarke AK, Mason CB, Chen Z-Y, Hugghins SY, Husic HJ, Samuelsson G and Moroney JV (1997a) A novel alpha-type carbonic anhydrase associated with the thylakoid membrane in Chlamydomonas reinhardtii is required for growth in ambient air. Plant Physiol 114: 215 (supplement) Karlsson J, Spear E, Mason CB, Moroney JV, Samuelsson G and Husic HD (1997b) Carbonic anhydrase is associated with the thylakoid membranes in Chlamydomonas reinhardtii. Plant Physiol 114: 215 (supplement) Keeney DR, Sahrawat KL, Adams SS (1985) Carbon dioxide concentration in soil: Effects on nitrification, denitrification and associated nitrous oxide production. Soil Biol Biochem 17: 571–573 Kuchitsu K, Tsuzuki M and Miyachi S (1988a) Changes in starch localization within the chloroplast induced by changes in concentration during growth of Chlamydomonas reinhardtii: Independent regulation of pyrenoid starch and stromal starch. Plant Cell Physiol 29: 1269–1278 Kuchitsu K, Tsuzuki M and Miyachi S (1988b) Characterization of the pyrenoid isolated from unicellular green alga Chlamydomonas reinhardtii: Particulate form of RuBisCO protein. Protoplasma 144: 17–24 Kuchitsu K, Tsuzuki M and Miyachi S (1991) Polypeptide composition and enzyme activities of the pyrenoid and its concentration in unicellular green algae. regulation by Can J Bot 69: 1062–1069 Lacoste-Royal G and Gibbs SP (1987) Immunocytochemical localization of ribulose-1,5-bisphosphate carboxylase in the pyrenoid and thylakoid region of the chloroplast of Chlamydomonas reinhardtii. Plant Physiol 83: 602–606 Manuel LJ and Moroney JV (1988) Inorganic carbon accumulation by Chlamydomonas reinhardtii. New proteins are made during adaptation to low Plant Physiol 88: 491–496 Marcus Y, Volokita M and Kaplan A (1984) The location of the transporting system for inorganic carbon and the nature of the form translocated in Chlamydomonas reinhardtii. J Exp Bot 35: 1136–1144 Marek LF and Spalding MH (1991) Changes in photorespiratory

546 enzyme activity in response to limiting in Chlamydomonas reinhardtii. Plant Physiol 97: 420–425 Mason CB, Manuel LJ and Moroney JV (1990) A new chloroplast protein is induced by growth on low in Chlamydomonas reinhardtii. Plant Physiol 93: 833–836 McKay RML and Gibbs SP (1989) Immunocytochemical localization of ribulose 1,5-bisphosphate carboxylase/ oxygenase in light-limited and light-saturated cells of Chlorella pyrenoidosa. Protoplasma 149: 31–37 McKay RML and Gibbs SP (1991) Composition and function of pyrenoids: cytochemical and immunocytochemical approaches. Can J Bot 69: 1040–1052 McKay RML, Gibbs SP and Vaughn KC (1991) RuBisCo activase is present in the pyrenoid of green algae. Protoplasma 162: 38–45 Miyachi S, Tsuzuki M, Maruyama I, Gantar M and Miyachi S concentration during growth on the (1986) Effects of i n t r a c e l l u l a r structure of Chlorella and Scenedesmus (Chlorophyta). J Phycol 22: 313–319 Moroney JV, Husic HD and Tolbert NE (1985) Effects of carbonic anhydrase inhibitors on inorganic carbon accumulation by Chlamydomonas reinhardtii. Plant Physiol 79: 177–183 Moroney JV, Tolbert NE and Sears BB (1986a) Complementation analysis of the inorganic carbon concentrating mechanism of Chlamydomonas reinhardtii. Mol Gen Genet 204: 199–203 Moroney JV, Wilson BJ and Tolbert NE (1986b) Glycolate metabolism and excretion by Chlamydomonas reinhardtii. Plant Physiol 82: 821–826 Moroney JV, Kitayama M, Togasaki RK and Tolbert NE (1987a) Evidence for inorganic carbon transport by intact chloroplasts of Chlamydomonas reinhardtii. Plant Physiol 83: 460–463 Moroney JV, Togasaki RK, Husic HD and Tolbert NE (1987b) Evidence that an internal carbonic anhydrase is present in 5% grown and air-grown Chlamydomonas. Plant Physiol 84: 757–761 Moroney JV, Husic HD, Tolbert NE, Kitayama M, Manuel LJ and Togasaki RK (1989) Isolation and characterization of a mutant of Chlamydomonas reinhardtii deficient in the concentrating mechanism. Plant Physiol 89: 897–903 Nelson EB and Tolbert NE (1969) The regulation of glycolate metabolism in Chlamydomonas reinhardtii. Biochim Biophys Acta 184: 263–270 Palmqvist K, Sjöberg S and Samuelsson G (1988) Induction of inorganic carbon accumulation in the unicellular green algae Scenedesmus obliquus and Chlamydomonas reinhardtii. Plant Physiol 87: 437–442 Palmqvist K, Sundblad L-G, Wingsle G and Samuelsson G (1990) Acclimation of photosynthetic light reactions during induction of inorganic carbon accumulation in the green alga Chlamydomonas reinhardtii. Plant Physiol 94: 357–366 Palmqvist K, Ögren E and Lernmark U (1994a) The mechanism is absent in the green alga and Coccomyxa: a comparative study of photosynthetic light responses of Coccomyxa , Chlamydomonas reinhardtii and barley protoplasts. Plant Cell Environ 17: 65–72 Palmqvist K, Yu J-W and Badger MR (1994b) Carbonic anhydrase cells activity and inorganic carbon fluxes in low- and of Chlamydomonas reinhardtii and Scenedesmus obliquus. Physiol Plant 90: 537–547 Plumed MdP, Villarejo A, Rios Adl, Garcia-Reina G and mechanism in a Ramazanov Z (1996) The

Martin H. Spalding starchless mutant of the green unicellular alga Chlorella pyrenoidosa. Planta 200: 28–31 stress and Pronina NA and Borodin VV (1993) concentration mechanism: Investigation by means of photosystem deficient and carbonic-anhydrase-deficient mutants of Chlamydomonas reinhardtii. Photosynthetica 28: 515–522 Ramazanov Z and Cardenas J (1992) Involvement of photorespiration and glycolate pathway in carbonic anhydrase induction and inorganic carbon concentration in Chlamy­ domonas reinhardtii. Physiol Plant 84: 502–508 Ramazanov Z and Cardenas J (1994) Photorespiratory ammonium assimilation in chloroplasts of Chlamydomonas reinhardtii. Physiol Plant 91: 495–502 Ramazanov Z, Mason CB, Geraghty AM, Spalding MH and Moroney JV (1993) The low 36 kD protein is localized to the chloroplast envelope of Chlamydomonas reinhardtii. Plant Physiol 101: 1195–1199 Ramazanov Z, Rawat M, Henk MC, Mason CB, Matthews SW and Moroney JV (1994) The induction ofthe mechanism is correlated with the formation of the starch sheath around the pyrenoid of Chlamydomonas reinhardtii. Planta 195: 210–216 Raven JA (1997) mechanisms: A direct role for thylakoid lumen acidification? Plant Cell Environ 20: 147– 154 Rawat M and Moroney JV (1991) Partial characterization of a new isozyme of carbonic anhydrase isolated from Chlamy­ domonas reinhardtii. J Biol Chem 266: 9719–9723 Rawat M and Moroney JV (1995) The regulation of carbonic anhydrase and ribulose-1,5-bisphosphate carboxylase/ in Chlamydomonas oxygenase activase by light and reinhardtii. Plant Physiol 109: 937–944 Reed ML and Graham D (1977) Carbon dioxide and the regulation of photosynthesis: activities of photosynthetic enzymes and carbonate dehydratase (carbonic anhydrase) in Chlorella after growth or adaptation in different carbon dioxide concentrations. Aust J Plant Physiol 4: 87–98 Roberts CS and Spalding MH (1995) Post-translational processing of the highly processed, secreted periplasmic carbonic anhydrase of Chlamydomonas is largely conserved in transgenic tobacco. Plant Mol Biol 29: 303–315 Sheen J (1990) Metabolic repression of transcription in higher plants. Plant Cell 2: 1027–1038 Sheen J (1994) Feedback control of gene expression. Photosyn Res 39: 427–438 Spalding MH (1989) Photosynthesis and photorespiration in freshwater green algae. Aquatic Botany 34: 181–209 Spalding MH (1990) Effect of photon flux density on inorganic carbon accumulation and net exchange in a mutant of Chlamydomonas reinhardtii. Photosynth Res 24: 245–252 Spalding MH and Jeffrey M (1989) Membrane-associated polypeptides induced in Chlamydomonas by limiting concentrations. Plant Physiol 89: 133–137 Spalding MH and Ogren WL (1982) Photosynthesis is required system in Chlamy­ for induction of the domonas reinhardii. FEBS Lett 145: 41–44 Spalding MH, Spreitzer RJ and Ogren WL (1983a) Carbonic anhydrase deficient mutant of Chlamydomonas requires elevated carbon dioxide concentration for photoautotrophic

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Acquisition

growth. Plant Physiol 73: 268–272 Spalding MH, Spreitzer RJ and Ogren WL (1983b) Reduced inorganic carbon transport in a mutant of Chlamydomonas reinhardii. Plant Physiol 73: 273—276 Spalding MH, Critchley C, Govindjee and Ogren WL (1984) Influence of carbon dioxide concentration during growth on fluorescence induction characteristics of the green alga Chlamydomonas reinhardtii. Photosynth Res 5: 169–176 Spalding MH, Winder TL, Anderson JC, Geraghty AM and Marek LF (1991) Changes in protein and gene expression mechanism in wildduring induction of the type and mutant Chlamydomonas. Can J Bot 69: 1008–1016 Spencer KG, Kimpel DL, Fisher ML, Togasaki RK and Miyachi S (1983) Carbonic anhydrase induction in Chlamydomonas reinhardtii II. Requirements for carbonic anhydrase induction. Plant Cell Physiol 24: 301–304 Sültemeyer DF, Klock G, Kreutzberg K and Fock HP (1988) Photosynthesis and apparent affinity for dissolved inorganic carbon by cells and chloroplasts of Chlamydomonas reinhardtii grown at high and low concentrations. Planta 176: 256– 260 Sültemeyer DF, Miller AG, Espie GS, Fock HP and Canvin DT transport by the green alga Chlamydomonas (1989) Active reinhardtii. Plant Physiol 89: 1213–1219 Sültemeyer DF, Fock HP and Canvin DT (1990) Mass spectrometric measurement ofintracellular carbonic anhydrase cells of Chlamydomonas. Plant activity in high and low Physiol 94: 1250–1257 Sültemeyer DF, Amoroso G and Fock H (1995) Induction of intracellular carbonic anhydrases during the adaptation to low inorganic carbon concentrations in wild-type and ca-1 mutant cells of Chlamydomonas reinhardtii. Planta 196: 217–224 Sundblad LG, Palmqvist K and Samuelsson G (1986) An energy-

547 dependent, transient peak in the minute range decay of luminescence, present in cells of Scenedesmus obliquus. FEES Lett 199: 75–79 Suzuki K and Spalding MH (1989) Adaptation ofChlamydomonas reinhardtii mutants to Plant Physiol 90: 1195–1200 Suzuki K, Marek LF and Spalding MH (1990) A photorespiratory mutant of Chlamydomonas reinhardtii. Plant Physiol 93: 231– 237 Thielmann J, Tolbert NE, Goyal A and Senger H (1990) Two systems for concentrating and bicarbonate during photosynthesis by Scenedesmus. Plant Physiol 92: 622–629 Van K and Spalding MH (1997) Insertional mutagenesis of the adaptation in signal transduction pathway for low Chlamydomonas. Plant Physiol 114: 266 (supplement) Villand P, Eriksson M and Samuelsson G (1997) Regulation of level. Plant Physiol 114: 258 genes byenvironmental (supplement) Villarejo A, Martínez F, Plumed MdP and Ramazanov Z (1996a) concentrating mechanism in a starchThe induction of the less mutant of Chlamydomonas reinhardtii. Physiol Plant 98: 798–802 Villarejo A, Reina GG and Ramazanov Z (1996b) Regulation of polypeptides in Chlamydomonas the reinhardtii. Planta 199: 481–485 Williams TG and Turpin DH (1987) The role ofexternal carbonic anhydrase in inorganic carbon acquisition by Chlamydomonas reinhardtii at alkaline pH. Plant Physiol 83: 92–96 Winder TL, Anderson JC and Spalding MH (1992) Translational regulation of the large and small subunits of ribulose bisphosphate carboxylase/oxygenase during induction of the mechanism in Chlamydomonas reinhardtii. Plant Physiol 98: 1409–1414

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Chapter 29 Regulation of Starch Biosynthesis Steven G. Ball

Laboratoire de Chimie Biologique, Unité Mixte de Recherches du CNRS n°111,

Bâtiment C9, Cité Scientifique, 59655 Villeneuve d'Ascq, France

Summary I. Starch: Structure and Function A. An Overview of Starch Structure B. Are Algal Storage Polysaccharides Comparable to Vascular Plant Starches? II. The Starch Pathway A. An Overview of Starch Metabolism B. Enzymes of Starch Metabolism in Chlamydomonas C. Compartmentation of Starch Metabolism in Chlamydomonas D. Regulation of Starch Metabolism III. The Genetics of Starch Biosynthesis A. The Iodine Screen for Starch Defective Mutants B. Mutants Defective for the Supply of ADP-glucose C. Mutants Defective for Elongation D. Mutants with Unidentified Enzymological Defects E. Mutants Defective for Pre-Amylopectin Trimming IV. A Model Explaining the Biogenesis of the Plant Starch Granule

V. Future Prospects

Acknowledgments References

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Summary Transient or long-term storage of photosynthate in starch granules can be considered as the last step of eukaryotic photosynthesis. Storage of glucose into structures larger than the size of an individual bacterial cell is slowly uncovering as an exceedingly complex mechanism, which distinguishes the chloroplast from its ancestor prochloron or cyanobacterial-like cell. There is no question that starch biosynthesis has evolved from a pre-existing simpler bacterial glycogen synthesis pathway. However the number of enzymes involved in plant starch synthesis appears considerably higher. Chlamydomonas reinhardtii is now emerging as the most powerful model system to select for mutants defective in various aspects of granule biogenesis, degradation or overproduction. A full description ofthe eight loci reported to be involved is presented. A genetic demonstration is made of the involvement of the 3-PGA/Pi ratio in controlling the rates of polysaccharide synthesis in algae.

J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 549–567. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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The evidence for the respective functions ofthe starch synthases in the building of specific sub-structures ofthe granule is detailed. The selection ofstarchless C. reinhardtii mutants, in which macrogranular starch is replaced with disorganized glycogen-like structures, has paved the way for a deeper understanding ofplant amylopectin synthesis. A model is thus presented proposing the existence of pre-amylopectin, a branched precursor that is subsequently trimmed into an ordered structure. The trimming is proposed to relieve the physical constraints on the upper size limit imposed on glycogen granule biogenesis. An account of the compartmentation of glycolysis and of both the pentose-phosphate and the starch biosynthesis pathways is given. The relevance of this compartmentation with respect to starch synthesis regulation is discussed.

I. Starch: Structure and Function

A. An Overview of Starch Structure Starch is one of the most abundant biological polymers present in Earth’s biosphere. It remains the major supply of calories in both the human and animal diet and is becoming a major source of plant raw material for non-food purposes. One third of the annual US maize crop is devoted today to starch extraction. The polysaccharide is purified and processed further into over 600 commercial products. Apart from being an economically important commodity, starch constitutes also a major carbon fixation in the chloroplast and to some sink for extent can be considered as the end-product of eukaryotic photosynthesis. Because starch is solely made of glucose residues linked in position biologists have considered and branched in the polysaccharide to merely represent a variant form of glycogen deserving little attention for itself. Plant biology studies were indeed focused on carbon partitioning with few studies dealing with the starch pathway itself and even less on the biogenesis of the polysaccharide structure. Yet the picture that is emerging today is that of a polysaccharide with an extremely complex and ordered structure (for review see Manners, 1989). The complexity is further reflected by the multiplicity of enzymes reported to be involved, some of which were completely unexpected from the preexisting work on animal, fungal or bacterial glycogen (for reviews see Caspar, 1994; Ball, 1995; Nelson and Pan, 1995; Preiss and Sivak, 1996; Smith et al., Abbreviations: AGPase – ADP-glucose pyrophosphorylase; Am – amylose; Ap – amylopectin; BE – Branching Enzyme; CCM – Concentration Mechanism; d.p. – degree of polymerization; GBSS – Granule-Bound Starch Synthase; – wavelength of the maximal absorbance of the iodinepolysaccharide complex; 3-PGA – 3-phosphoglycerate; Pgm – phosphoglucomutase; SSS – Soluble Starch Synthase

1997) . Our present state of knowledge concerning starch structure is summarized in Fig. 1 that relates the organization of the granule to that ofthe primary and secondary structures of amylopectin. However amylopectin represents the major but not the only polysaccharide fraction in starch. Amylose, the minor fraction of plant starch granules, is not represented in Fig. 1 because the position ofthese molecules relative to amylopectin is too uncertain. Table 1 summarizes the comparative structure of amylose and amylopectin. No mutants were ever reported to selectively lack amylopectin while mutants without amylose have been described in many plant systems including C. reinhardtii. These mutants build wild-type amounts of granules with normal organization. We therefore suspect that the major aspects of starch granule biogenesis in plants will be explained when we will understand the more complex mechanisms underlying amylopectin synthesis. Eukaryotic algae are of particular relevance for studies dealing with starch synthesis. Starch polysaccharides are, indeed, not found in bacteria and fungi. Because C. reinhardtii is the only starchstoring unicellular organism intensively studied by geneticists, it offers a quite unique opportunity to understand the basic mechanisms of starch granule biogenesis. This of course requires that the basic structural organization and biosynthesis of green algal starch are identical to those found in vascular plants. Until recently very little information was available concerning this important issue. We will thus briefly review the evidence that we believe establish C. reinhardtii as the most powerful model system to understand and therefore manipulate starch biosynthesis in plants.

B. Are Algal Storage Polysaccharides Comparable to Vascular Plant Starches? In vascular plants, starch can be found in both the chloroplast of source tissues and the amyloplast of

Chapter 29

Starch Biosynthesis

storage organs. In leaf cells starch granules are generally small. They are synthesized during the light phase in order to generate by subsequent breakdown a regular supply of sucrose to sink tissues (for review see Preiss and Sivak, 1996). This breakdown occurs predominantly during the night but can also be triggered in the light according to the demand imposed by the carbon sink. Therefore in the chloroplast of the leaf cell, starch synthesis is tightly coupled to photosynthesis and ensures transient carbon storage to meet the asynchronous demands of the sink tissues. The carbon flows from the chloroplast

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to the cytosol to be subsequently exported through sucrose translocation. In this case, the time scale of a switch from synthesis to degradation is thus of only a few minutes to a few hours and the carbon stores must therefore remain instantly available. Starch structure and biosynthesis has evolved to meet these requirements and the name ‘transient starch’ has been coined to describe this polysaccharide. In sink tissues such as the cereal grain endosperm, the potato tuber or the pea seed embryo, high levels of sucrose are constantly supplied and stored in the non photosynthetic amyloplast in the form of very

552 large semi-crystalline granules as depicted in Fig. 1. The carbon thus flows from the cytosol into the amyloplast. This form of starch will be used by the next generation of plants several weeks to several months or years later upon germination of the seed or tuber. Moreover a distinct set ofenzymes are induced during germination to ensure rapid mobilization of this form of carbon store. The term ‘storage starch’ accurately describes this situation. It is this form of polysaccharide which has been turned into so many different uses by our modern economies. Because of this, structural analyses of starches have been largely confined to storage starches and were, mistakenly, compared directly to structures of polysaccharides extracted from eukaryotic algae under variable, often uncontrolled, growing conditions. The crystalline lattices of higher plant storage starches fall into three classes (for review see Imberty et al., 1991). The A crystalline lattice can be found in cereal endosperm starches while the more hydrated B type typifies tuber starches. The C type consists of a mixture ofA and B crystals and can be found in pea embryo storage starches. Few X-ray diffraction studies have been performed on eukaryotic algal starches (Meeuse and Kreuger, 1959). In these studies starches from both red and green algae fell into the B class while a number of other green algal starches displayed the A cereal type of diffraction pattern. The cytosolic location of starch granules in red algae together with the reported absence of amylose led authors to believe these polysaccharides to be completely different from vascular plant starches and have named them ‘floridean starch’ (Meeuse et al., 1960). Chlorophyceae, Charophyceae and green seaweed all were reported to carry both amylopectin and amylose fractions although in quite variable relative amounts (for review, see Craigie, 1974). Moreover algal starches in general seemed to display less crystallinity than their vascular plant counterparts (Meeuse, 1962) leading to the suspicion that these polysaccharides although related to vascular plant starches may not have an identical organization. These differences, at first, seem confirmed in C reinhardtii. The conspicuous presence of the pyrenoid surrounded by a typical starch sheath in the middle of the chloroplast is not found in vascular plants. Moreover the cytological location of starch availability deposition responds dramatically to in the alga. Both Kuchitsu et al. (1988) and Ramazanov et al. (1994) noted that under high concentrations (4%) in the presence of light most of

Steven G. Ball the starch could be found in the chloroplast stroma while underlimiting (air levels of 0.04%) starch deposition occured predominantly around the pyrenoid. In addition, the presence of acetate favored stromal starch synthesis and seemed to slow pyrenoidal starch synthesis. A general tight correlation was established between the induction of concentration mechanism and pyrenoidal the starch synthesis while regulation of stromal polysaccharide synthesis seemed to be independent, thus defining two distinct types of polysaccharides. Both Kuchitsu et al. (1988) and Ramazanov et al. (1994) suggested a function for pyrenoidal starch in the CCM. Mutants from Chlamydomonas defective for the STA7 locus (Table 2) were subsequently shown to display no alteration of their CCM (PinoPlumed et al., 1996). A similar result was obtained with Chlorella pyrenoidosa uncharacterized starchdefective mutants (Villarejo et al., 1996). We believe that the sites of starch synthesis arise as a consequence of the location of metabolically active Calvin cycle multienzyme complexes and does not directly participate in the function of the algal CCM per se. Therefore, we do not think that pyrenoidal and stromal starches depend on distinct biosynthetic enzymes or define truly different polysaccharides. Süss (1995) has found (by immunolocalization) that Calvin cycle multienzyme complexes together with Rubisco, Rubisco activase, ferredoxin-NADP reductase and ATP synthase were indeed located both at the surface ofthe pyrenoid tubule membranes and at the stromal side of thylakoid membranes. Moreover Süss confirmed that Rubisco was found in the pyrenoid in great abundance but distinguished this form of the enzyme which he calls type II Rubisco from the type I enzyme which is found together with other enzymes of carbon metabolism at the sites of starch synthesis. According to Süss, and type II Rubisco would be needed to bind transfer it to other Rubisco molecules untill it finally reaches the pyrenoid tubule membrane type I enzyme that will incorporate it into 3-PGA. It is tempting to speculate that it is through 3-PGA activation of ADP-glucose pyrophosphorylase, the rate limiting step of starch biosynthesis, that the location of starch synthesis and thereby that of the granules themselves fixation during are tightly coupled to the sites of photosynthesis. No studies dealing with the structure of starch in C. reinhardtii were available prior to our own work. However one of the very few detailed studies on the

Chapter 29

Starch Biosynthesis

structure of green algal storage polysaccharides can be traced back to 1972 by Hirst et al.. In this study, starch from another monocellular green alga of the order Volvocales (Haematococcus pluvialis) was investigated in reasonable detail. Hirst et al. concluded that H. pluvialis contained a storage polysaccharide linked and branched glucose made of residues. It had an interaction with iodine very similar (wavelength to that of higher plant starches. The of the maximal absorbance of the iodine-polysaccharide complex, as defined in section IIIa) measured for the entire fraction (595 nm) was very close to those of tuber or cereal endosperm storage starches (570–590 nm). Moreover they were able to convincingly separate an amylopectin and amylose fraction whose relative amounts and were identical to those of vascular plant storage starches. They were also able to equate average, exterior and interior chain-lengths with those of higher plant storage starches. In another older study (Olaitan and Northcote, 1962) performed on Chlorella pyren­ oidosa, another monocellular green alga, an unusually low 7% value was reported for the amylose fraction while Hirst et al. (1972) reported a three-fold higher value. Variation in amylose content seem to rather be the rule than the exception in green algae. Some species were even reported to lack amylose (Craigie, 1974). We believe that the evidence now gathered in Chlamydomonas clearly point to differences due to growth conditions rather than changes reflecting true modifications of polysaccharide structure. It is quite by chance that we chose nitrogen starvation as our routine culture conditions to analyze starch (Ball et al., 1990). These conditions were chosen because they lead to a 10 to 50 fold increase in final cellular starch content. The pronounced depigmentation of nitrogen-starved algae enabled us to select for mutants by screening directly iodine-stained cell patches on solid media. The starch extracted from nitrogenstarved, acetate-supplied C. reinhardtii cultures was subjected to intensive structural characterizations (Delrue et al., 1992; Fontaine et al., 1993; Maddelein et al., 1994; Libessart et al., 1995; Buléon et al., 1997). We conclude from all these studies that the starch extracted from nitrogen-starved C. reinhardtii cultures is virtually identical to cereal endosperm storage starches. Our criteria are as follows: 1) The storage polysaccharide from C. reinhardtii was shown by chemical methylation to contain

553 exclusively glucose residues.

linked and

branched

2) The branching ratio measured by chemical methylation for the amylopectin (Ap) fraction was of 1:20 while that measured for amylose (Am) was approximately of 1:100. 3) The relative amount of Am varied between 15 to 30%. 4) The molecular mass distribution ofAp and Am molecules was identical to that ofmaize endosperm storage starch and distinct from potato tuber starch. of 550 nm) 5) The iodine interaction of Ap ( and Am ( > 620 nm) were identical to vascular plant storage starches. 6) The chain-length distribution ofAp was shown to be trimodal and identical to vascular plant storage starches. 7) The proton and carbon NMR spectra ofAp and Am were identical to the vascular plant storage starches. 9) The wide-angle X-ray diffraction pattern of wild type C.reinhardtii starches were of the Atype confirming the identity with cereal endosperm starches. 10) The physicochemical properties of the algal starches assayed by DSC (Differential Scanning Calorimetry) were analogous to higher plant storage starches. C. reinhardtii after 5 days of nitrogen starvation is displayed in Fig. 2A and compared to log phase-grown nitrogen-supplied algae (Fig. 2B). It is evident that drastic modifications in the flow of carbon metabolism have occurred. In nitrogensupplied medium, in the presence of acetate and light, log phase cells accumulate modest amounts of starch around the pyrenoid and in the stroma. The bulk of the carbon is directed to the building of the new cellular material required for the rapid cell division that occurs under these optimal growth conditions. Cell division provides thus a powerful photosynthate sink yielding a net outflow of carbon from the plastid to the cytosol. During nitrogen

Steven G. Ball

554

amylose. Moreover the remaining amylopectin displayed an altered chain length distribution consisting of a relative increase both in short (<9 glucose residues) and extra long (>50) chains. Very recently, Van den Koornhuyse et al. (1996) have shown that starches extracted from mutant cells defective for the supply of the ADP-glucose substrate displayed during nitrogen starvation a structure identical to that of the polysaccharide purified from wild-type nitrogen-supplied cultures. Therefore storage starch becomes a phenocopy of the transient form of polysaccharide whenever the supply of substrate is lowered. We believe that these structural differences have important functional implications with respect to crystallinity of the polysaccharides and therefore to the availability of the carbon stores through degradation. It is possible that switches from one structure to another may be triggered by modifying simply the substrate supply without the need to induce a specific subset of enzyme activities. The required changes will be obtained through the differences in affinities ofthe various starch synthases of distinct structural specificity for their common ADP-glucose substrate. Similar differences were found between the structures of transient and storage starches in higher plants. In pea embryos, mutations leading to a decrease in ADP-glucose supply were also found to substantially decrease the relative amount of amylose (for review, see Smith et al., 1997). II. The Starch Pathway

A. An Overview of Starch Metabolism starvation cellular material such as proteins, RNAs or photosynthetic pigments are degraded and converted into storage molecules such as starch or lipids. There is therefore a net inflow of carbon from the cytoplasm into the non photosynthetic plastid. These two extreme physiological conditions mimic the differences that are witnessed in vascular plants between the plant leaf chloroplast and the non photosynthetic amyloplast from storage organs. Libessart et al. (1995) have witnessed drastic differences in the structure and composition of starches extracted from nitrogen-supplied and nitrogen-starved Chlamydomonas cells. By comparison with starved cells (storage starch), starch extracted from nitrogen-supplied cultures (transient starch) contained very little low molecular weight

Our present state of knowledge concerning starch metabolism is summarized in Fig. 3. It is presently clear that starch biosynthesis has evolved from the preexisting cyanobacterial glycogen synthesis pathway. Before we detail various aspects of green algal starch biosynthesis it is necessary to give a brief account of the basic reactions involved. These consist of synthesis of the ADP-glucose substrate, transfer of glucose to the growing polysaccharide chain (elongation) and branching: 1) Plastidial phosphoglucomutase (Pgm, EC 5.4.2.2) Glucose-6-P

Glucose-1-P.

Chapter 29

Starch Biosynthesis

555 Starch synthases transfer glucose from the glycosylnucleotide ADP-glucose to the nonreducing end of a growing linked glucan. These enzymes catalyze an elongation step and cannot prime the reaction. Starch synthases can be found either predominantly bound to the granule or in the soluble phase. It is now accepted that the major granule-bound starch synthase is the major, if not only, amylose biosynthetic enzyme. Three distinct types of starch synthases have been found in those species of vascular plants that were characterized in sufficient detail. Their relative contributions to amylopectin synthesis remain unknown. 4) Branching enzyme (EC 2.4.1.18)

While in bacteria phosphoglucomutase cannot be considered a step specific to glycogen synthesis only, the conversion of glucose-6-P to glucose-1P by Pgm is the sole documented function for the plastidial form of the plant enzymes and can thus be considered as the first step ofthe starch pathway. 2) ADP-glucose pyrophosphorylase (AGPase, EC 2. 7. 7. 27). Glucose-1-P + ATP

ADP-glucose + PPi

ADP-glucose pyrophosphorylase catalyzes the rate-limiting step of starch and bacterial glycogen synthesis. The bacterial enzyme is a homotetramer made of subunits ranging in the 50 to 55 kDa range. 3-PGA is the most efficient allosteric activator of the cyanobacterial AGPase while orthophosphate is the most potent inhibitor. The higher plant enzymes are heterotetramers harboring two distinct but related types of subunits in the same size ranges as those of the bacterial enzymes. The so-called small subunits are proposed to be the catalytic subunits while the large subunits are proposed to modulate the sensitivity to allosteric effectors. As with the cyanobacterial enzyme the plant AGPase is exquisitely sensitive to the 3-PGA to orthophosphate ratio. 3) Starch synthase (EC 2.4.1.21) ADP-glucose +

ADP +

linked glucan ofat least 12-16 residues Linear branched glucan with one branch point. These enzymes share many common features with other starch hydrolytic enzymes. The vascular plant enzymes display the barrel enzyme core structure which is common to amylases, debranchhydrolases. ing enzymes and many other Two distinct families are found in vascular plants. Mutations in the structural gene for the A (type II) family have been reported to alter amylopectin synthesis and lead to the high amylose phenotype. No functions have yet been found for enzymes of the B (type I) family. Starch catabolism in plants involves three distinct pathways including hydrolases, phosphorylases and lyases. The relative importance of the three pathways in the different tissues is still a matter of debate (for a review see Steup, 1988). Starch hydrolases transfer a glucosyl group to water by a general acid catalysis mechanism. Amylases are generally classified as according both to the optical properties or EC of cleavage products and to the endo ( 3.2.1.1) or exo-amylase ( EC 3.2.1.2) type of reaction. Because of the endo-type attack and its unique ability to work on the granule itself, is able to rapidly depolymerize starch and thus paves the way for complete degradation of the polysaccharide through the production of soluble Amylases unlike several other hydrolases linkage. are unable to cleave the Starch phosphorylase (E.C. 2.4.1.1) catalyzes the transfer of glucose from the non-reducing end of an

556 glucan to orthophosphate yielding glucose-1 -phosphate. Starch lyases (EC 4.2.2.-) have presently only been described in red algae and fungi (Yu and Pedersen, 1993; Yu et al., 1997). These enzymes produce 1,5-anhydrofructose from the non glucan. Neither reducing end of an lyases nor phosphorylases can degrade the linkage or bypass the branch point. Therefore complete degradation always does require to some extent the concerted action of hydrolytic enzymes linkage. able to cleave the

B. Enzymes of Starch Metabolism in Chlamydomonas While in depth characterizations ofpartially purified ADP-glucose pyrophosphorylase and starch synthase from Chlorella pyrenoidosa (Preiss and Greenberg, 1967; Sanwal and Preiss, 1967), as well as of phosphorylase and ADP-glucose pyrophosphorylase from Chlorella vulgaris (Nakamura and Imamura, 1983, 1985), had been previously reported, the first characterizations of enzymes of starch metabolism in C. reinhardtii can be traced back to 1984 (Levi and Gibbs, 1984). In this study phosphorylase, amylase, limit dextrinase (EC 3.2.1.3) and maltase ( EC 3.2.1.20), four enzymes thought to be involved in starch catabolism were detected in synchronously grown (12-h light/dark cycle) C. reinhardtii cells. Limit-dextrinase can be defined as linkage. an enzyme that can only hydrolyze the Limit-dextrinases and isoamylases are collectively known as debranching enzymes. Limit-dextrinase, also known as plant pullulanase can digest pullulan, a bacterial polysaccharide made of a regular succession of maltotriose chains linked together by linkages at the ends ofeach maltotriosyl residue. Isoamylase cannot debranch tightly spaced branches and thus will not hydrolyze pullulan. However it will cleave more efficiently both glycogen and amylopectin. The activities of both phosphorylase and amylase were characterized in greater detail and followed through the cell cycle. Levi and Gibbs (1984) were able to monitor a four to five-fold increase in both activities between the midlight and middark periods. Both enzymes had broad pH optima ranging between 6.5 to 7.5. However the activity of only the amylase displayed a sharp decrease above 7.6. This could be correlated with the in vivo stromal pH shifts from 7 to 8 known to occur during a transition from dark to light. These observations are consistent with

Steven G. Ball a function of both enzymes in starch degradation. The partially purified amylase was further characon the grounds of the kinetics terized as an of amylopectin degradation. Moreover the enzyme was shown to be heat labile at 55 °C (5 min), inhibited by 2 mM N-ethylmaleimide (NEM) and insensitive to 10 mM EDTA inhibition. These properties are shared with the spinach leaf amylase but differ from those of the enzymes involved in the degradation of storage starch during germination. An enzyme of similar properties was also partially purified by Mouille et al. (1996a) and shown to be a 53 kDa starch hydrolase. Moreover Mouille et al. (1996a) were able to identify the nature of the oligosaccharides produced by the enzyme. These consisted ofvery short highly branched glucans less than 12 glucose residues in overall size. Further characterization of the C. reinhardtii phosphorylase was performed by Ball et al. (1991) who reported a strong mixed type of inhibition of the enzyme by ADP-glucose (50% inhibition by 7 assayed at 1.3 mM Pi). This inhibition, which was also found in Chlorella vulgaris by Nakamura and Imamura (1983), distinguishes the algal phosphorylases from their land plant counterparts. Strong inhibition of phosphorylases by the substrate of starch synthesis also argues that the enzyme is involved solely in polysaccharide degradation. Soluble and granule-bound starch synthases were reported initially in C.reinhardtii by Kuchitsu et al. (1988) who noted a three to five-fold increase in total grown soluble starch synthase activity in 4% cells by comparison to air-adapted cultures. Two soluble starch synthases (SSSI and SSSII) were subsequently partially purified and characterized by Fontaine et al. (1993), Maddelein et al. (1994) and Buléon et al. (1997). Molecular masses of 115 and 75 kDa were respectively measured for SSSI and SSSII by zymogram in denaturing conditions and western blotting. SSSI favored elongation ofglycogen over amylopectin while SSSII displayed opposite preferences. Both enzymes were unable to prime the reaction even in the presence of 0.5 M citrate while SSSII was clearly inhibited and SSSI activated twofold under these conditions. This behavior distinguished the algal synthases from the vascular plant enzymes. Moreover to date no traces could be found of a third soluble starch synthase in C. reinhardtii cultures (Buléon et al., 1997). GBSSI from C. reinhardtii was characterized as a 76 kDa protein with N-terminal sequences similar to those

Chapter 29

Starch Biosynthesis

reported for higher plant enzymes (Delrue et al., 1992). The granule-bound enzyme displayed a 4 mM for ADP-glucose, a five to ten-fold higher value than those reported for the soluble enzymes, and unlike higher plant enzymes could not use UDPglucose. The optimal pH of both granule-bound and soluble enzymes is markedly alkaline (above 9) which fits the biosynthetic function of these enzymes. Branching enzymes were initially detected by Ball et al. (1990, 1991). However the first significant characterizations of branching enzymes in C. rein­ hardtii were reported by Fontaine et al. (1993). Two branching enzymes (BEI and BEII) were detected following anion-exchange chromatography in C. reinhardtii. Both ofthese activities branched potato amylose and produced a polysaccharide with a of 540 nm. However BEII turned out to branch amylose faster than BEI. While the numbering ofthe enzyme peaks reflect their elution order on the columns, their characterization is presently insufficient to relate them precisely to the vascular plant enzymes ofthe A or B families. However biochemical results obtained to date are sufficient to imply the existence of at least two types of enzyme activities. ADP-glucose pyrophosphorylase was reported and characterized in partially purified extracts by Ball et al. (1991) and was subsequently purified to homogeneity by Iglesias et al. (1994). The enzyme was shown to be a heterotetramer composed of two 53 kDa and two 51 kDa subunits which both crossreacted with antiserum directed against the spinach leaf enzyme (Iglesias et al., 1994). The enzyme was found to be activated 22 fold by 3-PGA and inhibited by orthophosphate. The activity is thus tightly regulated by the 3-PGA/Pi ratio as is the case for most ADP-glucose pyrophosphorylases from both vascular plants and cyanobacteria. Cytosolic and plastidic phosphoglucomutase activities of similar properties were also reported in C. reinhardtii (Klein, 1986). It is therefore presently clear from all these enzymological studies that all the critical activities reported to be involved in higher plant starch synthesis are found in C. reinhardtii in the same number of isoforms and with similar biochemical properties. These observations further strengthen the case of C. reinhardtii as an ideal microbial model system to understand the biogenesis of the higher plant starch granule.

557

C. Compartmentation of Starch Metabolism in Chlamydomonas A detailed compartmentation study was reported for enzymes of glycolysis and of the oxidative pentosephosphate pathway (Fig. 4) (Klein, 1986). For the activities of the first part of the glycolytic chain (from fructose-6-phosphate to triose phosphate), over 90% of the activity was shown to be associated with the plastid. For the enzymes required for the conversion of 3-PGA to pyruvate, over 95% of the activity was associated with the cytosolic fractions. Moreover 70% of both glucose-6-phosphate dehydrogenase and gluconate-6-phosphate was found in the plastid (Klein, 1986). Hexokinase activity was also detected in C. rein­ hardtii (Levi and Gibbs, 1984). While the compartmentation of the hexokinase was not studied, C. reinhardtii’s inability to grow on glucose supplied in the medium as a carbon source together with the plastid location of the upper glycolytic pathway and the presence of a plastid-located starch degradation pathway strongly suggest the need for a plastidic form of the enzyme. Two hexokinases were found associated to purified chloroplasts by Singh et al. (1993), both in spinach and Chlamydomonas. The major form was found in the chloroplast stroma while the minor form was assigned to the cytoplasmic side of the outer plastid membrane. Moreover the entry ofglucose into the plastid was largely dependent on the exogenous supply ofATP. According to Singh et al. (1993), this could be explained by the presence of a hexose-phosphate translocator that would be supplied with glucose-6-P by the outer membrane hexokinase. Phosphoglucoisomerase and phosphoglucomutase which are needed to metabolize glucose-1-P produced by starch phosphorolysis were found both inside and outside the chloroplast. As to the enzymes of starch metabolism per se, extra-plastidic forms of or phosphorylase were not detected (Levi and Gibbs, 1984; Klein, 1986). These observations are at variance with those reported for higher plant leaves but remain in perfect agreement with those reported for Dunaliella marina, another unicellular green alga of the order Volvocales (Kombrink and Wöber, 1980). In the case of D. marina compartmentation studies coupled to detailed activity and zymogram analyses of starch synthases, branching enzymes, phosphorylases and ADP-glucose pyrophosphorylase establish the starch pathway as exclusively intra-

558

plastidic. In C. reinhardtii classical algal transit peptide sequences were indeed encoded by the ADPglucose pyrophosphorylase large subunit cDNA (Van den Koornhuyse et al., 1996). The picture that is slowly emerging for unicellular green algae is that of an entirely plastidic location of starch metabolism. This was further confirmed by Kreuzberg et al. (1987) and by a recent study of the compartmentation of metabolite pools (Klock and Kreuzberg, 1990). The compartmentation of glycolysis displayed in Fig. 4 requires the presence of a phosphate translocator to shuttle triose-P and 3-PGA from the plastid to the cytosol. Such a translocator has indeed been found (Klein et al., 1983) in C. reinhardtii. Moreover metabolites exported by intact purified chloroplasts after 15 min illumination in the presence were conclusively shown to consist of of labeled 23% 3-PGA, 15% DHAP, 20% hexose monophosphates and 13% glycolate (Klein, personal communication). Therefore in addition to triose-P transported through the phosphate translocator, hexose phosphates can also be shuttled between the two cellular compartments.

Steven G. Ball

D. Regulation of Starch Metabolism Contradictory results have appeared concerning polysaccharide content or the rates of starch accumulation during the cell cycle of synchronized microalgal cultures. While the contradictions may be due to species or strain specific differences, they most probably originate from the very different techniques that were used to measure these rates. Of particular relevance are the observations made by Klein (1987) which point to the existence of two maxima, at the very beginning and end of the light phase, in the starch content of synchronized (12 h light/ 12 h dark) C. reinhardtii cells. While, as was found by several others, the starch slowly decreased at night, an even sharper decline was observed at the middle of the light phase. This decrease coincides with the minimum observed in independent measures of starch synthesis rates. In fact net degradation was observed when measuring these rates despite the presence of an overall constant rate of ´ fixation in the light phase. These results strongly suggest cell cycle control of starch degradation during the light phase.

Chapter 29 Starch Biosynthesis Spudich and Sager (1980) in an older report had also noted that starch degradation measured in darkness was greatly stimulated if the cells switched to the dark were taken at mid-G1 at the middle of the light phase. It would be of great interest to correlate these observations with measures of key enzyme and most activities such as phosphorylase, of all ADP-glucose pyrophosphorylase. The evidence obtained by Levi and Gibbs (1984) seems to rule out and phosphorylase activities an increase in at the middle of the light phase. This analysis should also take into account the evolution of the plastidic pools of orthophosphate, hexose monophosphates, ADP-glucose and 3-PGA during the light phase. It must be stressed that Stitt and Heldt (1981) as well as Kruger et al.(1983) have found similar results for isolated spinach chloroplasts. In C.reinhardtii, a net decrease in ADP-glucose concentration would rapidly trigger degradation by relieving the inhibition of phosphorylase by the nucleotide sugar. Several studies have appeared concerning starch breakdown under anaerobiosis in darkness (Klein and Betz, 1978; Gfeller and Gibbs, 1984; Kreuzberg and Martin, 1984). Of interest is the oscillatory fermentation and starch degradation that was noted and characterized by Kreuzberg and Martin (1984). After a switch to anaerobiosis and darkness the rate of starch degradation decreases and increases (oscillates) in a highly regular fashion. A 59 min mean period of starch degradation was measured with phosphofructokinase as a possible rate-limiting step. Because they do not detect significant amounts of glucose in their extracts, Kreuzberg and Martin (1984) believe that the major route for starch degradation during anaerobiosis in darkness requires phosphorylase followed by glycolysis. Ofcourse one cannot rule out a hydrolytic onset of starch degradation followed by phosphorolysis ofthe soluble dextrin. Degradation of starch in darkness through either glycolysis or the pentose phosphate pathway both yield reducing equivalents that have to be reoxidized within the plastid. While both chlororespiration and the activity of a plastidic malate/oxaloacetate shuttle have been proposed as possible mechanisms for regeneration ofoxidized NADP for continuous starch breakdown, Klöck and Kreuzberg (1987) suggest the involvement of a plastidic glycerol-3-phosphate dehydrogenase that reduces one DHAP into glycerol3-P for each 3-PGA generated through starch breakdown. Klöck et al. (1989) further prove that starch degradation from isolated chloroplasts in the

559 dark does not require oxygen. It must be stressed that most studies dealing with starch degradation deal more with establishing the routes of fermentation from starch and their regulation, which are out of the scope of this review, rather than the mechanisms of starch mobilization per se. The identity of the enzymes responsible for breakdown of the crystalline granules, the nature of the dextrin produced and the key metabolic signals triggering starch degradation have, in fact received very little attention. The study of granule degradation and its regulation in C. reinhardtii could prove invaluable to understand the mechanisms at work for starch degradation in vascular plants.

III. The Genetics of Starch Biosynthesis

A. The Iodine Screen for Starch Defective Mutants Nitrogen starved Chlamydomonas cells, especially when supplied with acetate, loose the bulk of their photosynthetic pigments and accumulate both lipids and starch (Ball et al., 1990; Ball et al., 1991; Libessart et al., 1995). Cells plated on solid media with acetate but limiting nitrogen appear as pale yellow patches prior to the first round of iodine vapor spraying. The fixed cells destain rapidly and bleach completely within a few hours. The white fixed patches can be kept indefinitely and restained at will with iodine vapors. The color appearing then is that of the pure iodine-polysaccharide interaction with very little or no interference due to other cellular material (see linked Color Plate 7). Iodine interacts with glucans by inserting in the hydrophobic cavity of the linear glucan helices. However these helices can be generated only if the chains are sufficiently long. This explains why iodine will not interact with linear chains under 12 glucose residues in length at 20 °C. Above this threshold value the interaction will yield complexes whose stability will depend on length (Banks et al, 1971). Moreover the color of the iodine glucan complex will evolve from a weak brown to a strong dark green color for the chains with more than 100 glucose residues. This explains why the iodine-polysaccharide complexes of glycogen (with very short chains), amylopectin (with medium size chains) and amylose (with extra-long chains) will stain respectively brown, purple and green. The wave-length of the maximal absorbence of the iodine-polysaccharide complexes

560

(the ) will range respectively between 450–490 nm (glycogen) to 530-560 nm (amylopectin) and 600-650 nm (amylose). The mix of the purple amylopectin with the greenish amylose yields the dark blue color that typifies starch. It so happens that the starch content of 5 to 10 d-old nitrogen-starved algae is optimal to detect both large and minute differences in color among thousands to millions of surviving colonies on a few petri dishes. Because a direct screen with iodine cannot be applied easily in higher plants, Chlamydomonas ranks first as to the ease with which mutants defective for the starch pathway can be detected. In vascular plants mutant screens rely mostly on selection of kernels or seed with abnormal shape or texture. A detailed account of the elegant work performed on the maize mutants defective for starch biosynthesis can be found in Nelson and Pan (1995). Subsequently it was found that starchless algae are perfectly viable thus facilitating even more the genetic dissection ofstarch metabolism. Four distinct phenotypes can be revealed in C. reinhardtii. Nitrogen-starved cell patches will stain yellow if the amount ofpolysaccharide falls beneath 5% of the wild-type content. Purple-red cell patches will appear upon loss ofamylose. A relative decrease in amylopectin content will yield the high-amylose olive-green phenotype. The dark blue phenotype characterizes the wild-type references.

B. Mutants Defective for the Supply of ADPglucose Mutations in three different loci (STA1, STA5 and

Steven G. Ball

STA6) lower the supply of ADP-glucose in the chloroplast. The X-ray induced sta1-1 (Ball et al., 1991) and the insertion alleles sta1-2::ARG7 and sta1-3::ARG7 (van den Koornhuyse et al., 1996) all lead to identical phenotypes (Table 2). Mutants of STA1 accumulate from 1 to 8% of the wild-type starch amount. Moreover ADP-glucose pyrophosphorylase has a lower response to 3-PGA. While the purifiedADP-glucose pyrophosphorylase displays a maximal 22-fold activation by 3-PGA (Iglesias et al., 1994), the mutant enzyme either partially purified (Ball et al., 1991), or purified to near homogeneity (van den Koornhuyse et al., 1996), shows at most a two-fold enhancement. Ofparticular relevance is the fact that the kinetics of the mutant enzyme and its sensitivity to orthophosphate are unaffected in the absence of 3-PGA (Ball et al., 1991; van den Koornhuyse et al., 1996). In addition, the total amount of activity measured for the mutants ranges between 50 to 100% ofthe wild-type. Therefore the phenotype of the mutants can be solely attributed to the lack of 3-PGA activation of the enzyme and brings in vivo demonstration of the importance of allosteric regulation of the enzyme. We believe that it is the remaining sensitivity of the mutant enzyme to the orthophosphate inhibition which is responsible for the collapse of starch synthesis. Van den Koornhuyse et al. (1996) have cloned and sequenced a cDNA corresponding to the large subunit of ADP-glucose pyrophosphorylase. The cDNA was used as probe to establish that STA1 indeed encoded an ADP-glucose pyrophosphorylase subunit gene (van den Koornhuyse et al., 1996). Recently insertional mutagenesis yielded a novel mutant

Chapter 29

Starch Biosynthesis

containing less than 0.1% total storage polysaccharide making it and the Arabidopsis thaliana adg1 defective plants (Lin et al., 1988) the most severely impaired starch mutants isolated to date (C. Zabawinski, personal communication). The sta6-1::ARG7 carrying mutant grew well but turned out to be sterile and paralyzed. The starchless mutants were completely devoid of ADP-glucose pyrophosphorylase activity. PEG induced fusions between the wall defective original mutant strain and protoplasts generated from a sta1-1 carrying strain yielded fully wild-type pseudo-diploids establishing trans complementation of all defects. The fusion product proved fertile in crosses and was used to produce triploid zygotes. Upon meiosis the aneuploid progeny was used to analyze the segregation of the defects. This analysis coupled to Southern blotting using the large ADP-glucose pyrophosphorylase subunit cDNA probe established the presence of 2 genes required for AGPase activity. STA1 is now proven to encode the large 53 kDa regulatory subunit. The sta1-1 allele was also proved either to be a major translocation or, more likely, a pericentric inversion ofa large chromosomal segment. The inversion (or translocation) occurred in the middle of the enzyme coding sequence. Therefore the phenotype of sta1-1 is explained by the absence of the large subunit and the formation either of fully active monomers or more likely of a homotetramer ofsmall subunits. This was confirmed by the isolation of insertional mutants with the very same phenotype (van den Koornhuyse et al., 1996). The most likely hypothesis to explain the extremely severe phenotype caused by the sta6-1::ARG7 allele is that it encodes the small 51 kDa catalytic subunit of the enzyme. Interestingly it is clear from the triploid zygote genetic analysis that sterility segregated independently from both STA6 and STA1. Sterility therefore does not come as a consequence ofimpaired mobility and loss of starch. Indeed Fletchner and Cirino (1992) had previously demonstrated that photosynthetic mutants with low starch storing capacity did not differ in their mating efficiency when compared to other such mutants with normal or high starch storing abilities. Paralysis however could still define a major pleiotropic effect due to the absence ofstarch. Interestingly the flagellae ofstrains carrying the sta6-1::ARG7 allele appeared normal under the microscope while motility could not even be rescued through the supply of acetate in the medium.

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A spontaneous mutant displaying a low starch phenotype was isolated by Bulté and Wollman. The mutant carried a single genetic defect at the STA5 locus and was subsequently found to be selectively defective for the major Chlamydomonas phosphoglucomutase (van den Koornhuyse et al., 1996). The phenotype seemed however less severe than that of the corresponding Arabidopsis (Caspar et al., 1985; for review see Caspar, 1994) and Nicotiana sylvestris (Hanson and McHale, 1988) mutants. Since Klein (1986) showed that the major C. reinhardtii phosphoglucomutase was compartmented in the chloroplast and since the whole starch pathway seems to be confined to the plastid (Kombrink and Wöber, 1980), we conclude that it is the plastidic form of the enzyme that is missing in the mutant. Mutants carrying sta5-1 accumulate from 4 to 20% of the normal starch amounts (van den Koornhuyse et al., 1996). We believe this reflects the activity of the minor cytosolic phosphoglucomutase and strongly argues that glucose-1-P is transported through the plastid membranes (Fig. 4). As mentioned above, sta1-1 and sta5-1 both display modified structures of their residual starches without showing any modification ofall other major enzymes involved in starch biosynthesis. We thereby conclude that ADP-glucose supply tightly controls the polysaccharide’s structure and propose that this occurs through differential effects on the balance of the active starch synthases. It is known, for instance, that GBSSI has a five-fold higher for ADP-glucose than the soluble starch synthases. Modifications in the ADP-glucose pool could then alter the structure because ofthe specialized functions displayed by the different starch synthases in the building ofdifferent granule substructures (van den Koornhuyse et al., 1996)

C. Mutants Defective for Elongation Mutants defective for amylose biosynthesis appear as red staining cell patches on solid nitrogen sprayed with iodine (Table 2). All mutants selected map to a single locus (STA2) for which 26 distinct UV and two insertional mutants were generated (Delrue et al., 1992; Maddelein et al., 1994). All mutants are characterized by an identical phenotype. They accumulate wild-type amounts of starch lacking both amylose and a moderately branched fraction that was named Ap II (amylopectin type II). This fraction is clearly related to the intermediate material that is

562 found in wild-type starches ofhigher plants. Moreover a small but significant difference was found in the long chain content ofthe major amylopectin fraction (Ap I). It was therefore concluded that STA2 controls both the biosynthesis ofamylose and the fine structure of amylopectin. Delrue et al. (1992) described two mutations that yielded allele-specific modifications of both the molecular mass of the major protein associated with the granule and of the for ADP-glucose of the major granule-bound starch synthase activity. Moreover the major 76 kDa protein bound to the granule displayed the classical N-terminal KTGGL sequence reported for starch synthases. Insertional mutants carrying the sta2-27::ARG7 allele were further shown to lack both GBSSI and the major 76 kDa starch-bound protein. Taken together these observations make a compelling case for STA2 being the structural gene for GBSSI. X-ray mutagenesis has yielded three distinct alleles of a single locus (STA3) that leads to 40 to 80% decrease in starch amounts (Fontaine et al., 1993). The residual starch is enriched in amylose and yields the characteristic olive-green phenotype of the highamylose class of mutants (Table 2). The overall defect thus consists ofa 90% decrease in amylopectin synthesis. The remaining amylopectin has a completely modified structure. The chain-length distribution of amylopectin shows a prominent maximum at d.p. 6 (degree of polymerization of 6) and a strong decrease in those chains ranging in size between 8 to 40 glucose residues (Fontaine et al., 1993). These make the bulk of the so-called amylopectin clusters (see Fig. 1 panels C and D). Since the clusters harbor the glucan double helices responsible for the crystallinity ofstarch we suspected that the basic organization ofamylopectin was altered in the mutants. Wide-angle X-ray diffraction experiments confirm these suspicions (Buléon et al., 1997). The overall crystallinity in sta3 mutants has virtually collapsed and switched to the B-type of crystalline lattice. This is correlated with an unusual melting behavior ofthe glucan double helices during DSC (Differential Scanning Calorimetry) experiments suggestive of the presence of shorter double helices. Moreover the shape and size distribution of the granules are altered. These profound and diverse modifications can be traced back to one single enzymological defect. All mutants were shown to lack SSSII (Soluble Starch Synthase II) activity. Fontaine et al. (1993) also

Steven G. Ball found a two-fold decrease of the activity in heterozygous mutants which suggests thatSTA3 could be the structural gene for the enzyme. Maddelein et al. (1994) conducted a detailed investigation of mutants defective at the STA3 and STA2 loci. Double mutant strains witnessed a collapse of starch biosynthesis despite the presence of fully active SSSI which accounts for half of the total soluble starch synthase activity. This suggests that SSSI cannot fully sustain amylopectin synthesis on its own. Moreover the 5% residual polysaccharide found in the double mutants was deeply modified. In fact, its structure does not truly qualify as starch but still adopts a dense macrogranular organization with altered shapes and sizes. The chain-length distribution of the double mutant polysaccharide still harbors the prominent maximum at d.p 6, but now consists solely of very short glucans. The X-ray diffraction pattern has switched from the B to the C type and has regained some crystallinity due to the absence of the amylose fraction. The results of Maddelein et al. (1994) also confirm the suspected function ofGBSSI in the synthesis of the long glucans associated with (and covalently bound to) amylopectin.

D. Mutants with Unidentified Enzymological Defects Another high amylose, X-ray generated mutant displayed a significant (from 40 to 60%) reduction in starch amount together with a strong and intriguing modification of the chain-length distribution of the residual starch (Libessart et al., 1995). The phenotype was clearly seen on storage starch but was hard to distinguish from wild-type transient starch when the polysaccharide was extracted from nitrogen-supplied mutant cells. Despite intensive efforts we were unable to find the enzymological defect in mutants of the STA4 locus. Further characterization will have to await tagging of STA4 by insertional mutagenesis. Equally intriguing is the recent finding of low starch mutants in the homothallic Chlamydomonas monoica reported by Rickoll et al. (1997). The authors convincingly showed the presence of a mutation in a single mendelian gene (STA-1, not to be confused with the STA1 locus from C. reinhardtii). Electron micrographs failed to reveal standard starch granules in the mutants. However the defective algae still contained a quite significant amount of starch with a more or less normal structure and buoyant density. We have found the major biosynthetic enzymes in

Chapter 29 Starch Biosynthesis Chlamydomonas monoica but have presently not uncovered the biochemical defect. We do not yet know how the material we have extracted relates to the amorphous material seen on the electron micrographs reported by Rickoll et al. (1997). If this material is starch then the C. monoica mutants will be the first oftheir kind reported in the plant-kingdom and therefore deserve particular future attention. STA-1could thus be quite uniquely involved in starch granule morphogenesis.

E. Mutants Defective for Pre-Amylopectin Trimming Insertional mutagenesis has generated a new class of mutants initially displaying both the low starch (staining yellow with iodine) and the high-amylose (staining olive-green with iodine) phenotypes. Mutants of STA7 (Mouille et al., 1996a) and STA8 (Mouille et al., 1996b) both lead to the production of a water-soluble polysaccharide (WSP). This fraction amounted to 5% ofwhat would have been the amount of starch in a wild-type strain. The water-soluble linked and polysaccharide consisted of branched glucans with a chain-length distribution, an iodine interaction ( of 490 nm) and NMR spectra analogous to animal, fungal or bacterial glycogen. The polysaccharide was thus named phytoglycogen by analogy with the maize su1 (sugary 1) mutation which leads to the accumulation of a similar fraction (Sumner and Somers, 1944; for review see Nelson and Pan, 1995). The phytoglycogens from sta7 and sta8 mutants differ with respect to their chain-length distributions. In addition to phytoglycogen, sta8 mutants accumulate high amylose (60%) granular starch amounting to 20% of the starch content from a wild-type strain. The high amylose phenotype is expected for a mutation acting predominantly on amylopectin synthesis. The sta7 mutants accumulated up to 0.5% granular material. This material consisted of virtually pure amylose with a maximum of 1% linkages. Again in sta7 mutants we suspect that the major defect remains in the amylopectin pathway and that the collapse of amylose synthesis comes as an aftermath of the disappearance of the minimum amount of granular structure required for GBSSI binding. The phenotype of sta8 mutants resembles that of the su1 mutation harbored by sweet corn while that of sta7 containing mutants remains to be described in higherplants. Sweet corn was shown to be defective

563 for a debranching enzyme activity (Pan and Nelson, 1984; James et al., 1995). Enzymological characterization ofthe 7 distinct sta7 alleles have uncovered a selective defect in a 88 kDa debranching enzyme that behaved mostly as an isoamylase and displayed very weak pullulanase activity. No other enzymological defect could be scored in the mutants thus establishing that polysaccharide debranching is mandatory to obtain significant amylopectin synthesis. While sta8 containing mutants displayed wild-type levels of the 88 kDa debranching enzyme, the purification behavior ofthe enzymes on columns was reported to be modified (Mouille et al., 1996b). This suggests that the product of the STA8 locus might be a subunit or other enzyme interacting with the 88 kDa debranching enzyme. The astonishing results obtained during the characterization of STA7 have laid the foundations of a model explaining the biosynthesis of plant amylopectin (Ball et al. 1996). In this model debranching enzymes are considered as trimming enzymes involved in ordering the branches produced in an intermediate of synthesis that was named preamylopectin (see below).

IV. A Model Explaining the Biogenesis of the Plant Starch Granule Until recently researchers have believed that the intrinsic properties of starch synthase and starch branching enzymes would be sufficient to explain starch granule biogenesis. Yet the branching enzymes expressed in vivo in E. coli, even in the presence of glycogen or starch synthases, are unable to yield the asymmetrical pattern ofbranches responsible for the continuous growth ofamylopectin (Guan et al., 1995). By opposition, a symmetrical pattern of branching will lead to an upper limit to the size that the unit granule will be able to reach before steric hindrance impairs further growth. Mathematical modeling predicts that the maximal diameter for a symmetrically branched polysaccharide with a 1 to 12 branching ratio would be 25 nm which is precisely what is measured for the unit particles of glycogen. There seems to have been a tremendous drive to overcome this limit in the plant kingdom. As usual this has not been in a logical fashion by de novo design of a novel optimal pathway, but rather as painting (the old canvas being cyanobacterial glycogen synthesis) by adding an additional color

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touch in the perpetual search for harmony. This was achieved as often is the case by putting old tools (debranching enzyme previously used in bacterial glycogen degradation) to new uses (glucan trimming) and subsequently optimized to retrieve the energy ‘lost’ by the ordering steps. The model proposed in Fig. 5 (Ball et al., 1996) for plant amylopectin synthesis explains how the asymmetrical distribution of branches amylopectin could be generated. The model is built by taking into account the phenotypes of the C. reinhardtii sta3 and sta7 mutations. Briefly, a previously synthesized amorphous lamella containing tightly spaced branches is being elongated by a specific soluble starch synthase (SSSII in Chlamydomonas). No branching occurs at this first step. We explained this (Ball et al., 1996) by assuming that branching is prevented because the branching enzymes need a minimal length for the glucan to adapt in their catalytic site. As soon as this length is attained, unordered branching and elongation (by SSSI) occur simultaneously. At the same time debranching enzymes will trim down the structure into the next amorphous lamella. This model was named the discontinuous synthesis model for amylopectin synthesis. It explains the amylopectin cluster constant size throughout the plant kingdom (Jenkins et al., 1993) and relates the size of the crystalline lamella and therefore of the amylopectin cluster to the minimal requirements of the branching enzymes. Quite interestingly it was subsequently found that maize BEI and BEII require respectively a minimum number of 16 and 12 glucose residues for branching

to occur. A glucan of 12 residues is of the very same size as that measured for the crystalline lamella of maize amylopectin. It is comforting to realize that mutants of maize defective for BEII and thus retaining only BEI witness an increase of the size of this lamella from that which characterizes a d.p. 12 glucan to that of d.p. 16 (Jenkins and Mc Donald, 1996). V. Future Prospects The reasons for the appearance of starch in photosynthetic eukaryotes remain mysterious. Did starch assume early on a function specific to these organisms that could not be taken in charge by glycogen? The size and density of starch granules make these compounds the fastest sedimenting structures. This property could have been put to use for gravity sensing. Indeed the starch-statolith theory of geotropism in higher plants states that it is sedimentation of the starch-filled amyloplast that induces the curving response in roots. Evidence for or against this hypothesis have accumulated over the years. Bean (1977) has noted a negative (positive being downwards) geotactic behavior in Chlamydomonas consisting of an upward swimming pattern in darkness. Since C. reinhardtii is not reported to use gas vesicles as floating devices, the alga must have some gravity sensing mechanism perhaps related to the one used by land plants for oriented cell growth. It will thus be of great interest to examine the geotactic behavior of the starch defective mutants.

Chapter 29 Starch Biosynthesis However, this study will be complicated by the paralysis noted for the sta6-1::ARG7 carrying starchless mutant. An inverse relation was indeed observed between chemical energy storage and cell motility. Paralyzed mutants containing defects in PF-1, PF-2, PF-7 and PF-18, were shown to accumulate a substantially higher level of starch (Hamilton et al., 1992). Whether paralysis comes as a direct consequence of sta6-1::ARG7 will need further work to prove that insertional mutagenesis has not lead to the deletion of a neighboring PF cistron. The most exciting short-term prospects will definitely consist of unraveling the glucan trimming pathway involved in amylopectin synthesis. We strongly believe that more genes involved in this pathway will be identified in Chlamydomonas. Finally the iodine-spray screening procedure can also be adapted to select for strains carrying mutations for starch overproduction (STO) or breakdown (STB). Characterization of such loci has only just begun.

Acknowledgments Research is a collective human effort. The author would thus like to thank all researchers mentioned in this work for their contribution to the understanding of starch biosynthesis in Chlamydomonas. The author would also like to acknowledge the help and insights of Dr. Uwe Klein and Karen Van Winkle-Swift for disclosing and discussing their unpublished results.

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565 Buléon A, Gallant DJ, Bouchet B, Mouille G, D’Hulst C, Kossmann J and Ball S (1997) Starches from A to C: Chlamydomonas reinhardtii as a model microbial system to investigate the biosynthesis of the plant amylopectin crystal. Plant Physiol 115: 949–957 Caspar T (1994) Genetic dissection of the biosynthesis, degradation, and biological functions ofstarch. In: Meyerowitz EM and Sommerville C (eds). Arabidopsis, pp. 913–936, Monograph 27. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York Caspar T, Huber SC and Somerville C (1985) Alterations in growth, photosynthesis, and respiration in a starchless mutant of Arabidopsis thaliana (L.) deficient in chloroplast phosphoglucomutase activity. Plant Physiol 79: 11–17 Craigie JS ( 1974) Storage products. In: Stewart WDP (ed) Algal Physiology and Biochemistry, pp 206–235. Blackwells, Oxford Delrue B, Fontaine T, Routier F, Decq A, Wieruszeski JM, van den Koornhuyse N, Maddelein, ML, Fournet B and Ball S (1992) Waxy Chlamydomonas reinhardtii: Monocellular algal mutants defective in amylose biosynthesis and granule-bound starch synthase accumulate a structurally modified amylopectin. J Bacteriol 174: 3612–3620 Fletcher V and Cirino R (1992) The effect of growth regimen, heat shock and intracellular starch storage on the mating efficiency of Chlamydomonas reinhardtii. Br Phycol 27: 3–9 Fontaine T, D’Hulst C, Maddelein ML, Routier F, MariannePepin T, Decq A, Wieruszeski, JM, Delrue B, van den Koornhuyse N, Bossu JP, Fournet B and Ball SG (1993) Toward an understanding of the biogenesis of the starch granule. Evidence that Chlamydomonas soluble starch synthase II controls the synthesis of intermediate size glucans of amylopectin. J Biol Chem 268: 16223–16230 Gfeller R and Gibbs (1984) Fermentative metabolism of Chlamydomonas reinhardtii. I. Analysis of fermentative products from starch in dark and light. Plant Physiol 75: 212– 218 Guan HP, Kuriki T, Sivak M and Preiss J (1995) Maize branching enzyme catalyzes synthesis of glycogen-like polysaccharide in glgB-deficient Escherichia coli. Proc Natl Acad Sci USA 92: 964–967 H a m i l t o n BS, Nakamura K and Roncari DAK (1992) Accumulation of starch in Chlamydomonas reinhardtii flagellar mutants. Biochem Cell Biol 70: 255–258 Hanson KR and McHale NA (1988) A starchless mutant of Nicotiana sylvestris containing a modified plastid phosphoglucomutase. Plant Physiol 88: 838–844 Hirst E, Manners D, and Pennie IR(1972) Part XXI – The molecular structure of starch-type polysaccharide from Haematococcus pluvialis and Tetraselmis carteriiformis. Carbohydr Res 22: 5–11 Iglesias AA, Charng YY, B a l l S and Preiss J (1994) Characterization of the kinetic, regulatory and structural properties of ADP-glucose pyrophosphorylase from Chlamy­ domonas reinhardtii. Plant Physiol 104: 1287–1294 Imberty A, Buléon A, Tran V and Pérez S (1991) Recent advances in knowledge of starch structure. Starch/Stärke 43: 375–384 James MG, Robertson DS and Meyers AM (1995) Characterization of the maize gene sugary, a determinant of starch composition in kernels. Plant Cell 7: 417–429 Jenkins PJ and Donald AM (1995) The influence of amylose on starch granule structure. Int J Biol Macromol 17: 315–321

566 Jenkins PJ, Cameron RE and Donald AM (1993) A universal feature in the starch granules from different botanical sources. Starch/Stärke 45: 417–420 Klein U (1986) Compartmentation of glycolysis and of the oxidative pentose-phosphate pathway in Chlamydomonas reinhardtii. Planta 167: 81–86 Klein U (1987) Intracellular carbon partitioning in Chlamy­ domonas reinhardtii. Plant Physiol 85: 892–897 Klein U and Betz A (1978) Fermentative metabolism of hydrogenevolving Chlamydomonas moewusii. Plant Physiol 61: 953– 956 Klein U, Chen C andGibbs M (1983) Photosynthetic properties ofchloroplasts from Chlamydomonas reinhardtii. Plant Physiol 72: 488–491 Klöck G and Kreuzberg K (1987) Sn-glycerol-3-phosphate is a product of starch degradation in isolated chloroplasts of Chlamydomonas reinhardtii. Z Naturforsch 42c: 567–569 Klöck G and Kreuzberg K (1991) Compartmented metabolite pools in protoplasts from the green alga Chlamydomonas reinhardtii: Changes after the transition from aerobiosis to anaerobiosis in the dark. Biochim Biophys Acta 1073: 410415 Klöck G, Sueltmeyer DF, Fock HP and Kreuzberg K (1989) Gas exchange in intact isolated chloroplasts from Chlamydomonas reinhardtii during starch degradation in the dark. Physiol Plant 75: 109–113 Kombrink E and Wöber G (1980) Identification and subcellular localization of starch-metabolizing enzymes in the green alga Dunaliella marina. Planta 149: 130–137 Kreuzberg K (1984) Starch formation via a formate producing pathway in Chlamydomonas reinhardtii, Chlorogonium elongatum and Chlorella fusca. Physiol Plant 61: 87–94 Kreuzberg K and Martin W (1984) Oscillatory starch degradation and fermentation in the green alga Chlamydomonas reinhardtii. Biochim Biophys Acta 799: 291–297 Kreuzberg K, Klöck G and Grobheiser D (1987) Subcellular distribution of pyruvate-degrading enzymes in Chlamydomonas reinhardtii studied by an improved protoplast fractionation procedure. Physiol Plant 69: 481–488 Kruger NJ, Bulpin PV and ap Rees T (1983) The extent of starch degradation in the light in pea leaves. Planta 157: 271–273 Kuchitsu K, Tsuzuki M and Miyachi S (1988) Changes of starch localization within the chloroplast induced by changes in concentration during growth of Chlamydomonas reinhardtii: Independent regulation of pyrenoid starch and stroma starch. Plant Cell Physiol 29: 1269–1278 Levi C and Gibbs M (1984) Starch degradation in synchronously grown Chlamydomonas reinhardtii and characterization of the amylase. Plant Physiol 74: 459–463 Libessart N, Maddelein ML, van den Koornhuyse N, Decq A, Delrue B and Ball SG (1995) Storage, photosynthesis and growth: The conditional nature of mutations affecting starch synthesis and structure in Chlamydomonas reinhardtii. Plant Cell 7: 1117–1127 Lin TP, Caspar T, Somerville C and Preiss J (1988) Isolation and characterization of a starchless mutant of Arabidopsis thaliana (L.) Heynh. lacking ADP-glucose pyrophosphorylase activity. Plant Physiol 86: 1131–1135 Maddelein ML, Libessart N, Bellanger F, Delrue B, D’Hulst C, van den Koornhuyse N, Fontaine T, Wieruszeski JM, Decq A and Ball SG (1994) Toward an understanding of the biogenesis

Steven G. Ball of the starch granule: Determination of granule-bound and soluble starch synthase functions in amylopectin synthesis. J Biol Chem 269: 25150–25157 Manners DJ (1989) Recent developments in our understanding of amylopectin structure. Carbohydr Polymers 11: 87–112 Meeuse BJD (1962) Storage products. In: Lewin RA (ed) Physiology and Biochemistry of Algae, pp 289–311, Academic Press, New York and London Meeuse BJD and Kreger DR (1959) X-ray diffraction of algal starches. Biochim Biophys Acta 35: 26-30 Meeuse BJD, Andries and Wood (1960) Floridean starch. J Exp Bot 11: 129–140 Mouille G, Maddelein ML, Libessart N, Talaga P, Decq A, Delrue B and Ball S (1996a) Phytoglycogen processing: A mandatory step for starch biosynthesis in plants. Plant Cell 8: 1353–1366 Mouille G, Colleoni C, Maddelein ML, Libessart N, Decq A, Delrue B and Ball S (1996b) Glucan trimming a novel mechanism that explains the asymmetric distribution of branches in amylopectin. In: Nakamura Y (ed) Regulation and Manipulation of Starch and Sucrose Metabolism in Plants, pp 27–32. National Institute of Agrobiological Resources, Tsukuba. Nakamura Y and Imamura M (1983) Characteristics of a glucan phosphorylase from Chlorella vulgaris. Phytochemistry 22: 835–840 Nakamura Y and Imamura M (198 5) Regulation ofADP-glucose pyrophosphorylase from Chlorella vulgaris. Plant Physiol 78: 601–605 Nelson OE and Pan D (1995) Starch synthesis in maize endosperms. Annu Rev Plant Physiol Plant Mol Biol 46:475– 496 Olaitan SA and Nothcote DH (1962) Polysaccharides of Chlorella pyrenoidosa. Biochem J. 82: 509–519 Pan D and Nelson OE (1984) A debranching enzyme deficiency in endosperms of the Sugary-1 mutants of maize. Plant Physiol 74: 324–328 Pino-Plumed MD, Villarejo A, de los Rios A, Garcia-Reina G concentrating mechanism and Ramazano v Z (1996) The in a starchless mutant of the green unicellular alga Chlorella pyrenoidosa. Planta 200: 28–31 Preiss J and Greenberg E (1967) Biosynthesis ofstarch in Chlorella pyrenoidosa: I purification and properties of the adenosine diphosphoglucose: glucosyl transferase from Chlorella. Arch Biochem Biophys 118: 702–708 Preiss J and Sivak MN (1996) Starch synthesis in sinks and sources. In: Zamski E and Schaffer AA (eds) Photoassimilate Distribution in Plants and Crops: Source-Sink Relationships, pp 63–96. Marcel Dekker Inc., New York Ramazanov Z, Rawat M, Henk MC, Mason CB, Matthews SW concentrating and Moroney JV (1994) The induction of the mechanism is correlated with the formation of the starch sheath around the pyrenoid of Chlamydomonas reinhardtii. Planta 195: 210–216 Rickoll W, Rehkopf D, Dunn C, Van Winkle-Swift, K (1998) The sta-1 mutation prevents assembly of starch granules in nitrogenstarved cells and serves as a useful morphological marker during sexual reproduction in Chlamydomonas monoica (Chlorophyceae). J Phycol 34: 147–151 Robin JP, Mercier C, Charbonnière R and Guilbot A (1974) Lintnerized starches. Gel filtration and enzymatic studies of

Chapter 29 Starch Biosynthesis insoluble residues from prolonged acid treatment of potato starch. Cereal Chem 51: 389–406 Sanwal GG and Preiss J (1967) Biosynthesis of starch in Chlorella pyrenoidosa: II regulation of ATP: glucose-1-phosphate adenyl transferase (ADP-glucose pyrophosphorylase) by inorganic phosphate and 3-phosphoglycerate. Arch Biochem Biophys 119: 454–469 Singh KK, Chen C, Epstein DK and Gibbs M (1993) Respiration of sugars in spinach (Spinacia oleracea), maize (Zea mays), and Chlamydomonas reinhardtii F-60 chloroplasts with emphasis on the hexose kinases. Plant Physiol 102: 587–593 Smith A, Denyer K and Martin C (1997) The synthesis of the starch granule. Ann Rev Plant Physiol Plant Mol Biol 48: 67– 87 Spudich JL and Sager R (1980) Regulation of the Chlamydomonas cell cycle by light and dark. J Cell Biol 85: 136–145 Steup M (1988) Starch degradation. In: Preiss J (ed) The Biochemistry of Plants. A Comprehensive Treatise, Vol 14, pp 255–296. Academic Press, San Diego Stitt M and Heldt HW (1981) Simultaneous synthesis and degradation of starch in spinach chloroplasts in the light. Biochim Biophys Acta 638: 1–11 Sumner JB and Somers GF (1944) The water soluble polysaccharide of sweet corn. Arch Biochem 4: 4–7

567 Süss KH, Prokhorenko I and Adler K (1995) In situ association of Calvin cycle enzymes, ribulose-l,5-biphosphate carboxylase/ oxygenase, ferredoxin reductase, and nitrite reductase with thylakoid and pyrenoid membranes of Chlamydomonas reinhardtii chloroplasts as revealed by immunoelectron microscopy. Plant Physiol 107: 1387–1397 Van den Koornhuyse N, Libessart N, Delrue B, Zabawinski C, Decq A, Iglesias A, Preiss J and Ball S (1996) Control of starch composition and structure through substrate supply in the monocellular alga Chlamydomonas reinhardtii. J Biol Chem 271: 16281–16287 Villarejo A, Martinez F, Pino-Plumed MD and Ramazanov Z concentrating mechanism in (1996) The induction of the a starchless mutant of Chlamydomonas reinhardtii. Physiol Plant 99: 293–301 Yu S and Pedersen M (1993) Alpha-1,4-glucan lyase, a new class of starch/glycogen degrading enzyme. I. Efficient purification and characterization from red seaweeds. Biochim Biophys Acta 1156: 313-320 Yu S, Chrisyensen TM, Kragh KM, Bojsen K and Marcussen J (1997) Efficient purification, characterization and partial amino acid sequencing of two alpha-1,4-glucan lyases from fungi. Biochim Biophys Acta 1339: 311-320

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Chapter 30

State Transition and Photoinhibition

Nir Keren and Itzhak Ohad

Minerva, Avron Even-Ari Center for Photosynthesis Research,

Silberman Institute of Life Sciences, Department of Biological Chemistry,

The Hebrew University of Jerusalem, Jerusalem, 91904 Israel

Summary I. Introduction A. Photosynthetic Electron Flow in C. reinhardtii B. Light Energy Absorption and Dissipation C. Chlorophyll a Fluorescence Measurements II. State Transition: The Phenomenon A. Relation Between the Plastoquinone Pool Redox Balance and State Transitions B. Redox Related Activation of the LHCII-Kinase: Role of Cytochrome bf C. Phosphorylation-Dependent Reversible Association of LHCII with PS II III . Light Stress: Photoinhibition and Recovery A. Mechanisms of PS II Photoinactivation 1. Acceptor-Side Photoinhibition 2. Photoinactivation by Limiting Light Intensities 3. Donor-Side Photoinhibition 4. UV Induced Photodamage B. D1 Cleavage as a Result of the Different Photoinhibitory Treatments 1. Role of the Binding Site in D1 Protein Degradation in vivo C. Degradation of Other PS II Proteins 1. The Nature of the Protease D. The Recovery Process 1. Synthesis and Reassembly of PS II Proteins 2. The Role of Chlorophyll a and

in the Reassembly of PS II IV. Concluding Remarks and Perspectives Acknowledgments References

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Summary Optimal utilization of absorbed light energy and avoidance of oxidative damage induced by excessive excitation (photoinhibition) constitute a major problem for photosynthetic oxygen evolving cells. Adaptation to transient changes in light absorption and energy utilization is achieved by regulation of photochemistry, and both radiative and nonradiative energy dissipation. Balancing the absorbed energy distribution between PS II and PS I is achieved by redox dependent, reversible phosphorylation of the LHCII antennae, resulting in their reversible coupling with the photosystem cores (state transition). This process optimizes linear electron flow and the cyclic electron flow dependent ATP synthesis. Back electron flow, recombination of the PS II primary charge separated pair, and formation of and occur at all light intensities. Generation of active oxygen species causes oxidative damage and induces degradation of PS II-proteins, particularly the J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 569–596. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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PS II-D1 protein. Excessive excitation induces a stepwise inactivation of PS II, affecting primarily its acceptor side components, thereby promoting charge recombination and oxidative damage. Infrequent excitation of PS II (‘low light’), promotes charge recombination driven by the equilibrium between the oxidized states In this process of the PS II donor side manganese cluster and the reduced semiquinone acceptor are regenerated and their recombination results in oxidative damage exposing the PS II proteins to proteolytic cleavage. De novo protein synthesis and reassembly of PS II compensate for photoinactivation and degradation of the D1 protein. This process involves dissociation of the residual PS II core proteins from the antenna complex, lateral migration from the appressed to the non-appressed lamellae and reassembly with precursor pD1 protein, newly synthesized by polyribosomes bound to non-appressed lamellae. Processing of pD1, light dependent reactivation of the donor side Mn cluster, lateral migration of the reactivated PS II core to the appressed lamellae, and its reassociation with LHCII regenerates active PS II units and complete the PS II photoinactivation and repair cycle. Thus, photoinactivation and repair of PS II activity are regulated at the metabolic and molecular levels and are modulated by environmental changes.

I. Introduction The ongoing effort to understand, in molecular terms, the response of the photosynthetic eukaryotic cell to a wide range of light intensities under various ambient temperature and nutrient conditions involves extended physiological, biochemical, biophysical and molecular studies, which have been reviewed periodically in the past (Barber and Andersson, 1992; Prasil et al., 1992; Ohad et al., 1994; Gal et al., 1997). Many of these studies were reinforced by using C. reinhardtii as a model organism. The classical genetic approach, combined with site-directed mutagenesis and transformation techniques allows one to obtain engineered mutations in the chloroplast encoded genes (see Chapters 4, Kindle; 8, Goldschmidt-Clermont). Thus C. reinhardtii provides an ideal vehicle for the study of the mechanism of cellular responses to changes in the ambient light intensity and quality. These studies address, among others, the following questions: what are the primary targets for light stress (i.e. the

chlorophyll-protein complexes) and what are the resulting changes in their properties; what is the machinery responsible for alleviation of excess energization; what is the mechanism of light-induced damage and repair; how are the irreversibly altered proteins or protein complexes specifically degraded and replaced; how are these processes regulated and at what levels of macromolecular metabolism, protein synthesis, processing and assembly of the membrane integral protein-pigment complexes. Photosynthetic cells can respond to short term changes in the environmental light conditions or adapt to long term variations (Melis et al., 1996). The aim of this chapter is to review in general lines the knowledge obtained by analyzing the response of the unicellular green alga C. reinhardtii to short term, limiting or excessive illumination, with particular emphasis on the modulation of energy distribution between the two photosystems (state transition) and the process of photoinhibition and recovery of photosynthetic activity.

A. Photosynthetic Electron Flow in C. reinhardtii Abbreviations: – singlet and triplet chlorophyll; DBMIB – 2,5–dibromo–3–methyl–6–isopropyl–p–benzoquinone; DCIP – 2,3 dichlorophenol indophenol; DCMU – diuron, 3–(3´4–dichlorophenyl)–1,1–dimethyl urea; DMBQ – 2,6-dimethyl benzoquinone; DPC – 1,5-diphenyl carbazide; – singlet and triplet oxygen respectively; OEE – the oxygen evolving enhancer proteins; – oxidized and excited sate of the primary electron donor of RCII, respectively; Pheo – pheophytin, the primary electron acceptor of RCII; – the first and second electron acceptors of PS II; RCII – photochemical reaction center II; – mixed RCII populations containing Mn clusters that have lost either 1 and 2, or 2 and 3 electrons respectively; –oxidized states of the PS II Mn cluster that have lost either2 or 3 electrons respectively; Yz – tyrosine 161 of the D1 protein, the electron donor to

The composition of the photosynthetic apparatus in C. reinhardtii in terms of the major thylakoid membrane complexes, their subunit composition, function and macro-molecular organization into appressed and non-appressed membrane domains, is comparable to that of the green plant chloroplast (Ohad et al., 1967; Harris, 1989; Chapter 14, Olive and Wollman). The so-called ‘Z- scheme’ that describes the path of electron flow from the water oxidation complex associated with Photosystem II (Chapter 16, Ruffle and Sayre) via the plastoquinone pool, cytochrome bf complex (Chapter 24, Wollman),

Chapter 30 State Transition and Photoinhibition plastocyanin (Chapter 32, Merchant), Photosystem I to NADP reduction (Chapter 17, Webber and Bingham; Nechushtai et al., 1996), provides an accepted and well-demonstrated basis for the bioenergetics of photosynthesis in this organism. The ATP-synthase of C. reinhardtii is similar to that of higher plants (Chapter 25, Strotmann et al.). Genetic and biochemical evidence indicate the presence of an additional membrane complex, the NADPH-dehydrogenase accounting for the phenomenon ofchlororespiration in C. reinhardtii (Godde and Trebst, 1980; Godde, 1982; Chapter 18, Redding and Peltier). The light harvesting antennae system of C. rein­ hardtii PS II has been characterized (Bassi and Wollman, 1991), and includes the chlorophyll a/b LHCII complex comprised offive major polypeptides (Bassi and Wollman, 1991). The antenna also include the CP43 and CP47 chlorophyll a binding proteins of the PS II core. PS I harvests light through the chlorophyll bound to the PsaA and PsaB core proteins and the Chl a/b LHC-I antenna (Chapter 17, Webber and Bingham; Ish-Shalom and Ohad, 1983; Nechushtai et al., 1996).

B. Light Energy Absorption and Dissipation The absorbed light energy is used for photochemistry or dissipated via heat or fluorescence emission processes (Govindjee, 1995; Lavergne and Briantais, 1996). The rate of the photochemical activity depends on the light intensity as well as on the internal state of the intermediate electron carrier pools and the final acceptors. Therefore, the term ‘excess light’, used to describe excitation overload of the photosynthetic apparatus, is a relative term that should be discussed in conjunction with the algal metabolic state. Non-radiative dissipation (heat emission) can be performed by both reaction center and antenna complexes, and the rate of this process can be modulated by transient changes in the membrane carotenoid composition due to de-epoxidation of and increase in the xanthophyll content of the membranes (Demmig-Adams and Adams, 1996). For efficient utilization of the absorbed light, both photosystems should be excited to generate equal rates of electron flow and thus balance the reduction of the PQ pool with its oxidation. Increase nm, absorbed primarily in the light intensity at by the PS II light harvesting complex LHCII,

571 containing a large relative amount ofChl b, or increase in the light intensity absorbed preferentially by PS I to 740 nm will create an imbalance resulting i in the dissipation of the excess absorbed light energy by the respective photosystem. Imbalance between the excitation of the photosystems may occur not only as result of the changes in the light quality but also due to changes in the availability of electron acceptors for each of the two photosystems. Dissipation of excess energy by increase in the fluorescence emission occurs mostly in PS II. At cryogenic temperatures, fluorescence emission increases considerably also in PS I (Gershoni et al., 1982). Light, while providing the energy required to extract electrons from water, to generate ATP and reduce carbon and nitrogen, also activates regulatory mechanisms for the optimization of the energy utilization and protective mechanisms aimed to alleviate over-excitation even at relatively low photon flux densities. The term ‘fluorescence quenching’ is widely used to indicate that processes such as photochemistry or heat dissipation compete with fluorescence emission. Energy dissipation in the reaction centers and antenna by mechanisms that are not related to the photochemical processes, are generally termed non-photochemical quenching (Krause and Weis, 1991; Lavergne i and Briantais, 1996). The fluorescence yield can be quenched (i. e. increase in heat dissipation) by energization of the membrane due to increase in Quenching of antenna fluorescence is based on reversible covalent modifications of the thylakoid membrane carotenoids, whereby consecutive de-derived violaxanthin epoxidations of the increase the content of antheraxanthin and zeaxanthin (xanthophyll cycle). This reductive deepoxidation process requires NADPH (DemmigAdams and Adams, 1996) and is triggered by an induced by excessive illumincrease in the ination (Endo and Asada, 1996). The antenna xanthophylls participate in the quenching ofantenna chlorophyll triplet states. In addition, antheraxanthin and zeaxanthin enhance thermal dissipation of the excited singlet state of antenna chlorophylls (Crofts andYerkes, 1994; Gilmore et al., 1995; Horton et al., 1996). The thermal dissipation of the absorbed light energy within the antenna reduces energy transfer to PS II and lowers the fluorescence emission as well as the formation of harmful triplet chlorophyll in RCII (Vass and Styring, 1993).

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Grossman and coworkers have isolated a nuclear C. reinhardtii mutant, npq1 and used the lor1 mutant to construct a double mutant npq1/lor1. The npq1 mutant is impaired in the de-epoxidation of violaxanthin to antheraxanthin and zeaxanthin. The lor1 mutant is impaired in the conversion of lycopene to -carotene, lutein and loroxanthin. Each of the above mutants could grow phototrophically and did not exhibit light sensitivity, although both exhibited a reduction in the NPQ response. However, the npq1/ lor1 double mutant was light sensitive and exhibited a drastically reduced NPQ (Niyogi et al., 1997a,b). These findings represent direct evidence for the involvement of the xanthophylls in the non-radiative dissipation of excess energy as well as the fact that xanthophylls derived from -carotene can substitute for those derived from -carotene. The enzymes involved in this process are membrane bound and can be activated in isolated thylakoid membranes, in the presence of NADPH. The possibility that the enzymes are closely associated with the PS II antenna has been proposed for vascular plants (Gruszecki and Krupa, 1993). Presently it is not known whether the xanthophyll cycle enzymes in C. reinhardtii are associated with a specific membrane complex. The term also includes energy dissipation in the

Nir Keren and Itzhak Ohad

reaction centers. The nature ofthis process is thought to be related to either pH dependent release of ions from the membrane-bound water-splitting complex, (Krieger and Weis, 1992) or to the generation of centers (Schreiber and Neubauer, 1990). The fluorescence yield is also modulated by changes in energy transfer from the LHCII-antenna to the PS II reaction centers and redistribution of absorbed energy between the two photosystems. The dissociation of the LHCII-antennae due to phosphorylation of the LHCII proteins diminishes the PS II absorption cross-section and thus reduces the total energy absorbed and the ensuing fluorescence. This process is termed ‘state transition’ and the lowering ofthe PS II fluorescence emission rate in this case is referred to as (Horton and Hague, 1988) This phenomenon will be dealt with in more detail in Section II. Distinction between the different mechanisms of energy dissipation can be monitored by kinetic fluorescence measurements. High energy quenching ( ) in the reaction centers is a fast process occurring in the millisecond to second time range. The xanthophyll cycle and state transition are enzymatic processes which occur in the minute time range. The

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phosphorylation of LHCII related to the state transition in C. reinhardtii cells and thylakoids is achieved in about 10–15 min of light exposure (Owens and Ohad, 1982). It is important to note that nonradiative processes account for most of the energy dissipation. Fluorescence quenching by membrane energization ( ) has a smaller contribution to light energy management, relative to the state transition component, in algae as compared to higher plants (Falkowski et al., 1986; Schreiber et al., 1995).

C. Chlorophyll a Fluorescence Measurements Fluorescence dynamics phenomena can be monitored by measurements ofchlorophyll fluorescence in vivo, a fast, non-invasive method for evaluation of photosynthetic electron transfer and energy distribution. In this section the main methods used for photosynthetic fluorescence measurements and the parameters derived from them will be described briefly. For information on the in vivo measurements of photosynthetic activity, see Chapter 22 (Joliot et al.). Measurements of fluorescence rise kinetics (fluorescence induction) are performed using a ) and continuous weak actinic light ( measuring the emitted fluorescence above The fluorescence induced by the rapid onset of the continuous illumination preceded by dark-adaptation of intact cells (time required for the light ‘on’ condition, about 1 ms) is shown in Fig. lA.A fast rise in the fluorescence parallels the rise of the actinic light intensity with the opening of the shutter. This initial fluorescence measured immediately (>1 ms) after the complete opening of the shutter, is an estimate of the fluorescence level. This fluorescence is ascribed to competition with energy trapping by active centers as well as to active PS II centers that are closed due to the presence of in its reduced state in darkness and to irreversibly nonfunctional centers (Lavergne and Briantais, 1996). The level of measured as in Fig. 1A may be higher (<10%) than the true level due to closure of some of the PS II centers by the increasing actinic light intensity as the shutter opens. For further details on the measurement of see Strasser et al. (1995) and Srivastava et al. (1995). The transient fluorescence intensity changes that follow the fluorescence are termed the Kautsky effect (Kautsky and Hirsch, 1931; Govindjee, 1995). In vivo, the fluorescence intensity rises (0.1–0.5 s) to a higher level that

573 depends on the actinic light intensity and decreases afterward to a steady state level, Fs. The transient fluorescence intensity rise, as shown in Fig. 1 A, can be observed only after dark adaptation for several minutes and may be related to the initiation of the carbon fixation activity. It is possible that the lightinduced activation of the ferredoxin-thioredoxn dependent Calvin cycle enzymes (see also Chapter 26, Jaquot et al.) and of the RuBP-carboxylase may contribute to the lag in the induction of the carbon fixation process (Chapter 28, Spalding). If cells that have reached the steady state of fluorescence are brought back to darkness and immediately re-exposed to the actinic light, the initial transient fluorescence rise is abolished. This transient can be re-established only after appropriate dark re-adaptation lowering the plastid level ofATP and thus lowering the carbon fixation efficiency. The steady-state fluorescence level is related to the steady-state of electron flow and represents the ratio of closed to open reaction centers. To obtain the maximal level of fluorescence (Fm), using the above experimental approach one which can be has to cause complete reduction of achieved by addition of an inhibitor of to electron flow such as DCMU or Atrazine (Fig. 1 A) The in vivo monitoring of the fluorescence parameters described above can also be achieved by pulsed amplitude-modulated measurements (PAM) (Schreiber et al., 1995). The system uses a modulated weak light beam to excite the sample and the resulting modulated fluorescence signal in the absence of, or following an increase in light excitation of the system by an actinic non-modulated light beam is recorded (Fig. 1B). This configuration permits superimposition of continuous blue, red or far red light excitation of the sample, inducing reduction or oxidation of the plastoquinone pool accordingly. This allows the measurement of the state transition phenomenon as well as all other fluorescence quenching states under a variety of illumination intensities, and excitation as time intervals. Furthermore, the level of measured using this method is more accurate due to the very low intensity ofthe measuring light that does not cause closure of PS II centers. The Fm level can be measured by a short and intense light pulse (millisecond to second range) that causes transient closure of all the reaction centers. In this case there is no need to add inhibitors of electron flow. Thus, repetition of the measurements and alteration of the experimental conditions on the same cell suspension is possible. High light pulses generate a maximal

574 fluorescence level that depends on the sum of the fluorescence quenching processes at the particular measuring time. The levels of fluorescence intensity measured by the above techniques are relative and depend on light intensity, chlorophyll concentration, geometry of the measuring apparatus and the detecting system. Therefore it is essential to normalize the results of fluorescence experiments to a constant, stable parameter. A widely used term for the description of fluorescence measurements data is the variable fluorescence parameter, Fv (Fv = Fm – Fo). This value is usually normalized to the Fo or Fm levels. The value for is calculated as Fm – Fs and the non photochemical quenching as the difference of the Fm level of dark adapted cells and the Fm level after exposure of the cells to actnic light for several minutes (Fig. 1B). Prolonged illumination permits the activation of the xanthophyll cycle and a further decrease in the level of Fo and Fm ( ) can be observed. One should note that the level of Fo and Fm may undergo changes in cells exposed to photoinhibitory light (Section III) and thus the calculated values of Fv/Fm or Fv/Fo under such conditions (widely used in this research field) can not be simply interpreted in terms of damage and its localization by comparing them to those of control cells. To identify the extent and sequence of the damaging events, it is useful to relate also to the actual measured values of the Fo and Fm parameters as well as to additional measurements of PS II partial electron transfer reactions.

II. State Transition: The Phenomenon State transition, first described by Bonaventura and Myers (1969) and Murata (1969) and its physiological implications in vascular plants have been discussed in several reviews (Fork and Satoh, 1986; Bennett, 1991; Allen, 1992; Andersson and Barber, 1994; Gal et al., 1997). In this section we shall briefly describe the state transition phenomena as they occur in eukaryotic photosynthetic organisms harboring Chl a/b antenna systems (Fig. 2). Cells in which the LHCII-antenna is associated with the PS II complex and transfers energy of the absorbed light to reaction center II, are referred to as cells in state I and are characterized by a high fluorescence emission level. Cells in which LHCII is dissociated from the PS II complex, referred to as

Nir Keren and Itzhak Ohad cells in state II, emit a lower fluorescence from the PS II complex. Transition from state I to state II occurs when the PS II mobile antenna dissociates from the complex following phosphorylation of the LHCII polypeptides. The activation of the protein kinase involved in the phosphorylation of LHCII is regulated by the redox state of the plastoquinone pool. Thus, cells in state I, exposed to light preferentially exciting theLHCII-PSIIcomplex (light ), which reduces the plastoquinone II, pool, will change to the state II condition. Alternatively, cells in state II in which LHCII is phosphorylated and dissociated from PS II complex, will change to state I when exposed to light ), preferentially exciting PS I (light I, which causes oxidation of the plastoquinol pool, deactivation of the LHCII-kinase and dephosphorylation of LHCII by a constitutively active phosphatase. Reassociation of the LHCII with PS II increases the absorption cross section of the PS II complex and thus the PS II fluorescence (Fig. 2). Photoacoustic studies demonstrate that the dissociated phospho-LHCII antenna may transfer energy to PS I following lateral migration from the appressed to the non-appressed thylakoid membrane domains (Delosme et al., 1996). Further support for an increase in the antenna size of PS I, following transition from state I to state II, can be found in the observation that the fluorescence emission of PS I increases relative to that of PS II, as tested by fluorescence measurements at 77K (Wollman and Delepelaire, 1984). Dynamic changes in electron flow from PS II to the final electron acceptor sink, at any given time, cause similarly related changes in the level of PS IIemitted steady state fluorescence and may serve as an ‘on-line’ monitor of whole chain electron flow in vivo. A rate-limiting step or a block of electron flow occurring at any point in the electron transfer chain past PS II, will induce the maximal fluorescence level attainable at that particular light intensity. Addition ofelectron flow inhibitors binding at the or site of PS II preventing the oxidation of inhibitors of the cytochrome bf complex such as DBMIB (Bennett et al., 1988; Gal et al., 1988), cause reduction of all the electron acceptors up to the inhibited step and an increase in the fluorescence to the Fm level. The Fm level will be achieved also if plastoquinol oxidation is blocked by absence of electron acceptors for the oxidation of NADPH. In C. reinhardtii, the mobile antenna is considerably

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State Transition and Photoinhibition

larger than that of vascular plants (Bassi and Wollman, 1991). The phosphorylation pattern of the LHCII polypeptides involved in state transition in C. reinhardtii differs form that ofthe vascular plants. In vascular plants, state transition is related to the phosphorylation of the 25 kDa and 27 kDa polypeptides equivalent to p11 and p16 polypeptides in C. reinhardtii. The polypeptides equivalent to CP29 and CP26 (p9 and p10 according to the above authors’ nomenclature) are phosphorylated in relation to the transition from state I to state II in C reinhardtii but not in the vascular plants.

A. Relation Between the Plastoquinone Pool Redox Balance and State Transitions Among all electron carriers forming the electron transfer chain, most are found in stoichiometric relations or close to that value, except for plastoquinone which appears in a ratio of 8–10 molecules per chain (Lavergne et al., 1992). The plastoquinone pool is a pivotal point in electron transfer dynamics,

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its redox state being directly related to the ratio of PS II/PS I electron flow. Accordingly, the state transition dynamics reflect changes in the redox state of the plastoquinone pool. It follows that any condition that affects the redox state of the plastoquinone pool, will alter the Chl fluorescence level. Evidence for this concept can be found in measurements of the fluorescence rise kinetics and state transition phenomena in mutants impaired in the activity of cytochrome bf, plastocyanin or PS I (Lemaire et al., 1986; Shochat et al., 1990; Gong and Ohad, 1991; Topf et al., 1992) or following addition of inhibitors of electron flow through cytochrome bf (Frid et al., 1992). Decreasing the availability of the terminal or which serve electron acceptors such as as carbon and nitrogen source for C. reinhardtii growth, respectively (Harris, 1989; see also Chapters 28, Spalding; 33, Fernandez et al.), will affect the redox state of the plastoquinone pool at the set light intensity and the related fluorescence level. In C. reinhardtii, chlororespiration due to the activity of the NADPH-plastoquinone-oxidoreductase, the

576 level of ATP and availability of redox power derived from mitochondrial respiration, may affect the level of NADPH in the plastid and the redox state of the PQ pool (Godde andTrebst, 1980; Godde, 1982; Endo and Asada, 1996). For discussion of chlororespiration, see Chapter 35, Bennoun. In darkness, the plastoquinone pool is oxidized and thus the cells are in state I (Wollman and Delpelaire, 1984). In dark incubated C. reinhardtii cells supplemented with acetate as a carbon source, the plastoquinol pool is significantly reduced (Endo and Asada 1996), and the cells are in state II. Accordingly, the phosphorylated state of C. rein­ hardtii LHCII may be maintained in darkness in vivo in such cells for hours (Owens and Ohad, 1982). Inhibition of the mitochondrial respiratory chain in light exposed cells result in reduction of the plastoquinone pool, forcing the cells into the maximal level of the state II condition (Bennoun, 1982; Gans and Wollman, 1995). At light intensities that saturate electron flow to the electron acceptor sink in vivo, the plastoquinone pool is mostly reduced, LHCII is phosphorylated, and the cells are in state II (Owens and Ohad 1982; Wollman and Delepelaire 1984). It should be noted that under these conditions most ofthe PS II reaction centers are closed (Fm level). Thus the observed fluorescence level at any given time will be affected by reduction, the redox state ofthe plastoquinone pool and LHCII phosphorylation. State transition in C. reinhardtii can be modulated also by changes in the osmolarity of the medium. Hyper-osmotic shock stimulates reduction of mitochondria/chloroplast intersystem electron carriers at the expense of NADH inducing transition to state II, and may increase cyclic electron flow (Endo et al., 1995).

B. Redox Related Activation of the LHCII-Kinase: Role of Cytochrome bf C. reinhardtii mutants lacking the cytochrome bf complex, in which the plastoquinone pool is fully reduced in light-exposed cells, are unable to phosphorylate LHCII. This strongly indicates that indeed this cytochrome complex is involved in the signal transduction path activating the LHCII-kinase (Lemaire et al., 1986; Wollman and Lemaire, 1988; Gal et al., 1990). C. reinhardtii mutants impaired in plastocyanin or PS I activity, and thus unable to oxidize the plastoquinol pool in the light, can

Nir Keren and Itzhak Ohad phosphorylate LHCII (Delosme et al., 1996). Supporting evidence for the concept that the cytochrome bf is involved in the signal transduction system activating the LHCII-kinase(s), was provided by use of cytochrome bf inhibitors such as DBMIB (Bennett et al., 1987; Gal et al., 1988). Since the CP43 and the D2 polypeptides of PS II as well as some of the LHCII polypeptides in C. reinhardtii are phosphorylated under redox control, despite the fact that cytochrome bf activity is impaired (Delepelaire and Wollman, 1985; Wollman and Lemaire, 1988), it is apparent that more than one kinase is involved in the process of phosphorylation of thylakoid membrane proteins and the related state transition. Participation of more than one kinase in the redox-dependent regulation of thylakoid membrane protein phosphorylation in C. reinhardtii was implied before and a model of a kinase cascade was suggested, where one kinase is activated by redox control, and this kinase activates other phosphorylation processes by phosphotransferase activity (Wollman and Lemaire, 1988). A more recent study of cytochrome bf dependent thylakoid membrane protein kinase(s) activation, using EPR and optical spectroscopy, revealed that the thylakoid membrane bound LHCII-kinase can be activated only when a plastoquinol molecule interacts with the plastoquinol oxidizing site (Qo site) of cytochrome bf in which all the electron carriers ofthe high potential path (the Rieske Fe-S center and cytochrome f) are reduced, thus preventing the oxidation of the plastoquinol (the kinase(s) activating mode). Reduction of the plastoquinone pool per se, while preventing its interaction with the Qo site by addition of DBMIB, does not activate the LHCII-kinase. Maintaining the cytochrome bf in its reduced form in absence of reduced plastoquinol had the same effect (Vener et al., 1995, 1997). The above data were obtained, using isolated spinach thylakoid membranes, by activation of the kinase in darkness following a short transient low pH treatment that caused reduction of the PQ pool, the cytochrome bf complex and plastocyanin. This kinase activation method was found to be operative also in Lemna perpusila wild type, but not in a mutant lacking active cytochrome bf complex. However, such experiments have not yet been performed with C. reinhardtii thylakoid membranes. The life time of a cytochrome bf complex in the kinase activating mode in vivo, under light driven steady state electron flow, must be very short

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State Transition and Photoinhibition

considering the kinetics of cytochrome bf oxidoreduction (Vener et al., 1997). The state transition process requires alternating interaction of the kinase with a cytochrome bf complex in this mode and with open phosphorylation sites of the PS II-LHCII complex. Thus, lateral mobility of the above complexes constitutes an important rate limiting factor for the state transition kinetics (Bennett, 1991; Allen, 1992; Drepper, 1993; Gal et al., 1997).

C. Phosphorylation-Dependent Reversible Association of LHCII with PS II The LHCII polypeptides are phosphorylated at their N-terminal stroma exposed segments, and as in higher plants, part of the phosphorylated LHCII, the mobile antenna, dissociates from the PS II complex. The dissociated LHCII-antenna may migrate laterally to the non-appressed membrane domains. This was demonstrated by revealing an increase in the relative amount of LHCII in the non-appressed membrane fraction, by direct visualization of the LHCII particle re-distribution in the appressed and non-appressed membranes of C. reinhardtii using the freeze-fracture technique (Olive et al., 1992; Chapter 14, Olive and Wollman), as well as by photoacoustic measurements of energy transfer (Delosme et al., 1996). As a result, the cross-section of the PS II antenna and the energy transfer to PS II decrease while energy transfer to PS I increases. Deactivation of the thylakoid protein kinase(s) due to the oxidation of the pool, concomitant with dephosphorylation of the LHCII polypeptides by membrane bound phosphatase(s) (Bennett, 1980; Silverstein et al., 1993), allows reassociation of the antenna with the PS II complex. One should note that presently information on the nature, identity and properties of the thylakoidmembrane bound phosphatase is very limited (Allen, 1992; Carlberg and Andersson, 1996) and practically nothing is known about it in C. reinhardtii. The phosphatase involved in the dephosphorylation of the phospho-D1 protein in Spirodella oligorrhiza was reported to be light stimulated (Elich et al., 1993). The phosphatase activity responsible for the dephosphorylation of the Dl protein and LHCII in spinach thylakoid membranes, appears to be redox independent (Silverstein et al., 1993). It is not clear whether the dephosphorylation of phosphoproteins of the C. reinhardtii thylakoid membranes is light activated or enhanced. The amount of phosphate incorporated per LHCII

577 polypeptide, as calculated from different published reports, does not exceed about 10–20% of the total potential phosphorylation sites during the state transition process (Gal et al., 1997). This estimation is compatible with the measured lowering of the energy transferred to PS II in higher plants (Horton and Ruben, 1992). In C. reinhardtii cells grown for several generations in the presence of radioactive phosphate and in continuous light, about one esterified phosphate group was reported per LHCII polypeptide (Owens and Ohad, 1982). Presently, it is widely accepted that phosphorylation of LHCII and state transition are cause/effect related. However, the question remains which ofthe two phenomena is the cause and which the effect. Clearly state transition is not occurring under conditions that prevent the activation of the LHCII kinase(s) (Section II.B). One could envisage that phosphorylation of LHCII follows the dissociation of the antenna from the PS II core that is induced by a redox controlled mechanism yet unidentified. Evidence for the phosphorylation being the first step in the process of state transition can be found in experiments in which the kinase was activated at low temperature. Under this condition the phospho-LHCII remained associated with the PS II complex and energy transfer was not impaired. However upon rising the temperature, lateral migration of the phosphorylated LHCII occurred and state transition, i.e lowering of the energy transfer to PS II ensued (Carlberg et al., 1992). The mechanism whereby phosphorylation induces dissociation of the LHCII mobile component from the PS II complex is not fully understood. The process may involve electrostatic repulsion and conformational changes, resulting from the phosphorylation of the exposed LHCII N-terminal segment (Allen, 1992; Gal et al., 1997). The exact structural information on both the LHCII N-terminal region, and the conformational changes induced in LHCII by its phosphorylation, are still lacking. Microaggregation of LHCII observed upon phosphorylation suggests conformational changes in the structure of the complex. This is also evident from measurements of circular dichroism, fluorescence emission, and other biochemical data (Horton et al., 1991; Barzda et al., 1996). Addition of p-benzoquinone to cells in state I or II, achieved by preillumination with light II or I respectively, stabilizes their initial state irrespective of further illumination conditions and the related changes in the

578 PQ pool redox state. This effect is ascribed to the cross-linking of LHCII, or to the prevention of its phosphorylation or dephosphorylation and thus the respective interaction with the PS II complex (Bulté and Wollman 1990). Two dimensional crystals of LHCII isolated from spinach thylakoid membranes show that the complex occurs in a trimeric form (Kühlbrandt et al., 1994). The N-proximal sequence of the LHCII apoprotein containing the phosphorylation sites is essential for the trimerization of light-harvesting Chl a/b complex (Hobe et al., 1995). Possibly, phosphorylation of the N-terminal segment may have a structural effect on the adjacent sequence thus affecting the trimerization process (Chapter 19, Hoober et al.). Phosphorylation of thylakoid membrane proteins may also induce a partial unstacking ofthe appressed membranes in isolated thylakoids of spinach (Staehelin and Arntzen, 1983; Georgakopoulos and Argyroudi-Akoyunoglou, 1994). A similar observation was made in C. reinhardtii, where unstacking was accompanied by redistribution ofthe cytochrome bf complex between the appressed and the nonappressed membrane domains. It was suggested that these phenomena are related to the activation of cyclic electron flow around PS I and, actually, state transition is regulated by the need for ATP (Vallon et al., 1991). This concept is important in the sense that state transition, by regulating the distribution of energy between the two photosystems, regulates the maintenance of the ATP level via modulation of cyclic electron flow (Bulté et al., 1990). Recently it was reported that LHCI chlorophyll a/b binding proteins of PS I from barley thylakoid membranes are phosphorylated at threonine and serine residues (Knoetzel et al., 1995). This phosphorylation occurs on the amino-terminal surface exposed segment. However, it is not yet known whether this phenomenon occurs under redox control and whether it may be related to state transition. The LHCI of C. reinhardtii has been characterized in terms of polypeptide composition, chlorophyll a/b binding and spectroscopic properties (Bassi et al., 1992). However, no evidence for the phosphorylation of these antenna proteins is yet available. In mutants lacking the PS I complex, LHCI antenna appears to be associated with, and transfer energy to, PS II (Delosme et al., 1996). The role of the light-induced phosphorylation of the RCII core CP 43 or D2 proteins in relation to the state transition phenomenon in C. reinhardtii is not

Nir Keren and Itzhak Ohad known. The possibility that the phosphorylation of the PS II core proteins CP 43 and D2 may affect the conformation of PS II so as to facilitate the dissociation of the mobile phosphorylated LHCII from the PS II complex should, however, not be discarded. Phosphorylation of PS II core proteins may contribute to the monomer/dimer transitions of PS II. Isolated spinach dimer PS II core complexes exposed to light stress convert to monomers in their non-phosphorylated state, but not when phosphorylated (Kruse et al., 1995). Monomerization of the PS II complex in intact spinach thylakoid membranes was recently described to be induced by exposure to excessive light also under nonphosphorylation conditions (Mor et al., 1997). The PsbH protein associated with PS II is also phosphorylated in a redox-dependent manner in spinach thylakoid membranes (Allen, 1992). The role ofthe phospho-PsbH and whether its phosphorylation in C. reinhardtii is redox dependent are not yet clear. According to recent findings PsbH phosphorylation may facilitate assembly of PS II and/or PS II dimerization (Summer et al., 1997). Phosphorylation of thylakoid membrane proteins, resulting in a reversible reduction ofthe PS II antenna size and an increase in that of PS I, besides its roles in controlling the distribution oflight energy between the two photosystems, activating cyclic electron flow and regulating the management of the ATP level, may serve an additional purpose, namely providing partial protection to the PS II complex against photoinactivation. The sensitivity of PS II to excessive light excitation is well established (Prasil et al., 1992) and was extensively investigated in Chlamy­ domonas (see the following section). III. Light Stress: Photoinhibition and Recovery Photoinhibition is defined as light dependent inactivation of the photosynthetic electron flow due to exposure of chloroplasts within living cells, isolated thylakoid membranes or submembrane particles to excessive light intensities. The photoinactivation process may be accompanied by degradation of the PS II core proteins, particularly, the RCII-D1 protein. Often the term ‘light stress’ is used to indicate photoinhibition when it refers to intact cells or whole plants and it has the inherent implication that the phenomenon is not restricted to inhibition of

Chapter 30

State Transition and Photoinhibition

photosynthesis, but it represents a more generalized, systemic effect on the organism. Occasionally the term high-light photoinhibition is used as well, to differentiate between the loss of PS II activity at high photon flux densities from that observed at ambient light intensities. The phenomenon of photoinhibition was originally described by Kok and Jones (Jones and Kok, 1966a,b), who established the wavelength dependence and the kinetics of photoinhibition under excessive light intensities, and defined its target as Photosystem II (then termed Photoact II). The relation between photoinhibition and degradation of the D1 protein was observed first in C. reinhardtii cells (Arntzen et al., 1984; Kyle et al., 1984; Ohad et al., 1984), and this system has played a major role in the study of photoinhibitory events in vivo. In C. reinhardtii cell suspensions in fresh growth medium, exposure to light intensities four- to six-fold higher than those required for saturation of electron flow (chlorophyll optical concentrations in the range of 20–30 ), path of 1–3 cm, 2–3 x photons about 40 induces rapid loss of PS II activity ( min). Following the initial observation of the relation between the loss of PS II activity in C. reinhardtii and the degradation ofthe D1 protein, it was proposed that photoinhibition in vivo is due to the light-induced degradation of the PS II-D1 protein. Recovery of activity was thus related to the de novo synthesis of the D1 protein and replacement of the degraded protein via reassembly with pre-existing PS II proteins that were not degraded or damaged during the photoinhibition process (Ohad et al., 1984). These findings set the basis for the concept of the PS II repair cycle that was further confirmed by studies with other algae (Nedbal et al., 1990; Setlik et al., 1990; Melis and Nemson, 1995) and vascular plants (Aro et al., 1993; Dannehl et al., 1995). Most of the literature concerning the mechanism of photoinhibition refers to the phenomenon induced by exposure of the experimental system to light equivalent to one- to three-fold higher than the maximal intensity provided by sunlight at sea level. As opposed to that, the low-light induced, rapid turnover of the PS II proteins, particularly D1 and D2, was considered as a phenomenon unrelated to the photoinactivation of PS II, possibly involving participation of several photoreceptors (Mattoo et al., 1981; Greenberg et al., 1989). However, recently it was shown that both the high-light induced photoinactivation and degradation of PS II proteins,

579 and the rapid degradation and resynthesis of these proteins in low light, involve oxidative damage to PS II induced by back electron flow via generation of singlet oxygen (Ohad et al., 1994; Keren et al., 1995; Keren et al., 1997). Measurements of PS I activity using methyl viologen as an electron acceptor do not as also demonstrated by EPR indicate loss of spectroscopy (Kyle et al., 1984). It was generally accepted that PS II is the target ofthe photoinhibition process and Photosystem I is not subjected to loss of activity as a result of exposure to high light. However, this concept is not necessarily correct as it oxidowas recently demonstrated that, while reduction activity persists in thylakoid membranes photoinhibited in vitro, electron flow via PS I to is drastically inhibited. When the rate of electron flow to the acceptor sink is limited relative to the rate of PS I excitation and the supply of electrons by PS II exceeds the PS I electron flow capacity, the PS I acceptor side appears to be damaged (Sonoike, 1995) and part of the PsaB protein is degraded (Sonoike, 1996). PS I photoinactivation was found to occur also in vivo in chill-sensitive plants exposed to low temperature (Terashima et al., 1994). This phenomenon was not yet examined in exposed C. reinhardtii cells. The stage of irreversible photoinactivation is preceded by an initial reversible stage characterized by a significant transient increase in the level of Fo fluorescence. This phenomenon can be ascribed to equilibrium, establishchanges in the ing a quasi-stable or long-lived state of which results in a temporary closure of PS II centers that can relax in cells exposed to low light and to a lesser extent even in darkness. The above interpretation is based on experimental results obtained with C. reinhardtii cells (Ohad et al., 1988; Etienne and Kirilovsky, 1992). More detailed experiments confirmed and extended this concept using isolated spinach thylakoid membranes (Hundal et al., 1990; Vass et al., 1992). The molecular mechanism leading to the observed reversible changes in the electron flow form to in vivo, is not yet fully understood. It can be attributed to several factors such as conformational changes in the PS II proteins or changes in their interaction due to alteration of the ionic composition of the plastid matrix and lumen. This phenomenon may be considered as part of the PS II down-regulation process (Prasil et al., 1992). High light exposure of C. reinhardtii cells transiently inactivates the LHCII-kinase and LHCII

580 phosphorylation but does not prevent the dissociation of the antenna from PS II. The kinase activity can be recovered in the absence of de novo protein synthesis and seems to be related to structural changes of the thylakoid membranes denoted by reversible changes in the size and distribution of the intramembrane particles as detected by the freeze-fracture technique (Schuster et al., 1986). Changes in the ionic composition of the plastid, such as depletion of bicarbonate (Govindjee, 1993; Demeter et al., 1995) which may occur under saturating illumination may affect the redox potential of the acceptor quinones Under such and alter the equilibrium conditions electron flow from PS II to the plastoquinone pool may be lowered. Release of the bound from the water oxidation complex by acidification of the thylakoid lumen may inactivate the PS II donor side (Krieger and Weis, 1992), as well as thus lowering increase the redox potential of reduction of and electron flow to the plastoquinone pool (Johnson et al., 1995). These phenomena, as well as state transition and activation of the xanthophyll cycle, down-regulate PS II activity (Demmig-Adams and Adams, 1996) and are considered to provide partial protection against irreversible photoinactivation and degradation of the PS II proteins. While the Fv/Fm ratio is linearly related to the oxygen evolution activity of isolated thylakoid membranes or chloroplasts over a wide range of light intensities (Krause and Weis 1991) this may not be the case in vivo, under conditions that may activate PS II-cyclic electron flow (Prasil et al., 1996). When these parameters are measured in vivo in photoinactivated C. reinhardtii cells, the loss of Fv/Fm precedes to some extent the loss of oxygen evolution activity. This could be due to the fact that possibly PS II may not be the rate limiting factor in oxygen evolution and carbon fixation under high light conditions in vivo. An increase in the Fo level due to any ofthe phenomena described above will affect the ratio Fv/Fo or Fv/Fm without resulting in a real loss of oxygen evolution under saturating light intensity during the measurements. In all the above cases, after an short period of photoinactivation, PS II activity may be recovered upon further incubation of the cells at low light intensities in vivo, without need for de novo protein synthesis (Ohad et al., 1988).

A. Mechanisms of PS II Photoinactivation In the following section the mechanism of the

Nir Keren and Itzhak Ohad irreversible photoinactivation of PS II under all light conditions will be discussed. The termphotoinhibition will be used to indicate loss of photosynthetic electron flow associated with the degradation ofRCII-D1(D2) and other PS II core proteins, which can be restored only following de novo plastid protein synthesis and reassembly of active PS II centers.

1. Acceptor-Side Photoinhibition Measurements of oxygen evolution activity of thylakoid membranes obtained from C. reinhardtii cells exposed to high light intensity, and using artificial PS II electron acceptors such as DMBQ, or site, indicate DCIP which function at or past the loss ofPS II activity. Use ofelectron donors bypassing the water oxidation complex such as DPC does not re-establish PS II activity indicating that the observed loss of electron transfer is not due to inactivation of the water splitting complex. However, PS II activity may be at least partially restored if silicomolybdate site (Kyle et is used as an electron acceptor at the al., 1984). Thus, photoinactivation of C. reinhardtii cells affectsprimarily, orat least initially, the acceptor side of PS II but not the primary photochemistry. This is indicated also by measurements of the changes quinone binding site using PS II-herbicide in the binding tests (Kyle et al., 1984). These results, as well as further thermoluminescence studies of the and related charge recombination and the oxido-reduction activity in persistence of the photoinactivated cells, confirmed the involvement of the PS II acceptor side in initiating the photoinhibiton process (Ohad et al., 1988). Following further exposure of the cells to high light, the charge separation activity ofPS II is irreversibly inactivated as well and is followed by the degradation ofthe PS II proteins. Oxidative damage to PS II is induced by singlet oxygen generated by the interaction of ground state formed by back electron flow oxygen ( )with and charge recombination between the cation radical of the primary electron donor of PS II and the anion (Vass and radical of the primary acceptor, Styring, 1993). Charge recombination prevails under high light intensities at which charge separation rates and resulting electron flow exceed the capacity ofthe plastoquinone pool to accept electrons from PS II. A similar condition occurs when the oxidation of the plastoquinol pool is rate limiting even at low light intensities as is the case in mutants impaired in cytochrome bf, plastocyanin or PS I activity (Shochat

Chapter 30

State Transition and Photoinhibition

et al., 1990; Gong and Ohad, 1991; Zer et al., 1994; Hollinderbaumer et al., 1997). The ratio forward/ backward electron flow in PS II in vivo is therefore a function of the excitation rate, i.e. light intensity and energy transfer from the antenna, the activity of the acceptor side of PS II, and the capacity of the photosynthetic apparatus to oxidize the plastoquinol pool. For a given light intensity, this rate will be modulated by any factor affecting the rate ofelectron flow to the final electron acceptor sink. Therefore, back electron flow and the resulting damage to PS II increases when the carbon fixation reaches its maximal rate, a situation that can occur even at subsaturating light intensities if other environmental factors such as availability of carbon dioxide, nutrients and temperature are suboptimal. According to this scheme, mutations impairing the equilibrium will affect the ratio of forward/backward electron flow and the light sensitivity of the cells to photoinhibition. This phenomenon was also reported in other cells such as cyanobacteria (Kirilovsky et al., 1989; Ohad et al., 1990; Kirilovsky and Etienne, 1991). Alteration ofthe above equilibrium is induced in wild-type cells exposed to high light, under otherwise optimal growth conditions, as a result of extensive reduction of the PQ pool lowering the occupancy of the site by PQ. Damage to PS II induced by this situation in vivo alters the site, generating a stable reduced (Vass et al., 1992; Kirilovsky et al., 1994) that promotes back electron flow (closed centers, Lavergne and Briantais, 1996). It follows that photo-incactivation and degradation of PS II proteins may occur at all light intensities. However, in vivo, de novo synthesis of the RCIID1 (D2) proteins allows replacement ofthe degraded proteins and thus PS II activity may be maintained. As long as the potential rate of PS II recovery exceeds the rate of photoinactivation and protein degradation, no significant loss of oxygen evolution and carbon fixation may be measured in vivo (Kyle et al., 1984; see below).

2. Photoinactivation by Limiting Light Intensities The initiation of PS II photoinactivation by exposure to excessive light intensities in vivo occurs as a result of ‘closing’ the reaction centers, i.e. when the rate of reduction of the plastoquinone pool exceeds that of is mostly reduced. A its oxidation and thus different situation occurs at low, sub-saturating light intensities. In this case, the probability ofconsecutive photon events in the same PS II decreases with

581 decreasing light intensity. Decay of the unstable states in the population of RCII to the stable conditions (Rutherford et al., 1982) may occur via intermediate formation of the primary charge Recombination may separated state generate or a harmful with a ratio close to 1:3. The of at ambient temperature in C. reinhardtii is about 5 s (Keren et al., 1995). Thus, the longer the time interval between consecutive excitations, the higher the probability for charge recombination and oxidative damage. Evidence in support of this hypothesis was provided by experiments in which C. reinhardtii cells were exposed to trains of single-turnover light flashes consisting of one to six flashes per train (5 Hz within the flash train), each of the flash trains being delivered at dark time intervals equivalent to the of the recombination time. Under these conditions, one could predict that extensive charge recombination and related degradation of the D1 protein will occur in cells exposed to series of trains of one, but not of two flashes, as indeed observed experimentally (Keren et al., 1995). The fact that absorption of twice as much light energy, delivered in the series of two flashes, has no damaging effect as compared to that induced by single flashes, agrees with the hypothesis that charge recombination (Ohad et al., 1994), and not the excitation per se, (Park et al., 1996; Tyystjaervi and Aro, 1996) is the primary event leading to the PS II damage. An additional prediction of the above hypothesis is that the relative quantum efficiency (damage per charge separation) of the PS II photoinactivation and D1 protein degradation should rise with the length ofthe dark interval between single flashes, thus increasing the probability for charge recombination of the sates. Moreover, no damage is expected ifthe same number of flashes is given at a high frequency (10 Hz), promoting double reduction of the quinone and forward electron flow, thus avoiding charge recombination. Since the photoinactivation of PS II and D1 protein degradation are ascribed to oxidative damage via generation of singlet oxygen, no loss of PS II and degradation of the protein should occur if the flashes are delivered under anaerobic conditions. All these predictions were verified in experiments carried out with isolated spinach thylakoids (Keren et al., 1997). Similar results have been obtained in C. reinhardtii cells exposed to a series of2000 single flashes delivered at a dark interval of 40 s as well as in intact pea leaves and cyanobacteria (unpublished). One can conclude that the series of events leading

582 to PS II photoinactivation under low light intensities states are: 1) Back electron flow form to the pair via re-generation of the intermediate in darkness; 2) Charge recombination with a high probability to form triplet chlorophyll; 3) Interaction with oxygen; 4) Oxidative damage of the following the formation of singlet oxygen. In the process of back electron flow from prevails in cells exposed serves as an intermediate. continuously to light when the electron flow to is blocked metabolically, by mutations, or by addition site (closed centers). of herbicides binding at the Following rapid consecutive primary charge separations, at high light intensities, the state is formed and rapid charge recombination occurs. Such a state cannot be obtained in the process of a back reaction due to low rates of PS II excitation. In this case the charge separated pair is formed by the chemical oxidation of via the reduction of the D1 protein tyrosine 161 residue acting as the electron The reduction of Pheo may occur via donor to (Diner and Babcock, 1996) and the oxidation of states and are generated during the charge recombination process. Surprisingly, despite the fact that the recombination process is the same, the efficiency of the damage per charge recombination formed in the presence of DCMU is of the pair about half as compared to that obtained in absence of DCMU. The highest efficiency is obtained when serves as the electron donor for the back electron or Partial protection flow as compared to by DCMU against photodamage induced by excessive light can be explained if one considers that formed by charge the lifetime of the ) recombination in the presence of is shorter and therefore the chances for interaction with ground state oxygen are reduced (van Mieghem et al., 1995). However this situation can not occur in PS II exposed to low excitation rates as is the case in low light exposed cells when serves as the electron donor for the back reaction process (Zer et al., 1995). Presently no clear explanation can be offered for this phenomenon. One should note that the relative damage efficiency of the back electron flow via recombination of the states occurring at low light, exceeds by at least one order of magnitude that obtained by back electron flow in closed reaction centers occurring at excessive illumination, both in vivo (Keren et al., 1995) and in vitro (Keren et al., 1997). This phenomenon can be ascribed to the higher

Nir Keren and Itzhak Ohad chances for singlet oxygen generation in the former case. However, in absolute terms, the chances for the formation of the singlet oxygen are relatively low ( /photon absorbed, Tyystjaervi and Aro, may be 1996; Keren et al., 1997), since the quenched by interaction with ground state RCII -carotenes. The possibility that back electron flow may assume different routes besides generation of the should also be considered. One such alternative route This can be offered by the cytochrome cytochrome is an intrinsic component of RCII (Whitmarsh and Pakrasi, 1996; Prasil et al., 1996) encoded by the psbE and psbF genes, which are part ofthe same operon in higher plants but are located in different regions of the plastid chromosome in C. reinhardtii (Mor et al., 1995). One of the intriguing is that under physiological features of cytochrome conditions, it exists in a high or a low potential form which differ by about 300 mV. The slow turnover of indicates that it cannot participate in cytochrome the linear electron flow of PS II and it has been suggested that it is involved in secondary electron transport serving to protect PS II against excess light (Barber and De Las Rivas, 1993; Poulson et al., 1995; Prasil et al., 1996; Mor et al., 1997). Establishing redox conditions in isolated thylakoid membranes compatible with the oxidation of the low decreases potential form of cytochrome considerably the rate of PS II photoinhibition (Nedbal et al., 1992).

3. Donor-Side Photoinhibition So far we have considered the mechanism of PS II photoinactivation due to back electron flow from the acceptor side in a complex that possesses an active electron donor side. A different situation may however occur when the activity of the PS II donor side is may be still reduced by impaired. In this case However, if electron donation from the S-states of is impaired, the cation radical the Mn cluster to persists and following a subsequent charge separation is formed as well. These highly the cation radical reactive radicals may oxidize the protein or pigment bed of the complex and inactivate it, followed by degradation of RCII proteins. In the case of donor side photoinactivation the quantum efficiency ofthe damage is much higher than that resulting from the singlet oxygen generated by back electron flow on the acceptor side since the above cation radicals will

Chapter 30

State Transition and Photoinhibition

be formed after only a few charge separation events (Eckert et al., 1991). In cells exposed to high light intensities the maintained by the high rate of electron flow may ions which are bound to the cause release of the water oxidation complex and are essential for its activity. This possibility is indicated by the acidification ofthe thylakoid lumen in C. reinhardtii cells exposed to high light (Topf et al., 1992). The ‘donor side’ photoinhibition is not expected to occur naturally under low light conditions but can be induced by directed mutations (Minagawa et al., 1996). One should note that both high light or low light exposure induce irreversible photoinactivation of PS II. In the first case the inactivation of either the acceptor or donor side of PS II becomes a limiting factor in electron flow before the inactivation of primary charge separation in RCII. In the second case an active electron acceptor and donor side are a prerequisite for the generation of the oxidative damage via back electron flow from the acceptor side to the donor side of PS II. In this case the promoting condition is the limiting rate of PS II excitation (Keren et al., 1995). The mechanisms of photoinactivation of PS II described so far are based on the accepted idea that /Pheo are the primary electron-donor acceptor pair in PS II. Recently, based on spectroscopic analysis of PS II cores isolated from C. reinhardtii thylakoid membranes and exposed to high light, it was proposed that photoinactivation is due to destruction of a specific Chl of RCII, apparently one of the accessory chlorophylls, which according to the authors may be the actual primary RCII electron donor in the process ofcharge separation in PS II (Bumann and Oesterhelt, 1995). The possibility that this phenomenon is the primary event in the oxidative damage and photoinactivation in vivo remains to be further investigated

4. UV Induced Photodamage Photosystem II is sensitive to ultraviolet light and is rapidly photoinactivated even by low fluency of UVB radiation. One should also consider that UVB radiation accompanies visible light in nature and is emitted by many artificial light sources. According to Edelman and coworkers, UVB irradiation damages the PS II complex in Spirodela oligorrhiza, possibly due to absorption by the reduced acceptor side

583 (Greenberg et al., 1989). semiquinones, Recently, experimental evidence indicates that the Mn cluster of the PS II donor side may be the primary target of the UVB light induced damage in higher plant thylakoid membranes (Friso et al., 1995; Vass et al., 1996). Visible light induces a more extensive degradation of the D1 protein as compared to that of the D2 protein (Zer and Ohad, 1995). The degradation of the D2 protein is, however, enhanced by exposure to UVB light relative to that of the D1 protein (Jansen et al., 1996), possibly due to its binding of acceptor. Since quinone reduction following the charge separation under continuous illumination is accompanied by Mn oxidation, it is difficult to determine which of the sites is responsible for the damage. Under laboratory conditions, use of inappropriate light sources, with high relative emission in the UVB spectral region, may induce donor side photoinactivation even at moderate light intensities.

B. D1 Cleavage as a Result of the Different Photoinhibitory Treatments A distinct difference in the photoinactivation mechanism of PS II impaired in the acceptor or donor side activity, is evident from the analysis of the D1 protein degradation pattern. The initial primary cleavage site occurs on the matrix side of the exposed segment interconnecting the transmembrane D-E loops ofthe D1 protein, close to the binding niche (Trebst, 1986). This location is inferred from hydropathy profiles of the D1 protein and the similarity of the D1, D2 proteins with the L, M proteins of the bacterial RC (Trebst, 1987; Sobolev and Edelman, 1995), for which the 3-D structure was resolved by X-ray crystallography (Michel and Deisenhofer, 1988). This region of the D1 protein is assumed to be in close contact with the analogous loop of the D2 protein which is believed to cover the D1 protein segment and is the first to be exposed to proteolytic degradation in isolated thylakoid membranes (Tietjien et al., 1993). The pattern of D1 protein degradation products in vitro, following photoinactivation under limiting PS II activity on the acceptor side (Shipton and Barber, 1992; Andersson et al., 1994), is similar to that obtained in vivo in low-light exposed Spirodela oligorrhiza. This observation was facilitated greatly by the specific radioactive labeling of the D1 protein under low light intensities. This could be due to

584 retarded onset of the turnover of the other PS II core proteins under these conditions, and thus the initial D1 protein degradation products could be identified by their radio-labeling in pulse-chase experiments. The degradation products included fragments of 29 kDa, 23.5 kDa and 17 kDa (Greenberg et al., 1987). Such degradation products have not yet been observed in C. reinhardtii cells (Wettern et al., 1983), possibly due to the high light intensity used in most experiments. At high light intensity a more complex pattern of protein labeling is generated during the pulse and this hinders the identification of labeled peptides resulting from the initial breakdown of the D1 protein. Attempts to use immunodetection for the identification of degradation products of the D1 protein in C. reinhardtii failed, possibly due to the low specificity of the available polyclonal antibodies toward the resulting degradation products. A different pattern of D1 degradation products is obtained following photoinactivation under donor side limitation of electron flow. An initial cleavage site appears to be located close to the C-terminal side of the D1 protein, compatible with the putative Mn cluster binding side (De Las Rivas et al., 1992), similar to that induced by UVB irradiation of thylakoids. These observations support the concept that the mechanisms of the two photoinactivation processes differ in terms of the identity of the primary source of damage and its possible location within PS II. Presently, experimental evidence related to the cleavage site of the D1 protein is not available for C. reinhardtii.

1. Role of the Binding Site in D1 Protein Degradation in vivo The occupancy of the site by various ligands may modulate the exposure of the cleavage site to the putative protease(s) induced by photoinhibition of PS II. The occupancy of this site by the natural ligand, PQ, appears to allow the light-induced cleavage of the D1 protein in vivo, while complete in light exposed reduction of the PQ pool to cells prevents the degradation of the D1 protein, despite the irreversible inactivation of PS II. This situation is obtained in mutants that cannot oxidize by light-dependent electron flow due to inactivation of cytochrome bf, plastocyanin or PS I (Shochat et al., 1990; Gong and Ohad, 1991). However, lowering of the light intensity and thus of the rate of electron flow from the residually active

Nir Keren and Itzhak Ohad PS II population, allows reoxidation of the plastoquinol pool by ambient oxygen. This treatment induces rapid degradation of the D1 protein in the high-light pretreated mutant cells (Zer et al., 1994). The above observations could indicate that degradation of the RCII-D1 protein in the high-light exposed mutants was already triggered or modified irreversibly, as also indicated by the concomitant loss of PS II activity. However, access of the protease(s) to the cleavage site could be still enhanced site by PQ, or prevented by the occupancy of the or artificial ligands. Binding of different ligands may affect the conformation of the site, so as to influence the exposure of the cleavage site to the protease. The light-induced degradation of D1 in C. rein­ hardtii cells is retarded by urea- and triazine-derived herbicides which compete with the natural ligand, plastoquinone (Gong and Ohad, 1991; Zer and Ohad, 1995). However, phenol-type herbicides, which compete with PQ binding at this site and block PS II electron flow to the PQ pool, do not confer protection against the light-induced cleavage of the D1 protein in Spirodela oligorrhiza (Jansen et al., 1993). Preliminary results obtained with C. reinhardtii cells did not indicate a clear difference in the protection against light-induced D1 protein degradation between the above herbicides (Zer and Ohad, 1995). The effect of DCMU on the photoinactivation and PS II core protein degradation was studied in more detail in C. reinhardtii cells. Addition of DCMU to cell suspensions exposed to high light intensities did not prevent the irreversible PS II photoinactivation which was only slightly retarded. The degradation of the D1, D2, and CP43, but not that of the CP 47 proteins was temporarily inhibited. The degradation of the photoinactivated RCII-D1 protein in the highlight treated cells in the presence of DCMU was initiated following further exposure ofthe cells even to low light or in darkness (Zer and Ohad, 1995). Support for the concept that ligands occupying the site may affect the D1 protein conformation and thus promote or prevent exposure of the cleavage site to proteolysis can be found in experiments in which DCMU inhibited cleavage of the D1 protein by trypsin (Trebst et al., 1988; Zer and Ohad, 1995). The initial pattern of D1 protein cleavage induced by light exposure is similar to that induced by trypsin in thylakoid membranes (Trebst et al., 1988). The retardation of the D1 protein degradation in cells

Chapter 30

State Transition and Photoinhibition

exposed to light in presence ofDCMU, is comparable to the partial protection by DCMU against tryptic cleavage of the D1 protein in thylakoids isolated from C. reinhardtii cells. At the stage at which DCMU did not further protect the D1 protein from degradation in vivo in cells exposed to photoinhibitory light, cleavage of the residual D1 protein was no longer prevented if thylakoids obtained from such cells were exposed to trypsin in the presence of DCMU in vitro. These results indicate that the binding site is gradually conformation of the altered prior to its exposure to the cleavage system (Zer and Ohad, 1995). Based on the above observations, it has been proposed that a gradual series of changes occur in light-exposed PS II, resulting in the modification of its acceptor side, loss of affinity to DCMU at the site and exposure of the site to cleavage. One should note that the binding of various ligands to the niche of PS II is not identical in terms of their interaction with the various amino acid residues forming the hydrophobic quinone binding domain (Ohad et al., 1990; Trebst, 1991; Sobolev and Edelman, 1995). Electron microscopy analysis of single particles of isolated spinach PS II(LHCII) complexes indicates that this complex assumes a dimeric form, and it is implied that this is the natural state of PS II in vivo (Boekema et al., 1995; Hankamer et al., 1996). Exposure to photoinhibitory conditions appears to induce monomerization of the complex (Mor et al., 1997) and it is possible that part of the light-induced exposure of the D1 protein to the proteolytic activity may be related to this process. Monomerization of PS II also seems to lower the electron flow activity of PS II (Hankamer et al., 1997).

C. Degradation of Other PS II Proteins The light-induced degradation of the PS II proteins was initially considered to be specific for the D1 protein (Mattoo et al., 1981). However, with increase in the light intensity the RCII-D2 protein turns over as well, although at a lower rate as compared to that of the D1 protein. The study of this phenomenon was initially hindered by the limited SDS-PAGE resolution of the two proteins and was made possible using immunodetection by specific antibodies (Schuster et al., 1988). Furthermore, it also became apparent that besides the RCII-D1 and D2 proteins, both the CP43 and the CP47 antenna proteins of the PS II core,

585 products of the psbB and psbC genes of the plastid genome, are degraded as well in isolated spinach thylakoids exposed to excessive light intensities (Andersson and Barber, 1994). A detailed study of the light-induced degradation of these PS II core proteins in C. reinhardtii cells revealed that the degradation ofthe D1 protein precedes that ofthe D2 protein by about an hour and then proceeds at comparable rates in cells exposed to light intensity equivalent to 5 fold that required to saturate photosynthesis (Zer et al., 1995). The rates of CP43 and CP 47 protein degradation were somewhat lower than those of the D1 protein degradation. These results strongly indicate that the process of lightinduced degradation of the PS II core proteins is complex and may occur in several steps. It is possible that partial RCII-D1 protein degradation affects the exposure of other PS II core proteins to the degradation process (Zer et al., 1995). Other PS II core protein degradation products have not been identified. The possibility that in Dunaliela salina the degradation of the D1 protein occurs in vivo, via the formation of an aggregated intermediate form of about 160 kDa, was recently proposed by Melis and Nemson (1995). During photoinactivation and degradation of the D1 protein in vitro, aggregation of D1-D2 heterodimer and cross-linking of the D1 protein to cytochrome have been reported in spinach thylakoids (Barbato et al., 1991; Barbato et al., 1992; Shipton and Barber 1992; Andersson et al., 1994). However, the significance and relevance of these findings to the mechanism of the D1 protein degradation in C. reinhardtii cells in vivo are not known. While D1 protein phosphorylation in higher plants was demonstrated to regulate its light-induced degradation (Aro et al., 1993; Dannehl et al., 1995), the D1 protein is not phosphorylated in C. reinhardtii (de Vitry et al., 1991), and in mosses (Rintamäki et al., 1995). However the RCII-D2 protein is phosphorylated in C. reinhardtii (de Vitry et al., 1991) as well as in vascular plants. It is possible that changes in the conformation of PS II in these organisms, induced by phosphorylation of the D2 protein, regulate the exposure of the cleavage site of the altered RCII-D1 protein to proteolysis. Phosphorylation of the PS II core proteins appears to affect the site, as indicated by a lowering of the binding affinity for site directed herbicides (Shochat et al., 1982; Giardi et al., 1992).

586

1. The Nature of the Protease The nature ofthe protease(s) involved in PS II protein degradation is not known. Based on the initial observation that the D1 protein is degraded in cells exposed to intense light (Ohad et al., 1985), as well as in isolated C. reinhardtii thylakoid membranes (Reisman and Ohad, 1986), it was suggested that the protease(s) involved in this process are membrane bound. This observation was further supported by findings obtained using spinach thylakoid membranes (Andersson et al., 1994). Since addition ofATP was not required for this process, it appears that ClpP proteases (Adam, 1996) do not participate, at least not in the initial cleavage process. Furthermore, degradation of the D1 protein occurs also in isolated RCII particles and it was proposed that the D1 protein itself may have proteolytic activity (Andersson, 1996). The question arises as to whether light is required for the proteolytic process per se and if it can be involved in the activation of a protease rather than in the induction of a change in the substrate D1 protein which may allow its cleavage as discussed above. Experiments in which this question was addressed in intact C. reinhardtii cells (Kirilovsky et al., 1990; Zer et al., 1994), as well as in vitro (Hundal et al., 1990; Kirilovsky and Etienne, 1991), indicate that once the PS II(RCII) has been modified in a yet unknown way by exposure to light, the degradation of the proteins can continue in darkness. The observation that light-induced loss of PS II activity can be obtained at low temperatures in both intact cells and isolated thylakoid membranes while the PS II protein degradation is inhibited and can be initiated by raising the temperature even in darkness (Hundal et al., 1990; Kirilovsky and Etienne, 1991) demonstrate that light per se is not required for the cleavage process. Series of single turnover light flashes delivered at 40 s dark intervals induce significant D1 protein degradation in dark incubated C. reinhardtii cells, while series of two such flashes spaced at 200 ms and delivered at similar dark intervals have no effect on the D1 protein (Keren et al., 1995). These results cannot be reconciled with the idea that light activates the protease(s) rather than induces the exposure of the substrate. The fact that different sites are cleaved, according to the respective mechanisms of photoinactivation, may also be considered as an indication that the degradation process is initiated by alteration of the

Nir Keren and Itzhak Ohad substrate. However, so far no information is available on the specific chemical nature of the substrate modification. The putative protease(s) responsible for the lightinduced D1 protein degradation is considered to be nuclear encoded as no sequence homologous to a protease was identified in the plastid genome sequences available to date (Shimada and Sugiura, 1991) except ClpP (Adam, 1996). The light-induced degradation of the D1 protein in phosphate-starved C. reinhardtii cells is inhibited by cycloheximide (Trebst and Sollbracht, 1996). This observation was interpreted in terms of activation of a phosphatase(s) which may regulate the activity ofa nucleus encoded unstable phosphorylated protease.

D. The Recovery Process The stable integration of the newly synthesized pD1 precursor protein is one of the limiting steps in the reassembly of PS II (Adir et al., 1990). It is possible one of the four major that cytochrome components of RCII that appears to remain stable while the RCII-D1 and D2 proteins are degraded, may serve as a docking site or anchor for the assembly of the PS II core proteins. As such, it may play a crucial role in the process of light-induced PS II protein turnover and recovery of PS II activity in all light conditions A schematic representation of the PS II photoinactivation, D1 protein degradation and recovery processes is shown in Fig. 3. The enhanced synthesis of the D1 protein during the recovery process involves activation ofthe pD1 precursor protein translation by membrane bound polyribosomes associated mostly with the non-appressed membrane domains (Chapter 12, Hauser et al.). This is followed by the insertion of the nascent peptide into the membrane and its integration together with the pre-existing PS II core proteins, that were not degraded following PS II photoinactivation, into an active PS II complex. The re-assembly of RCII requires also availability of Chl a and -carotene. Activation of the donor side of the reassembled complex occurs only after processing of the carboxyl end segment of pD1. However, the binding ofmanganese, to form the active Mn cluster, requires a light activation step (Rova et al., 1996). The presence of a pD1 protein in RCII prevents the formation of the manganese cluster responsible for the water oxidation process (Rutherford et al., 1988). The translation of the pD1 protein was reported to be

Chapter 30

State Transition and Photoinhibition

activated by light in C. reinhardtii cells (Danon and Mayfield, 1994; Mayfield et al., 1994). Detailed information on the light-dependent redox control of the psbA mRNA translation and the mRNA degradation is found in Chapters 10 (Stern and Drager) and 12 (Hauser et al.).

1. Synthesis and Reassembly of PS II Proteins Short radioactive pulse-chase experiments and localization of the newly synthesized protein in C. reinhardtii cells exposed to excessive light, have demonstrated that pD1 is found initially in the nonappressed thylakoid membranes, where it is associated already after 2 min with the core proteins, D2 and possibly cytochrome as well. Processing of the pD1 precursor protein occurs already at this stage in the reassembly of RCII. Within several minutes the processed D1 protein appears already in a larger complex found in both non-appressed and appressed thylakoid membrane domains, followed by a further increase in complex size that can be found only in the appressed membranes (Adir et al., 1990) and could represent the dimeric form of the

587

PS II core. A similar pattern of assembly of pD1 into the PS II complex, following its synthesis by run-off translation in vitro, was described in more details in isolated spinach thylakoid membranes (van Wijk et al., 1996). The rapid processing of the pD1 protein is essential for the stability ofthe reassembled PS II core. The reassembled PS II core containing the pD1 protein supports charge separation and reduction of the acceptor quinones. However, in the absence of an active electron donor, the reassembled complex is highly sensitive to light (see donor side photoinhibition, Section III.A.3). The danger in forming active PS II complexes containing the ‘suicidal’ pD1 component is real, as mutants impaired in the processing protease present highly light-sensitive PS II complexes that are capable of charge separation and reduction of acceptors in Scenedesmus obliquus (Gong and Ohad, 1995) and cyanobacteria (Anbudurai et al., 1994). The processing protease has been identified in cyanobacteria as an exo-carboxypeptidase which is a product of the ctpA gene (Anbudurai et al., 1994). The gene sequence shows homology to the retinoid binding protein ofmammals

588 and the penicillin binding protein of E. coli. The processing protease gene has been identified also in spinach (Inagaki et al., 1996) and recently the D1 processing protease Scenedesmus obliquus was purified to homogeneity and the gene was identified (Trost et al., 1997). No information is yet available on the nature of this protease in C. reinhardtii. Among all plastid-encoded PS II proteins, the D1 protein is the only one synthesized as a precursor and one could consider that this property may have a selective value. Site-directed mutants in which the extended C-terminal peptide present in the pD1 protein has been deleted in C. reinhardtii do not show a phenotype different from the wild type when grown under normal light conditions (Schrader and Johanningmeier, 1992). However, no information is yet available on the selective value of the D1 protein extension in the process of PS II assembly during membrane biogenesis, relative to rapid D1 protein turnover induced by excessive illumination. The assembly of a PS II complex possessing an active oxygen-evolving electron donor side requires the binding of the nuclear encoded 17 kDa, 23 kDa and 33 kDa OEE proteins (oxygen evolution enhancer proteins). These proteins are located in the thylakoid lumen and remain bound to the lumenal side of the PS II complex in C. reinhardtii cells during the photoinactivation process as long as the D1 protein is not degraded. Upon degradation of the D1 protein the OEE proteins are released into the lumenal space, but are not degraded for up to 24 h even if the synthesis of D1 protein and reassembly of the PS II complex are inhibited. The proteins are thus available and rebind to the re-assembled PS II core as soon as this complex is formed (Eisenberg-Domovich et al., 1995). Mutants defective in the OEE 23 kDa protein in C. reinhardtii cells exhibit light sensitivity similar to mutants lacking the pD1 processing protease (Rova et al., 1996). The rate of de novo synthesis of D1 protein in C. reinhardtii increases with an increase in light intensity. The increase in the rate of D1 protein labeling under such conditions is ascribed to the light-induced degradation and resynthesis (turnover) of the protein. The question arises as to what the relation between these two processes might be. The photons rate ofD1 protein labeling at 20 in C. reinhardtii cells that have been pre-exposed to photons light intensities from 20 to 1000 for a short period, increases proportionally to the light intensity during the pre-exposure period (Ohad

Nir Keren and Itzhak Ohad et al., 1988). This phenomenon indicates that the increase in the translation and stable incorporation ofthe D1 protein into PS II complexes is not directly related to the light intensity during the translation process, but to the extent of light-induced degradation of the D1 protein. The availability of the residual PS II proteins may facilitate the fast and stable reassembly of the pD1 protein into new RCII. Blocking the translation of plastid proteins by chloramphenicol or lincomycin does not affect the rate of D1 protein degradation induced by high light (Schuster et al., 1988; Shochat et al., 1990; Zer et al., 1994). The degradation of the protein is not directly related to its replacement in C. reinhardtii cells, as evident by the findings that its light-induced degradation in isolated thylakoid membranes is not significantly different form that observed in vivo. The reported lowering of D1 protein degradation by translation inhibitors in cyanobacteria (Komenda and Barber, 1995), was not observed in C. reinhardtii cells (Shochat et al., 1995). However, the opposite is true, namely, the degradation of the D1 protein is a pre-requisite for the enhanced translation and stabilization of the D1 protein by assembly with the remaining components ofPS II. This is demonstrated by experimental results obtained with mutants impaired in cytochrome bf, plastocyanin or PS I activity in which the plastoquinone pool is completely reduced in cells exposed to high light and the D1 protein degradation is retarded (Section III.B.1). Pulse-labeling of these cells during the high light exposure does not show rapid synthesis and accumulation of newly synthesized D1 protein, in contrast to the wild-type cells in which the D1 protein is rapidly degraded and replaced under similar light conditions. However, rapid D1 labeling and accumulation occurs in the mutant cells when the light intensity is lowered, allowing reoxidation ofthe plastoquinone pool and degradation of the damaged RCII-D1 protein (Zer et al., 1994). These results indicate that accumulation ofstable newly synthesized D1 protein is possible only if the protein is integrated into a PS II complex. The capability to synthesize the D1 protein and, possibly, the other PS II complex proteins during the photoinhibition process is gradually reduced as the time of exposure and light intensities increase. Radioactive labeling of cells that have been exposed photons to light intensities of 2–4 × for about 90 min and have lost about–80% of their PS II activity, is strongly diminished as compared with

Chapter 30 State Transition and Photoinhibition that obtained at the onset of the high light exposure, even if the cell suspension is supplemented with acetate as a source of respiration-derived energy. Recovery of protein synthesis activity occurs only after further incubation of the cells for 1–3 h at low light intensity and parallels the recovery of PS II activity. This loss of protein synthesis capacity of the plastids is not related to the loss of electron flow via PS II, since inhibition of PS II activity by DCMU does not affect PS II protein synthesis in cells supplemented with acetate as a carbon source (S. Shochat and I. Ohad, unpublished). The photoinactivation of PS II occurs gradually during the high light exposure and, as a result, electron transfer from may also be inhibited while and PS I to possibly cyclic electron flow activity is not affected. The amount of residual active PS II can still reduce plastoquinone sufficiently to prime the onset of cyclic PS I electron flow (Heber et al., 1978; Topf et al., 1992). At high light intensity electron flow may induce across the thylakoid membranes. a high Photoacoustic measurements in C. reinhardtii cells indicated that photochemical energy storage, possibly related to PS I cyclic electron flow, persisted in photoinhibited cells (Canaani et al., 1989). In the presence of ammonium nitrate as a nitrogen source the proton gradient is dissipated by accumulation of in the thylakoid lumen followed by chloride and water uptake and an excessive swelling of the thylakoid membranes ensues. It is possible that under these conditions the plastid translation activity is hindered and it is resumed only upon lowering the light intensity and slow recovery of the normal thylakoid membrane structural organization. Mutants impaired in cyclic electron flow activity do not exhibit this phenomenon (Topf et al., 1992).

2. The Role of Chlorophyll a and -Carotene in the Reassembly of PS II Besides lowering the rate of D1 protein synthesis, additional factors may limit the recovery of PS II activity. Reassembly of PS II requires Chl a and carotene serving not only the function of electron quenchers but also a major structural donors or role in the assembly of the complex. The fate of the six Chl a molecules and -carotene of RCII during the turnover of the PS II proteins is not known. The D1 protein may be synthesized and accumulated to small amounts, following photoinhibition ofthe light grown C. reinhardtii y-1 mutant

589 if the cells are further incubated in darkness, a condition that prevents chlorophyll synthesis in these cells (Zer et al., 1994). Therefore, pre-existing Chl was available to rebind during the reassembly of RCII. The source of this Chl is not known. The Chl a of the damaged RCII, if liberated following RCII disassembly and protein degradation, could be extremely harmful due to its photo-dynamic activity especially at the photoinhibitory light intensity. Since free Chl can diffuse in the lipid bilayer, the damage induced by free interaction with oxygen should be delocalized and non specific, and one could expect damage to many different membrane and stromal components of the chloroplast besides that induced to PS II. Yet this is not the case. Thus, either the RCII chlorophylls remain bound to some proteins, possibly the PS II core antenna or LHCII that can quench the state by interaction with the antenna carotenes, or they can be rapidly broken down to non-harmful degradation products. The Chl required for the reassembly of the newly synthesized RCII proteins may be taken from preexisting Chl-proteins. This possibility was demonstrated in experiments in which the C. reinhardtii y-1 mutant cells grown in darkness, and thus lacking the thylakoid membrane system (Gershoni et al., 1982), were allowed to green in the light in presence of chloramphenicol preventing synthesis of the plastid translated PS II protein. These cells accumulated large amounts of the LHCII protein-chlorophyll complexes but no reaction centers. However, the missing PS II proteins could be synthesized and photosynthetic activity could be reestablished, at least partially, if the cells were washed free of chloramphenicol and further incubated in darkness or in the light in the presence of cycloheximide, conditions which prevent further chlorophyll synthesis (Gershoni et al., 1982). These experiments demonstrate that not only the few Chl molecules required for the assembly of RCII, but also the large amount of Chl required for the assembly of the PS II core antenna, CP43 and CP 47, as well as for the formation of PS I, could be taken from pre-existing Chl proteins (Gershoni et al., 1982). Chlorophyll transfer from pre-existing antenna complexes to newly formed reaction centers was also reported in Chlorella pyrenoidosa cells (Lavintman et al., 1981) and vascular plants (Dannehl et al., 1996). The discovery that the early light-induced protein (ELIP), homologous to the Cab gene product (Grimm et al., 1989), is induced under light-stress conditions

Nir Keren and Itzhak Ohad

590 (Adamska et al., 1993), could indicate that this protein may be a Chl scavenger during the fast turnover of the PS II proteins. Carotenogenesis induced by light stress in Dunaliella salina is accompanied by induction of CbR, an ELIP like protein, which was proposed to act as a Chl or possibly xanthophyll-binding protein (Levy et al., 1993). The possibility that ELIP-like proteins may be ‘scavengers’ ofreleased carotenes and chlorophylls during the rapid turnover of the PS II core proteins is attractive; however, it requires further experimental support. ELIPs have not yet been identified in Chlamy­ domonas. The -carotenes needed for the reassembly of PS II can possibly be supplied by the membrane bound carotene pool which in C. reinhardtii cells may also occur in the form of -carotene globules (Ohad et al., 1969). Depletion of the carotene pool may prevent the recovery of PS II following photoinhibition. This phenomenon was elegantly demonstrated by Trebst and Depka (1997), who showed that preincubation of C. reinhardtii cells in the presence of the carotenogenesis inhibitor norflurazon, acting on phytoene desaturase, prevents the recovery of PS II and the re-accumulation of D1 protein during photoinhibition.

IV. Concluding Remarks and Perspectives In this chapter we have described mechanisms that serve C. reinhardtii in its response to changes in the ambient light intensity and spectral characteristics. These mechanisms are not, by any means, the only ones available for that free-living, motile, photoautotrophic alga which dwells in fresh water ponds. Motile unicellular algae may just as well swim away from or toward the light, until reaching the optimal environment for their physiological needs. These may change with the cell cycle state, availability of nutrients and temperature as well as with the transient presence of toxic levels of various chemicals that may increase or decrease with the changes in the fresh water supply (rain) or its evaporation. C. reinhardtii is endowed with an ‘eye-spot’ consisting of an array of -carotene globules within the single cup-shaped chloroplast, adjacent to the plastid envelope, close to the plasma membrane (Ohad et al., 1967; Ohad et al., 1969). The shade produced by the eye spot may create an asymmetry center in terms of the light direction. Apparently the

eye-spot seems to be involved in the control of phototaxis. A special rhodopsin-like light sensor (chlamyrhodopsin) located in the plasma membrane is considered to regulate the coupling oflight direction and intensity to the phototactic response and serves as part of the signal transducing system controlling the flagellar movement (Lawson and Satir 1994; Deininger et al., et al., 1995; Pazour et al., 1995; Rueffer and Nultsch, 1997). This ability, that is absent from sessile oxygen evolving photosynthetic organisms, may have contributed to the development of interactions between the control of the state transition and repair cycle of photoinhibited photosystems and the phototrophic response. The experimental designs, thus far, have not permitted an integrative approach, for studying the physiological response ofcells exposed to varying light conditions compatible with those encountered in nature. One could expect that in the future such experimental systems will be developed and will shed light on the entire chain of events that interconnect the cellular responses to ambient light as well as their relative contribution to C. reinhardtii well-being.

Acknowledgments We are grateful to Drs. Tsafrir S. Mor (presently, Boyce Thomson Institute, Cornell Univ.), Susana Shochat, Alma Gal, Christof Franke and Yael Eisenberg-Domovich (HUJI), for their critical reading of the manuscript and for their suggestions. We acknowledge the support by the Sonderforschungsbereich 184 in cooperation with R. G. Herrmann and W. Rüdiger, by the German-Israeli Science Foundation in cooperation with L. Eichacker and W. Rüdiger, München; by the Human Frontiers of Science Program; the Israel National Science Foundation and by the Minerva, Avron, Even-Ari Research Center.

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595 Chlamydomonas reinhardtii: Role of Photosystem II reaction center and phosphorylation of the high affinity binding site. Biochim Biophys Acta 1681: 21–31 Shochat S, Adir N, Gal A, Inoue Y, Mets L and Ohad I (1990) Photoinactivation of Photosystem II and degradation of the D1 protein are reduced in a cytochrome less mutant of Chlamydomonas reinhardtii. Z Naturforsch 45c 395–401 Shochat S, Zer H and Ohad I (1995) Chloroplast translation activity is not required for light dependent degradation of the D1 protein in Chlamydomonas reinhardtii. In: Mathis P (ed) Photosynthesis: from Light to Biosphere, Vol IV, pp 295–298. Kluwer Academic Publishers, Dordrecht Silverstein T, Cheng L and Allen JF (1993) Chloroplast thylakoid protein phosphatase reactions are redoxindependent and kinetically heterogeneous. FEBS Lett 334: 101–105 Sobolev V and Edelman M (1995) Modeling the quinone-B binding site of the Photosystem-II reaction center using notions of complementary and contact-surface between atoms. Proteins Structure Function and Genetics 21: 214–225 Sonoike K (1995) Selective photoinhibition of Photosystem I in isolated thylakoid membranes from cucumber and spinach. Plant Cell Physiol 36: 825–830 Sonoike K (1996) Degradation of psaB gene product, the reaction center subunit of Photosystem I, is caused during photoinhibition of Photosystem I: possible involvement of active oxygen species. Plant Sci 115: 157–164 Srivastava A, Strasser RJ and Govindjee (1995) Polyphasic rise of chlorophyll a fluorescence in herbicide resistant D1 mutants of Chlamydomonas reinhardtii. Photosynth Res 43: 131–141 Staehelin LA and Arntzen CJ (1983) Regulation of chloroplasts membrane function: protein phosphorylation changes the spatial organization of membrane components. J Cell Biol 97: 1327–1337 Strasser RJ, Srivastava A and Govindjee (1995) Polyphasic chlorophyll a fluorescence transient in plants and cyanobacteria. Photochem Photobiol 61: 32–42 Summer EJ, Schmid VHR, Bruns BU and Schmidt GW (1997) Requirement for the H-phosphoprotein in Photosystem II of Chlamydomonas reinhardtii. Plant Physiol 113: 1359–1368 Terashima I, Funayama S and Sonoike K (1994) The site of photoinhibition in leaves of Cucumis sativus L. at low temperature is Photosystem I, not Photosystem II. Planta 193: 300–306 Tietjien KG, Draber W, Goossnes J, Jansen JR, Kluth JF, Schindler M, Wroblowsky H-J, Hilp U and Trebst A (1993) Binding of -binding niche of Phototriazines and triazones in the system II. Z Naturforsch 48c: 205–212. Topf J, Gong H, Timberg R, Mets L and Ohad I (1992) Thylakoid membrane energization and swelling in photoinhibited Chlamydomonas cells is prevented in mutants unable to perform cyclic electron flow. Photosynthesis Research 32: 59–69 Trebst A (1986) The topology of the plastoquinone and herbicide binding peptides of Photosystem II in the thylakoid membranes. Z Naturforsch 41c: 240–245 Trebst A (1987) The three-dimensional structure ofthe herbicide binding niche on the reaction center polypeptides of Photosystem II. Z Naturforsch 42c: 742–750 Trebst A and Depka B (1997) Role of carotene in the rapid turnover and assembly of Photosystem II in Chlamydomonas reinhardtii. FEBS Lett 400: 359–362

596 Trebst A and Sollbracht E (1996) Cycloheximide retards high light driven D1 protein degradation in Chlamydomonas reinhardtii. Plant Sci 115: 191–197 Trebst A, Depka B, Kraft B and Johanningmeier U (1988) The site modulates the conformation of the Photosystem II reaction center polypeptides. Photosynth Res 18: 163–177 Trost JT, Chisholm DA, Jordan DB and Diner B A (1997) The D1 C terminal processing protease of Photosystem II from Scenedesmus obliquus: Protein purification and gene characterization in wild type and processing mutants. J Biol Chem 272: 20348–20356 Tyystjarvi E and Aro E-M (1996) The rate constant of photoinhibition, measured in lincomycin-treated leaves, is directly proportional to light intensity. Proc Natl Acad Sci USA 93: 2213–2218 Vallon O, Bulté L, Dainese P, Olive J, Bassi R and Wollman FA (1991) Lateral redistribution of cytochrome complexes along thylakoids membranes upon state transition. Proc Natl Acad Sci USA 88: 8262–8266 van Mieghem F, Brettel K, Hillmann B, Kamlowski A, Rutherford AW and Schlodder E (1995) Charge recombination reactions in Photosystem II. 1. Yields, recombination pathways and kinetics of the primary pair. Biochemistry 34: 4798–4813 van Wijk KJ, Andersson B and Aro E-M (1996) Kinetic resolution of the incorporation of the D1 protein into Photosystem II and localization of assembly intermediates in thylakoid membranes of spinach chloroplasts. J Biol Chem 271: 9627–9636 Vass I and Styring S (1993) Characterization of chlorophyll triplet states in Photosystem II sequentially during photoinhibition. Biochemistry, 32: 3334–3341 Vass I, Styring S, Hundal T, Koivuniemi A, Aro E-M and Andersson B (1992) Reversible and irreversible intermediates during photoinhibition of Photosystem II: Stable reduced species promotes chlorophyll triplet formation. Proc Natl Acad Sci USA 89: 1408–1412 Vass I, Sass L, Spetea C, Bakou A, Ghanotakis DF and Petrouleas V (1996) UV-B-induced inhibition of Photosystem II electron transport studied by EPR and chlorophyll fluorescence.

Nir Keren and Itzhak Ohad I m p a i r m e n t of donor and acceptor side components. Biochemistry 35: 8964–8973 Vener AV, van-Kan PJM, Gal A, Andersson B and Ohad I (1995) Activation/deactivation cycle of redox–controlled thylakoid protein phosphorylation. J Biol Chem 270: 25225–25232 Vener A, van Kan PJ, Rich PR, Ohad I and Andersson B (1997) Plastoquinol at the Qo-site of reduced cytochrome b/f mediates signal transduction between light and thylakoid phosphorylation. Thylakoid protein kinase deactivation by a single turnover flash. Proc Natl Acad Sci USA 94: 1585–1590 Wettern M, Owens JC and Ohad I (1983) Role of thylakoid polypeptide phosphorylation and turnover in the assembly and function of Photosystem II. Methods Enzymol 97: 554–567 Whitmarsh J, and Pakrasi HB (1996) Form and function of In: Ort DR, and Yocum CF (eds) Oxygenic cytochrome Photosynthesis: The Light Reactions, pp 249–264. Kluwer Academic Publishers, Dordrecht Wollman F-A and Delepelaire P (1984) Correlation between changes in light energy distribution and changes in thylakoid membrane polypeptide phosphorylation in Chlamydomonas reinhardtii. J Cell Biol 98: 1–7 Wollman F-A and Lemaire C (1988) Studies on kinase-controlled state transitions in Photosystem II and mutants from Chlamydomonas reinhardtii which lack quinone-binding proteins. Biochim Biophys Acta 933: 85–94 Zer H and Ohad I (1995) Photoinactivation of Photosystem II induces changes in the photochemical reaction center II abolishing the regulatory role of the site in the D1 protein degradation. Eur J Biochem 231: 448–453 Zer H, Prasil O and Ohad I (1994) Role of plastoquinol oxidoreduction in regulation of photochemical reaction center II D1 protein turnover in vivo. J Biol Chem 269: 17670–17676 Zer H, Keren N and Ohad I (1995) The regulatory role of the site in the RCII-D1 protein degradation is gradually lost following exposure of Chlamydomonas cells to low light. In: Mathis P (ed) Photosynthesis: from Light to Biosphere, Vol 4, pp 307–310. Kluwer Academic Press, Dordrecht

Chapter 31 Synthesis of Metalloproteins Involved in Photosynthesis:

Plastocyanin and Cytochromes

Sabeeha Merchant Department of Chemistry and Biochemistry, University of California, Los Angeles, CA 90095-1569, U.S.A.

Summary I. Introduction A. Metalloproteins in the Photosynthetic Apparatus B. Biosynthesis of Metalloproteins 1. Metal-Responsive Gene Expression in Cyanobacteria and Green Algae 2. Post-Translational Assembly of Metallocofactor-Containing Proteins 3. Metal Metabolism II. Copper-Responsive Synthesis of Plastocyanin and Cytochrome

A. Plastocyanin 1. Properties and Function 2. Genetic Information and Biosynthesis in Algae B. Regulation of Plastocyanin Accumulation 1. mRNA Accumulation in Algae and Cyanobacteria 2. Degradation in C. reinhardtii C. Cytochrome 1. Properties and Function 2. Genetic Information and Biosynthesis D. Regulation of Cytochrome Synthesis 1. Copper as the Signal 2. Transcriptional Regulation 3. Response to Iron and Heme E. Other Responses to Copper-Deficiency 1. Copper Transport 2. Coproporphyrinogen Oxidase F. Adaptation to Copper-Deficiency III. Genetic Analysis of Chloroplast Metalloprotein Assembly A. The c-type Cytochromes 1. The ccs Mutants 2. The Ccs Factors a. CcsA b. Ccs1 3. A Novel Pathway in the Chloroplast B. The b-Heme(s) of the f Complex C. Plastocyanin D. Approaches for the Dissection of Other Assembly Pathways IV. Conclusions Acknowledgments References

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Summary The introduction of a metal cofactor broadens the catalytic repertoire ofa protein catalyst by facilitating certain types of chemical reactions. The chemistry of the cofactor depends upon the type of coordinating ligands, and also upon the geometry ofthe metal-containing active site. Metalloprotein biosynthesis requires 1) mechanisms of metal acquisition and transport to the sub-cellular compartment where metal delivery to the active site of the metalloprotein can occur, and 2) mechanisms of coordinate regulation of polypeptide synthesis with cofactor synthesis (for organic cofactors like heme) or assimilation (for inorganic cofactors like the cluster). The copper-responsive accumulation of copper-containing plastocyanin and heme-containing cytochrome in Chlamydomonas reinhardtii has served as a model for studies of regulation of metalloproteins by metals because the simple growth requirements of Chlamydomonas facilitate studies of metal metabolism. In a fully copper-supplemented medium, plastocyanin accumulates in C. reinhardtii but Cyt does not. As the medium is depleted of copper mechanisms for adaptation to copper-deficiency are induced. These include degradation of plastocyanin, transcriptional activation of Cyc6 and Cpx1 genes (encoding Cyt and coprogen oxidase, respectively), and induction of copper transport and cupric reductase activity. Studies of chloroplast metalloprotein assembly in Chlamydomonas have focused primarily on the heme proteins, with the c-type cytochromes being the best studied examples. The occurrence and distribution in the genome databases of Ccs genes required for chloroplast c-type cytochrome maturation suggests that they represent a third family of cytochrome maturation components, distinct from the Cyt heme lyases of mammalian and fungal mitochondria and also distinct from the components identified in most proteobacteria and plant mitochondria. Approaches for the analysis of the assembly of other cofactor-containing chloroplast proteins are discussed.

I. Introduction

A. Metalloproteins in the Photosynthetic Apparatus Metalloproteins are defined broadly as proteins with metal-containing cofactors. The metal cofactor may be liganded only by functional groups provided by amino acid side chains from the polypeptide, as in plastocyanin, carbonic anhydrase or the Zncontaining transcription factors; the metal cofactor may be a component of an organic prosthetic group, as in the chlorophyll and heme proteins; or the metal may associate with inorganic ions to form a stable cluster, as in the ferredoxins. Each of these types of metalloproteins is found in the photosynthetic apparatus (Table 1). Metalloproteins are abundant in photosynthetic membranes as they are in all energy transducing membranes owing to the catalytic properties of metal cofactors in reactions involving electron transfer, oxygen chemistry and photochemistry. Since the function ofthe metalloprotein is Abbreviations: – acid; – b-heme of cytochrome on the lumen side; – b-heme of cytochrome on the stromal side; Ccs genes – cytochrome c synthesis genes; coprogen – coproporphyrinogen; – Cu – copper-deficient; +Cu –

copper-sufficient; CuREs – copper-responsive elements; Cyt – cytochrome

critically dependent upon the cofactor, consideration of the biosynthetic pathway of the protein must include the process of cofactor metabolism, cofactorprotein assembly and cofactor-dependent gene expression. While these aspects have been emphasized for the study ofthe biosynthesis ofchlorophyllbinding proteins, there are very few studies of the cofactor-dependent synthesis of other types of metalloproteins in photosynthetic organisms.

B. Biosynthesis of Metalloproteins 1. Metal-Responsive Gene Expression in Cyanobacteria and Green Algae Two systems for the study of metal-dependent synthesis of metalloproteins are provided by the reciprocal iron-dependent synthesis of ferredoxin vs. flavodoxin in cyanobacteria and the reciprocal copper-dependent synthesis of plastocyanin vs. cytochrome in some cyanobacteria and green algae. Ferredoxin and flavodoxin are functionally interchangeable soluble electron transfer proteins. Ferredoxin is the preferred catalyst in iron-replete media; in iron-deficient media flavodoxin synthesis is induced (Hutber et al., 1977; Sandmann and Malkin,

1983). The importance of this adaptation is

underscored by the observation that the growth of

Chapter 31

Biosynthesis of Metalloproteins

599 1997). The focus of this chapter is on the copperdependent synthesis of plastocyanin and cytochrome in Chlamydomonas.

2. Post-Translational Assembly of Metallocofactor-Containing Proteins

photosynthetic organisms in nature can be limited by iron availability (Martin and Fitzwater, 1988), and its relevance in natural populations is apparent from the demonstration that flavodoxin abundance in marine diatoms is determined by iron availability in the ocean (LaRoche et al., 1996). The interested reader is referred to a previous volume in this series for molecular details of the iron-responsive pathway in cyanobacteria (Straus, 1994). Plastocyanin and cytochrome form another pair of interchangeable photosynthetic electron transfer catalysts in organisms that have genetic information for both proteins—in copper-supplemented (+Cu) media, plastocyanin is abundant while in copperdeficient (–Cu) media, cytochrome is abundant (Wood, 1978; Sandmann et al., 1983; Sandmann, 1986). Krogmann and coworkers have suggested that this switch is important in natural populations of photosynthetic microorganisms where copper becomes a limiting trace element during an algal is abundant in such bloom. Indeed, cytochrome populations but when the organism is cultured in the laboratory in supplemented medium, the capacity for plastocyanin synthesis is evident (Ho et al., 1979). A discussion of the copper-responsive pathway in cyanobacteria and other algae can be found in recent reviews of plastocyanin and cytochrome function and biosynthesis (Morand et al., 1994; Merchant,

For most proteins of the photosynthetic apparatus, assembly of the apoprotein and cofactor is just one of many events that must occur to generate a functional electron transfer catalyst from a newly-translated polypeptide. In general, holoprotein formation tends to be a near-terminal step in the biosynthetic pathway of most metalloproteins. The in vivo process occurs rapidly and shows high substrate specificity, both for the polypeptide and the cofactor—features which are indicative of biologically catalyzed reactions. The distinctive cell biology and metabolism of the chloroplast suggests that metalloprotein assembly pathways in this organelle might exhibit some unique characteristics compared to analogous ones in other organisms. Since genetic approaches have been particularly useful in illuminating metalloprotein assembly in bacteria and mitochondria, it is expected that Chlamydomonas will be ideal for the dissection of such pathways in the chloroplast. This chapter will detail the present state of knowledge of plastocyanin, b- and c-type cytochrome assembly as deduced from studies with Chlamydomonas.

3. Metal Metabolism A third topic of relevance to metalloprotein biosynthesis in the chloroplast is metal uptake, sequestration and delivery to the site of metalloprotein assembly, and also the pathway and mechanism of metallocofactor biosynthesis. With the exception of the tetrapyrrole pathway, which is well-studied (Chapter 20, Timko), metal and metallocofactor metabolism in photosynthetic organisms has not been as intensively studied as it has in other organisms (e.g. Hausinger et al., 1990; Dean et al., 1993; de Silva et al., 1996). The mechanisms involved in the synthesis of iron-sulfur centers in chloroplasts and cyanobacteria is just beginning to be unraveled (reviewed in Merchant and Dreyfuss, 1998), but while Chlamydomonas has proven to be an excellent model for the delineation of molybdocofactor biosynthesis (Chapter 33, Fernandez et al.), it has not yet been exploited for studies of metallocofactor metabolism in the context of chloroplast protein assembly.

600 II. Copper-Responsive Synthesis of Plastocyanin and Cytochrome

A. Plastocyanin

1. Properties and Function Plastocyanin is a small (98 amino acids in C. reinhardtii) copper protein, Em ~ 370 mV, which functions in photosynthesis to transfer electrons from cytochrome f to PSI (reviewed by Boulter et al., 1977;Redinbo et al., 1994;Gross, 1996). It is referred to as a ‘blue’ copper protein owing to the spectroscopic properties of its copper-binding site, viz. a high extinction coefficient around 600 nm for the oxidized The protein was protein isolated originally from Chlorella ellipsoidea, and subsequently from a number of other photosynthetic organisms including Chlamydomonas (Gorman and Levine, 1966a). Several algal plastocyanins have been purified and the structures of three of these have been determined by application of X-ray crystallographic or NMR methods (Kunert et al., 1976; Yoshizaki et al., 1981, 1989; Moore et al., 1988; Collyer et al., 1990; Nakamura et al., 1992; Redinbo et al., 1993). The role of plastocyanin in photosynthesis was deduced by measurements of photosynthetic activity (whole cell fixation, Hill reaction, light-induced absorbance changes) in wildtype vs. mutant ac208 (re-named pcy1-ac208; Table 2) which carries a frame-shift mutation in the Pcy1 gene encoding pre-apoplastocyanin (Gorman and Levine, 1965; 1966c; Levine and Gorman, 1966; Quinn et al., 1993); this role was subsequently supported by demonstration of plastocyanin function in in vitro electron transfer assays with sonicated thylakoid membrane preparations (Hauska et al., 1971).

2. Genetic Information and Biosynthesis in Algae The protein is encoded in the nuclear genome as a precursor (Merchant and Bogorad, 1986; Li and Merchant, 1992; Nakamura et al. 1992; Quinn et al., 1993) and transported post-translationally (Howe and Merchant, 1993; Lawrence and Kindle, 1997) to the thylakoid lumen where it functions (Hauska et al., 1971; Haehnel et al., 1981). Although the vascular plant genes (PetE) are intron-less, the algal Pcy1 genes are interrupted (Quinn et al., 1993; Nakamura

Sabeeha Merchant et al., 1997; J. Quinn and S. Merchant, manuscript in preparation). An intron in the Scenedesmus obliquus and Pediastrum boryanum sequences occurs at exactly the same position as does the single intron in the Chlamydomonas Pcy1 gene; S. obliquus has a second intron which is unique to that organism. Plastocyanin is the most abundant copper protein in green algae because these organisms do not contain a Cu-Zn superoxide dismutase as do plants (Hewitt, 1983; Sakurai et al., 1993). The copper at the active site of plastocyanin is, naturally, essential for the redox function of the protein in photosynthesis and the protein is not expected to be functional in copperdeficient photosynthetic cells. Many green algae, including C. reinhardtii, remain photosynthetically competent nevertheless because they substitute a soluble, heme-containing cytochrome under these conditions (see below).

B. Regulation of Plastocyanin Accumulation

1. mRNA Accumulation in Algae and Cyanobacteria Plastocyanin does not accumulate in copper-deficient cultures of many green algae. The mechanisms of regulation are quite varied. In Scenedesmus obliquus, the abundance of mRNA encoding plastocyanin is reduced in copper-deficient cultures (Li and Merchant, 1992). The reduction in template is assumed to result in reduced synthesis, and hence, reduced accumulation of the protein in copperdeficient cells. The mechanisms contributing to differential copper-responsive accumulation of Pcy1 mRNA in S. obliquus is not yet known. In another alga, Pediastrum boryanum, Pcy1 mRNA accumulates in both copper-supplemented and copperdeficient cells but is not translatable in copperdeficient cells (Nakamura et al., 1997). The mRNA isolated from copper-deficient cells was noted to be shorter than that from copper-supplemented cells. The difference in size was attributed to the absence of the 5´ region (including part of the protein coding sequence) in the RNA from copper-deficient cells, which accounts for lack of synthesis of plastocyanin under these conditions. The mechanism by which the two transcripts are generated is not yet known, but the phenomenon certainly represents a novel example of metal-responsive gene expression (see also Section II.E.2).

Chapter 31

Biosynthesis of Metalloproteins

2. Degradation in C. reinhardtii In Chlamydomonas, the Pcy1 mRNA is abundant in phototrophically- or heterotrophically-grown cells. It is likely that the amount of mRNA template is in excess of that required for producing plastocyanin at the level required for photosynthesis. For instance, when strain pcy1-ac208 is rescued by transformation with the wild-type Pcy1 gene, transformants that accumulate only low levels of the Pcy1 mRNA (few percent relative to wild-type) can accumulate plastocyanin to levels similar to that found in the wild-type strains (e.g. Quinn et al., 1993). Thus, template abundance is not a major factor in determining the amount of plastocyanin in C. rein­ hardtii. The abundance of the mRNA is independent of the copper concentration of the medium; the pre-protein is synthesized and processed to the mature form at the same rate in copper-deficient and copper-sufficient cells (Merchant and Bogorad, 1986a,b). However, in copper-deficient cells, the protein is degraded in the thylakoid lumen whereas in copper-supplemented

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cells, it assembles with copper and accumulates (Fig. 1). Differential accumulation of plastocyanin in C. reinhardtii in response to copper is a consequence of the different half-life of the protein in copper-deficient vs. copper-replete cells. The apoprotein is thermodynamically de-stabilized and more protease-susceptible relative to the holoprotein, and this is one feature which contributes to the rapid degradation of plastocyanin in copper-deficient cells (Li and Merchant, 1995). The protease responsible for plastocyanin degradation has not been identified; it is likely to be lumen-localized and its activity is proposed to be increased in copper-deficient cells. By removing a major copper-binding protein, the cell would be ensuring copper availability for the biosynthesis and function of other essential copper enzymes, such as cytochrome oxidase.

C. Cytochrome

1. Properties and Function Cytochrome

is also a small (90 amino acids in

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C. reinhardtii), lumen-localized redox-active (Em ~ 370 mV for C. reinhardtii) protein. In earlier work, the protein had been named according to the absorption maximum of the band of the reduced protein. Thus, it has been called cytochrome or depending on the spectroscopic properties or

Sabeeha Merchant

of the particular preparation (Gorman and Levine, 1966b; Wood, 1977). However, the proteins from various algae and cyanobacteria are functionally and now structurally related; the name cytochrome defines this class of soluble c-type cytochromes (Pettigrew and Moore, 1987).

Chapter 31

Biosynthesis of Metalloproteins

Cytochrome was originally not distinguished from cytochrome f owing to the very similar spectroscopic properties of the two proteins (Gorman and Levine, 1966b;Kunert et al., 1976). The fact that the mutant strain ac206 (now named ccs1-ac206) lacked both proteins supported the argument that the two cytochromes were one and the same, although it is now known that the strain exhibits a pleiotropic cytochrome assembly defect (Section III.A.1). Wood (1977) purified both proteins from Chlamydomonas and demonstrated that cytochrome and cytochrome f were distinctly different proteins. He discovered that cytochrome was synthesized only under certain conditions in C. reinhardtii—either in stationary phase cultures that were not well-aerated or in copperdeficient cultures when plastocyanin function was compromised. On the basis of the reciprocal pattern of accumulation of cytochrome vs. plastocyanin as a function of copper availability in the medium, a number of researchers suggested that cytochrome served as a back-up for plastocyanin (Wood, 1978; Bohner et al., 1980) and this was supported by several lines of evidence, including: 1) its location in the thylakoid lumen (Wildner and Hauska, 1974); 2) its ability to substitute for plastocyanin in vitro in plastocyanin-depleted thylakoid membrane preparations (Hauska et al., 1971); 3) its similar physical properties—Em and pI—compared to plastocyanin (Ho and Krogmann, 1984); and 4) its occurrence in all photosynthetic algae and cyanobacteria that lack plastocyanin (Sandmann et al., 1983). The protein has been purified from several genera of algae and is biochemically well-characterized (Gorman and Levine, 1966b;Yakushiji, 1971;Bohme and Pelzer, 1982; Campos et al., 1993). The structure of Chlamydomonas Cyt reveals that it is a typical member of the soluble c-type cytochrome family; with a series the protein is predominantly and tight turns enveloping the heme of (Kerfeld et al., 1995).

2. Genetic Information and Biosynthesis In C. reinhardtii, cytochrome is encoded by a single nuclear gene (Merchant and Bogorad, 1987; Hill et al., 1991). As for plastocyanin, the protein is synthesized in precursor form and transported posttranslationally to the chloroplast via a lumen-targeting pathway (Howe and Merchant, 1993; see chapter 13, Perret et al.). In organisms that contain genetic information for both plastocyanin and cytochrome

603 the latter protein does not accumulate except under copper-deficient conditions, or occasionally, under microaerobic or anaerobic culture conditions (Wood, 1978; Bohner et al., 1980).

D. Regulation of Cytochrome

Synthesis

1. Copper as the Signal Cytochrome functions only as a replacement for plastocyanin—it is not synthesized unless plastocyanin function is compromised by copper-deficiency (Hill and Merchant, 1992). To test whether cytochrome synthesis occurs in response to plastocyanin deficiency (perceived, perhaps, via the redox state of the electron transfer chain) or directly in response to copper ion deprivation, Merchant and Bogorad (1987b) examined Cyt accumulation in a C. reinhardtii mutant that could not synthesize the plastocyanin polypeptide (strain pcy1-ac208). Accumulation of cytochrome remained copperresponsive in the mutant despite the absence of plastocyanin—pcy1mutants are, therefore, acetaterequiring in copper-supplemented medium but not in copper-deficient medium.

2. Transcriptional Regulation The copper-responsive regulation of the algal Cyc6 gene has been studied only for C. reinhardtii where it is under transcriptional regulation by copper (Hill et al., 1991; Merchant et al., 1991; Fig. 1). Cyc6 is not expressed in copper-replete cells (<1 mRNA per cell); in copper-deficient cells, the abundance of the Cyc6 mRNA is proportional to the plastocyanin deficiency resulting from lack of copper (Hill and Merchant, 1992). The Cyc6 mRNA accumulates to an abundance of several hundred molecules per cell when the gene is maximally induced. Copperresponsive elements (CuREs) associated with the Cyc6 gene are localized to a 90 nucleotide sequence in close proximity to the start site of transcription (Quinn and Merchant, 1995). These sites appear to be targets for transcriptional activators which must function downstream of the putative copper sensor in the signal transduction pathway. Site-directed mutagenesis indicates that GTAC forms the core of a CuRE, but the element is not characterized beyond that (J. Quinn, unpublished). Analysis of the CuREs in the context of various reporter gene constructs indicates that they account for all the copper-

604 responsive properties of the Cyc6 gene. The rapid and metal-selective response of the Cyc6 gene suggests that it may be useful for the design of chemically-regulated promoters for algal cells. The synthesis of plastocyanin and cytochrome has been examined also in C. smithii which has genetic information for both proteins, and the same pattern of reciprocal expression is observed (J. Quinn and S. Merchant, unpublished). On the other hand, C. mundana, which was isolated from sewage lagoons, lacks the capacity to synthesize plastocyanin (Wood, 1978). Accordingly, cytochrome accumulates in C. mundana independently of copper availability. This is true also for other algae and cyanobacteria which lack the genetic information for plastocyanin synthesis.

3. Response to Iron and Heme The Cyc6 gene does not respond at the transcriptional level to cellular iron status (K. Hill, unpublished). On the other hand, Cyt accumulation does depend on heme availability, since heme is a substrate for holoprotein formation (see below). In addition, heme (or a tetrapyrrole pathway intermediate) might serve also to regulate translation of the Cyc6 mRNA (Howe and Merchant, 1994a). This suggestion is based on the reduced synthesis of Cyt in cells depleted of heme by treatment with gabaculine (an inhibitor of synthesis), but the identity of the tetrapyrrole regulator and the mechanisms involved have not been studied. The absence of hem mutants of Chlamydomonas makes it difficult to distinguish between heme vs. tetrapyrrole pathway intermediates as regulators. The effect may not be specific to the translation of heme-containing proteins, since some reduction of plastocyanin and Rubisco small subunit synthesis was also noted. The synthesis of cytochrome f is also reduced in gabaculine-treated cells (e.g. Howe et al., 1995; Kuras et al., 1997), but the mechanisms involved in the chloroplast are expected to be different from those operating outside the organelle.

E. Other Responses to Copper-Deficiency 1. Copper Transport Chlamydomonas cells exhibit an extraordinary capacity for copper uptake. For instance, changes in

Sabeeha Merchant gene expression are evident in response to provision of copper at concentrations as low as a few nanomolar (Merchant et al., 1991). Also, copper-deficient cells can deplete all measurable copper from their growth medium even when it is provided in complex with a chelator (Hill et al., 1996). Measurement of copper uptake by Chlamydomonas indicates the presence of system in both – Cu a high affinity and +Cu cells; however, –Cu cells have a ten- to twenty-fold increased capacity for copper uptake relative to +Cu cells, perhaps owing to increased expression of genes encoding transporter components. The copper transporter has not yet been identified, but it is possible that it may be associated with a cupric reductase, as has been proposed for other organisms. Indeed, copper-deficient cells exhibit up to two-fold increased cupric reductase activity relative to copper-sufficient cells (Hill et al., 1996).

2. Coproporphyrinogen Oxidase Comparison of proteins synthesized by +Cu vs. –Cu cells revealed a 39 kD soluble protein, the synthesis of which was increased in copper-deficient relative to copper-sufficient cells (Merchant and Bogorad, 1986a). This protein was subsequently identified as the enzyme Coproporphyrinogen (coprogen) oxidase and its activity was shown to be several-fold greater in copper-deficient vs. copper-sufficient cells (Hill and Merchant, 1995). Northern analysis indicated a corresponding increase in the abundance of Cpx1 transcripts (encoding coprogen oxidase). Coprogen oxidase functions in tetrapyrrole biosynthesis and its increased activity in – Cu cells can be rationalized on the basis of an increased demand for heme in cells Coprogen that are synthesizing cytochrome oxidase is also induced in other organisms when there is an increased demand for heme. The induction cannot be mimicked by treatment with inhibitors of the tetrapyrrole pathway (e.g. gabaculine), suggesting that the increase in coprogen oxidase occurs in response to copper as the signal rather than to feedback from the tetrapyrrole pathway. The expression of genes encoding early enzymes of the acid tetrapyrrole pathway (such as dehydratase and glutamate semi-aldehyde aminotransferase) are not induced in –Cu cells—the response of the Cpx1 gene may well be unique. Coprogen oxidase is encoded by a single nuclear gene, Cpx1, in Chlamydomonas. The gene produces

Chapter 31

Biosynthesis of Metalloproteins

two transcripts, a longer form synthesized in both –Cu and +Cu cells, and a shorter form which is synthesized only in –Cu cells. The size difference is attributed to a difference in the length of the 5´ untranslated region. Therefore, the reading frame remains intact in both forms, unlike the situation for Pcy1 of P. boryanum (Section II.B.1). The abundance of the short form of Cpx1 is up to 35-fold higher under copper-deficient conditions relative to the long form, and this is attributed to transcriptional activation via CuREs (J. Quinn, unpublished). The significance of the two forms of Cpx1 mRNA is not yet known, but it makes one wonder whether the generation of alternate mRNA species with distinct 5´ ends may be a common regulatory mechanism in the copperresponsive signal transduction pathway of green algae. Analysis of the time-course, metal selectivity and metal sensitivity of the various responses to copperdeficiency suggest that a common regulatory pathway controls each of the above copper-responsive processes in C. reinhardtii including activation of Cyc6 and Cpx1 gene transcription, increased cupric reductase and copper transport, and increased plastocyanin degradation.

F. Adaptation to Copper-Deficiency The function of the regulatory pathway is viewed as an adaptation to copper-deficiency which allows the organism to survive in face of a changing supply of an essential, yet potentially toxic, micronutrient in its growth environment. While the stimulation of copper uptake is an immediate and predictable response, the substitution of a heme protein for a copper protein is a more long term and novel adaptation. Besides plastocyanin, Chlamydomonas contains at least two other copper proteins, cytochrome oxidase and urate oxidase. A substitute for cytochrome oxidase is not evident because severely copper-deficient cells are unable to grow heterotrophically in the dark—conditions where cytochrome oxidase function is essential (J. Quinn and S. Merchant, unpublished). The replacement of can be viewed plastocyanin with cytochrome therefore as a mechanism for ensuring re-distribution of copper from the photosynthetic apparatus to the essential respiratory complex.

605 III. Genetic Analysis of Chloroplast Metalloprotein Assembly

A. The c-type Cytochromes In the c-type cytochromes, the heme group is covalently attached to the polypeptide by thioether linkage between cysteinyl residues of a CxxCH sequence on the apoprotein and the vinyl side chains at C3 and C8 of the porphyrin ring. Consequently, the possibility that cytochrome assembly might be catalyzed has long been appreciated and the pathway of c-type cytochrome maturation has been studied in various organisms, including Chlamydomonas (reviewed by Gonzales and Neupert, 1990; Howe and Merchant, 1994b; Kranz and Beckman, 1995; Thony-Meyer, 1997).

1. The ccs Mutants Chlamydomonas contains two c-type cytochromes: Cyt f of the complex (Chapter 14, Olive and (discussed Wollman) and soluble cytochrome above). The post-translational pathway for maturation of cytochrome was determined by in vivo radiolabeling experiments which established the following sequence in vivo—pre-apocyt intermediate-apocyt mature apocyt holocyt —and placed the heme attachment reaction in the thylakoid lumen (Howe and Merchant, 1993; 1994a). Thus, as for other c-type cytochromes, assembly occurs on the p-side of the energy transducing membrane, and as for most metalloproteins, holoprotein formation is a near terminal step in the biosynthetic pathway. The ability to monitor the biosynthetic pathway in vivo by application of radiolabeling techniques facilitated the definition of heme attachment mutants from a collection of acetaterequiring strains (Howe and Merchant, 1992; Xie et al., 1998). The pathway of cytochrome maturation in these strains, called ccs for c-type cytochrome synthesis, is normal with respect to the rate of synthesis and processing of the pre-apoprotein, but the apoprotein is not converted to the holoform to any appreciable extent. The ccs strains are pleiotropically deficient in both chloroplast c-type cytochromes. Since cytochrome f is encoded in the plastid genome (petA) and cytochrome in the nuclear genome (Cyc6), the post-translational steps involving translocation across

606 the thylakoid membrane and heme attachment are the only steps that could be common in their biosynthesis. While translocation across the thylakoid membrane is a process that is shared with other lumen proteins (Chapter 13, Perret et al.), heme attachment is required only for the c-type cytochromes. The recognition of the pleiotropic Cyt minus/Cyt f-minus phenotype therefore simplified the screen for additional candidate ccs strains (Table 2). Sixteen such strains have been placed into five complementation groups—ccsA corresponding to the plastid ccsA gene, and CCS1 through CCS4 corresponding to four nuclear genes (Xie et al., 1998). Since some of the loci are represented by only one mutant strain, it is possible that the pathway has not yet been saturated by mutagenesis and additional loci might yet be discovered.

2. The Ccs Factors

a. CcsA The plastid ccsA gene, which rescues all three uniparental mutants ct34, ct59 and B6, corresponds to an open reading frame (formerly called ycf5) found in all plastid genomes characterized to date (Reith, 1995). CcsA had been proposed to be a cytochrome biogenesis protein on the basis of limited sequence similarity with Ccl1, a Rhodobacter capsulatus protein required for the synthesis of all c-type cytochromes in the bacterial periplasm (Beckman et al., 1992). Indeed, inactivation of the C. reinhardtii open reading frame led to a nonphotosynthetic strain whose phenotype was attributed to a pleiotropic cytochrome deficiency (Xie and Merchant, 1996). The C-terminal region of the protein is the most highly conserved and includes a functionally important sequence motif, WGxxWxWDxxE, found originally in Ccll of Rb. capsulatus. CcsA is highly hydrophobic and probably spans the membrane multiple times. The WGxxWxWDxxE motif is predicted to lie on the lumen-side of the membrane which is consistent with a role for this sequence in some part of the heme attachment reaction..

b. Ccs1 The Ccs1 gene was identified on the basis that it was disrupted in a cytochrome -deficient strain—abf3 (Inoue et al., 1997). The authors recognized a

Sabeeha Merchant pleiotropic c-type cytochrome deficiency in this strain and determined on the basis of further biochemical characterization that abf3 was blocked at the step of heme attachment. Although abf3 was not analyzed genetically, the fact that the cloned gene rescued all ccs1 mutants (B. Dreyfuss and S. Merchant, unpublished) but not other ccs strains indicated that the cloned gene corresponded to the CCS1 locus. Candidate homologues of Ccs1, encoded by ycf44 (for hypothetical chloroplast frame) occur in Synechocystis sp. 6803 and the plastid genomes of several algae, which suggests that, like CcsA, Ccs 1 functions in all plastids. Analysis of the sequence indicates that Ccs 1 is a membrane protein—but its specific biochemical function has not yet been deduced. The availability of ccs1 mutants which can be rescued by the cloned gene opens the door to functional analysis of this novel protein. Ccs1 is absent in ccsA mutants, suggesting that CcsA may form a complex with Ccs1 in vivo (B. Dreyfuss, unpublished) The abundance of ccsA and Ccs1 mRNAs is very low compared to the abundance of the petA or Cyc6 mRNAs, which suggests that the gene products are low abundance proteins: this is consistent with a catalytic role of Ccs factors in the assembly of the petA and Cyc6 gene products. A number of functions may be envisioned for the Ccs factors. One possibility is that the membrane-associated components may serve as subunits of a heme transporter which would deliver heme from its site of synthesis to the lumen side of the thylakoid membrane. Some of the other subunits of the putative Ccs complex might serve as substrate (apoprotein/heme) chaperones or as cytochrome c/heme lyases. By analogy to the thioredoxin-like proteins which are required for sequential oxidation and reduction of apocytochromes in the bacterial periplasm, one might propose that similar reactions might be catalyzed by the chloroplast Ccs factors. Alternatively, an oxidoreductase might be required to maintain the heme iron in the reduced state.

3. A Novel Pathway in the Chloroplast Previously, two pathways for cytochrome biogenesis were known. One pathway, discovered originally by genetic analysis of cytochrome c and biogenesis in Saccharomyces cerevisiae and Neurospora crassa, operates in fungal, mammalian, Drosophila and C. elegans mitochondria and consists of the Cyt c­

Chapter 31

Biosynthesis of Metalloproteins

607

and lyases (reviewed by Gonzales and Neupert, 1990). Another, discovered originally by analysis of bacterial cytochrome c-deficient mutants, operates in many proteobacteria—including rhizobia, Rhodobacter spp., E. coli, Pseudomonas spp.—and plant mitochondria, and consists of a multicomponent cytochrome assembly complex in the energy transducing membrane (reviewed by Kranz and Beckman, 1995; Thony-Meyer, 1997). Analysis of the distribution of CcsA and Ccs1 homologues in the genome databases indicates that they define a third, novel pathway for c-type cytochrome biogenesis which operates not only in plastids and cyanobacteria but also in a subset of bacterial species including Bacillus subtilis, Mycobacterium tuberculosis, Mycobacterium leprae, Neisseria spp. and Helico­ bacter pylori (discussed in Xie et al., 1998). Candidate homologues of CcsA and Ccs1 are encoded in an operon-like arrangement in some of these bacteria, suggesting that the other open reading frames in the operon may correspond to products encoded at the remaining CCS loci. The putative Ccs factors appear to be essential in bacteria and they have not been subject to functional analyses in these organisms. The possibility of combining genetic and biochemical methodologies makes Chlamydomonas an ideal experimental system for the study of the Ccs pathway of cytochrome biogenesis.

the following pathway for sequential heme insertion, which occurs post-translationally and independently of inter-subunit associations: apocyt + heme holocyt heme-dependent intermediate + heme (Kuras et al., 1997; see also chapter 24, Wollman). The elucidation of this pathway allowed the identification of ccb strains among a collection of cytochrome -deficient mutants (Kuras et al., 1997). The phenotype of the ccb strains mimicked exactly the phenotype of strains carrying site-directed mutations at the ligands, and the block could therefore be assigned to the step where the heme-dependent intermediate assembles with heme to yield holocytochrome . Genetic analysis classified the mutants to four nuclear loci (Table 2), but the genes have not yet been cloned and the functions of the gene products can only be speculated upon. These include cofactor processing (delivery, chaperoning), apoprotein processing or catalysis of heme insertion. The catalyzed assembly of b-type cytochromes has not been studied in any system; thus, the further characterization of the Chlamydomonas pathway will undoubtedly lead to novel insights into cofactor-protein assembly. Since the association of the b-hemes with the protein was thought to be non-covalent, the need for catalysis had not been appreciated until the identification of the ccb mutants.

B. The b-Heme(s) of the

C. Plastocyanin

f Complex

The definition of other assembly defective mutants has not been as straightforward because it can be difficult to distinguish the phenotype of a cofactorinsertion defect from a defect associated with a cofactor-independent step (such as subunit-subunit association). In general, unassembled photosynthetic complexes are degraded, especially in Chlamydomonas chloroplasts, regardless of the nature of the defect. Thus, a c-heme insertion mutant does not accumulate any of the subunits of the cytochrome complex, and this phenotype is not significantly different from a b-heme insertion mutant (Kuras et al., 1997; discussed below). Nevertheless, specific defects can be distinguished if the assembly pathways can be monitored in vivo in wild-type cells so that intermediates are visualized. For cytochrome maturation, Wollman and co-workers exploited pulseradiolabeling techniques, in combination with sitedirected mutagenesis of cofactor binding ligands, to identify biosynthetic intermediates and to deduce

A similar approach, namely screening nonphotosynthetic strains for specific assembly defects, led to the identification of two loci required for plastocyanin biosynthesis—PCY1, corresponding to the Pcy1 gene encoding pre-apoplastocyanin and PCY2 (Li et al., 1996). In the pcy2-1 mutant, plastocyanin is synthesized and processed as in wildtype cells, but the apoprotein accumulates at the expense of the holoprotein (Fig. 1). The accumulation of apoplastocyanin is consistent with in organello studies using plant chloroplasts, which established that copper assembles with apoplastocyanin in the thylakoid lumen after processing by the thylakoid peptidase (Li et al., 1990). Studies of copper-protein assembly in other organisms indicates that the pcy2 phenotype might be attributable to a defect in copper delivery to the lumen or copper delivery to the active site of plastocyanin (Zumft et al., 1990; Chen et al., 1993; Glerum et al., 1996). Nevertheless, the possibility that the phenotype results from post-

608 assembly de-stabilization of holoplastocyanin cannot be ruled out at present.

D. Approaches for the Dissection of Other Assembly Pathways Several strategies are being employed for the study of metalloprotein assembly in photosynthetic organisms (reviewed by Merchant and Dreyfuss, 1998). One approach exploits genome sequence information in the databases to identify candidate genes (and hence gene products) involved in the pathway of interest on the basis of relationships to well-characterized proteins of known function. In this situation, the gene products can be tested for function in the pathway of interest if a biochemical assay for metalloprotein assembly is available, and reverse genetic approaches can be applied to test for relevance of that function to the in vivo biosynthetic pathway. Assembly factors that function in the chloroplast can also be cloned on the basis of their ability to complement mutations in homologous genes in other organisms. This approach has been applied to the study of molybdenum cofactor biosynthesis (e.g. Stallmeyer et al., 1995) and has also been particularly useful for the cloning of various metal transporters of plants (e.g. Eide et al., 1996). The third approach, involving screening for mutants with assembly defects followed by the identification of the wild-type loci and assembly factors, has enormous potential because it can lead to the discovery ofnovel factors (such as the Ccs proteins) or reveal the existence of previously-undefined pathways (such as for holocyt synthesis). Furthermore, the availability of mutant strains can simplify functional studies by site-directed mutagenesis. Chlamydomonas is an excellent model system for such an approach. The key to identifying mutants of interest is the recognition of a cofactor assembly defect: when apoproteins accumulate as in strain pcy2-1 (Li et al., 1996) or the nia mutants of tobacco (Gabard et al., 1987; Chapter 33, Fernandez et al.), the assignment is simple, but this is generally not the case for proteins of the photosynthetic apparatus. Many of the cofactors in the photosynthetic complexes are integral to their structure, and cofactorbinding mutants often do not accumulate cofactordepleted versions of the complex of interest (e.g. Whitmarsh et al., 1991, and ccb or ccs mutants discussed above). In this case, assembly defects are deduced when the mutation: 1) is not in the genes

Sabeeha Merchant encoding the various polypeptide constituents ofthe complex; and 2) does not affect the expression of individual subunits of a complex (e.g. Voelker and Barkan, 1995). In vitro systems for the assay of assembly, or the ability to monitor assembly in vivo, are pre-requisites for the more precise localization of the defect to a particular assembly step involving cofactor insertion. The Chlamydomonas system is particularly suited for the development of in vivo assays owing to the facility with which radiolabeling experiments can be executed.

IV. Conclusions Chlamydomonas has served as a useful model for studying metal-responsive gene expression and metalloprotein assembly in the context of assembly of the photosynthetic apparatus. The copperdependent regulation of plastocyanin and cytochrome remains one of the best characterized adaptations to a trace element deficiency. However, the regulatory molecules involved in sensing cellular copper status and responding to it have not been identified. The use of genetic approaches for dissecting this regulatory pathway holds great promise and needs to be exploited. Iron metabolism is another topic of great interest. C. reinhardtii cells display dramatic responses to iron starvation, including de-greening and changes in gene expression; however, little is known about the molecular basis of adaptation. The recent discovery ofan iron-regulated transferrin-like protein in Dunaliella salina suggests that studies of iron metabolism in green algae is bound to lead to new findings (Fisher et al., 1997). Likewise, the rich repertoire of non-photosynthetic mutants and the facilitywith which they can be generated and analyzed ensures that this experimental system will continue to contribute to our understanding of metalloprotein assembly processes.

Acknowledgments The work in my laboratory in this area has been supported by grants from the NIGMS ofthe NIH and the US Department of Agriculture. I thank the members of my laboratory for their contributions to the work discussed in this chapter, and Catherine deVitry, Mats Eriksson, Jeffrey Moseley and Jeanette Quinn for their comments on the manuscript.

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Biosynthesis of Metalloproteins

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Sabeeha Merchant of photosynthetic metalloproteins. Annu Rev Plant Physiol Plant Mol Biol 49: 25–51 Merchant S, Hill K, Kim JH, Thompson J, Zaitlin D and Bogorad L (1990) Isolation and characterization of a complementary DNA clone for an algal pre-apoplastocyanin. J Biol Chem 265: 12372–12379 Merchant S, Hill K and Howe G (1991) Dynamic interplay between two copper-titrating components in the transcriptional EMBO J 10: 1383–1389 regulation of Cyt Moore JM, Case DA, Chazin WJ, Gippert GP, Havel TF, Powls R and Wright PE (1988) Three-dimensional solution structure of plastocyanin from the green alga Scenedesmus obliquus. Science 240: 314–317 Morand LZ, Cheng RH, Krogmann DW and Ho KK (1994) Soluble electron transfer catalysts of cyanobacteria. In Bryant DA (ed) The Molecular Biology of Cyanobacteria, pp 381– 407. Kluwer Academic Publishers, Dordrecht Nakamura M, Yamagishi M, Yoshizaki F and Sugimura Y (1992) The syntheses of plastocyanin and cytochrome c-553 are regulated by copper at the pre-translational level in a green alga, Pediastrum boryanum. J. Biochem. 111: 219–224 Nakamura M, Yoshizaki F, Sugimura Y (1997) Regulation by copper of the expression of plastocyanin in a green alga, Pediastrum boryanum. Plant Physiol 114 (suppl.):136 Okamoto Y, Minami Y, Matsubara H and Sugimura Y (1987) Studies on algal cytochromes VI: Some properties and amino acid sequence of cytochrome from a green alga, Bryopsis maxima. J Biochem 102: 1251–1260 Pakrasi HB, de Ciechi P and Whitmarsh J (1991) Site directed mutagenesis of the heme axial ligands of cytochrome b559 affects the stability of the Photosystem II complex. EMBO J 10: 1619–1627 Pettigrew GW and Moore GR (1987) Cytochromes c. Biological Aspects. Springer-Verlag, New York Quinn JM and Merchant S (1995) Two copper-responsive elements associated with the Chlamydomonas Cyc6 gene function as targets for transcriptional activators. Plant Cell 7: 623–638 Quinn JM, Li HH, Singer J, Morimoto B, Mets L, Kindle K and Merchant S (1993) The plastocyanin-deficient phenotype of Chlamydomonas reinhardii Ac-208 results from a frame-shift mutation in the nuclear gene encoding pre-apoplastocyanin. J Biol Chem 268: 7832–7841 Redinbo MR, Cascio D, Choukair MK, Rice D, Merchant S and Yeates TO (1993) The 1.5-Å crystal structure of plastocyanin from the green alga Chlamydomonas reinhardtii. Biochemistry 32: 10560–10567 Redinbo MR, Yeates TO and Merchant S (1994) Plastocyanin: Structural and functional analysis. J Bioenerg Biomembr 26: 49–66 Reith M (1995) Molecular biology of rhodophyte and chromophyte plastids. Annu Rev Plant Physiol Plant Mol Biol 46: 549–575 Sakurai H, Kusumoto N, Kitayama K and Togasaki RK (1993) Isozymes of superoxide dismutase in Chlamydomonas and Purification of one of the major isozymes containing Fe. Plant Cell Physiol 34: 1133–1137 Sandmann G (1986) Formation of plastocyanin and cytochrome c-553 in different species of blue-green algae. Arch Microbiol 145: 76–79 Sandmann G and Malkin R (1983) Iron-sulfur centers and

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activities of the photosynthetic electron transport chain in irondeficient cultures of the blue-green alga Aphanocapsa. Plant Physiol 73: 724–728 Sandmann G, Reck H, Kessler E and Boger P (1983) Distribution of plastocyanin and soluble plastidic cytochrome c in various classes of algae. Arch Microbiol 134: 23–27 Stallmeyer B, Nerlich A, Schiemann J, Brinkmann H and Mendel RR (1995) Molybdenum co-factor biosynthesis: The Arabidopsis thaliana cDNA cnx1 encodes a multifunctional two-domain protein homologous to a mammalian neuroprotein, the insect protein Cinnamon and three Escherichia coli proteins. Plant J 8: 751–762 Straus NA (1994) Iron deprivation: Physiology and gene regulation. In Bryant DA (ed) The Molecular Biology of Cyanobacteria, pp 731–750. Kluwer Academic Publishers, Dordrecht Thony-Meyer L (1997) Biogenesis of respiratory cytochromes in bacteria. Microbiol Mol Biol Rev 61: 337–376 Voelker R and Barkan A (1995) Nuclear genes required for posttranslational steps in the biogenesis of the chloroplast cytochrome f complex in maize. Mol Gen Genet 249: 507– 514 Wildner GF and Hauska G (1974) Localization of the reaction site of cytochrome 552 in chloroplasts from Euglena gracilis. Arch Biochem Biophys 164: 136–144 Wood PM (1977) The roles of c-type cytochromes in algal

611 photosynthesis. Eur J Biochem 72: 605–612 Wood PM (1978) Interchangeable copper and iron proteins in algal photosynthesis. Eur J Biochem 87: 9–19 Xie Z and Merchant S (1996) The plastid-encoded ccsA gene is required for heme attachment to chloroplast c-type cytochromes. J Biol Chem 271: 4632–4639 Xie Z, Culler D, Dreyfuss BW, Kuras R, Wollman F-A, GirardBascou J and Merchant S (1998) Genetic analysis of chloroplast c-type cytochrome assembly: One chloroplast locus and at least four nuclear loci are required for heme attachment. Genetics 148: 681–692 Yakushiji E (1971) Cytochromes: Algal. Methods Enzymol 23: 364–368 Yoshizaki F, Sugimura Y and Shimokoriyama M (1981) Purification, crystallization, and properties of plastocyanin from a green alga, Enteromorpha prolifera. J Biochem 89: 1533–1539 Yoshizaki F, Fukazawa T, Mishina Y and Sugimura Y (1989) Some properties and amino acid sequence of plastocyanin from a green alga, Ulva arasakii. J Biochem 106: 282–288 Zumft WG, Viebrock-Sambale A and Braun C (1990) Nitrous oxide reductase from denitrifying Pseudomonas stutzeri. Genes for copper-processing and properties of the deduced products, including a new member of the family of ATP/GTP-binding proteins. Eur J Biochem 192: 591–599

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Chapter 32 Responses to Deficiencies in Macronutrients John P. Davies and Arthur R. Grossman The Carnegie Institution of Washington, Department of Plant Biology, 260 Panama Street, Stanford, CA 94305, U.S.A.

Summary I. Introduction II. Nutrients in the Environment A. Sulfur B. Phosphorus III. Specific Responses A. Transport Processes 1. Sulfate Transport 2. Phosphate Transport 3. Ammonium Transport 4. Inorganic Carbon Transport B. Accessing Alternative External Sources of Nutrients 1. Sulfur 2. Phosphorus 3. Nitrogen 4. Inorganic Carbon C. Accessing Internal Stores of Nutrients 1. Sulfur 2. Phosphorus 3. Nitrogen 4. Inorganic Carbon IV. Common Responses A. Cell Division B. Nutrient Stress and Photosynthesis 1. The ‘Light’ Reactions 2. Non-Photochemical Quenching 3. The ‘Dark’ Reactions C. Respiration and Nutrient Stress V. Model Integrating the Responses to Nutrient Deprivation VI. Regulation of the Responses to Nutrient Deprivation A. Sulfur B. Phosphorus C. Nitrogen D. Inorganic Carbon VII. Identification of Mutants Deficient in the Acclimation to Nutrient Deprivation Acknowledgments References

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J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 613–635. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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Summary All organisms are able to acclimate to less than optimal environmental conditions. In this chapter we discuss how Chlamydomonas reinhardtii acclimates to deficiencies in macronutrients. The physiological changes that occur during macronutrient starvation can be classified as specific and common responses. Specific responses occur during limitation for a particular nutrient, while common responses occur during limitation for any nutrient. Specific responses include changes in nutrient transport characteristics and induction of nutrient scavenging systems. General responses include decreases in cell division, photosynthesis and respiration. The decrease in photosynthesis has been shown to be essential for surviving severe sulfur-stress as a mutant unable to decrease photosynthesis cannot survive these conditions. We propose a model of how specific and general responses to nutrient limitation are regulated and discuss how mutants deficient in these responses fit into this model.

I. Introduction The ability of an organism to adapt to changes in its environment is essential for both optimal growth and survival. The acclimation response is complex and highly regulated. It must integrate the monitoring of the environment with control over genetic and physiological responses. In this chapter, we will discuss the strategies that Chlamydomonas reinhardtii uses to acclimate to deficiencies in macronutrients. However, where information is lacking we will discuss acclimation mechanisms of other organisms. We will focus primarily on the responses to growth in medium lacking sulfur and phosphorus, but because there are conceptual similarities in the acclimation to deficiencies in nitrogen and carbon, we will discuss briefly the responses to these deficiencies. The chapters on nitrogen metabolism (Chapter 33, Fernández et al.) and the carbon concentrating mechanism (Chapter 28, Spalding) deal with these topics more fully. Many of the responses of C. reinhardtii to nutrient deficient conditions are similar to those exhibited by Abbreviations: APS – 5´ adenosyl-3´ phosphosulfate; Ars – Arylsulfatase; CCM – carbon concentrating mechanism; DHAP – dihydroxyacetone phosphate; FBP – fructose-1,6-bisphosphate; quantum efficiency of photosynthesis; G3P – glycer– concentration at which velocity is aldehyde-3-phosphate; half-maximal; LHCI – light harvesting complex of Photosystem I; LHCII – light harvesting complex of Photosystem II; MAP – mitogen activated protein; NCS – 4-nitrocatecholsulfate; NPQ – non-photochemical quenching; PAPS – 3´-phosphoadenosyl-5´ phosphosulfate; PCR – photosynthetic carbon reduction; PEP – phosphoenol pyruvate; PK – pyruvate kinase; PS I – Photosystem I; PS II – Photosystem II; Rubisco – ribulose bisphosphate carboxylase/oxygenase; RuBP – ribulose bisphosphate; TCA–tricarboxylic acid; –maximal velocity; – 5-bromo-4-chloro-3-indolyl phosphate; –5-bromo4-chloro-3-indolyl sulfate

vascular plants and other soil dwelling microorganisms. Here, we will review physiological and metabolic changes that accompany nutrient limitation, the ways in which these changes are regulated, and the sensors that perceive the nutrient status ofthe environment. Thephysiological responses to nutrientdeprivation can be classified as those that occur in response to limitation for a specific nutrient (specific responses), or any nutrient (common responses). The specific responses include alterations in transport processes that enable cells to import the limiting nutrient more efficiently (Badger et al., 1980; Franco et al., 1987; Quesada et al., 1993; Palmqvist et al., 1994; Yildiz et al., 1994); (Shimogawara and Grossman, unpublished) and the induction of scavenging enzymes that enable the cell to access alternative sources of the limiting nutrient. These scavenging enzymes are either exported from the cell to mobilize alternative sources of nutrients by degrading compounds in the environment (Lien and Schreiner, 1975; Loppes et al., 1977; Patni et al., 1977; Nagy et al., 1981; Kimpel et al., 1983; de Hostos et al., 1988; Muñoz-Blanco et al., 1990; Quisel et al., 1996), or maintained within the cell to recycle nutrients by degrading proteins, RNA, and lipids (Siersma and Chiang, 1971; Moroney et al., 1987; Husic et al., 1989; Plumley and Schmidt, 1989; Ferreira and Teixeira, 1992; Löffler et al., 1992;Siderius et al., 1996). The common responses include ceasing cell division (Lien and Knutsen, 1973; Pringle and Hartwell, 1981; Davies et al., 1996), accumulating starch (Ball et al., 1990), and slowing photosynthesis (Badger et al., 1980; Peltier and Schmidt, 1991; Davies et al., 1996; Wykoff et al., 1997) and respiration (Theodorou and Plaxton, 1993; Gauthier and Turpin, 1994). In the broad sense, nutrient deprivation causes the diversion of energy use away

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Nutrient Stress

from growth by inhibiting cell division, and at least during the initial stages of nutrient limitation, toward the accumulation of storage compounds such as starch. Persistent exposure to nutrient-deficient conditions can result in a nearly complete loss of photosynthetic and respiratory activities. The responses to nutrient-deficient conditions are highly regulated (Lien and Schreiner, 1975; de Hostos et al., 1988; Davies et al., 1994; Quisel et al, 1996). When an essential nutrient falls below a critical level for sulfate) (de Hostos et (e.g. approximately al., 1988), the cells sense the limitation and express a specific set of genes that helps them alter their physiology and acclimate to the deficiency. We will discuss the recent advances in our understanding of how C. reinhardtii senses and responds to the nutrient status of the environment, and the experimental strategies that have been critical for making these advances.

II. Nutrients in the Environment

A. Sulfur Sulfur is an essential nutrient present in proteins, lipids, intermediary metabolites and components of electron transport chains. Sulfur is most commonly imported into the cell as the sulfate anion (Beil et al., 1996). Once inside the cell, sulfate is activated by ATP sulfurylase to form 5´ adenosyl-3´-phosphosulfate (APS), which can be transferred directly to a thiosulfate reductase and reduced by sulfite reductase (Schmidt, 1973), or APS can be phosphorylated by APS kinase to form 3´-phosphoadensine-5´-phosphosulfate (PAPS). The sulfate in PAPS may be transferred to a carrier and reduced by sulfite reductase. The reduced sulfur is then used in the biosynthesis of amino acids and other sulfur containing compounds (Schmidt, 1986). Although the free sulfate anion is the preferred source of sulfur for most organisms, it is often not the most common form in the environment. In fact, inorganic sulfate is only a minor (1–15%) component of the sulfur in many soils (David et al., 1982; Autry and Fitzgerald, 1990; Whalen and Warman, 1996b). Most of the soil sulfur is covalently associated with organic molecules; the predominant forms are sulfate esters (C-O-S bonds) and sulfonates (C-S bonds) (Stanko-Golden and Fitzgerald, 1991; Houle and Carignan, 1992). Figure 1 illustrates the cycling of sulfur in the

615 environment. The soluble sulfate within the soil solution is in a dynamic state and influenced by several factors. Sulfur is added to the soil as sulfate through acid rain, and salts through fertilization, organic sulfur through the decomposition of organic matter (Freney et al., 1975; Chapman, 1987; StankoGolden and Fitzgerald, 1991). can be oxidized to sulfate by the atmosphere (Benner et al., 1992) and organic sulfur converted into inorganic sulfate by enzymes in the soil (Strickland and Fitzgerald, 1983; Fitzgerald and Watwood, 1985). Free sulfate may be taken up by plants and microbes, leach through the soil matrix, adsorb onto soil particles, or become covalently bound to organic molecules in the soil (Strickland et al., 1986; Dhamala and Mitchell, 1995). Radioactive tracer studies have demonstrated that introduced sulfate is rapidly converted into sulfate esters and sulfonates (Swank et al., 1984; Strickland et al., 1986; Dhamala and Mitchell, 1995), forms of sulfur not imported by most organisms. These molecules can be converted into higher molecular weight complexes (Whalen and Warman, 1996a), or broken down through the action of extracellular degrading enzymes (Strickland and Fitzgerald, 1983; Fitzgerald and Andrew, 1985; Fitzgerald and Watwood, 1985; Lou and Warman, 1992; Whalen and Warman, 1996a). The higher molecular weight complexes must be broken down to smaller forms before they are accessible to the sulfate ester and sulfonate degrading enzymes. The sulfur in sulfate esters and sulfonates differ both in the atoms to which they are bonded and their oxidation state. In sulfate esters the sulfur is bonded through an oxygen to a carbon, and the oxidation state of the sulfur is +6. The sulfate can be removed from the carbon backbone by a hydrolysis reaction catalyzed by a sulfatase. The released sulfate may then be transported into the cell (Fitzgerald and Strickland, 1987; Lou and Warman, 1992; Whalen and Warman, 1996b). In sulfonates the sulfur is bonded directly to a carbon atom and the oxidation state of the sulfur is +4. It is not clear how sulfonates are metabolized, but there is evidence from studies with E. coli that it is not necessary to oxidize the sulfur to sulfate prior to its use since ATP sulfurylase and APS kinase are not required for growth on sulfonates. However, sulfite reductase is required for the use of sulfonates, suggesting that E. coli uses the partially reduced state of sulfur in sulfonates in the normal reductive pathway (Uria-Nickelsen et al., 1993). One prominent sulfonate present in photo-

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synthetic organisms is the sulfolipid of the thylakoid membranes, 6-sulfoquinovosyl diacylglycerol, which may play an important role in photosynthetic electron transport (Sato et al., 1995a; Chapter 2l,Trémolières). The sulfonated moiety of the sulfolipid can be present at relatively high levels in the soil. Strickland and Fitzgerald (Strickland and Fitzgerald, 1983) have demonstrated that the sulfur in 6-sulfoquinovose, the sugar sulfonate of this sulfolipid, can be recycled in -sulfoquinovose is incubated in the soil. When the soil, the radio-labeled sulfur is converted into both inorganic sulfate and organic sulfate esters in a time dependent manner. The reuse of this sulfur can be inhibited by antibiotics, indicating that these reactions are mediated by bacteria (Strickland and Fitzgerald, 1983). The enzymes that catalyze the synthesis of sulfate esters and sulfonates, as well as the synthesis and degradation of the higher molecular weight compounds, have not been well characterized. In contrast, several sulfate ester degrading enzymes (sulfatases) from a variety of different organisms

John P. Davies and Arthur R. Grossman

(Klebsiella nidulans, Pseudomonas C12B, Neuro­ spora crassa, Chlamydomonas reinhardtii, Asper­ gillus nidulans) have been isolated and characterized. The synthesis of these sulfatases is regulated by the availability of the free sulfate anion in the environment (Scott and Metzenberg, 1970; Apte et al., 1974; Lien and Schreiner, 1975;de Hostos et al., 1988; Murooka et al., 1990; Beil et al., 1996). In many soils, there is a clear inverse correlation between the level of free sulfate and arylsulfatase activity (Stanko and Fitzgerald, 1990; Whalen and Warman, 1996a). Hence, elevated arylsulfatase activity is indicative of low levels offree sulfate in the soil. Because sulfur is often bonded to organic molecules and cannot be recycled until it is freed by inducible or derepressible sulfur releasing enzymes, these enzymes and the processes that regulate their expression are critical for the functioning of the sulfur cycle and the health of the ecosystem. Limitations in accessible forms of sulfur are likely to influence the composition of ecosystems and may limit plant productivity in agricultural settings. In

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Nutrient Stress

several studies, the application of sulfur-containing fertilizers has resulted in improved crop yields (Mahler and Maples, 1987; Warman and Sampson, 1994). For example, the growth of wheat on low sulfur soils resulted in lower yields and diminished quality of grain compared with wheat grown on the same soils fertilized with sulfate. In general, seed yield and quality are more sensitive to sulfur limitation than plant growth and development. However, severe sulfur deprivation can lead to considerable stunting of growth (Mahler and Maples, 1987). In recent years, sulfur-limited crop yields have been detected. In the past, high levels of sulfur were inadvertently added to soils through the application of sulfate contaminated fertilizers and precipitation of acid rain (Cole and Johnson, 1977; Johnson et al, 1982; David et al., 1988; Mac Donald et al., 1991). However, because the purity of fertilizers has improved and acid rain has diminished, there has been a notable rise in the number of soil environments that are limited for sulfur (Marschner, 1995). Monitoring the sulfur status of soils will become increasingly important to insure optimal crop yields.

617 soluble phosphate is limiting. Adsorbed phosphates are in equilibrium within the soil solution (Jungk et al., 1993) and calcium-phosphates can be mobilized by carboxylates (malate and citrate) that are synthesized by plants and released into the rhizosphere when available phosphate in the environment is low (Gerke et al., 1994). Extracellular organic phosphorus is mobilized via the action of hydrolytic enzymes which include acidic and alkaline phosphatases, and phytases (Beck et al., 1989; Tarafdar and Marschner, 1995; Seeling and Hungk, 1996). These enzymes, which catalyze the hydrolysis of phosphate from organic compounds, are induced in both plants and microbes when the soluble phosphate levels become low (Jungk et al., 1993; Joner and Jakobsen, 1995; Seeling and Hungk, 1996). It has been estimated that the activity of soil phosphatases generates approximately one third of the phosphorus that is used by field grown barley plants (Jungk et al., 1993). As with sulfur, the ability of organisms to induce phosphate-releasing enzymes is a critical element of the phosphorus cycle and maintenance of the ecosystem.

B. Phosphorus III. Specific Responses Phosphorus is a macronutrient present in many essential biological compounds including nucleic acids, lipids and intermediary metabolites. Phosphorus is present both in the intracellular and extracellular environment as the free phosphate anion, precipitated phosphate salts, phosphate esters and phytates (Ron Vaz et al., 1993; Pant et al., 1994). Soluble phosphate is the form most often taken up by plants and microbes. However, much of the phosphate present in the soil solution is tightly adsorbed to soil particles, complexed with calcium and iron, orpresent as organic phosphates; these forms are not directly available to plants and microbes (Jungk et al., 1993). Yet, organisms must be able to access these complex forms of phosphate since the level of soluble phosphate in the soil is very often too low to support growth and reproduction. For example, in the arable lands of Europe, the soluble soil phosphate concentration is approximately (or which is only about 3% of the level required for optimal production by grain crops (Jungk et al., 1993). The adsorbed phosphate, insoluble phosphate salts, phosphate esters, and phytates in the soil can all be converted to soluble phosphate when the level of

A. Transport Processes 1. Sulfate Transport One of the first changes that can be measured as an organism becomes limited for a nutrient is the induction of a more efficient system for the transport of that nutrient into the cell. In most cases, both the affinity and capacity to import the limiting nutrient increases. For example, when C. reinhardtii is transferred from sulfur-replete to sulfur-lacking medium, the for sulfate transport decreases by over 5-fold from to while the increases by approximately ten-fold from 20 fmol of to 200 fmol sulfate sulfate and can be A change in the detected within 30 min of transferring the cells from replete to deficient conditions, although it takes about 6 h to attain steady state transport characteristics. Based on the characteristics of sulfate transport in the light and dark and the use of various inhibitors that block the formation of ATP or relax a trans­ membrane pH gradient, it was demonstrated that the uptake of sulfate is an energy dependent process that

618 is driven by a proton motive force (Yildiz et al., 1994). These characteristics are similar to sulfate transport in both vascular plants and Saccharomyces cerevisiae where sulfate is imported along with protons. The S. cerevisiae system is particularly well characterized; it has been shown that under low sulfur conditions the sulfate is transported by a sulfate/ proton symporter that requires a pH gradient across the plasma membrane. Three protons are imported and one potassium exported for every sulfate ion that is taken up (Roomans et al., 1979). A S. cerevisiae strain with a mutation in a gene encoding a sulfate transporter requires high levels of sulfate for growth (Smith et al., 1995b). It was previously reported that for the high and low affinity sulfate transport the and respectively, and systems were that the high affinity sulfate transport activity is blocked by inhibitors of cytoplasmic protein synthesis such as cycloheximide, which indicates that de novo protein synthesis is required for its expression (Breton and Surdin-Kerjan, 1977). Similarly, de novo protein synthesis is required for expression of the high affinity sulfate transport activity in C. reinhardtii (Yildiz et al., 1994). The genes encoding a number of high affinity sulfate transporters, including those of S. cerevisiae (Smith et al., 1995b), Stylosanthese hamata (Smith et al., 1995a), Arabidopsis thaliana [Genbank Accession No. D85416] and Hordeum vulgare [Genbank Accession No. U52867] have been isolated. Based on the gene sequences, the transporters have 12 putative membrane spanning helices and appear to be related to other anion transporters (Sandal and Marcker, 1994; Smith et al., 1995b).

2. Phosphate Transport Phosphate levels in the soil generally range from 0.65 to (Clarkson and Scattergood, 1982), suggesting that a high affinity phosphate transporter is required for the uptake of free phosphate under most growth conditions. Indeed, a phosphate for phosphate of approximately transporter with a is present in C. reinhardtii grown in either phosphate replete or deficient medium (Shimogawara and Grossman, unpublished). In contrast to the for phosphate transport increases almost 20the fold, from 4 fmol for unstarved cells, to 70 fmol for phosphorus starved cells. These data suggest that the starvation of C. reinhardtii for phosphorus results in increased synthesis of a single

John P. Davies and Arthur R. Grossman phosphate transport system (or that there are two distinct systems with the same affinity for the phosphate anion). During phosphorus limitation of for vascular plants there is also an increase in the phosphate uptake with no change in the (Shimogawara and Usuda, 1995).

3. Ammonium Transport Ammonium is the preferred source of nitrogen for C. reinhardtii, although it can also use nitrate, nitrite and several organic compounds including some amino acids (Franco et al,, 1987; Quesada et al., 1993; Chapter 33, Fernández et al.). C. reinhardtii appears to have two independent systems for transporting ammonium, a low affinity system that is constitutively expressed and a high affinity system that is only expressed when the concentration of ammonium falls below a critical level (Franco et al., 1988). Mutants disrupted in the functioning of these transport systems have been identified by selecting for cells resistant to methylammonium. One mutant, ma1, appears to be devoid of the constitutively expressed low affinity ammonium transporter (Franco et al., 1987), while the other, ma2, lacks the inducible high affinity ammonium transporter (Franco et al., 1988).

4. Inorganic Carbon Transport C. reinhardtii imports inorganic carbon as or bicarbonate in an energy dependent manner (Sültemeyer et al., 1991) and uses it in the dark reactions of photosynthesis. The concentration of dissolved inorganic carbon that half saturates photosynthesis is about 20 times lower in cells grown (0.03%) than in high (5%) (Badger in low and et al., 1980). Low -grown cells have bicarbonate transport systems with much higher -grown affinities for their substrates than high cells (Badger et al., 1980; Sültemeyer et al., 1989; Sültemeyer et al., 1991; Palmqvist et al., 1994; Chapter 28, Spalding). At this time the locations of the inorganic carbon transport systems are not known. Some models propose that transport occurs at both the plasma membrane and the chloroplast envelope (Sültemeyer et al., 1991), while others propose that an active transport system may be located only on the crosses the plasma chloroplast envelope and that membrane passively (Moroney et al., 1987; Palmqvist et al., 1994). In summary, C. reinhardtii cells exposed to low

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Nutrient Stress

levels of sulfate, phosphate, ammonium and inorganic carbon induce more efficient systems for transporting the limitingnutrient.The inducedsulfate,ammonium and inorganic carbon transport systems have a higher affinity for the limiting nutrient than the constitutively expressed systems, while the induced phosphate transport system does not. Furthermore, all of the induced transport systems have a higher capacity for importing the limiting nutrient.

B. Accessing Alternative External Sources of Nutrients 1. Sulfur Although organisms have preferred sources for specific nutrients, many also have the ability to use alternative sources of a limiting nutrient. For example, Pseudomonas putida can use both sulfate and certain sulfate esters such as 4-nitrocatecholsulfate (NCS), as sources of sulfur. When P. putida is grown on medium containing both sulfate and NCS, it depletes the medium of the former prior to using the latter. The sulfate is readily metabolized while a sulfatase is required to release sulfate from NCS. This sulfatase must be synthesized, processed, and secreted from the cell to access sulfate esters, and therefore it is energetically more costly to use the sulfur in sulfate esters (Beil et al., 1996). However, when the preferred source of a sulfur is not available, the ability to synthesize enzymes that hydrolyze sulfate esters can be advantageous. When C. reinhardtii is starved for sulfur, several newly synthesized polypeptides accumulate in the periplasmic space (de Hostos et al., 1988). One of these proteins, an arylsulfatase (Ars), is capable of cleaving sulfate from aromatic sulfate esters (Lien and Schreiner, 1975; Schreiner et al., 1975;de Hostos et al., 1988). The other proteins have not been characterized but could be alkylsulfases, sulfonatases, choline sulfatases, or sulfoquinovose degrading enzymes. The C. reinhardtii Ars has been purified to homogeneity. It is a 68 kDa glycoprotein located in the periplasmic space. Ars activity accumulates in cells placed in medium containing limiting levels of sulfate; concentrations of sulfate in excess of prevent Ars accumulation (de Hostos et al., 1988). Two cDNAs encoding Ars activity have been cloned (de Hostos et al., 1989; Davies and Grossman, unpublished information) and are designated Ars1 and Ars2. These cDNAs are almost identical in their

619 coding regions, and appear to be coordinately regulated (Davies and Grossman, unpublished). Since the amounts of Ars activity, protein, and mRNA are correlated, it appears that expression of the Ars genes is controlled at the level of transcription (de Hostos et al., 1989; Davies et al., 1994). Arylsulfatases are classified as one of two types based on their substrate inhibitor sensitivities. Type I arylsulfatases are inhibited by cyanide but not by sulfate or phosphate. This class has a low affinity for the chromogenic substrates p-nitrophenyl sulfate and NCS. Type II arylsulfatases are inhibited by sulfate and phosphate but not by cyanide, and have a relatively high affinity for NCS but not p-nitrophenyl sulfate (Dodgson and Spencer, 1957; Lien and Schreiner, 1975). The C. reinhardtii Ars is a type I enzyme. It appears to function as a homodimer and is stable in solution for at least 3 h with no loss of activity (Lien and Schreiner, 1975). The protein has at least three O-linked oligosaccharides and is synthesized as a precursor protein with a signal sequence that is cleaved as it is exported from the cell; the enzyme is localized to the periplasmic space or associated with the cell wall (de Hostos et al., 1988). This extracellular location allows the enzyme to hydrolyze soluble sulfate esters in the medium, thereby releasing free sulfate that can be assimilated by the cell. The only known substrates of the C. reinhardtii enzyme are aromatic sulfate esters; the sulfated polysaccharides and proteins of the C. reinhardtii cell wall do not appear to serve as substrates for this enzyme (de Hostos et al., 1988).

2. Phosphorus Phosphatases are prominent enzymes in the soil that are synthesized by many organisms when phosphate is limiting. These hydrolytic enzymes often have a broad substrate specificity that allows organisms to access a diverse range of phosphate esters in the soil. C. reinhardtii has at least four or five phosphatases that are located in different compartments ofthe cell and that have different catalytic characteristics (Loppes et al., 1977; Patni et al., 1977; Nagy et al., 1981 ;Quisel et al., 1996). Three of these phosphatases appear to reside in the periplasmic space where they can encounter substrates in the soil solution (Loppes and Matagne, 1973; Loppes and Deltour, 1975; Matagne and Loppes, 1975). The periplasmic phosphatases are classified by their acidic, neutral and alkaline pH optima. The periplasmic acid

620 phosphatase represents a minor component of the external phosphatase activity and appears to increase to some extent when the cells are deprived of phosphate. A neutral periplasmic phosphatase was detected by Loppes et al. (1977) but not by Patni et al. (1977) or Quisel et al. (1996); it may be a strain specific enzyme. The Loppes group has identified mutants that do not express this neutral phosphatase activity (Loppes, 1976b; Loppes et al., 1977). The majority of phosphatase activity in the periplasmic space has an alkaline pH optimum (Quisel et al., 1996). This periplasmic alkaline phosphatase activity is highly induced when the growth medium is depleted of phosphorus. Quisel et al. have fractionated the activity into two separate phosphatases, both of which have pH optima above 9 (Quisel et al., 1996). Most of the alkaline phosphatase activity (85–90%) has a pH -dependent and is associated optimum of 9.5, is with a glycoprotein of approximately 190 kDa. This 190 kDa enzyme has an affinity for a variety of naturally occurring substrates including nucleosidephosphates and aromatic phosphate esters. Its for these compounds ranges from 100–200 The other inducible alkaline phosphatase (2–10% of the activity) has a pH optimum of 9.0, does not require divalent cations and has a strong preference for arylphosphates (Quisel et al., 1996).

3. Nitrogen C. reinhardtii prefers to use ammonium as its source of nitrogen, but it can synthesize proteins that enable the use of alternative sources of nitrogen when ammonium is not present. It is not uncommon for ammonium to be limiting in the environment, and the ability to obtain nitrogen from other sources is advantageous (Chapter 33, Fernández et al.). C. reinhardtii cells are able to grow on nitrate, nitrite, certain amino acids, urea, purines, and purine derivatives as a sole source of nitrogen. These compounds may be imported directly, or deaminated in the extracellular environment. Nitrate, nitrite, urea, arginine, andurate (a purine derivative) can be directly imported by C. reinhardtii from the medium (Kirk and Kirk, 1978; Pineda et al., 1984; Galván et al., 1996), but most amino acids are deaminated outside the cell and the released ammonium is then transported into the cell (Muñoz-Blanco et al., 1990). Several transporters and enzymes that enable the alga to use these compounds as a source of nitrogen

John P. Davies and Arthur R. Grossman have been characterized. The presence of high affinity nitrate, nitrite (Galván et al., 1996) and arginine transporters have also been demonstrated (Kirk and Kirk, 1978). An amino acid oxidase capable of deaminating amino acids has been purified from C. reinhardtii by two groups. Both groups found that the enzyme is located in the periplasmic space and has a molecular weight of approximately 60 kDa. However, the measured for the enzyme isolated by the two groups were greater than ten fold different (Piedras et al., 1992; Vallon et al., 1993). It is not clear if there are two different periplasmic amino acid oxidases in the same cell, or if the two strains used for the isolation of the enzyme have somewhat different enzymes.

4. Inorganic Carbon Carbon dioxide is imported preferentially by C. reinhardtii relative to bicarbonate. When cells are exposed to low levels of a periplasmic carbonic anhydrase which facilitates the interconversion of bicaronate and is induced (Kimpel et al., 1983). This enzyme, along with the high affinity transporter, enhances the ability of the organism to concenimport and fix inorganic carbon when trations are low. The carbonic anhydrase is a glycosylated enzyme which functions as a heterotetramer composed of two large and two small polypeptides that are generated by proteolytic cleavage of a single primary translation product (Kamo et al., 1990; Funke et al., 1997). High levels of carbonic anhydrase are synthesized when the cells in the light (Spalding are exposed to low levels of and Ogren, 1982; Spencer et al., 1983). Two genes encoding carbonic anhydrase have been cloned and the predicted amino acid sequences of the carbonic anhydrase polypeptides are nearly identical. Expression of these genes is differentially regulated, while Cah1 is expressed in the light in low (Fujiwara Cah2 is expressed in the dark in high et al., 1990).

C. Accessing Internal Stores of Nutrients Mobilizing internal stores of an essential nutrient does not allow growth to continue over an extended period of time, but it may help the organism to survive and acclimate to a nutrient deficient environment.

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Nutrient Stress

1. Sulfur A specific sulfur storage compound is not known to exist within cells; however, the degradation of proteins and lipids that are nonessential during sulfur deprivation may supply the cells with a limited amount ofsulfur. While the sulfur-stress induced degradation of cellular structures and molecules has not been examined carefully in C. reinhardtii, there have been studies ofthese processes in other organisms. Sulfurdeprived Lemna minor degrades many of its intracellular proteins, but appears to degrade preferentially ribulose bisphosphate carboxylase/ oxygenase (Rubisco), which has an especially high number of sulfur-containing amino acids. In cells grown on sulfur-replete medium, the Rubisco pool constitutes a substantial sulfur reserve. The amount of sulfur present in the chloroplast in the form of Rubisco is estimated to be equivalent to 50 mM (Ferreira and Teixeira, 1992). The cyanobacterium Synechococcus sp. PCC 7942 degrades its lightharvesting phycobilisomes to recover sulfur from cysteine and methionine in times of sulfur limitation (Collier and Grossman, 1992, 1994). Photosynthetic membranes also contain significant quantities of sulfur in the form of the sulfolipid, 6-sulfoquinovosyl diacylglycerol (Chapter 21, Trémolières). This lipid comprises approximately 5% of the lipid in a C. reinhardtii cell (Sato et al, 1995a) or 12.5% of the lipid in the thylakoid membranes (Sato et al., 1995b). It is not known whether this lipid is degraded and if so whether the released sulfur is reused when C. reinhardtii is deprived of sulfur.

2. Phosphorus Polyphosphates are present in many organisms including Chlamydomonas species. These compounds may serve as an internal store of phosphate that can be mobilized rapidly when external sources of phosphate decline. Consistent with this idea is the observation that the capacity for the synthesis of polyphosphates by Chlamydomonas eugametos increases when the organism is starved for phosphate; C. eugametos appears to be primed to replace the polyphosphates that are lost during growth in phosphate-deficient medium (Siderius et al., 1996). The ribosomes and the cellular RNAs represent other relatively large pools of internal phosphate. Ribonucleaase activity increases in plants during

621 phosphate limited growth (Löffler et al., 1992; Dodds et al., 1996). At least five ribonucleases are induced in phosphate-starved tomato cells. Three ofthese are located in the vacuole, one in the cytosol, and one is exported from the cell. Some of these nucleases appear to be involved in scavenging phosphate from intracellular RNA. When tomato plants are starved for phosphate, the level of cytoplasmic RNA, particularly rRNA, decreases (Löffler et al., 1992). The phospholipids of the thylakoid membranes may also be used as a source of phosphate during phosphate limited growth. The cyanobacterium Synechocystis sp. strain PCC 6803 degrades its phospholipids during phosphorus limitation; the abundance of phospholipids in the thylakoid membrane drops from approximately 16% to 7% (Güler et al., 1996).

3. Nitrogen C. reinhardtii cells deprived of nitrogen degrade most of their highly abundant proteins. One of the most noticeable e°ffects of nitrogen limitation is the bleached appearance associated with the loss oflight harvesting pigments and proteins (Plumley and Schmidt, 1989). Nitrogen-limited C. reinhardtii cells also degrade most of their Rubisco (Plumley and Schmidt, 1989) and ribosomal proteins (Siersma and Chiang, 1971). Degradation of these very abundant proteins allows cells to recycle the amino acids into proteins more suited for survival when the cells are starved for nitrogen. The recycled amino acids can be incorporated into transporters and enzymes that help scavenge nitrogen from alternative sources in the environment.

4. Inorganic Carbon The chloroplast of C. reinhardtii contains a carbonic anhydrase that catalyses the reversible dehydration of bicarbonate to for use by Rubisco (Moroney et al., 1987; Husic et al., 1989). This activity, more abundant in low (0.03%) than high (5%) -grown cells (Sültemeyer et al., 1995), increases the local concentration of around Rubisco, thereby increasing the fixation reaction. This carbonic anhydrase activity is essential for growth in low and is thought to be associated with a levels of protein having a molecular mass of 45 kDa (Husic et al., 1989). A mutant of C. reinhardtii, designated

622 ca1, is deficient in the chloroplastic carbonic anhydrase (Katzman et al., 1994), and requires for photoautrophic growth elevated levels of (Spalding et al., 1983a; Funke et al., 1997).

IV. Common Responses

A. Cell Division Nutrient deprivation prevents cell cycle progression (Lien and Knutsen, 1973; Pringle and Hartwell, 1981). The mechanism of cell cycle regulation in C. reinhardtii has not been elucidated. However, regulatory processes that control the cell cycle of S. cerevisiae have been identified. Since the mechanisms controlling cell cycle progression appear to be highly conserved (Sherr, 1994), the regulatory processes that occur in S. cerevisiae are also likely to function in C. reinhardtii. Mutants of S. cerevisiae defective in the regulation of cell cycle control during nutrient limitation have been isolated and characterized (Toda et al., 1987; Wilson and Tatchell, 1988; Costigan and Snyder, 1994). The analysis of these mutants has led to the identification of two distinct control processes. One process, involving a cAMP-dependent protein kinase pathway, modulates cell cycle progression in response to carbon, nitrogen and sulfur levels (Toda et al., 1987; Wilson and Tatchell, 1988). The second, a mitogen activated protein (MAP) kinase pathway, has been shown to affect cell cycle arrest in response to nitrogen levels (Costigan and Snyder, 1994). The cAMP-dependent protein kinase regulates cell cycle progression by controlling cyclin proteolysis in response to the level of cAMP within the cell (Barral et al., 1995). In nutrient replete conditions cAMP concentrations are high and cell division proceeds, while in nutrient deficient conditions cAMP levels drop and cell division stops (Eraso and Gancedo, 1985). cAMP levels are mediated by a RAS (a small GTP binding protein) controlled adenylate cyclase and two phosphodiesterases which degrade cAMP. The ability of RAS to activate adenylate cyclase is thought to be controlled by receptors that sense the nutrient status of the cell (Kataoka et al., 1984; Cannon and Tatchell, 1987; Toda et al., 1987). Two mutations in S. cerevisiae, bcy1 and pde2, which affect cAMP-dependent protein kinase activity and disrupt nutrient-dependent cell cycle control, also cause sensitivity to nutrient limitation (Toda et al., 1987; Wilson and Tatchell, 1988). BCY1 encodes a negative regulator of the cAMP-dependent protein

John P. Davies and Arthur R. Grossman kinase. A bcy1 strain has constitutive cAMPdependent protein kinase activity and proceeds through the cell cycle even when nutrients are limiting and cAMP levels are low (Toda et al., 1987). PDE2 encodes a high affinity phosphodiesterase that normally degrades cAMP (Sass et al., 1986). Therefore, a pde2 mutant has high cAMP levels, which result in high levels of cAMP dependent protein kinase activity and cell cycle progression even under nutrient-deficient conditions (Toda et al., 1985; Wilson and Tatchell, 1988). Both bcy1 and pde2 mutations are lethal when cells are starved for an essential nutrient; the cells starve to death because they proceed through the cell cycle when nutrients are not available. The MAP kinase pathway appears to function independently of the cAMP-activated protein kinase. The slk1 mutant of S. cerevisiae, which is deficient in a MAP kinase is also unable to arrest cell division when grown in nitrogen deficient medium (Costigan and Snyder, 1994). Mutations in genes of C. reinhardtii that regulate similar pathways are likely to disrupt control of the cell cycle in response to nutrient status, but these have not yet been identified.

B. Nutrient Stress and Photosynthesis When C. reinhardtii is deprived of an essential nutrient there is a marked decrease in photosynthetic activity (Badger et al., 1980; Spalding et al., 1983c; Peltier and Schmidt, 1991; Plumley and Schmidt, 1989; Wykoff et al., 1997). This decrease appears to be essential for the survival ofthe alga during sustained conditions of nutrient deprivation. The sac1 mutant of C. reinhardtii is unable to decrease photosynthetic evolution during sulfur-limited growth and dies when the cells are maintained in the light and deprived of sulfur. Sulfur-starved sac1 cells can be saved if they are placed in the dark or treated with the herbicide DCMU in the light following imposition of sulfurdeficient conditions. These data suggest that the decrease in photosynthetic activity observed during sulfur stress is, at least in part, an active process controlled by the Sac1 gene product. The regulated decrease in photosynthetic electron transport appears to be part of the process of acclimation to sulfur deficient conditions and probably occurs during deficiency of other nutrients (Davies et al., 1996).

1. The ‘Light’ Reactions The light reactions of photosynthesis convert

Chapter 32

Nutrient Stress

absorbed light energy in to chemical energy in the form of ATP and NADPH (Golbeck, 1992; Hope, 1993;Vermaas, 1993; Grossman et al., 1995, Chapter 16, Ruffle and Sayre; Chapter 17, Webber and Bingham; Chapter 24, Wollman). Nutrient limitation causes a change in the activity of the light reactions of photosynthesis. C. reinhardtii cells grown in sulfurdeficient medium for 1 day or in phosphorus-deficient medium for 4 days exhibit a 75% decrease in maximal evolution. Measurement of photosynthetic electron transport uncoupled from water splitting, ATP reduction, demonstrates that synthesis and the maximal rate of photosynthetic electron transport also decreases by about 75% during sulfur or phosphorus deprivation. The decrease in photosynthetic electron flow is a consequence of a 20% decrease in the number of active PS II reaction centers (possibly by photoinhibition) as well as a 50% increase in the number of PS II centers that are unable to reduce the secondary quinone ( nonreducing centers). Neither the inactive PS II centers nor the non-reducing centers are productive in evolution. In addition to the decrease in active PS II centers non-reducing centers, a state and increase in transition occurs during nutrient limitation (D. D. Wykoff, J. P. Davies and A. R. Grossman, unpublished). State 1 is defined as the condition when LHCII transfers energy to PS II reaction centers efficiently, and state 2 is when less energy is transferred to PS II (and possibly more is transferred to Photosystem I (PS I)) (Allen et al., 1981; Wollman and Delepelaire, 1983; Delepelaire and Wollman, 1985; Allen, 1992; Chapter 30, Keren and Ohad). Nutrient stressed cells are in state 2 and less of the absorbed light energy is directed to PS II relative to PS I; this increases the production of ATP relative to NADPH. Although a state transition does not cause a evolution in decrease in the quantum yield of saturating light, it does cause a decreased quantum yield at subsaturating light levels. Mechanistically, state transitions are thought to be caused by a detachment of LHCII from PS II reaction centers, and appear to be regulated by the reduction state of the photosynthetic electron transport chain (Allen et al., 1981; Allen, 1992; see Chapter 30, Keren and Ohad). In sulfur- and phosphorus-limited cells the photosynthetic electron transport chain is highly reduced, anabolic processes are slowed, NADPH is not used rapidly and the major terminal electron acceptor of the light reactions, is limiting. This causes a decrease in the flow of electrons through

623 the photosynthetic electron transport system and results in a highly reduced PQ pool. The reduced PQ pool signals the activation of a kinase that phosphorylates LHCII, causing it to detach from the PS II reaction center (Allen et al., 1981; Wollman and Delepelaire, 1983; Delepelaire and Wollman, 1985; Allen, 1992; Chapter 30, Keren and Ohad). Changes in photosynthetic electron transport similar to those described for sulfur and phosphorus deprivation also occur during nitrogen limitation. Nitrogen-stressed C. reinhardtii evolves at a lower rate than non-stressed cells (Peltier and Schmidt, 1991). The marine algae Dunaliella tertiolectra, Thalassiosira pseudonana, T. weisflogii, Skeletonema capstatum and Isochrysis galbana exhibit lower rates of evolution when they are nitrogen-limited, also. Like sulfur- and phosphorus-limited C. reinhardtii, nitrogen-limited D. tertiolectra and T. weissflogii accumulate inactive PS II centers and nonreducing centers (Kolber et al., 1988). Furthermore, like sulfur- and phosphorus-starved C. reinhardtii, nitrogen-limited C. reinhardtii cells tend to be in state 2 (i.e. the LHCII are not attached to PS II) (Peltier and Schmidt, 1991) which results in diminished PS II and elevated PS I activity at subsaturating light levels (Berges et al., 1996). However, in contrast to sulfur- and phosphoruslimited C. reinhardtii, nitrogen limitation causes a 50–60% decline in the number of PS II reaction centers per cell in all the algae thus far examined (Kolber et al., 1988; Herzig and Falkowski, 1989; Plumley and Schmidt, 1989; Berges et al., 1996). cause some of the same changes Low levels of in photosynthesis that are observed when cells are starved for sulfur, phosphorus or nitrogen. C. levels (0.03%) reinhardtii cells grown atambient have a lower rate of evolution than cells grown in (5%) (Badger et al., 1980; Spalding et elevated limitation, as when cells al., 1983a,b). Severe are grown in medium that has been purged with free air, causes a decrease in the rate of electron to these cells are also in state 2 transfer from (Demeter et al., 1995). In summary, sulfur, phosphorus, nitrogen, and deprivation decrease photosynthetic activity by causing the accumulation of inactive PS II centers as well as non-reducing PS II centers. Nutrient limitation does not decrease PS I activity and cells appear to maintain cyclic photophosphorylation. Also, nutrient limitation causes a transition of the cells into state 2, thereby causing a decrease in the quantum efficiency of photosynthetic evolution at

624

subsaturating levels oflight and increasing the relative rate of production of ATP to NADPH. The additional ATP may be used to support the induced nutrient scavenging systems induced during nutrient stress. A striking difference between the effects of limitation for different nutrients is that sulfur or phosphorus limitation causes a 20% decrease in active PS II reaction centers while nitrogen deprivation causes a loss of 50–60% of active centers. Also, although sulfur, phosphorus, and nitrogen may not be present is ubiquitous in the in particular environments, environment and cells would be unlikely to ever experience a complete absence of it. Therefore, it is unclear if there are adaptive responses to environments completely devoid of While many of the changes in photosynthesis can be explained by control processes inherent in the physiology and biochemistry of photosynthesis, there appears to be a component of the nutrient-stress induced decrease in photosynthesis that is regulated directly by the nutrient status of the cell. The sac1 mutant of C. reinhardtii is unable to decrease photosynthesis when it is starved for sulfur (Davies et al., 1996). Examination of its photosynthetic characteristics indicates that, unlike wild-type cells, the sac1 strain does not show a decrease in PS II activity nor does it accumulate non-reducing centers in sulfate deficient medium. However, like wild-type cells, it is in state 2, indicating that the photosynthetic electron transport chain is highly reduced (D. D. Wykoff, J. P. Davies and A. R. Grossman, unpublished). Thus, the effect on photosynthesis that appears to be specifically regulated by the nutrient status ofthe environment is the decrease in PS II activity caused by a loss of active PS II centers and the accumulation of nonreducing centers. Because inactive PS II centers and non-reducing centers accumulate during other types of nutrient stress, there may be a common mechanism responsible for eliciting these specific changes.

2. Non-Photochemical Quenching The amount of light absorbed by a photosynthetic organism can be significantly more than the amount required to saturate photosynthesis. When photosynthesis is saturated, over-excitation of the photosynthetic apparatus could lead to the production of triplet chlorophyll and both singlet oxygen and toxic oxygen radicals (Demmig-Adams and Adams,

John P. Davies and Arthur R. Grossman

1992). Although photosynthetic organisms can, to some extent, detoxify active oxygen species with the aid of superoxide dismutase and ascorbate peroxidase, high levels of these radicals can saturate and overwhelm the protective processes (Asada, 1994). Photosynthetic organisms prevent the production of active oxygen species by limiting the amount of energy that reaches the PS II reaction center. Excess energy absorbed by the chlorophyll molecules of LHCII can be dissipated as heat through a process measured as non-photochemical quenching of chlorophyll fluorescence (NPQ) that is thought to be mediated by the carotenoids, zeaxanthin and antheraxanthin (Demmig-Adams and Adams, 1992). Lutein, an integral xanthophyll in LHCII, may also contribute to NPQ (Niyogi et al., 1997). Photosynthesis in nutrient-deficient cells is saturated at lower light levels than in non-limited cells (D. D. Wykoff, J. P. Davies and A. R. Grossman, unpublished). These cells absorb light in excess of their photosynthetic capacity at lower levels of light than cells grown in replete medium, and therefore, must dissipate more energy as heat. When D. bardawil and D. salina cells are starved for sulfur for 2 days and grown in moderate light, NPQ is enhanced, zeaxanthin and antheraxanthin levels increase, and Cbr, a protein proposed to bind these carotenoids, accumulates (Levy et al., 1992, 1993). These changes are thought to enhance NPQ and thereby protect the algae from damage due to excessive light (Levy et al., 1993; Braun et al., 1996). No large increase in NPQ or the xanthophyll pigments was observed in C. reinhardtii cells starved for sulfur for one day or phosphorus for four days (D. D. Wykoff, J. P. Davies and A. R. Grossman, unpublished). This difference may be the result of the light conditions under which the experiments were performed. The nutrientdeprived C. reinhardtii cells were incubated in light that did not saturate photosynthesis. Perhaps under more intense light or a longer period of starvation, NPQ and the xanthopylls play an important role in the acclimation of C. reinhardtii to nutrient limitation.

3. The ‘Dark’ Reactions During the dark reactions of photosynthesis, NADPH and for other reductive and ATP are used to fix and anabolic processes (Fig. 2). The dark reactions of photosynthesis are affected by nutrient stress since there is a dramatic alteration in the flux of metabolites through the metabolic pathways of the cell. During

Chapter 32

Nutrient Stress

sulfur, phosphorus and nitrogen deficiencies the cell cycle is inhibited (Lien and Knutsen, 1973; Davies et al., 1996) and there is a decrease in the need for proteins, lipids, and complex carbohydrates that are required to support cell growth. During nutrient limitation feedback inhibition limits the flux of fixed carbon into amino acids, lipids and nucleic acids, which results in the reduced export of PGA generated from the Photosynthetic Carbon Reduction (PCR) cycle. Starch biosynthesis is induced because PGA accumulates; PGA activates ADP-glucose pyrophosphorylase, which catalyses the rate limiting step in starch biosynthesis (Preiss, 1982; Ball et al., 1991; Chapter 29, Ball). The capacity of the plastid to accumulate starch is finite and starch synthesis eventually stops even though the precursors remain high in the chloroplast (Caspar et al., 1991). Without sinks for the reduced carbon, intermediates of amino acid, lipid and starch biosynthesis accumulate. The accumulation of phosphorylated metabolites leads to the sequestration of phosphate, which also depletes

625 the chloroplast of PCR cycle intermediates. Eventually, carbon fixation becomes limited by the rate of RuBP regeneration (Brooks, 1986; Jacob and Lawlor, 1992; Jacob and Lawlor, 1993; Rao and Terry, 1995).

C. Respiration and Nutrient Stress Cells grown in sulfur-, phosphorus- and nitrogendeficient medium all exhibit a significant decrease in the rate of respiration. This decrease appears to be caused by low levels of cellular ADP (Gauthier and Turpin, 1994), a substrate needed for both glycolysis and oxidative phosphorylation (Theodorou and Plaxton, 1993). Nutrient-starved cells have low levels of ADP because cell division and anabolic processes stop, and the ATP that accumulates is not rapidly recycled (Gauthier and Turpin, 1994). The reduced ADP concentration leads to a decrease in the rate of glycolysis and oxidative phosphorylation because reactions catalysed by pyruvate kinase (PK) (Duff et

626 al., 1989b;Theodorou et al., 1991) and ATP synthase are inhibited (Bryce et al., 1990). PK catalyzes the conversion of phosphoenol pyruvate (PEP), ADP, and into pyruvate and ATP. Pyruvate enters the TC A cycle where its oxidation to is coupled to the synthesis of NADH, and ATP. Many photosynthetic organisms appear to have alternative pathways for the synthesis of pyruvate that are not inhibited by low concentrations ofADP. PEP carboxylase catalyzes the formation of oxaloacetate, which upon reduction to malate is transported into the mitochondrion and enters the TCA cycle (Duff et al., 1989b). The vacuolar enzyme, PEP phosphatase catalyzes the conversion of PEP to pyruvate and inorganic phosphate (Duff et al., 1989a; Theodorou et al., 1991) and this pyruvate can be used by mitochondria to fuel the TCA cycle. However, when the green alga Selenastrum minutum is grown in medium devoid of nitrogen or phosphorus, it accumulates high levels of PEP, indicating that the overall use of PEP has diminished (Gauthier and Turpin, 1994) and that the turnover of PEP by PEP carboxylase and PEP phosphatase is not very rapid when compared to the reaction catalyzed by PK. The level of oxidative phosphorylation also decreases substantially during nutrient limitation because ADP levels in the cell are low and thereby limit the ATP synthase for substrate. Because the ATP synthase cannot function, a substantial builds up across the inner mitochondrial membrane (Bryce et al., 1990) and slows electron transport. Some electron flow can continue in a reduced ADP environment because of the presence of an alternative oxidase. This oxidase, present in many organisms (Siedow and Berthold, 1986; Rychter and Mikulska, 1990; Rychter et al., 1992), including C. reinhardtii (Weger and Dasgupta, 1993), allows the transfer of electrons from the reduced ubiquinone pool to molecular oxygen without the generation of a or ATP synthesis, and is not inhibited by low levels of ADP (Weger and Dasgupta, 1993). Thus, C. reinhardtii has the ability to perform some oxidative electron transport under nutrient-deficient conditions. This alternative oxidase may serve to eliminate high potential electrons during adverse conditions.

V. Model Integrating the Responses to Nutrient Deprivation Outlined in Fig. 3 is a model which explains how

John P. Davies and Arthur R. Grossman C. reinhardtii regulates its responses to nutrientdeficient conditions. There is evidence suggesting that separate sensors exist for each macronutrient; the sensors would enable the cell to monitor the nutrient status of the environment. These sensors may be on the surface of the cell where they can monitor the amount of the nutrient in the medium, or they may be inside the cell where they can sense the concentration of specific metabolites. For example, the cell could monitor the level of sulfate in the medium, or the intracellular concentrations of cysteine and/or O-acetylserine, the immediate precursor of cysteine. No matter how the nutrient status is monitored, the perception of nutrient deficiency induces a signal that activates cellular responses. Physiological studies suggest that there are two types of responses; one response has evolved to help in the acquisition of the limiting nutrient (i.e. the induction of hydrolytic enzymes and transport processes) while another, more general response, helps tune the physiology of the cell to the dramatic decrease in growth and cell division that accompanies nutrient limitation (i.e. decreased cell division, respiration and photosynthesis). For each of the different nutrient limitations, there may be signal transduction pathways that control the specific and common responses that diverge at some point after the sensor. This implies that there are signal transduction elements that are specific for both types of responses. There may also be a convergence of the signaling pathways that control the common responses to nutrient limitation. This is inferred from the finding that during sulfur, phosphorus, nitrogen and inorganic carbon starvation, the decrease in photosynthetic electron transport displays similar characteristics for each nutrient limiting condition (suggesting a similar mechanism for decreasing photosynthesis). The model predicts the existence of a number of different classes of genes that control the acclimation of C. reinhardtii to nutrient-limited growth. For example, there should be genes that govern both the specific and common responses to a specific limitation; examples of this would include genes encoding nutrient sensors. There should also be genes that control only the specific responses, such as those encoding transcription factors that regulate the synthesis of specific hydrolytic enzymes, and genes that control only the common responses, such as those that cause photosynthesis to decline during any of a number of different nutrient limitations. In the

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following section, we will discuss the progress that has been made in identifying and analyzing mutants that affect the acclimation of cells to nutrient deficiencies, and how their phenotypes are indicative of the different classes of genes mentioned above. We will also discuss experimental approaches which may be used to further test and refine this model.

VI. Regulation of the Responses to Nutrient Deprivation

A. Sulfur Genetic dissection of the regulation of the response of C. reinhardtii to sulfur limitation has been attempted by screening for mutants that exhibit aberrant expression of arylsulfatase activity (Davies et al., 1994) which is normally detected only during sulfur stress (Lien and Schreiner, 1975; de Hostos et al., 1988). Arylsulfatase activity is easily assayed using the chromogenic substrate 5-bromo-4-chlorothe hydrolysis of which 3-indolyl sulfate, results in blue product. This assay has been used to

627

screen for mutants that either cannot induce arylsulfatase activity when the cells are deprived of sulfur, or that express it constitutively in sulfur replete medium (Davies et al.; 1994). Mutants of both types have been isolated and designated sulfur acclimation (sac) mutants. The sac1 and sac2 mutants are deficient in arylsulfatase activity in sulfur-deficient medium; sac1 shows no activity and sac2 shows severely diminished activity. The sac3 mutant expresses arylsulfatase in replete medium. All three of these mutations cause aberrations in the accumulation of the other sulfur-stress induced periplasmic proteins, and the sulfur-stress induced uptake of the sulfate anion (Davies et al., 1994). The sac 1 mutant is also unable to decrease photosynthetic electron transport during sulfur limitation (Davies et al., 1996). However, it responds normally to phosphorus- and nitrogen-deficient conditions. These data indicate that the sac1 lesion is in a gene controlling both the specific and common responses to sulfur limitation; and that this lesion affects the acclimation of cells only to sulfur stress, and not nitrogen, phosphorus or carbon limitation. The Sac1 gene product plays a central role in the

628 acclimation of C. reinhardtii to sulfur stress. While the phenotype of the sac1 mutant indicates that Sac1 has a regulatory role, the predicted amino acid sequence of the protein is similar to that of a class of ion transporters (Davies and Grossman, unpublished). The deduced amino acid sequence of Sac1 and the phenotype of the sac1 mutant show some analogies to the SNF3 gene product and the phenotype of the snf3 mutant of S. cerevisiae. The amino acid sequence of Snf3p is similar to those of hexose transporters (Celenza et al., 1988), and s n f 3 mutants do not exhibit high affinity hexose uptake in response to low glucose (Coons et al., 1995). However, the SNF3 gene product does not function as a hexose transporter but regulates the production of the transporters. Snf3p has been proposed to function in sensing the extracellular hexose concentration (Liang and Gaber, 1996). Sac1 may act similarly; it may sense extracellular sulfate levels by binding sulfate, and when the concentration of sulfate drops below a critical level, Sac1 may initiate a signaling process which triggers the acclimation response. The Sac2 and Sac3 genes appear to function downstream of Sac1. Mutations in Sac2 and Sac3 affect only the specific responses to sulfur limitation. A mutation in Sac1 is epistatic to one in Sac2, indicating that these two genes may function in the same signaling pathway. There is no clear epistatic relationship of mutations in Sac1 and Sac3, or Sac2 and Sac3, and it is not clear how Sac3 controls the specific responses to sulfur stress (Davies et al., 1994).

B. Phosphorus Mutants of C. reinhardtii deficient in the derepressible neutral and alkaline phosphatases have been isolated by screening colonies with the chromogenic substrate naphthylphosphate. There are at least 3 loci, exemplified by the mutant strains pd2, pd3, and pd24, that affect expression of the inducible neutral phosphatase (Loppes, 1976a; Loppes et al., 1977). Strains carrying the pd2, pd3, and pd24 mutations are all thought to have a lesion in a gene that regulates expression of the derepressible neutral phosphatase (Loppes et al., 1977). Bachir et al. (1996) identified four allelic mutations at the DA locus that alter alkaline phosphatase activity. Mutations in Da prevent accumulation of the 190 kDa alkaline phosphatase, but expression of the other phosphate-stress induced periplasmic proteins is not altered. Mutations in Da

John P. Davies and Arthur R. Grossman are probably either in the gene encoding the alkaline phosphatase or a specific regulator of its expression (Bachir et al., 1996). Because it does not appear that mutations in any of these genes affect the common responses to nutrient deprivation, they probably act downstream of the branch between the specific and common responses to phosphate limitation.

C. Nitrogen C. reinhardtii mutants affecting acclimation to nitrogen-deficient conditions have been isolated by selecting for cells resistant to chlorate or methylammonium which are toxic analogues of nitrate and ammonium, respectively (Sosa et al., 1978; Franco et al., 1987; Franco et al, 1988; Prieto and Fernández, 1993; Schnell and Lefebvre, 1993; Chapter 33, Fernández et al.). The nit2 and nit8 (also called nar2) mutants are resistant to chlorate. Nit2 encodes a regulator of nitrate reductase activity (Schnell and Lefebvre, 1993) while Nit8 is essential for the functioning of the nitrate transporters (Galván et al., 1996). The methylammonium resistant mutants, ma1 (Franco et al., 1987) and ma2 (Franco et al., 1988) have defects in the low affinity and high affinity ammonium transport systems, respectively. The lesions in these mutants may be in the ammonium transporters or in genes that regulate their synthesis. Further analysis of these mutants is necessary to distinguish between the possibilities.

D. Inorganic Carbon Mutants in the carbon concentrating mechanism, CCM, of C. reinhardtii have been identified by screening for cells that require high levels of (5%) for photoautotrophic growth (Chapter 28, Spalding). Mutations in four complementation groups have been found. Two of these, pmp1 and ca1, affect different parts of the CCM. Cells with pmp1 mutations are unable to induce the more efficient inorganic carbon uptake system (Spalding et al., 1983a, 1983b), while ca1 affects the activity of the chloroplastic carbonic anhydrase (Spalding et al., 1983a; Sültemeyer et al., 1995, Funke et al., 1997). Another mutant, cia5, cannot synthesize a number of different activities associated with CCM, and appears to have a lesion in a gene involved in the regulation of the (Moroney et responses of C. reinhardtii to low al., 1989; Burow et al., 1996). However, it is not clear if the cia5 mutation affects only the specific or both

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the common and specific responses to carbon limitation. The other known high -requiring mutant, cia4, is not allelic to pmp1, ca1 or cia5, but it has not been well characterized.

VII. Identification of Mutants Deficient in the Acclimation to Nutrient Deprivation Mutations affecting both regulatory and enzymatic activities in various nutrient acclimation processes have been isolated. Further characterization of the these mutants and cloning and characterization of the mutated genes will provide insights into the biochemical and regulatory processes involved in acclimation of photosynthetic organisms to nutrient deficient environments. To get a clearer understanding of the acclimation processes and its regulation, more mutants should be isolated and characterized. Multiple mutants in each complementation group need to be identified using the screens described above, but more novel screens and selection processes should also be used for the identification of new mutants. Below we describe methods for isolating mutants. Mutants deficient in their ability to induce high affinity nutrient transporters can be selected by using toxic compounds that are taken into the cell by these transporters. For example, a mutagenized population can be starved for a nutrient to induce the high affinity transporter, these cells could then be exposed to the toxic compound for a defined period of time and plated onto complete medium. Conditions can be defined such that cells that efficiently import the toxic compound die, while those less effective in import of the compound survive. These cells may have lesions in genes that encode or control expression of the high affinity transporter. Selenate and chromate can be imported by the high affinity sulfate transport system, incorporated into the amino acids cysteine and methionine, and kill cells. These compounds have been used in other systems to isolate mutants with lesions in genes encoding high affinity sulfate transporters and proteins that regulate the expression and activity of these transporters (Marzluf, 1970; Chen and Metzenberg, 1974; Smith et al., 1995b). Similarly, arsenate (Thiel, 1988) and (Shimarogowa, personal communication) have been used to select for cells that are unable to increase phosphate transport during phosphorus-limited growth. Additionally, chlorate and methylammonium

629 have been used to select for mutants deficient in induced nitrate (Sosa et al., 1978; Prieto and Fernández, 1993; Schnell and Lefebvre, 1993) and ammonium transport (Franco et al., 1987; Franco et al., 1988). Mutants of Chlorella ellipsoidea deficient induced inorganic carbon transport have in low been identified by incubating colonies derived from and then exposing a mutagenized population to the colonies to film to determine relative amounts of imported (Matsuda and Colman, 1996). A similar screen could be performed with C. reinhardtii. Screening for mutants displaying aberrant expression of hydrolytic enzymes has been useful for isolating strains deficient in acclimation processes. Mutants abnormal for the sulfur-stress-induced expression ofarylsulfatase have been identified using (Davies et al., 1994) the chromogenic reagent or related compounds (Metzenberg and Ahlgren, 1970). Napthylsulfate is also a suitable substrate for screening for Ars activity (Ohresser, et al., 1997). (D. D. Wykoff, J. Quisel Similar substrates, and A. R. Grossman, unpublished) and napthylphosphate (Loppes, 1976a; Loppes et al., 1977; Bachir et al., 1996) have been used to screen for mutants with abnormal expression of the phosphatases. Easily assayed substrates for amino acid oxidase or carbonic anhydrase have not been identified. However, mutants disrupted in the induced expression of genes encoding these proteins could be identified using a chimeric gene containing the promoter of these genes fused to the Ars reporter gene. Several groups have constructed chimeric genes containing promoters from other genes fused to the Ars2 coding sequences. Transgenic cultures containing these genes express arylsulfatase activity and transcription of the introduced genes have been regulated by the promoter sequences (Davies et al., 1992; Davies and Grossman, 1994; Quinn and Merchant, 1995; Jacobshagen et al., 1996; Periz and Keller, 1997; Ohresser et al., 1997). Similar fusions could be made with promoters from genes that are transcribed only during a specific type of nutrient stress. Strains containing these chimeric genes could be mutagenized and screened for aberrant expression from the inducible promoter by assaying for arylsulfatase activity. Mutants deficient in the common responses to nutrient deprivation may be identified by screening for strains that cannot stop cell cycle progression or that are unable to decrease photosynthesis in response to nutrient stress. Mutants that are unable to stop cell cycle progression during nutrient stress will die when

630 deprived of an essential nutrient. These mutants can be identified by replica plating cells on complete medium and medium lacking in an essential nutrient. After incubation for an extended time on the medium devoid of the specific nutrient, the limiting nutrient could be added to the plates to promote cell growth. Cells unable to survive this treatment could be collected from the nutrient replete plates and tested for cell cycle control during nutrient limitation. Mutants that are unable to decrease photosynthesis during nutrient stress may be identified by their aberrant fluorescent characteristics using video imaging devices that monitor fluorescence (Chapter 22, Bennoun and Béal); one such device is described by Niyogi et al. (1997). This device can capture fluorescence images of colonies grown on medium lacking in a specific nutrient deficient medium and the images can be quantified by a computer to provide measurements of various parameters of photosynthesis, such as the quantum efficiency During sulfur- and phosphorus-limited growth of wild-type cells the quantum efficiency of PS II declines, but sulfur starved sac1 cells maintain a high these cells can be easily distinguished from wild-type cells. Mutants deficient in the common responses to nutrient limitation may be in a gene encoding a sensor or signal transduction component, or in genes that affect cell cycle progression or PS II activity. Mutants that cannot regulate cell cycle progression or photosynthetic activity during nutrient limitation may be sensitive to deficiencies in various nutrients, and would be extremely interesting to analyze in detail. They may also provide clues to how C. reinhardtii integrates the perception of nutrients levels with the regulation of the cell cycle and photosynthesis.

Acknowledgments The authors thank Elena Casey, David Kehoe, Rakefet Schwarz and Dennis Wykoff for reading and providing helpful suggestions on the manuscript, and Kathi Bump for helping to format it. We are also very grateful to the United States Department of Agriculture and the Carnegie Institution of Washington for supporting our work. This is CIW publication number 1340.

John P. Davies and Arthur R. Grossman

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Loppes R and Matagne RF (1973) Acid phosphatase mutants in Chlamydomonas: isolation and characterization by biochemical, electrophoretic and genetic analysis. Genetics 75: 593–604 Loppes R, Braipson J, Matagne RF, Sassen A and Ledoux L (1977) Regulation of the neutral phosphatase in Chlamy­ domonas reinhardi: An immunogenetic study of wild-type and mutant strains. Biochem Genet 15: 1147–1157 Lou G and Warman PR (1992) Enzymatic hydrolysis of ester sulphate in soil organic matter extracts. Biol Fertil Soils 14: 112–115 MacDonald NW, Burton AJ, Jurgensen MF, McLaughlin JW and Mroz GD (1991) Variation in forest soil properties along a Great Lakes air pollution gradient. Soil Sci Soc Am J 55: 1709–1715 Mahler RJ and Maples RL (1987) Effect of sulfur additions on soil and the nutrition of wheat. Commun Soil Sci Plant Anal 18:653–673 Marschner H (1995) Mineral Nutrition of Higher Plants, pp 229– 312. Academic Press, San Diego Marzluf GA (1970) Genetic and metabolic controls for sulfate metabolism in Neurospora crassa: Isolation and study of chromate-resistant and sulfate transport-negative mutants. J Bacteriol 102: 716–721 Matagne RF and Loppes R (1975) Isolation and study of mutants lacking a derepressible phosphatase in Chlamydomonas reinhardi. Genetics 80: 239–250 Matsuda Y and Colman B (1996) A new screening method for algal photosynthetic mutants. Plant Physiol 110: 1283–1291 Metzenberg RL and Ahlgren SK (1970) Mutants of Neurospora deficient in aryl sulfatase. Genetics 64: 409–422 Moroney JV, Husic HD, Tolbert NE, Kitayama M, Manuel LJ and Togasaki RK (1989) Isolation and characterization of a mutant of Chlamydomonas reinhardtii deficient in the concentrating mechanism. Plant Physiol 89: 897–903 Moroney JV, Togasaki RK, Husic HD and Tolbert NE (1987) Evidence that an internal carbonic anhydrase is present in 5% -grown and air-grown Chlamydomonas. Plant Physiol 84: 757–761 Muñoz-Blanco J, Hidalgo-Martinez J and Cardenas J (1990) Extracellular deamination of L-amino acids by Chlamydomonas reinhardtii cells. Planta 182: 194–198 Murooka Y, Ishibashi K, Yasumoto M, Sasaki M, Sugino H, Azakami H and Yamashita M (1990) A sulfur- and tyramineregulated Kiebsiella aerogenes operon containing the arylsulfatase (atsA) gene and the atsB gene. J Bacteriol 172: 2131–2140 Nagy AH, Erdos G, Beliaeva NN and Gyurjan I (1981) Acid phosphatase isoenzymes of Chlamydomonas reinhardtii. Mol Gen Genet 184: 314–317 Niyogi KN, Bjorkmann O and Grossman AR (1997) Chlamy­ domonas xanthophyl1 cycle mutants identified by video imaging of chlorophyll fluorescence quenching. Plant Cell 9: 1369 1380 Ohresser M, Matagne RF and Loppes R (1997) Expression of the arylsulfatase reporter gene under the control of the NIT1 promoter in Chlamydomonas reinhardtii. Curr Genet 31:264– 271 Palmqvist K, Yu J-W and Badger MR (1994) Carbonic anhydrase activity and inorganic carbon fluxes in low- and high- cells of Chlamydomonas reinhardtii and Scenedesmus obliquus. Physiol Plant 90: 537-547

633 Pant HK, Edwards AC and Vaughn D (1994) Extraction, molecular fractionation and enzyme degradation of organically associated phosphorus in soil solutions. Biol Fertil Soils 194: 196–200 Patni NJ, Dhawale SW and Aaronson S (1977) Extracellular phosphatases of Chlamydomonas reinhardi and their regulation. J Bacteriol 13 0:205–211 Peltier G and Schmidt GW (1991) Chlororespiration: An adaptation to nitrogen deficiency in Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 88: 4791–4795 Periz G, and Keller LR (1997) DNA elements regulating altubulin gene induction during regeneration of eukaryotic flagella. Mol Cell Biol 17: 3858–3866 Piedras P, Pineda M, Muñoz J and Cardenas J (1992) Purification and characterization of an L-amino-acid oxidase from Chlamydomonas reinhardtii. Planta 188: 13–18 Pineda M, Fernandez E and Cardenas J (1984) Urate oxidase of Chlamydomonas reinhardtii. Physiol Plant 62: 453–457 Plumley FG and Schmidt GW (1989) Nitrogen-dependent regulation of photosynthetic gene expression. Proc Natl Acad Sci USA 86: 2678–2682 Preiss J (1982) Regulation of the biosynthesis and degradation of starch. Annu Rev Plant Physiol 33: 431–454 Prieto R and Fernández E (1993) Toxicity of and mutagenesis by chlorate are independent of nitrate reductase activity in Chlamydomonas reinhardtii. Mol Gen Genet 237: 429–438 Pringle JR and Hartwell LH (1981) The Saccharomyces cerevisiae cell cycle. In: Strathern JN, Jones EW and Broach JR (eds) The Molecular Biology of the Yeast Saccharomyces cerevisiae: Life cycle and inheritance, pp 97–142. Cold Spring Harbor Laboratory, Cold Spring Harbor Quesada A, Galván A, Schnell R, Lefebvre PA and Fernández E (1993) Five nitrate assimilation-related loci are clustered in Chlamydomonas reinhardtii. Mol Gen Genet 240: 387–394 Quinn JM and Merchant S (1995) Two copper-responsive elements associated with the Chlamydomonas Cyc6 gene function as targets for transcriptional activators. Plant Cell 7: 623–638 Quisel J, Wykoff D and Grossman AR (1996) Biochemical characterization ofthe extracellular phosphatases produced by phosphorus-deprived Chlamydomonas reinhardtii. Plant Physiol 111:839–848 Rao IM and Terry N (1995) Leafphosphate status, photosynthesis, and carbon partitioning in sugar beet. Plant Physiol 107:1313– 1321 Ron Vaz MD, Edwards AC, Shand CA and Cresser MS (1993) Phosphorus fractions in soil solution: Influence of soil acidity and fertiliser additions. Plant and Soil 148: 175–183 Roomans GM, Kuypers GAJ, Theuvenet APR and Borst-Pauwels GWFH (1979) Kinetics of sulfate uptake by yeast. Biochim Biophys Acta 551: 197–206 Rychter AM, Chauveau M, Bomsel J-L and Lance C (1992) The effect of phosphate deficiency on mitochondrial activity and adenylate levels in bean roots. Physiol Plant 84: 80–86 Rychter AM and Mikulska M (1990) The relationship between phosphate status and cyanide-resistant respiration in bean roots. Physiol Plant 79: 663–667 Sandal NN and Marcker KA (1994) Similarities between a soybean nodulin, Neurospora crassa sulphate permease II and a putative human tumour suppressor. Trends Biochem Sci 19: 19

634 Sass P, Field J, Nikawa J, Toda T and Wigler M (1986) Cloning and characterization of the high-affinity cAMP phosphodiesterease of Saccharomyces cerevisiae. Proc Natl Acad Sci USA 83: 9303–9307 Sato N, Sonoike K, Tsuzuki M and Kawaguchi A (1995a) Impaired Photosystem II in a mutant of Chlamydomonas reinhardtii defective in sulfoquinovosyl diacylglycerol. Eur J Biochem 234: 16–23 Sato N, Tsuzuki M, Matsuda Y, Ehara T, Osafune T and Kawaguchi A (1995b) Isolation and characterization of mutants affected in lipid metabolism of Chlamydomonas reinhardtii. Eur J Biochem 230: 987–993 Schmidt A (1973) Sulfate reduction in a cell-free system of Chlorella. The ferredoxin dependent reduction of a proteinbound intermediate by a thiosulfonate reductase. Arch Microbiol 93: 29–52 Schmidt A (1986) Regulation of sulfur metabolism in plants. Progr Bot 48: 133–150 Schnell RA and Lefebvre PA (1993) Isolation of the Chlamydomonas regulatory gene NIT2 by transposon tagging. Genetics 134: 737–747 Schreiner Ø, Lien T and Knutsen G (1975) The capacity for arylsulfatase synthesis in synchronous and synchronized cultures of Chlamydomonas reinhardtii. Biochim Biophys Acta 384: 180–193 Scott WA and Metzenberg RL (1970) Location of aryl sulfatase in conidia and young mycelia of Neurospora crassa. J Bacteriol 104:1254–1265 Seeling B and Hungk A (1996) Utilization of organic phosphorus in calcium chloride extracts of soil by barley plants and hydrolysis by acid and alkaline phosphatases. Plant and Soil 178: 179–184 Sherr CJ (1994) G1 phase progression: Cycling on cue. Cell 79: 551–555 Shimogawara K and Usuda H (1995) Uptake of inorganic phosphate by suspension-cultured tobacco cells: Kinetics and regulation of starvation. Plant Cell Physiol 36: 341–351 Siderius M, Musgrave A, van den Ende H, Koerten H, Cambier P and van der Meer P (1996) Chlamydomonas eugametos (Chlorophyta) stores phosphate in polyphosphate bodies together with calcium. J Phycol 32: 402–409 Siedow JN and Berthold DA (1986) The alternative oxidase: A cyanide-resistant respiratory pathway in higher plants. Physiol Plant 66: 569–573 Siersma PW and Chiang K-S (1971) Conservation and degradation of cytoplasmic and chloroplast ribosomes in Chlamydomonas reinhardtii. J Mol Biol 58: 167–185 Smith FW, Ealing PM, Hawkesford M and Clarkson DT (1995a) Plant members of a family of sulfate transporters reveal functional subtypes. Proc Natl Acad Sci USA 92: 9373–9377 Smith FW, Hawkesford MJ, Prosser IM and Clarkson DT(1995b) Isolation of a cDNA from Saccharomyces cerevisiae that encodes a high affinity sulphate transporter at the plasma membrane. Mol Gen Genet 247: 709–715 Sosa FM, Ortega T and Barea JL (1978) Mutants from Chlamydomonas reinhardtii affected in their nitrate assimilation capability. Plant Sci Lett 11: 51–58 Spalding MH and Ogren WL (1982) Photosynthesis is required for induction of the -concentrating system in Chlamy­ domonas reinhardtii. FEBS Lett 145: 41–44 Spalding MH, Spreitzer R and Ogren WL (1983a) Carbonic

John P. Davies and Arthur R. Grossman anhydrase-deficient mutant of Chlamydomonas reinhardtii requires elevated carbon dioxide concentration for photoautotrophic growth. Plant Physiol 73: 269–272 Spalding MH, Spreitzer RJ and Ogren WL (1983b) Genetic and -concentrating system of physiological analysis of the Chlamydomonas reinhardtii. Planta 159: 261–266 Spalding MH, Spreitzer RJ and Ogren WL (1983c) Reduced inorganic carbon transport in a -requiring mutant of Chlamydomonas reinhardtii. Plant Physiol 73: 273–276 Spencer KG, Kimpel DL, Fisher ML, Togasaki RK and Miyachi S (1983) Carbonic anhydrase in Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 24: 301–304 Stanko KM and Fitzgerald JW (1990) Sulfur transformations in forest soils collected along an elevational gradient. Soil Biol Biochem 22: 213–216 Stanko-Golden KM and Fitzgerald JW (1991) Sulfur transformations and pool sizes in tropical forest soils. Soil Biol Biochem 23: 1053–1058 Strickland TC and Fitzgerald JW (1983) Mineralization of sulphur in sulphoquinovose by forest soils. Soil Biol Biochem 15: 347–349 Strickland T, Fitzgerald JW and Swank WT (1986) In situ measurements of sulfate incorporation into forest floor and soil organic matter. Can J For Res 16: 549–553 Sültemeyer D, Miller AG, Espie GS, Fock H and Canvin DT (1989) Active transport by the green alga Chlamydomonas reinhardtii. Plant Physiol 89: 1213–1219 Sültemeyer DF, Fock HP and Canvin DT (1991) Active uptake of inorganic carbon by Chlamydomonas reinhardtii: Evidence for simultaneous transport of and and characterization of active transport. Can J Bot 69: 995–1002 Sültemeyer D, Amoroso G and Fock H (1995) Induction of intracellular carbonic anhydrases during the adaptation to low inorganic carbon concentrations in wild-type and ca-1 mutant cells of Chlamydomonas reinhardtii. Planta 196: 217–224 Swank WT, Fitzgerald JW and Ash JT (1984) Microbial transformation of sulfate in forest soils. Science 223: 182–184 Tarafdar JC and Marschner H (1995) Dual inoculation with Aspergillus fumigatus and Glomus mosseae enhances biomass production and nutrient uptake in wheat (Triticum aestivum L.) supplied with organic phosphorus as Na-phytate. Plant and Soil 173:97–102 Theodorou ME, Elrifi IR, Turpin DH and Plaxton WC (1991) Effects of phosphorus limitation on respiratory metabolism in the green alga Selenastrum minutum. Plant Physiol 95: 1089– 1095 Theodorou ME and Plaxton WC (1993) Metabolic adaptations of plant respiration to nutritional phosphate deprivation. Plant Physiol 101: 339–344 Thiel T (1988) Phosphate transport and arsenate resistance in the cyanobactcrium Anabaena variabilis. J Bacteriol 170: 1143– 1147 Toda T, Uno I, Ishikawa T, Powers S, Katoaka T, Broek D, Cameron S, Broach J, Matsumoto K and Wigler M (1985) In yeast, RAS proteins are controlling elements of adenylate cyclase. Cell 40: 27–36 Toda T, Cameron S, Sass P, Zoller M, Scott JD, McMullen B, Hurwitz M, Krebs EG and Wigler M (1987) Cloning and characterization of BCY1, a locus encoding a regulatory subunit ofthe cyclic AMP-dependent protein kinase in Saccharomyces cerevisiae. Mol Cell Biol 7: 1371–1377

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Uria-Nickelsen MR, Leadbetter ER and Godchaux III W (1993) Sulphonate utilization by enteric bacteria. J Gen Microbiol 139:203–208 Vallon O, Bulte L, Kuras R, Olive J and Wollman F-A (1993) Extensive accumulation of an extracellular L-amino-acid oxidase during gametogenesis of Chlamydomonas reinhardtii. Eur J Biochem 215: 351–360 Vermaas W (1993) Molecular-biological approaches to analyze Photosystem II structure and function. Annu Rev Plant Physiol Plant Mol Biol 44: 457–481 Warman PR and Sampson HG (1994) Effect of sulfur additions on the yield and elemental composition of canola and spring wheat. J Plant Nutr 17: 1817–1825 Weger HG and Dasgupta R (1993) Regulation of alternative pathway respiration in Chlamydomonas reinhardtii (Chlorophyceae). J Phycol 29: 300–308 Whalen JK and Warman PR (1996a) Arylsulfatase activity in soil

635 and soil extracts using natural and artificial substrates. Biol Fertil Soils 22: 373-378 Whalen JK and Warman PR (1996b) Examination of ester sulfates in Podzolic and Regosolic soils using an immobilized arylsulfatase reactor. Biol Fertil Soils 23: 64–69 Wilson RB and Tatchell K (1988) SRA5 encodes the cyclic AMP phosphodiesterase of Saccharomyces cerevisiae. Mol Cell Biol 8: 505–510 Wollman F-A and Delepelaire P (1983) Correlation between changes in light energy distribution and changes in thylakoid membrane polypeptide phosphorylation in Chlamydomonas reinhardtii, J Cell Biol 98: 1–7 Yildiz F, Davies JP and Grossman AR (1994) Characterization of sulfate transport in Chlamydomonas reinhardtii during sulfur-limited and sulfur-sufficient growth. Plant Physiol 104: 981-987

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Chapter 33 Nitrogen Assimilation and its Regulation

Emilio Fernández, Aurora Galván and Alberto Quesada*

Departamento de Bioquímica y Biología Molecular e Instituto Andaluz de Biotecnología, Facultad

de Ciencias, Universidad de Córdoba, E–14071 Córdoba, Spain

Summary I. Introduction. Pathways for Nitrogen Assimilation in Chlamydomonas II. Assimilation of Ammonium A. Ammonium Transport B. The Glutamine Synthetase/Glutamate Synthase Pathway C. Glutamate Dehydrogenases III. Assimilation of Amino Acids A. Amino Acid Transport and Deamination B. Transaminase Activities IV. Assimilation of Purines A. Uptake Systems for Purines B. Enzymes for Purine Assimilation C. Urea Assimilation V. Assimilation of Nitrate and Nitrite A. Nitrate Assimilation Mutants 1. Nitrate Reductase Structural Mutants 2. Molybdopterin Cofactor Mutants 3. Regulatory Mutants B. Nitrate and Nitrite Transport 1. Nitrate and Nitrite Uptake Activities 2. Nitrate and Nitrite Transporter Genes and Their Expression C. Nitrate Reduction 1. Nitrate Reductase Enzyme 2. Nitrate Reductase Gene 3. Nitrate Reductase Gene Expression D. Nitrite Reduction 1. Nitrite Reductase Enzyme 2. Nitrite Reductase Gene and Expression VI. Concluding Remarks Acknowledgments References

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address: Departamento de Biología Experimental, Facultad de Ciencias Experimentales, Universidad de Jaén, 23071-Jaén,

Spain

J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 637–659. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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Summary Nitrogen is available in the environment in different redox states and in a wide variety of nitrogenous compounds. Algal cells are capable of assimilating nitrogen from many of these molecules into, ultimately, the amino group of glutamate. In this chapter, recent findings on nitrogen assimilation pathways in Chlamydomonas reinhardtii are reviewed. Specific transport systems are required to take up nitrogenous compounds, though most amino acids are deaminated extracellularly. Ammonium is the preferred nitrogen source. The presence of ammonium both represses and inhibits the utilization of alternative nitrogen compounds. Purines are assimilated phototrophically and catabolized to generate urea and then ammonium by the standard aerobic pathway that operates in other organisms. The nitrate assimilation pathway has been dissected at the molecular level and a useful collection of mutants is now available. The mutants are affected at various steps: nitrate/nitrite transport, nitrate and nitrite reduction, biosynthesis of molybdopterin cofactor, and regulation of the pathway. At least six genes which might encode structural elements for this pathway are clustered together within a 45 kb genomic region. They show coordinated regulation with a similar response to different nutritional or environmental effectors. Regulatory elements responsible for molecular mechanisms of control, including sensing of nitrate and ammonium and the transduction of these signals, are starting to be defined. More research is required for resolution of the details.

I. Introduction. Pathways for Nitrogen Assimilation in Chlamydomonas In recent years, the nitrogen assimilation pathways in plants have received a lot of attention by many groups whose efforts are directed at the molecular definition of pathway components and regulatory mechanisms. This is specially true for nitrate assimilation, since nitrate is a preferred nitrogen source for plants and accounts for most of the nitrogen incorporated into living matter. The massive use of fertilizers to increase crop yield is leading to serious environmental and public health problems (Guerrero et al., 1981; Hoff et al., 1994; Crawford, 1995). Microbial activity in the soil readily converts nitrogen fertilizers into nitrate, which is easily leached out and accumulates in ground waters, resulting in their increasing contamination. The development of molecular techniques in Chlamydomonas and its Abbreviations: AAO – amino acid oxidase; AAT – aspartate amino transferase; ABC – ATP-binding cassette; AST – alanine amino transferase; fd – ferredoxin; GDH – glutamate dehydrogenase; GS – glutamine synthetase; GOGAT– glutamate synthase; HANT – high affinity nitrate transporter / HANiT high affinity nitrite transporter; LAO – L-amino acid oxidase; MFS – major facilitator superfamily; MoCo – molybdopterin cofactor; MoCoCP – MoCo Carrier Protein; MSX – methionine-D,Lsulfoximine; MVH – reduced methyl viologen; NR – nitrate reductase; NiR – nitrite reductase; OG – 2-oxoglutarate; NT – nitrate transporter; NiT – nitrite transporter; PLP – pyridoxal phosphate; POT – proton-dependent oligopeptide transporter; PPT – phosphinothricin

amenability for biochemical, physiological and genetic approaches has made this algal system a suitable model for the much more complex systems of higher plants. Interesting reviews on nitrogen assimilation in cyanobacteria, algae and plants have been published (Guerrero et al., 1981; Syrett, 1981; Harris, 1989; Fernández and Cárdenas, 1989; Flores and Herrero, 1994; Hoff et al., 1994; Crawford, 1995). In the present chapter we will focus on recent data on nitrogen assimilation in Chlamydomonas reinhardtii. A general scheme representing the major pathways for nitrogen assimilation in this alga is shown in Fig. 1. Ammonium plays a central role as a preferred nitrogen source that causes repression of alternative pathways of nitrogen assimilation when reduced carbon is non-limiting (Fernández and Cárdenas, 1989). Ammonium is incorporated into C-skeletons by the glutamine synthetase/glutamate synthase (GS/GOGAT) cycle in the form of Lglutamate which is the major nitrogen donor for several biosynthetic pathways (e.g. tetrapyrrole biosynthesis; Chapter 20, Timko). Organic nitrogen sources such as most amino acids, urea or purines can be assimilated by C. reinhardtii to yield ultimately ammonium. In fact, amino acids, except arginine which is transported as is, are deaminated extracellularly. Oxidized forms of nitrogen (nitrate and nitrite) are utilized efficiently after reduction to ammonium. Specific transport systems are required for each different nitrogen source.

Chapter 33 Nitrogen Assimilation and its Regulation

II. Assimilation of Ammonium C. reinhardtii cells assimilate ammonium preferentially over any other alternative nitrogen source such as nitrate, nitrite, urea, purine, or amino acids. This strategy is more economical for the cell because it allows reduced synthesis of enzymes for alternative pathways and also requires less energy per nitrogen assimilated into organic form (Florencio and Vega, 1983a; Pineda et al., 1987; Harris, 1989). Ammonium is also generated intracellularly from photorespiration, protein turnover and nucleic acid catabolism. This ammonium is re-captured via the nitrogen assimilation pathway (Hipkin et al., 1982; Cullimore and Sims, 1980). In C. reinhardtii, the main route for ammonium incorporation into carbon skeletons seems to be the GS/GOGAT cycle (Fig. 1; Cullimore and Sims, 1981a,b).

A. Ammonium Transport The first step in assimilation of extracellular is ammonium is transport. Although ammonia a weak base and can permeate the cell membrane at high pH, uptake studies indicate that ammonium transport is a highly regulated carrier-mediated

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process in C. reinhardtii. Kinetic characterization of wild type and methylammonium-resistant mutants (see Section VA.3) has revealed the existence of two specific carriers for ammonium (or methylammonium) transport (systems 1 and 2). Both systems are regulated by ammonium (Franco et al., 1987a, 1988a,b). In addition, methylammonium (ammonium) uptake shows circadian rhythms (Byrne et al., 1992). System 1 shows a high Km and a high Vmax, and system 2 has a low Km and a low Vmax for the transport of both ammonium and methylammonium. System 1 is constitutive and responsible for ammonium transport at high concentrations, whereas system 2 is ammonium repressible and responsible for providing ammonium to the alga when ammonium is present at low concentrations in the medium and system 1 operates inefficiently (Franco et al., 1988b). None of these carriers has been identified at the molecular level in C. reinhardtii. However, ammonium transporters have been isolated from Saccharo­ myces cerevisiae (Marini et al., 1994), Coryne­ bacterium glutamicum (Siewe et al., 1996), Arabidopsis thaliana (Ninnemann et al., 1994) and tomato (Lauter et al., 1996). Hydrophobicity analysis of the proteins (Mep1p and Mep2p in S. cerevisiae) reveals a typical transmembrane topology. The

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ammonium transporters of fungi, plants and bacteria are highly similar at the level of primary amino acid sequence (above 26 % identity) (Siewe et al., 1996). The conservation of the sequence together with the preferential expression of the tomato gene (Leamt1) in root hairs exposed to ammonium emphasizes the importance of ammonium transport systems (Lauter et al., 1996).

B. The Glutamine Synthetase/Glutamate Synthase Pathway The major pathway in plants for ammonium incorporation into carbon skeletons is the GS/GOGAT cycle which supplies glutamate (Miflin and Lea, 1976; Cullimore and Sims, 1981a,b,c; McNally et al., 1983). GS catalyzes the formation of glutamine from ammonium and glutamate in an ATP-dependent reaction. As in vascular plants, two different GS isoforms have been identified in C. reinhardtii, one (GS1) is located in the cytosol and the other (GS2) is located in the chloroplast (Florencio and Vega, 1983b; Fischer and Klein, 1988). Both isozymes have been purified and shown to have a similar octameric structure (Florencio and Vega, 1983b) (Table 1). Recently, cDNAs for both enzymes have been cloned from C. reinhardtii and the deduced amino acid sequences show a high similarity (above 70%) with the corresponding proteins from vascular plants, supporting the proposed cytosolic and chloroplastic distribution for GS1 and GS2, respectively (Chen and Silflow, 1996). GS enzymes are regulated tightly by environmental factors like light and carbon or nitrogen supply. In addition, it has been proposed that GS activity might control, through ammonium derivatives, the nitrate assimilation pathway (Cullimore and Sims, 1981a; Hoff et al., 1994). Transcriptional and post-transcriptional control mechanisms have been shown to regulate C. reinhardtii GS expression. GS can be inactivated reversibly by dark or by ammonium treatment (Cullimore and Sims, 1981b). The enzyme can be reactivated in vivo by reilluminating the cell culture (Cullimore and Sims, 1981b) or in vitro with thioredoxins, proteins which are involved in the maintenance of enzyme thiol groups in an active and reduced form (Florencio et al., 1993; Chapter 26 by Jacquot et al.). The specific activation of GS1 by cytoplasmic thioredoxin h or GS2 by chloroplastic thioredoxin m suggests that redox modulation of GS by thioredoxins takes place in response to appropriate

environmental conditions (Florencio et al., 1993). Analysis of GS transcript abundance and GS activity shows different regulation patterns by nitrate or ammonium for GS1 vs. GS2 (Fischer and Klein, 1988; Chen and Silflow, 1996). The GS1 transcript increases in cells grown in nitrate and decreases in cells grown in ammonium, while the GS2 transcript is expressed constitutively with respect to nitrate or ammonium supply (Chen and Silflow, 1996). These data are in agreement with the decrease in total GS activity and the increase in the GS2:GS1 ratio when cells are transferred from nitrate to ammonium (Fischer and Klein, 1988). On the other hand, adaptation from high to low results in an increase of 30% in the activity of the chloroplastic GS2 isoenzyme in C. reinhardtii cells (Ramazanov and Cárdenas, 1994). Since significant changes are not observed in the abundance of the transcript, this modulation is not effected via transcriptional regulation (Chen and Silflow, 1996). GOGAT catalyzes the transfer of the amide group from glutamine to 2-oxoglutarate in a reaction which requires reducing equivalents. Two molecules of glutamate, the substrate of GS, are produced (Lea and Miflin, 1975). Two GOGAT isoenzymes, one which is specific for reduced ferredoxin as the electron donor and another which is specific for NADH, have been purified and characterized from C. reinhardtii (see Table 1; Cullimore and Sims, 1981c; Galván et al., 1984; Márquez et al., 1984; Márquez et al., 1986b). Though the two enzymes have been suggested to occur in different intracellular compartments, ferredoxin-GOGAT in the chloroplast and NADHGOGAT in the cytosol (Márquez et al., 1984, 1986b), Fischer and Klein (1988) find all the NADH-GOGAT and all the ferredoxin-GOGAT exclusively in the chloroplast. Thus, in C. reinhardtii, only the plastid would contain a complete GS/GOGAT cycle for ammonium assimilation. Genes encoding GOGAT enzymes from C. reinhardtii have not been cloned so far, and studies of GOGAT gene expression are therefore lacking. Fd-GOGAT and NADH-GOGAT are probably not both fully active at the same time. Their activities may well be strongly dependent on the supply of their specific electron donors. Ferredoxin-GOGAT increases in the light in synchronous and asynchronous cultures, and the activity is also higher in ammonium than in nitrate media (Cullimore and Sims, 1981b; Márquez et al., 1986a; Martínez-Rivas et al., 1991). Light does not increase NADH-GOGAT activity suggesting that

Chapter 33 Nitrogen Assimilation and its Regulation

this enzyme is unimportant or plays only an auxiliary role in photosynthetic ammonium assimilation (Martínez-Rivas et al., 1991).

C. Glutamate Dehydrogenases NAD(P)H-glutamate dehydrogenase (GDH) enzymes, which catalyze the reversible reductive amination of 2-oxoglutarate to give glutamate, have been identified in Chlamydomonas reinhardtii but their role in nitrogen assimilation is still controversial (Cullimore and Sims, 1981b; Fischer and Klein,

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1988; Muñoz-Blanco et al., 1989; Muñoz-Blanco and Cárdenas, 1992). Purification and characterization of three NAD(P)H-GDH isoenzymes named GDH 1, GDH2 and GDH3 have been reported (Muñoz-Blanco et al., 1989; Moyano et al., 1992a). In contrast to Fischer and Klein (1988) who found GDH activity exclusively in the chloroplast, Moyano et al. (1992b) report a mitochondrial localization for the three isoenzymes, which are differentially regulated under different trophic and environmental conditions (Muñoz-Blanco et al., 1989; Moyano et al., 1995). It has been proposed that GDH1 could

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participate in ammonium assimilation, whereas GDH2 and GDH3 would have an anaplerotic role in the tricarboxylic acid cycle by producing 2oxoglutarate (Moyano et al., 1995).

III. Assimilation of Amino Acids The ability of C. reinhardtii to use different amino acids as nitrogen sources is strongly strain-dependent (Harris, 1989). A number of enzymes, such as transaminases and an L-amino acid oxidase (LAO), and specific transport systems are involved in amino acid assimilation. Indeed, in C. reinhardtii cells, mutagenic treatment can significantly modify the amino acid utilization pattern of isolated mutants. These modifications include the ability to use amino acids in the absence of acetate, the use of additional amino acids as a nitrogen source, or the restriction in the spectrum of amino acid utilization (Prieto and Fernández, 1993).

A. Amino Acid Transport and Deamination Extracellular amino acids can be used as a nitrogen source by C. reinhardtii but, with the exception of arginine, their assimilation is strongly dependent on acetate supply (Muñoz-Blanco et al., 1990). In fact, arginine is the only amino acid which is actively transported into the cells by a specific transport system. The transport system is induced under nitrogen starvation and is repressed by ammonium (Kirk and Kirk, 1978). Amino acids other than arginine are deaminated extracellularly, yielding the corresponding 2-oxoacid plus ammonium which enters the cell readily and is incorporated by its specific assimilation pathway (Muñoz-Blanco et al., 1990). In acetate-containing media, the oxoacids produced by deamination are not utilized, which implies that, with the exception of arginine, the organic skeletons of amino acids cannot be used as carbon sources by C. reinhardtii (Muñoz-Blanco et al., 1990). The enzyme responsible for amino acid deamination has been isolated and characterized as an L-amino acid oxidase, but there exists controversy about its molecular size and subunit composition. One work reports on a 470 kDa native enzyme consisting of eight similar-sized subunits (Piedras et al., 1992) and another on a 900–1300 kDa native enzyme containing 13–16 identical subunits (Vallon et al., 1993). The purified enzyme catalyzes

deamination of all amino acid tested, with the exception of cysteine and proline (Vallon et al., 1993). However, the range of amino acid nitrogen utilization by a particular C. reinhardtii strain is restricted to 12 amino acids: asparagine, glutamine, arginine, lysine, alanine, valine, leucine, isoleucine, serine, methionine, histidine and phenylalanine (Muñoz-Blanco et al., 1990). LAO is induced after nitrogen starvation in C. reinhardtii cultures containing a source of organic carbon (Piedras et al., 1992; Vallon et al., 1993). The enzyme has been localized by immunochemical methods to the extracellular compartment confined by the cell wall. Accordingly, it accumulates in the culture medium of a cell wall-less mutant, and during the mating reaction (Vallon et al., 1993). The assimilation of some extracellular amino acids by C. reinhardtii is related to the existence of specific transport systems and transaminase enzymes. This mechanism was suggested to explain resistance/sensitivity to methionine-D,L-sulfoximine (MSX) and phosphinothricin (PPT), which are irreversible inhibitors of GS (Franco et al., 1996a,b). One MSX-resistant strain from C. reinhardtii has been proposed to express a new MSX/2-oxoglutarate transaminase activity and, in contrast to wild-type strains, to grow with MSX as the sole nitrogen source (Franco et al., 1996a). The high rate at which such mutants arise is interpreted to imply a gain-of-function, i.e. the activation of a cryptic gene. In contrast, the wild-type strain from C. reinhardtii is resistant to PPT which is not transported by the cells and can be deaminated to some extent by LAO. Thus, PPT supports cell growth when it is present as the only nitrogen source (Franco et al., 1996b).

B. Transaminase Activities When inorganic nitrogen is supplied to C. reinhardtii, the GS/GOGAT cycle (and eventually the GDH aminating activity) leads to the production of glutamate, a key molecule in the coupling of the metabolic pathways of amino acids with those of carbohydrates and lipids (Torchinsky, 1987; Lam et al., 1996). Two aminotransferase activities have been characterized from the alga, the L-aspartate and Lalanine aminotransferases (AAT and AST, respectively; Muñoz-Blanco et al., 1988; Laín-Guelbenzu et al., 1990, 1991; Chen et al., 1996). AAT catalyzes the reversible transfer of the amino group between aspartate and glutamate, and AST between alanine

Chapter 33 Nitrogen Assimilation and its Regulation

and glutamate. AAT and AST activities increase when C. reinhardtii cells are starved for nitrogen or carbon, and when an organic carbon source is being assimilated (Muñoz-Blanco et al., 1988; Cárdenas et al., 1990). Under these conditions, the transamination reactions have to play a relevant role in the control of the internal amino acid pool, providing nitrogen and/ or carbon skeletons to keep cells functioning (MuñozBlanco et al., 1988; Cárdenas et al., 1990; Lam et al., 1996). In addition, AAT activity is closely related to the aspartate/malate shuttle, which is the main system for translocating reducing equivalents from the chloroplast and mitochondria to the peroxisomes or the cytosol (Halliwell, 1981; Journet et al., 1981). This result suggests a linkage between control of the intracellular redox potential and transaminase activities (Muñoz-Blanco et al., 1988). The AAT enzyme from C. reinhardtii is a dimer of identical 65 kDa subunits, contains a pyridoxal phosphate (PLP) cofactor and is able to transaminate, though to a lower extent, the amino group from phenylalanine, tyrosine, serine, and alanine (Laín-Guelbenzu et al., 1990). AST has also been purified from Chlamydomonas, and consists of two identical 45 kDa subunits containing a PLP-cofactor but, in contrast to AAT, AST is highly specific for its amino acid substrates (Laín-Guelbenzu et al., 1991). A cDNA clone encoding AST has been isolated among several mRNA from C. reinhardtii. The cDNA-encoded protein has a molecular mass of 58 kDa (Chen et al., 1996). Antibodies raised against an

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AST protein from barley cross-react with a inducible polypeptide of 53 kDa from C. reinhardtii. Under these conditions, the physiological role of AST seems to be to supply amino groups from glutamate to glyoxylate in the C2 photorespiration reactions (Chen et al., 1996).

IV. Assimilation of Purines Purines are important constituents of nucleic acids, ATP and other coenzymes, metabolic regulators and activated metabolites. Studies on purine assimilation and its metabolism have been carried out in a few green algae (Vogels and Van der Drift, 1976; Antia et al., 1991), including especially C. reinhardtii (Pineda and Cárdenas, 1996). This organism can utilize adenine, guanine, hypoxanthine, urate, allantoin or allantoate as the only nitrogen source under phototrophic growth conditions, which suggests that the standard aerobic pathway of purine degradation of higher plants, animals and many microorganisms (Fig. 2) is also operative in this alga (Pineda et al., 1984). In fact, uptake of purines has been studied and enzymes for this pathway have been identified, purified and biochemically characterized (Fernández and Cárdenas, 198la; Pérez-Vicente et al., 1988; Alamillo et al., 1991; Pineda et al., 1994; Piedras, 1995). Their most significant properties are shown in Table 1.

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A. Uptake Systems for Purines

B. Enzymes for Purine Assimilation

In C. reinhardtii, adenine, guanine, hypoxanthine, xanthine and urate are taken up by high affinity transport systems which show saturable and hyperbolic kinetics with Ks at the micromolar range (Pineda and Cárdenas, 1996). Purine transport activity is repressed by ammonium and its expression depends on protein synthesis. Adenine and guanine seem to be taken up by the same transporter which can be induced by either of these purines. This transport system shows identical kinetic properties for adenine and guanine and a typical competitive cross-inhibition pattern (Lisa et al., 1995). However, hypoxanthine and xanthine use different transport systems to enter C. reinhardtii cells (Pérez-Vicente et al., 1991). These two systems are induced by either purine but to very different extents. Hypoxanthine uptake is affected only by adenine and guanine; the transport of these two purines is in turn inhibited competitively by hypoxanthine with a similar Ki of 1.3 micromolar (Pérez-Vicente et al., 1991; Lisa et al., 1995). Xanthine and urate inhibit, to a small extent, both adenine and guanine uptake. However, xanthine uptake is only inhibited competitively by urate and vice versa, which suggests that both purines are transported by a common system different from that responsible for adenine, guanine and hypoxanthine uptake (Fig. 1; Lisa et al., 1995). Utilization of xanthine as the sole nitrogen source involves the formation ofa transient, intracellularpool ofxanthine (up to 20% of the total xanthine supplied). Electrondense material can be observed in cytoplasmic vacuoles from xanthine-grown cells, which indicates that xanthine is accumulated intracellularly in part as a solid material (Pérez-Vicente et al., 1995). Urate uptake does not require induction by urate, since urate uptake components are expressed in nitrogenfree media. Ammonium added to the medium inhibits urate uptake without affecting urate oxidase activity (Pineda et al., 1987). Inhibition of GS with Lmethionine-DL-sulfoximine in cells taking up urate actively causes ammonium excretion accompanied by rapid inhibition of urate uptake, which indicates that ammonium itself is a negative effector of urate uptake activity. Ammonium has also been proposed as a co-repressor of the urate uptake system (Pineda et al., 1987).

Some of the enzymes for purine assimilation in C. reinhardtii have been purified to electrophoretic homogeneity and characterized (Table 1). Xanthine dehydrogenase (330 kDa) catalyzes oxidation of hypoxanthine to xanthine and xanthine to urate, and does not use oxygen as an electron acceptor (PérezVicente et al., 1988, 1992). An interesting phenomenon of irreversible inactivation of xanthine dehydrogenase by incubation with xanthine or has been hypoxanthine in the absence of reported (Pérez-Vicente et al., 1992). NADH and NADPH, the electron donors for the diaphorase activity associated with xanthine dehydrogenase (Pérez-Vicente et al., 1987), also cause the same inactivation whichaffectsprimarily themolybdenum center. It has been proposed that sequential and uninterrupted electron flow from xanthine to is needed to maintain the enzyme in an active form (Pérez-Vicente et al., 1992). Xanthine dehydrogenase is repressed by ammonium and does not require xanthine for its induction, since high activities are found in cells incubated in urea, adenine, urate, guanine, xanthine or nitrogen-free media (PérezVicente et al., 1988). Urate oxidase (uricase) from C. reinhardtii is a homotetramer of 31 kDa subunits containing one copper per subunit (Alamillo et al., 1991). C. reinhardtii uricase is repressed by ammonium and induced by urate. However, the constitutive amounts of uricase protein determined with specific polyclonal antibodies from cells grown in different media suggests that the regulation by ammonium occurs by modulation of enzyme activity (Pineda et al., 1984; Alamillo, 1992). Extracts from C. reinhardtii cells grown with allantoin as the sole nitrogen source are able to catalyze hydrolysis of allantoin and allantoic acid. Allantoinase and allantoicase are two separate enzymes. Their association with membranes is differenct, since allantoinase, and not allantoicase, was solubilized after deoxycholate treatment. These enzymes have been suggested to be located in different cellular compartments (Piedras et al., 1995). Allantoinase seems to be a constitutive enzyme, since significant activity is detected in cells grown with any nitrogen source whereas allantoicase is not detected in nitrate or ammonium-grown cells (Piedras et al., 1995).

Chapter 33 Nitrogen Assimilation and its Regulation

C. Urea Assimilation Urea can be assimilated by C. reinhardtii readily when present in the culture medium. It is first transported actively (Williams and Hodson, 1977) and then stored in the chloroplast in form of a metabolically inactive pool. Urea catabolism occurs from a different metabolic pool (Dagestad et al., 1981). The compartmentation of urea into a metabolically inactive storage form is similar to that reported for xanthine (Pérez-Vicente et al., 1995). In C. reinhardtii, urea is hydrolyzed to ammonia by the enzyme complex ATP:urea amidolyase (Leftley and Syrett, 1973). This complex consists of two enzyme activities: urea carboxylase, which catalyzes the ATPdependent condensation of urea and bicarbonate to yield allophanate, and allophanate lyase (Whitney and Cooper, 1973; Hodson et al., 1975). ATP:urea amidolyase is repressed by ammonium and is induced by urea or acetamide in the absence of ammonium (Hodson et al., 1975; Semler et al., 1975).

V. Assimilation of Nitrate and Nitrite Three independent steps can be distinguished in nitrate assimilation: i) the import of nitrate by the cells mediated by specific nitrate transporters (NT), ii) the reduction of nitrate to nitrite catalyzed by nitrate reductase (NR), and iii) the reduction of nitrite to ammonium catalyzed by nitrite reductase (NiR) (Fig. 1). Complementary approaches, genetic, molecular and physiological, are allowing us to identify and isolate genes related to nitrate assimilation, and to understand their function in the C. reinhardtii system. The definition of transport mechanisms and their regulatory features, and the elucidation of molecular mechanisms for regulation of this pathway are important objectives because of the agronomic relevance of nitrogen metabolism in crop plants (Hoff et al., 1994; Crawford, 1995).

A. Nitrate Assimilation Mutants The isolation and characterization ofmutants deficient in the nitrate assimilation pathway is a powerful tool to define the structural and regulatory elements of the route of nitrate metabolism, and also to understand the particular and individual function of each component. In fungi and plants, mutant strains

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deficient in NR activity have been isolated routinely by positive selection in media containing chlorate and a neutral nitrogen source such as urea (Cove, 1976; Toby and Kemp, 1977; Müller and Grafe, 1978; Nichols and Syrett, 1978; Sosa et al., 1978). That NR deficiency leads to a chlorate-resistance phenotype has been explained generally by assuming that chlorate, not toxic by itself, is converted into the toxic compound chlorite by the catalytic activity of NR (Arberg, 1947; Liljestrom and Aberg, 1966; Solomonson and Vennesland, 1972). Nevertheless, some chlorate-resistant mutants have NR activity and some NR deficient mutants are sensitive to chlorate (Cove, 1976; Nichols and Syrett, 1978). Further, most C. reinhardtii chlorate-resistant mutants isolated by Nichols and Syrett (1978) are able to grow with nitrate only if acetate is absent from the medium. The relationship between chlorate resistance and nitrate assimilation has not been explained. The frequency with which nitrate assimilation mutants are obtained among a population of chlorate-resistant strains depends on the strain used for mutagenesis, the medium in which the selection was performed and the concentration of chlorate used for selection. Recently, it has been noted that chlorate is both toxic and mutagenic to C. reinhardtii cells, since chlorate treatment leads to a decrease in cell survival, and the proportion of mutants recovered increases with increasing chlorate concentrations or time of treatment. These chlorate-resistant mutants exhibit a modified pattern of purine or amino acid utilization, and they also exhibit a different profile of resistance to a number of chemicals such as methylammonium, sulphanilamide or arsenate with respect to the wild type (Prieto and Fernández, 1993). Chlorate toxicity in C. reinhardtii seems to be dependent primarily on the NT or nitrite transporter (NiT) system, whose expression requires a functional pathway-specific regulatory locus NIT2, and functional NRT2 loci encoding NT/NiT systems and which are clustered with the NR structural gene Nia1 (formerly named nit1; see Table 2 and Section V.C). Chlorate toxicity and chlorate-resistance phenotypes are independent of the existence of an active NR enzyme, but depend on the function of at least seven loci unrelated to the nitrate assimilation pathway (Prieto and Fernández, 1993). These loci are thought to control the efficiency of the nitrate transporter for chlorate indirectly . The conclusion ofthese experiments is that chlorate enters the cells by the NT/NiT systems and, once inside,

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promotes general mutagenesis and cell toxicity after its conversion to chlorite, which is not necessarily catalyzed by NR. Chlorate-inhibited growth by NRindependent mechanisms has also been reported in Nicotiana tabacum (Müller and Grafe, 1978). Since NR-deficient mutants are routinely selected by using chlorate, they have necessarily to be chlorate resistant, but mutations in other genes might also be selected in chlorate-containing media (Prieto and Fernández, 1993). These observations could explain the apparently contradictory findings concerning chlorate-resistance and nitrate reductase not only in C. reinhardtii, but also in other organisms.

1. Nitrate Reductase Structural Mutants C. reinhardtii mutants deficient in NR activity fall into three phenotypic categories, as shown in Table 2: i) those lacking NAD(P)H-NR activity and either displaying terminal-NR activity or not, but possessing an active molybdopterin cofactor (MoCo); they are defective in the NR structural gene Nia1, ii) those lacking NR, NiR, NT and NiT activities; they are defective in the regulatory locus NIT2, and iii) those lacking the MoCo common to NR and other molybdoenzymes (of which only xanthine dehydrogenase has been identified in C. reinhardtii); they are defective at the NIT3, NIT4, NIT5/NIT6, or NIT7 loci (Fernández and Cárdenas 1989; Galván et al., 1992; Aguilar et al., 1992c). The results of in vivo genetic complementation tests between mutant strains in the first phenotypic category in combination with in vitro complementation tests for reconstituted NR activity from extracts of those strains have led to an interesting structural model for nitrate reductase (Fernández and Cárdenas, 1981a, 1982b; Fernández and Matagne, 1984, 1986). It was proposed that NR consists of two kinds of subunits: one of 45 kDa having NAD(P)Hcytochrome c reductase (diaphorase) activity, and another of 67 kDa with no activity but responsible for terminal-NR activity. Both ‘subunits’ would be encoded by the Nia1 gene which thus could be dicistronic (Fernández and Cárdenas, 1982b, 1983a,b; Fernández and Matagne, 1984, 1986; Franco et al., 1984a). NAD(P)H-NR activity can be restored in vitro by mixing enzyme preparations from two nia1 mutants, each having one of the two NR ‘subunits’ intact. Hybrid NR enzymes with restored activity are formed efficiently in diploid cells from appropriate nia1 haploid mutants that have each of the ‘subunits’

intact (Fernández and Matagne, 1986). However, characterization of the cloned Nia1 gene indicates that this gene is monocistronic and expresses a single 3.4 Kb transcript (Fernández et al., 1989). Furthermore, NR has been purified to electrophoretic homogeneity as a homodimer of 210 kDa. Thus the original two-subunit structural model has been modified. The 105 kDa NR subunits are very prone to proteolytic cleavage to yield two halves of 52 kDa (Kalakoutskii and Fernández, 1995). The efficient complementation between C. reinhardtii Nia1 mutants is therefore proposed to result from the ability of defective subunits to assemble into a heterologous functional NR(Fernández and Matagne, 1986). Intragenic complementation among Nicotiana plumbaginifolia Nia1 mutants has also been reported (Cherel et al., 1990).

2. Molybdopterin Cofactor Mutants MoCo-deficient mutants of C. reinhardtii that are defective in nitrate assimilation define five unlinked genes Nit3, Nit4, Nit5, Nit6, and Nit7 (Table 2; Fernández and Matagne, 1984; Aguilar et al., 1992c). Mutants that carry nit5 or nit6 alleles are particularly interesting, since they are phenotypically wild type with respect to nitrate utilization. They are referred to as cryptic MoCo mutants (Fernández and Matagne, 1984). Strain 21gr (nit5) and (nit6) are resistant to high concentrations of molybdate or tungstate (A. Llamas, K. Kalakoutskii and E. Fernández, unpublished; Fernández and Aguilar, 1987). These phenotypes have not been reported in other eukaryotic systems. Strains carrying mutations at the NIT4 or NIT5/NIT6 loci lack MoCo, but they recover the ability to grow in nitrate-containing media when high concentrations of molybdate (15 mM) are present. These loci have been proposed to encode proteins required for molybdate processing, i.e. transport, storage, insertion of molybdate into the organic moiety ofthe cofactor, processing of inserted molybdate, etc. (Fernández and Matagne, 1984; Fernández and Aguilar, 1987). Any of these mutants might correspond to the molybdate-repairable Cnx1 mutant from Arabidopsis thaliana, whose corresponding cDNA was cloned recently (Stallmeyer et al., 1995). Mutants defective at NIT3 or NIT7 loci are not rescued by high concentrations of molybdate. These loci are proposed to be involved in molybdopterin biosynthesis in C. reinhardtii (Fernández andAguilar,

Chapter 33 Nitrogen Assimilation and its Regulation

1987; Aguilar et al., 1992c). It seems that the ability of NR subunits to assemble into an enzyme complex depends on the organic moiety of the molybdenum cofactor (Johnson, 1980; Mendel et al., 1981). In agreement with this assumption, molybdate processing mutants from C. reinhardtii (nit4, nit5/nit6) have NR subunits assembled into a 220 kDa complex with only diaphorase activity. Interestingly, in nit3 and nit7 strains, NR subunits are still able to assemble to a certain extent, which suggests that these subunits might interact either by themselves in the absence of MoCo or in the presence of some molybdopterin precursor (Fernández and Aguilar, 1987; Aguilar et al., 1992c). A simple and reliable method has been set up to measure molybdopterin cofactor and some of its precursors. An oxidation product of molybdopterin cofactor (or the precursors) is produced and this can be quantitatively measured in cell-free extracts of C. reinhardtii (Aguilar et al., 1992a).This MoCo oxidation product was detected in extracts from nit3, nit4, nit5/nit6 and nit7 mutants at amounts similar to those found in the wild type extracts, which suggests that feed-back inhibition ofthe MoCo biosynthetic pathway is taking place even in the absence of active MoCo (Aguilar et al., 1992a). The presence of a MoCo oxidation product in nit4 and nit5/nit6 mutants is not surprising, since they can grow with nitrate in the presence of high molybdate. However, the existence of the MoCo oxidation product in nit3 and nit7 mutants suggests that they contain either molybdopterin or a close precursor of it that cannot be repaired by molybdate (Aguilar et al.,

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1992a). Thus, the nit3 and nit7 strains would be defective in a final step for molybdopterin biosynthesis or in the assembly or stabilization of the molybdenum-molybdopterin complex. MoCo is produced constitutively in C. reinhardtii, though its amounts are maximum in nitrate-grown cells and during the exponential phase of growth (Fernández and Cárdenas, 1981b; Aguilar et al., 1991). There is no correlation between the synthesis of MoCo and that of NR, as described for N. tabacum (Mendel et al., 1984; Aguilar et al., 1991). MoCo species are distributed mostly in two pools, namely a pool from which MoCo is able to reconstitute directly NR from apo NR of the Neurospora crassa nit1 mutant, and a pool in which MoCo is tightly bound to molybdoenzymes and only releasable by heat treatment(Aguilar et al., 1991, 1992b). Active MoCo can exist either in a low molecular weight form or bound to a 50 kDa protein named MoCo Carrier Protein (MoCoCP). This MoCoCP is able to transfer directly MoCo to apoNR only if both proteins are in physical contact (Aguilar et al., 1992b). The first MoCoCP was described in Escherichia coli (Amy and Rajagopalan, 1979). This 40 kDa E. coli protein releases MoCo very easily in contrast to the protein from C. reinhardtii. A protecting or stabilizing role of MoCo by the C. reinhardtii CP has been found (C. P. Witte, R. R. Mendel and E. Fernández, unpublished), as reported for E. coli (Amy and Rajagoplan, 1979). The C. reinhardtii CP has also been proposed to play a role in MoCo insertion into apomolybdoenzymes (Aguilar et al., 1992b). Recently, a

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MoCoCP with similar properties to the C. reinhardtii MoCoCP was purified and characterized from Vicia faba bean seeds (Kalakoutskii and Fernández, 1997).

3. Regulatory Mutants Mutant strains deficient at the NIT2 locus lack not only NR activity but also NiR and nitrate and nitrite uptake activities, though some nit2 strains show a leaky phenotype in nitrate or nitrite media (Nichols and Syrett, 1978; Fernández and Cárdenas, 1982a; Galván et al., 1992). These mutants still possess ammonium-repressible enzymes for the purine degradative pathway (Fernández and Cárdenas, 1981b; Pineda et al., 1984, 1987); so, it has been proposed that Nit2 is a regulatory gene specific for nitrate assimilation (Fernández and Cárdenas, 1989). Mutations at NIT2 are recessive to the wild-type NIT2 allele suggesting a positive control of Nit2 on the expression of nitrate assimilation genes (Fernández and Matagne, 1986). The NIT2 locus has been tagged with the C. reinhardtii Gulliver transposon, and the corresponding gene has been isolated (Ferris, 1989; Schnell and Lefebvre, 1993). The nit2 mutant phenotype can be complemented by transformation with genomic clones containing the regions surrounding the transposon insertion point. The DNA corresponding to the putative gene hybridizes to a 6 kb mRNA, and the nit2 mutant expresses an mRNA with an aberrant size (Schnell and Lefebvre, 1993). Interestingly, the Nit2 mRNA is down- regulated by ammonium, and accumulates in nitrogen-free media. The fact that nitrate assimilation related genes are regulated by both ammonium and Nit2, and that Nit2 is itself ammonium-regulated suggests that ammonium is mediating the metabolic repression of the nitrate assimilation pathway by controlling the synthesis of the Nit2 mRNA (Schnell and Lefebvre, 1993). Nit2 is the only regulatory gene for nitrate assimilation identified so far in plants. In addition, a major nitrogen regulatory gene, different from Nit2, has been postulated to mediate general repression by ammonium in C. reinhardtii (Fernández and Matagne, 1986). Two new putative regulatory genes, named Nrg1 and Nrg2, have been identified by insertional mutagenesis in C. reinhardtii (Prieto et al., 1996). Only Nrg1 is tagged by the transforming DNA; the nrg2 mutant might have appeared in cells where reversion or homologous recombination of the

selectable marker gene has occurred (Prieto et al., 1996). Mutations in these loci are recessive and unlinked. The nrg mutants show the phenotype of chlorate sensitivity in the presence of ammonium, which would result both from the expression of NT/ NiT transporters under repression conditions and from the activity of NT/NiT transporters in the presence of the inhibitor ammonium (Prieto et al., 1996). In fact, these mutants express low but detectable amounts of nitrate assimilation-related enzyme activities in ammonium containing media. However, the ammonium transporter activities are unaltered and show ammonium repression ofxanthine utilization like the wild-type strain. The reciprocal epistatic effect of the interactions between nit2 and nrg1 or nrg2 mutant alleles suggests that Nrgl and Nrg2 mediate directly or indirectly (through Nit2, for instance) their effects on structural genes for nitrate assimilation. Alternatively, Nrg genes could be involved in sensing the intracellular concentration of nitrogen metabolites. A defect in Nrg function would allow nitrate transport and expression of Nit2 and other structural genes under repression conditions (Prieto et al., 1996). Recently, spontaneous and insertional mutants with defective ammonia repression of the Nia1 gene have been isolated. They are allelic and define a new locus FAR1 (Zhang and Lefebvre, 1997). These mutants show similar phenotypes to Nrg deficient strains. The sequence and pattern of expression of Nrg1 and Nrg2 will, together with analogous studies of Nit2, provide considerable information about the molecular mechanisms controlling nitrate assimilation gene expression. Methylammonium, a non-metabolizable ammonium analogue for C. reinhardtii cells, has been used to isolate spontaneous mutants with an altered regulation pattern for nitrate reduction (Section II.A; Franco et al., 1987a). In nitrate medium containing methylammonium, nitrate uptake and reduction activities are inhibited and wild-type cells show a very poor growth (Franco et al., 1984b, 1987a). In methylammonium-resistant mutants isolated in nitrate medium, these enzyme activities are released from inhibition and the strains are therefore able to grow. These mutants define two loci MA1 and MA2, which do not encode regulatory genes but encode components for each of the two ammonium (methylammonium) transport systems (Section II.A). In mutant 2170 defective in the carrier 1, both NR and NiR are released from ammonium repression in

Chapter 33 Nitrogen Assimilation and its Regulation

ammonium plus nitrate containing media (Franco et al. 1987a, 1988a). Thus, ammonium effects on nitrate assimilation depend on its ability to enter the cells, and its further accumulation or metabolism.

B. Nitrate and Nitrite Transport Most of the research in the pathway has focused for years on the nitrate reducing enzymes. However, the first point of control ought to be at the level of transport so that the amount of nitrate taken up can be regulated to parallel the ability of the cell to metabolize it. Thus, excretion ofreduced nitrogenous compounds, which represents a waste of energy in terms of nitrate uptake and reduction to generate the reduced form, would be avoided (Fernández and Cárdenas, 1989; Hoff et al., 1994; Crawford, 1995).

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activities are blocked in the dark and by ammonium, (Florencio and Vega, and both activities require 1983a; Córdoba et al., 1987; Galván et al., 1991; Quesada et al., 1993, 1994; Galván et al., 1996). iv) Proteins involved in nitrite transport in the wild type show a rapid turnover (Galván et al., 1991). Since nitrite is reduced to ammonium in the chloroplast (Fischer and Klein, 1988), it has to cross two barriers, the plasma membrane and the two chloroplast envelope membranes. An efficient NiT could exist in the chloroplast inner envelope membrane, since intracellular nitrite, in contrast to nitrate, can not be detected in C. reinhardtii (Quesada et al., 1994; M.T. Navarro, A. Galván and E. Fernández, unpublished). The HANiT described above seems to operate at the plasmalemma.

1. Nitrate and Nitrite Uptake Activities

2. Nitrate and Nitrite Transporter Genes and Their Expression

Physiological studies have been performed to characterize nitrate and nitrite uptake systems in C. reinhardtii. In response to nitrate, a large number of soluble and membrane-bound polypeptides are synthesized in C. reinhardtii, which demonstrates the high complexity of this pathway even in such a simple eukaryotic organism (Watt et al., 1993). Nitrate uptake shows biphasic kinetics in C. reinhardtii, corresponding to high and low affinity transport activities (Watt et al., 1992). The synthesis of a membrane-bound polypeptide of 21 kDa and a pI 5, which occurs in response to nitrate provided at 1.1 mM concentration, is related to the low affinity transport activity (Watt et al., 1992). Low and high affinity nitrate transport activities have also been described in vascular plants (Glass, 1988; Siddiqi et al. 1990; Ulrich , 1992; Tsay et al., 1993; Wang and Crawford, 1996). Three high affinity nitrate/nitrite transporter (HANT/HANiT, respectively) systems have been characterized at the biochemical and molecular level in C. reinhardtii (see below). The characteristics of these transporters can be summarized as follows: i) nitrate and nitrite are transported by different specific systems with Ks in the micromolar range, though there exists also a bispecific transporter for both anions (Galván et al., 1996); ii) the NT and NiT are ammonium repressible, induced optimally in nitrate media, and under the control of the regulatory gene Nit2 (Galván et al., 1992; Quesada et al., 1994; Quesada and Fernández, 1994). iii) the NT and NiT

The NR-encoding NIA1 locus from C. reinhardtii is included in a cluster of at least six genes related to nitrate assimilation, formerly named Nar genes (Quesada et al., 1993; A. Quesada, I. Gómez and E. Fernández, unpublished). Transcripts from these genes are repressed by ammonium and positively controlled by the Nit2 gene product. Four genes are present within a 32 Kb genomic region of Nia1 (Quesada et al., 1993), including Nar1 (close to the 3´-end of Nia1), Nar2, Nrt2;1 (formerly nar3) and Nrt2;2 (formerly nar4; in this order from the 5´-end of Nia1) (see Fig. 3, Caboche et al., 1994). Several strains, selected among chlorate-resistant mutants, have deletions and/or rearrangements in the gene cluster and lack nitrate uptake activity and the ability to accumulate nitrate when it is provided at micromolar concentrations in the medium. Thus, the genes in the cluster are suggested to correspond to a HANT system. Two genes within the cluster, Nrt2;1 and Nrt2;2 show strong cross-hybridization, suggesting that they are related and perhaps originated from a gene duplication event (Quesada et al., 1993). The HANT genes have been identified among the Nar genes by the construction oftrue nitrate transport mutants (Quesada et al., 1994). These mutants correspond to strains with a Nar gene deletion which have a Nia1 copy unlinked to the cluster. The unlinked Nia1 gene was introduced into the Nar gene deletion strain by a genetic cross to a Nia1 transformant (Quesada et al., 1994). The Nar gene deletion mutants show NR activity but lack nitrate transport and are

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incapable of growing on nitrate-containing media. NT mutants are complemented by transformations with plasmids carrying Nrt2;1 plus Nar2 or Nrt2;2 plus Nar2, but not with any of these genes alone (Quesada et al., 1994). Complementary DNA sequences from Nrt2; 1 and Nrt2;2 genes show strong identity at both the DNA and the deduced amino acid sequence level. The encoded polypeptides have hydrophobic profiles which predict 12 transmembrane-spanning domains, supporting their involvement in solute transport (Quesada et al., 1994). Nrt2;l and Nrt2;2 proteins show a high similarity (up to 40% at the amino acid level) with CRNA, the NT-encoding gene from Aspergillus nidulans (Unkles et al., 1991), which is clustered together with the NR- and NiR-encoding genes from the fungus (Johnstone et al., 1990). However, the C. reinhardtii gene cluster contains two homologous genes encoding the putative transmembrane proteins involved in the nitrate transport activity, plus the Nar2 gene which is

necessary for having a fully functional uptake system (Quesada et al., 1994). Co-expression of Nrt2;1 and Nar2 in transformed strains leads to the recovery of HANT activity, accompanied by an increase in the HANiT activity. In contrast, co-expression of Nrt2;2 and Nar2 leads only to the recovery ofHANT activity (Galván et al., 1996). The kinetic parameters of HANT activities among strains transformed with the two gene combinations are slightly different, and these data indicate that Nrt2;l and Nar2 are components of a bispecific high affinity transporter for both nitrate and nitrite, whereas Nrt2;2 and Nar2 are components of a HANT (Galván et al., 1996). Nrt2-related cDNAs have been isolated from vascular plants (Trueman et al., 1996b; Quesada et al., 1997). Proteins CrnA and Nrt2 conform to a cluster of highly conserved transporters within the major facilitator superfamily (MFS), which also includes antibiotic resistance and solute transport proteins from prokaryotes and eukaryotes. A

Chapter 33 Nitrogen Assimilation and its Regulation

consistent similarity is also found between Nrt2 and NarK, a nitrate/nitrite anti-porter from Escherichia coli (Trueman et al., 1996a). Nrt2 sequences are unrelated to the cyanobacterial NT which belongs to the ABC-transporter superfamily (Omata et al., 1993), nor to the low affinity NT encoded by the CHL1 gene from Arabidopsis, which is a member of the POT (proton-dependent oligopeptide transporter) superfamily (Tsay et al., 1993; Trueman et al., 1996a). So far, there is no information about the functional expression of vascular plant NRT2 proteins, nor about the existence of Nar2-related proteins in other organisms. The Nar2 gene has been ‘knocked-out’ by homologous recombination upon transformation with an interrupted cDNA construction, which results in gene replacement (Nelson and Lefebvre, 1995). The isolated mutant does not grow on nitrate nor nitrite media, and the corresponding genetic locus has been named NIT8, following the original nomenclature. Two possibilities have been proposed for NAR2/ (NIT8) function: one is that Nar2 is a structural component of the nitrate uptake system, which associates functionally with each ofNrt2; 1 or Nrt2;2 (Quesada et al., 1994; Galván et al., 1996); and the other that Nar2 is a regulatory component of the nitrate assimilation pathway since the nit8 phenotype has been associated with a defect in nitrite growth (Nelson and Lefebvre, 1995). Very recently, Nar2 (Nit8) and Nrt2;1 cDNAs from C. reinhardtii have been expressed in oocytes from Xenopus, and significant electrogenic nitrate transport was achieved only when both transcripts were expressed together (A. Miller, personal communication). This finding strongly supports a structural role for Nar2 though an additional regulatory function cannot be ruled out. Excepting Nrt2;2, Nar genes are expressed in a co-regulated manner with Nia1. Transcripts of Nrt2;2 accumulate inversely with those of Nrt2; 1 and are more abundant after long periods of incubation in nitrate-containing or nitrogen-free media (Quesada and Fernández, 1994). Nar gene transcripts are upregulated in NR mutants when compared to the wild type, which further extends the postulated regulatory role ofNR to all Nar genes (Quesada and Fernández, 1994; Section V.C.3). No function has been reported for the Nar1 gene of the nitrate gene cluster (Quesada et al., 1993). Under the growth conditions used, there is apparently no effect of Nar1-gene deletion on nitrate or nitrite utilization (Quesada et al., 1993, 1994; Galván et al.,

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1996). Clustering of Nar1 with other co-regulated genes indicates that it could correspond to an as yet unidentified uptake activity or an enzyme of the pathway (Quesada et al., 1993; Quesada and Fernández, 1994). On the other hand, strains with a deleted nitrate gene cluster show a high affinity and nitrite-specific uptake activity, which is downregulated by ammonium (Quesada et al., 1993; Galván et al., 1996; Navarro et al., 1996). This fact suggests that there exists a transporter protein encoded by a gene not yet identified. Interestingly, a new Nrt2-related gene has been cloned recently from C. reinhardtii (Nrt2;3), and its expression has been related to the HANiT activity (A. Quesada, J. Hidalgo and E. Fernández, unpublished). The C. reinhardtii Nrt2 gene sequences and those of recently identified homologue genes in barley (Trueman et al., 1996b) have been used to design a general system for cloning related sequences from plants, which has allowed the isolation of Nrt2­ homologue sequences from Nicotiana (Quesada et al., 1997) and spinach (Quesada and Fernández, unpublished results). Nrt2 genes from vascular plants are expressed mainly in the root system, and are negatively-regulated by reduced-nitrogen compounds, such as glutamine, and positively by the supply of nitrate (Trueman et al., 1996b; Quesada et al., 1997). The genetic modification of nitrate transporter expression is now possible, and may represent a tool to optimize nitrate assimilation in crop plants.

C. Nitrate Reduction After nitrate uptake, the next step of the pathway is the reduction of nitrate to nitrite catalyzed by NR. This enzyme is subject to transcriptional, and posttranscriptional control, and its activity can be reversibly inactivated (Fernández and Cárdenas, 1989; Hoff et al., 1994; Crawford, 1995).

1. Nitrate Reductase Enzyme The biochemical and molecular properties of C. reinhardtii NR are similar to those reported in other green algae and vascular plants (Fernández and Cárdenas, 1989; Hoff et al., 1994; Zhou and Kleinhofs, 1996). NR transfers two electrons from NAD(P)H to nitrate via three redox centers (FAD, heme and MoCo). The prosthetic groups are located in three functional domains separated by two short

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hinge regions: the MoCo region in the N-terminal domain, the heme region in the middle, and the FAD region at the C-terminal end (Zhou and Kleinhofs, 1996). The NR enzyme complex shows two partial activities referred to as diaphorase (NAD(P)Hcytocrome c reductase) and terminal-NR ( NR or reduced viologen dye-NR) which involve different protein domains (Franco et al., 1984a; Hoff et al., 1994). NR from C. reinhardtii and higher plants is a homodimer with subunits of 110 kDa (Hoff et al., 1994; Kalakoustskii and Fernández, 1995). Reducing equivalents for nitrate reduction are supplied by the oxidative pentose phosphate pathway. Indeed, Glucose-6-phosphate dehydrogenase activity increases in C. reinhardtii cells transferred from ammonium to nitrate or to N- free media (Hipkin and Cannons, 1985; Huppe and Turpin, 1996). This increase is accompanied by the appearance of novel Glucose-6-phosphate dehydrogenase isoforms (Huppe and Turpin, 1996). Localization of the NR protein in green algae and higher plants is generally accepted to be cytosolic (Fischer and Klein, 1988; Hoff et al., 1994). It has to be pointed out that in Hansenula anomala reversible inactivation of NADPH-NR activity by heat shock correlates with NR association with mitochondrial membranes (Siverio et al., 1993). In Neurospora crassa, immunolocalization experiments suggest that NR is bound to the plasma membrane (Roldán et al., 1982). Whether or not association of NR to membranes is a general feature is not known. The physiological significance of membrane association in the context of regulation of nitrate assimilation also needs further study. As a key component of the nitrate assimilation pathway, NR activity is tightly regulated by environmental factors — mostly light, nitrogen and carbon availability. These factors seem to act by modifying transcription, post-transcriptional processes or enzyme activity (Fernández and Cárdenas, 1989). The presence of ammonium prevents NR expression; NR expression is high only when nitrate is in the medium (Fernández et al., 1989, Quesada and Fernández, 1994). Addition of ammonium to cells growing in nitrate causes a decrease in NR activity due to inhibition of de novo NR synthesis, reversible inactivation ofNR, and irreversible loss of previously synthesized enzyme (Guerrero et al., 1981; Franco et al., 1987b; Fernández and Cárdenas, 1989). In C. reinhardtii, inactivation of NR by ammonium has been proposed to occur by means of a redox

interconversion process (Herrera et al., 1972; Florencio and Vega, 1982). This inactive form ofNR seems to signal its subsequent degradation (Franco et al., 1987b). Reactivation of inactive NR occurs by oxidation in vitro with oxidants such as ferricyanide and in vivo with nitrate (Córdoba et al., 1985). Blue light can also reactivate the C. reinhardtii inactive NR form in situ (Azuara and Aparicio, 1983). When NR is constitutively expressed from the cabII-1 gene promoter in the presence of ammonium, Nia1 transcript but not NR activity is observed (Blankenship and Kindle, 1992). In this case, NR is synthesized mostly in an inactive form which can be reactivated by ferricyanide or nitrate, suggesting that nitrate plays an important role in post-transcriptional regulation (Navarro et al., 1996).

2. Nitrate Reductase Gene The C. reinhardtii NR gene (Nia1, formerly named nit1) was cloned by heterologous hybridization with a barley NR cDNA (Fernández et al., 1989). The genomic fragment containing Nia1 was mapped and found to be closely linked to a NR mutation (nit1­ 305). The nit1 strain was in addition complemented by particle-gun mediated transformation with the wild-type Nia1 gene. This work represented one of the first reports of stable nuclear transformation in this alga; another was the complementation of arg7 strains with the cloned ARG7 locus (Fernández et al., 1989; Kindle et al., 1989; Debuchy et al., 1989; Kindle, 1990; see chapter by Kindle). The integration of the introduced DNA occurs mainly by nonhomologous recombination, although the use of truncated Nia1 constructs to rescue Nia1 NR-deficient strains allows for the detection of homologous recombination events (Sodeinde and Kindle, 1993). The Volvox carteri NR locus (nitA) has been isolated by using Nia1 from C. reinhardtii as a hybridization probe (Gruber et al., 1992). NR sequences from both organisms show a high degree ofamino acid sequence conservation, and they are also conserved with respect to the higher plant and fungal NRs (Fernández et al., 1989; Gruber et al., 1992; Zhou and Kleinhofs, 1996). Sequence alignment reveals three conserved domains involved in the binding of FAD, heme, and MoCo cofactors. The positions of eight introns are identical between the two algal NR genes, but the C. reinhardtii gene has five additional introns which do not exist in nitA, and another two are slightly displaced in nitA (Gruber et al., 1992; Zhou and

Chapter 33 Nitrogen Assimilation and its Regulation

Kleinhofs, 1996). Like the C. reinhardtii Nia1 gene, nitA has been used to set up reliable techniques for stable nuclear transformation of V. carteri (Schiedlmeier et al., 1994).

3. Nitrate Reductase Gene Expression The analysis of Nia1 gene expression shows that accumulation of its mRNA is controlled by ammonium, nitrate, light and the Nit2 regulatory gene product (Fernández et al. 1989; Quesada and Fernández, 1994). Ammonium represses Nia1 expression, and nitrogen starvation causes derepression. Nia1 transcripts accumulate faster and to a higher extent under derepressing conditions in proportion to increasing nitrate concentrations in the media (Quesada and Fernández, 1994). Light is also an important factor for efficient expression of Nia1, but the supply of acetate in the dark partially relieves the light requirement for expression. Thus, it has been suggested that the energy status of the cells regulates the expression of Nia1 (Quesada and Fernández, 1994). Both Nia1 mRNA accumulation and NR activity (the remaining partial activity) were up-regulated in various NR-deficient mutants which are derepressed in nitrogen-free media compared to the wild-type strain, and it was suggested therefore that NR activity and/or structure regulates its own mRNA expression (Fernández and Cárdenas, 1982a; Quesada and Fernández, 1994). The expression of other enzyme activities (including NiR, HANiT) and Nar gene transcript abundance are also up-regulated in N- free medium in C. reinhardtii NR mutants (Galván et al., 1992; Quesada and Fernández, 1994). This regulatory effect has been assigned to NR itself since C. reinhardtii NiR mutants show wild type regulation (M. T. Navarro, A. Galván, E. Fernández, unpublished), in contrast to the NiR mutants isolated with an antisense strategy in Nicotiana tabacum (Vaucheret et al., 1992). The Nia1 coding sequence has been used also as a reporter gene by placement under control of the nitrogen-constitutive and light-induced promoter sequence of the CabII-1 gene from C. reinhardtii (Blankenship and Kindle, 1992). Transformants can be obtained by complementation of nit1 strains or even nit1 nit2 strains on nitrate media with the chimeric constructs. Nia1 mRNAs are transcribed from the CabII-1 promoter in ammonium containing medium, and transcripts are released from Nit2 control

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in transgenic nit2 strains. These facts indicate that ammonium and Nit2 regulate Nia1 gene expression in wild-type cells by acting at the transcriptional level on the Nia1 promoter (Blankenship and Kindle, 1992). In contrast, nitrate induced the enhanced expression of transcripts from the CabII–1:Nia1 construct (Navarro et al., 1996), suggesting that the positive effect of nitrate on Nia1 mRNA accumulation (Quesada and Fernández, 1994) is probably occurring at a post-transcriptional level (Navarro et al., 1996). Constitutive expression of Nia1 transcripts from the CabII–1:Nia1 constructs in transformed strains (Blankenship and Kindle, 1992) allows release of HANT and HANiT activities from ammonium inhibitory effects, indicating that NR expression is changing the regulation of NT genes, even though NR is mostly in an inactive form and the NT mRNAs are not detectable in ammonium containing media (Navarro et al., 1996). The Nia1 gene promoter has been analyzed by fusing it to the arylsulfatase (Ars) gene used as a reporter (Ohresser et al., 1997). About 1 kb of the Nia1 promoter sequence drives the ammoniumregulated expression of Ars, but no differences are found between the induction ofthe chimericNia1:Ars gene when nitrate or nitrite are supplied to the cells, or under nitrogen starvation conditions. In addition, no Ars- expressing transformants are obtained in a nit2 genetic background, which is in agreement with the transcriptional control by both ammonium and the Nit2 gene product of Nia1 gene expression (Blankenship and Kindle, 1992; Ohresser et al., 1997), and with post-transcriptional regulation by nitrate (Navarro et al., 1996; Ohresser et al., 1997). Deletion analysis revealed that several DNA regions within the Nia1 promoter have a strong effect on Nia1:Ars gene expression. The region between –282 and –198 (from the putative Nia1 transcription starting point) is required for expression after ammonium removal; the region from –342 to –282 contains a transcription repressor sequence; and finally various deletions from the 5´ end of the 1 kb Nia1 promoter produced a progressive decrease of Ars activity (Ohresser et al., 1997). An additional regulatory mechanism for expression of the NR-gene might occur at the mRNA maturation level, since several introns from the V. carteri nitA gene increase the efficiency of transformation with this selectable marker (Gruber et al., 1996). When a construct containing the nitA cDNA fused to its own promoter is used in transformation experiments, the

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frequency at which transformants arose was much lower than that obtained with the intact nitA gene. If intron 1, or introns 9 plus 10, are re-integrated into the cDNA construct, the transformation frequency was greatly enhanced (Gruber et al., 1996).

D. Nitrite Reduction Nitrite reduction is the last step of the nitrate assimilation pathway leading to ammonium production in a six-electron step catalyzed by NiR, which uses reduced ferredoxin (fd) as an electron donor (Vega et al., 1980; Campbell and Kinghorn, 1990).

1. Nitrite Reductase Enzyme The C. reinhardtii fd-NiR is a nucleus-encoded chloroplastic enzyme (Fischer and Klein, 1988; Quesada et al., 1997a). The holoenzyme is a 63 kDa monomer with two redox centers: a siroheme and a cluster (Romero et al., 1987; Campbell and Kinghorn, 1990; A. Quesada, I. Gómez and E. Fernández, unpublished). Immunological studies with specific polyclonal antibodies against either NiR or fd-GOGAT from C. reinhardtii show that the antibodies are cross-reactive, which indicates that they share common antigenic determinants located at the fd-binding domain (Romero et al., 1988, Pajuelo et al., 1993). NiR activity shows a similar regulation pattern as does NR activity and requires the absence of ammonium, and the presence of light and nitrate for maximum levels (Fernández and Cárdenas, 1982; Galván et al., 1991). Like the other structural elements of the pathway NiR is also subject to control by NR itself (Galván et al., 1992). NiR is a stable enzyme with a longer half-life than NR or NiT (Galván et al., 1991; Pajuelo et al., 1995).

2. Nitrite Reductase Gene and Expression Two NiR-deficient mutants have been identified among strains that carry a deletion in the Nar genes. Their genetic analysis shows that the defective locus is closely linked to the Nia1 gene (Quesada et al., 1993). By cloning up to 45 kb of DNA in the vicinity of the Nia1 gene, two new genes were detected by RNA hybridization analysis (A. Quesada, I. Gómez and E. Fernández, unpublished) (see Fig. 3). One of them (Nii1, see below) co- regulates with the Nar genes, being repressed by ammonium, positivelycontrolled by nitrate and the Nit2 gene product, and

down-regulated by NR activity. In addition, there is a positive effect of light on expression of the Nii1 gene, which is partially substituted in the dark by the supply ofacetate. The newly identified gene is deleted or reorganized in the two NiR-deficient mutants which do not express the corresponding mRNA. The NiR-deficient phenotype is complemented by transformation with a genomic fragment which includes the RNA hybridizing region, indicating that the corresponding gene is probably the NiR-encoding structural gene from C. reinhardtii (Nii1). In fact, Nii1 cDNA encodes a protein that shows high conservation with vascular plant and cyanobacterial NiRs. The deduced Nii1 amino acid sequence is preceded by a putative chloroplast-transit peptide, whose structure is in agreement with an established consensus for plastid protein targeting in C. rein­ hardtii (Franzén et al., 1990; A. Quesada, I. Gómez and E. Fernández, unpublished). NiR structure is very similar among cyanobacteria, algae and vascular plants. This includes four conserved cysteines involved in the binding of the enzyme cofactors, siroheme and a redox center, and a positively-charged region that probably corresponds to the domain interacting with ferredoxin (Campbell and Kinghorn, 1990). This conservation of protein function among the three main classes of aerobic and photosynthetic organisms is a unique feature among nitrate assimilation genes. In fact, nitrate (nitrite) transporter genes from plants have a different evolutionary origin from those of cyanobacteria (Section V.B.2; Omata et al., 1993). However, an Nrt2 sequence in the cyanobacterium Oscillatoria chalybea has recently been described, but its function remains unknown (Unman et al., 1996). NR structure from plants is not related to that of cyanobacteria which is closer to the enterobacterial NR (Campbell and Kinghorn, 1990; Unthan et al., 1996). The chloroplast localization of plant NiRs might provide an explanation for why evolutionary pressure has not affected NiR enzyme structure to the same extent as it has for cytosolic or transmembrane proteins (Campbell and Kinghorn, 1990; Luque et al., 1993; A. Quesada, I. Gómez and E. Fernández, unpublished).

VI. Concluding Remarks Chlamydomonas reinhardtii, a relatively simple organism in structural and metabolic terms, is able to assimilate efficiently nitrogen from many different

Chapter 33 Nitrogen Assimilation and its Regulation

nitrogenous compounds. This efficiency is first ensured by specific transport systems that operate with substrate concentrations in the micromolar range, and that seem to depend on other membrane systems such as proton-ATPases, cation channels or pumps. These auxiliary systems will require future research to understand transport processes in depth. The characterized nitrate assimilation pathway in C. reinhardtii has allowed us to define the structural elements participating in the process. However, only four regulatory genes have been reported: Nit2, Far1, Nrg1, and Nrg2. Their function is still far from clear and more genes for the sensing of ammonium and nitrate should be characterized and their functions studied. Molecular and metabolic signals for the cross-talk among nitrogen metabolism elements from the chloroplast and the cytoplasm, and between Nand C-metabolism are still open questions.

Acknowledgments We thank Dr. Anthony Miller for sharing unpublished data, and Dr. Manuel Pineda for criticisms on the manuscript. Work from author’s laboratory is supported in part by grants from the BIOTEC program of the European Union (BIO2CT930400), DGICYT, Spain (PB95-0554-CO2-01), and Junta de Andalucía, Spain (PAI, Grupo CVI-0128).

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of nitrate in Chlamydomonas reinhardii. Phytochemistry 21: 1195–1200 Florencio FJ and Vega JM (1983a) Utilization of nitrate, nitrite and ammonium by Chlamydomonas reinhardii. Planta 158: 288–293 Florencio FJ, and Vega JM (1983b) Separation, purification and characterization of two isoforms of glutamine synthetase from Chlamydomonas reinhardii. Z Naturforsch 38c: 531–538 Florencio FJ, Gadal P and Buchaman BB (1993) Thioredoxinlinked activation of the chloroplast and cytosolic forms of Chlamydomonas reinhardtii glutamine synthetase. Plant Physiol Biochem 31: 649–655 Flores E and Herrero A (1994) Assimilatory nitrogen metabolism and its regulation. In: Bryant DA (ed) The Molecular Biology of Cyanobacteria, pp 487–517. Kluwer Academic Publishers, Dordrecht Franco AR, Cárdenas J and Fernández E (1984a) Heteromultimeric structure on the nitrate reductase complex of Chlamydomonas reinhardii. EM BO J 3: 1403–1407 Franco AR, Cárdenas J and Fernández E (1984b) Ammonium (methylammonium) is the corepressor of nitrate reductase in Chlamydomonas reinhardii. FEES Lett 176: 453–456 Franco AR, Cárdenas J and Fernández E (1987a) A mutant of Chlamydomonas reinhardtiialtered in the transport of ammonium and methylammonium. Mol Gen Genet 206: 414–418 Franco AR, Cárdenas J and Fernández E (1987b) Involvement of reversible inactivation in the regulation of nitrate reductase enzyme levels in Chlamydomonas reinhardtii. Plant Physiol 84: 665–669 Franco AR, Cárdenas J and Fernández E (1988a) Regulation by ammonium ofnitrate and nitrite assimilation in Chlamydomonas reinhardtii. Biochim Biophys Acta 951: 98–103 Franco AR, Cárdenas J and Fernández E (1988b) Two different carriers transport both ammonium and methylammonium in Chlamydomonas reinhardtii. J Biol Chem 263: 14039–14043 Franco AR, Diaz ME, Pineda M and Cárdenas J (1996a) Characterization of a mutant of Chlamydomonas reinhardtii that uses L-methionine-S- sulfoximine and phosphinothricin as nitrogen sources for growth. Plant Physiol 110: 1215–1222 Franco AR, López-Siles FJ and Cárdenas J (1996b) Resistance to phosphinothricin (glufosinate) and its utilization as a nitrogen source by Chlamydomonas reinhardtii. Appl Environ Microbiol 62:3834–3839 Franzén LG, Rochaix JD and Heijne G (1990) Chloroplast transit peptides from the green alga Chlamydomonas reinhardtii share features with both mitochondria and higher plant chloroplast presequences. FEBS Lett 260: 165–168 Galván A, Córdoba F, Cárdenas J and Fernández E (1991) Regulation of nitrite uptake and nitrite reductase expression in Chlamydomonas reinhardtii. Biochim Biophys Acta 1074: 6– 11 Galván A, Cárdenas J and Fernández E (1992) Nitrate reductase regulates expression of nitrite uptake and nitrite reductase activities in Chlamydomonas reinhardtii Plant Physiol 98: 422–426 Galván A, Quesada A and Fernández E (1996) Nitrate and nitrite are transported by different specific transport systems and by a bispecific transporter in Chlamydomonas reinhardtii. J Biol Chem 271:2088–2092 Galván F, Márquez A and Vega JM (1984) Purification and molecular properties of ferredoxin-glutamate synthase from

Chapter 33 Nitrogen Assimilation and its Regulation

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Mendel RR, Buchanan RJ and Wray JL (1984) Characterization of a new type of molybdenum cofactor-mutant in cell cultures of Nicotiana tabacum. Mol Gen Genet 195: 186–189 Miflin PJ and Lea P (1976) The pathway ofnitrogen assimilation in plants. Phytochemistry 15: 873–885 Moyano E, Cárdenas J and Muñoz-Blanco J (1992a) Purification isozymes of L-glutamate and properties of three dehydrogenase of Chlamydomonas reinhardtii. Biochim Biophys Acta 1119: 63–68 Moyano E, Ramazanov Z, Cárdenas J and Muñoz-Blanco J (1992b) Intracellular localization of three L-glutamate dehydrogenase isozymes from Chlamydomonas reinhardtii. Plant Physiol 100: 1575–1579 Moyano E, Cárdenas J and Muñoz-Blanco J (1995) Involvement - glutamate dehydrogenase isoenzymes in carbon of and nitrogen metabolism in Chlamydomonas reinhardtii. Physiol Plant 94: 553–559 Müller AJ and Grafe R (1978) Isolation and characterization of cell lines of Nicotiana tabacum lacking nitrate reductase. Mol Gen Genet 161:67–76 Muñoz-Blanco J and Cárdenas J (1992) Changes in glutamate dehydrogenase activity of Chlamydomonas reinhardtii under different trophic and stress conditions. Plant Cell Environ 12: 173–182 Muñoz-Blanco J, Laín-Guelbenzu B and Cárdenas J (1988) Characterization of an L-aspartate aminotransferase activity in Chlamydomonas reinhardtii. Physiol Plant 74: 433–439 Muñoz-Blanco J, Moyano E and Cárdenas J (1989) Glutamate dehydrogenase isozymes of Chlamydomonas reinhardtii. FEMS Microbiol Lett 61: 315-318 Muñoz-Blanco J, Hidalgo-Martínez J and Cárdenas J (1990) Extracellular deamination of L-amino acids by Chlamydomonas reinhardtii cells. Planta 182: 194–198 Navarro MT, Prieto R, Fernández E and Galván A (1996) Constitutive expression of nitrate reductase changes the regulation ofnitrate and nitrite transporters in Chlamydomonas reinhardtii. Plant J 9: 819–827 Nelson JAE and Lefebvre PA (1995) Targeted disruption of the NIT8 gene in Chlamydomonas reinhardtii. Mol Cell Biol 15: 5762–5769 Nichols GL and Syrett PJ (1978) Nitrate reductase deficient mutants of Chlamydomonas reinhardii. Isolation and genetics. J Gen Microbiol 108: 71–77 Ninnemann O, Jauniaux JC and Frommer WB (1994) Identification of a high-affinity transporter from plants. EMBO J 13: 3464–3471 Ohresser M, Matagne RF and Loppes R (1997) Expression of the arylsulphatase gene under the control of the nit1 promoter in Chlamydomonas reinhardtii. Curr Genet 31:264–271 Omata T, Andriesse X and Hirano A (1993) Identification and characterization of a gene cluster involved in nitrate transport in the cyanobacterium Synechococcus sp PCC 7942. Mol Gen Genet 236: 193–202 Pajuelo E, Borrero JA and Márquez A (1993) Immunological approach to subunit composition offerredoxin-nitrite reductase from Chlamydomonas reinhardtii. Plant Sci 95: 9–21 Pajuelo E, Pajuelo P, Clemente MT and Márquez A (1995) Regulation of the expression of ferredoxin-nitrite reductase in synchronous cultures of Chlamydomonas reinhardtii. Biochem Biophys Acta 1249:72–78 Pérez-Vicente R, Cárdenas J and Pineda M (1991) Distinction

between hypoxanthine and xanthine transport in Chlamy­ domonas reinhardtii. Plant Physiol 95: 126–130 Pérez-Vicente R, Pineda M and Cárdenas J (1987) Occurrence of an NADH-diaphorase activity associated with xanthine dehydrogenase in Chlamydomonas reinhardtii. FEMS Microbiol Lett 43: 321–325 Pérez-Vicente R, Pineda M and Cárdenas J (1988) Isolation and characterization of xanthine dehydrogenase from Chlamy­ domonas reinhardtii. Physiol Plant 72: 101–107 Pérez-Vicente R, Alamillo JM, Cárdenas J and Pineda M (1992) Purification and substrate inactivation of xanthine dehydrogenase from Chlamydomonas reinhardtii, Biochim Biophys Acta 1117: 159–166 Pérez-Vicente R, Burón MI, González-Reyes JA, Cárdenas J and Pineda M (1995) Xanthine accumulation and vacuolization in Chlamydomonas reinhardtii cells. Protoplasma 186: 93–98 Piedras P (1995) Metabolismo de los ureidos alantoina y alantoato en Chlamydomonas reinhardtii. PhD Dissertation. Universidad de Córdoba, Córdoba Piedras P, Pineda M, Muñoz J and Cárdenas J (1992) Purification and characterization of an L-amino acid oxidase from Chlamydomonas reinhardtii. Planta 188: 13–18 Piedras P, Cárdenas J and Pineda M (1995) Solubilization and extraction ofallantoinase and allantoicase from the green alga Chlamydomonas reinhardtii. Phytochem Anal 6: 239–243 Pineda M and Cárdenas J (1985) The urate uptake system in Chlamydomonas reinhardtii. Biochim Biophys Acta 820: 95– 99 Pineda M and Cárdenas J (1996) Transport and assimilation of purines in Chlamydomonas reinhardtii. Sci Mar 60: 195–201 Pineda M, Fernández E and Cárdenas J (1984) Urate oxidase of Chlamydomonas reinhardii. Physiol Plant 62: 453–457 Pineda M, Cabello P and Cárdenas J (1987) Ammonium regulation of urate uptake in Chlamydomonas reinhardtii. Planta 171: 496–500 Pineda M, Piedras P and Cárdenas J (1994) A continuous spectrophotometric assay for ureideglycolase activity with lactate dehydrogenase or glyoxylate reductase as coupling enzyme. Anal Biochem 222: 450–455 Prieto R and Fernández E (1993) Toxicity and mutagenesis by chlorate are independent of nitrate reductase activity in Chlamydomonas reinhardtii. Mol Gen Genet 237: 429–438 Prieto R, Dubus A, Galván A and Fernández E (1996) Isolation and characterization oftwo new regulatory mutants for nitrate assimilation in Chlamydomonas reinhardtii obtained by insertional mutagenesis. Mol Gen Genet 251: 461–471 Quesada A and Fernández E (1994) Expression of nitrate assimilation related genes in Chlamydomonas reinhardtii. Plant Mol Biol 24: 185–194 Quesada A, Galván A, Schnell RA, Lefebvre PA and Fernández E (1993) Five nitrate assimilation-related loci are clustered in Chlamydomonas reinhardtii. Mol Gen Genet 240: 387–394 Quesada A, Galván A and Fernández E (1994) Identification of nitrate transporter genes in Chlamydomonas reinhardtii. Plant J 5: 407–419 Quesada A, Krapp A, Trueman L, Daniel-Vedele F, Fernández E, Forde BG and Caboche M (1997) PCR-identification of a Nicotiana plumbaginifolia cDNA homologous to the high affinity nitrate transporters ofthe crnA family. Plant Mol Biol 34:265–274 Ramazanov Z andCárdenasJ (1994)Photorespiratoryammonium

Chapter 33 Nitrogen Assimilation and its Regulation

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Chapter 34 Mitochondrial Genetics Claire Remacle and René F. Matagne

Genetics of Microorganisms, Department of Plant Biology, B22,

University of Liège, B-4000 Liège, Belgium

Summary 661

I. Introduction 662

II. Mitochondrial Genome 662

A. Structure, Size and Copy Number 662

B. Gene Content 662

C. Replication 663

D. Transcription and Processing 664

E. Translation 664

III. Mitochondria and the Electron Transport Chain

664 IV. Mutations Affecting the Mitochondrial Genome

665 A. Lethal Minute Colony Mutants 665

B. Obligate Photoautotrophic Respiratory-Deficient Mutants 665

C. Myxothiazol-Resistant Mutants 667

668

V. Transmission of Mitochondrial Genes in Meiotic Zygotes VI. Transmission of Mitochondrial Genes in Vegetative Zygotes and Mapping of Mitochondrial Mutations by

669

Recombinational Analysis 669

A. Inheritance of Mitochondrial Genes in Vegetative Zygotes B. Mapping Mutations by Recombinational Analysis 670

671

VII. Mitochondrial Transformation 672

Acknowledgments 672

References

Summary Chlamydomonas reinhardtii is now becoming a useful model for the study of mitochondrial genetics in photosynthetic organisms. The linear 15.8 kb mitochondrial genome of the alga has been completely sequenced and all the genes have been identified. Although Chlamydomonas probably cannot survive in the absence of the mitochondrial genome, the inactivation of certain mitochondrial genes is not lethal for the cell. The lack of a functional cob (apocytochrome b) or cox1 (subunit 1 ofcytochrome c oxidase) gene prevents cell division in the dark but has little effect on photoautotrophic growth. A deletion encompassing the cob and nd4 (subunit 4 of NADH dehydrogenase) genes still allows the cells to grow in the light. Specific amino acid changes in the apocytochrome b sequence, conferring resistance to inhibitors (myxothiazol, mucidin) of complex III, have also been characterized. The mitochondrial mutations, as well as the RFLP that distinguishes two interfertile strains, have permitted the demonstration that in crosses, the meiotic progeny issued from the zygotes most often inherit the mitochondrial DNA from the mating-type minus parent. The mitochondrial DNA of mating-type plus origin is slowly eliminated during the maturation and the meiotic divisions of the zygospore. This contrasts with the parent; elimination of the transmission pattern observed for the chloroplast genome (transmission from the chloroplast DNA very soon after zygote formation). J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 661–674. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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Claire Remacle and René F. Matagne

The rare zygotes which divide mitotically and give rise to diploid progeny transmit mitochondrial genomes from both parents. In these diploids, recombination events occur between mitochondrial markers and the recombination frequency is dependent on the physical distance between the markers. As the recombination frequency is ca. 3.2% per kb, mitochondrial mutations less than 5–7 kb apart can be genetically mapped by recombinational analysis. One case of mitochondrial transformation has also been reported in Chlamydomonas. Future experiments in that line will allow the evaluation of the consequences of DNA modifications made in vitro and the analysis of the phenotypic changes caused by the presence of novel gene sequences in the mitochondrial genome.

I. Introduction Because Saccharomyces cerevisiae has the unique property to survive in the absence of mitochondrial DNA, yeast mitochondria represent a genetic system that is particularly amenable to experimental analysis. The numerous mitochondrial mutations characterized in baker’s yeast have allowed extensive genetic analysis, including recombination and more recently, transformation methodologies (Costanzo and Fox, 1990; Gillham, 1994). Few mitochondrial mutations causing an aberrant phenotype independently of the nuclear background have been characterized in higher plants. Maternallyinherited mutations, corresponding to deletions in mitochondrial genes coding for subunits of cytochrome c oxidase, subunits of NADH dehydrogenase or ribosomal proteins, have been identified in maize. The mutant plants exhibit poor growth, reduced vigor and pale striping on leaves (NCS or non-chromosomal stripe mutants). All NCS mutants still contain at least 50% of non-mutated mitochondrial DNA copies, and this heteroplasmy has been suggested to account for plant viability. Only the defective pale-green sectors are near-homoplasmic for mitochondrial mutation (Newton, 1995). Recently, two CMS (cytoplasmic male sterility) mutants with reduced respiration have been characterized in Nicotiania sylvestris. Both are nearhomoplasmic for a deletion in the nad7 (subunit 7 of NADH dehydrogenase) gene, indicating that the complex I defect is not lethal for the plant (Pla et al., 1995) Abbreviations: bp – base pair; cob – apocytochrome b; cox – cytochrome c oxidase; dk – dark; dum – dark uniparental minus; kb – kilobase pairs; – mating-type plus; MT – – mating-type minus; nd – NADH dehydrogenase; ORF – open reading frame; PCR – polymerase chain reaction; RFLP – restriction fragment length polymorphism; rtl – reverse transcriptase like; SHAM – salicylhydroxamic acid

In Chlamydomonas reinhardtii, mitochondrial genetics has progressed considerably in the last ten years and this organism can now be used as a model system for plant cells to investigate mitochondrial gene function. The small (15.8 kb) mitochondrial genome of C. reinhardtii has been sequenced completely and all the genes residing in the organelle have been identified. Several mutants inactivated in respiratory complexes or resistant to complex III inhibitors have been characterized, and various methods permitting the genetic analysis of the mitochondrial genome are now available. These recent developments will be the focus ofthe present chapter. A review on the same topics has been published recently by Boynton and Gillham (1996).

II. Mitochondrial Genome

A. Structure, Size and Copy Number The mitochondrial DNA of C. reinhardtii was first characterized by Ryan et al. (1978) from a mitochondrial pellet. It has a buoyant density of 1.706 g/ml in CsCl, a melting temperature of 87.9 °C in standard saline citrate and a kinetic complexity of daltons. Electron microscopy studies have shown that 99% of the molecules are linear or, more rarely (less than 1%), open or closed circular. The linear and circular molecules are all within the range The genome is present in about 50 of copies per cell and represents 0.6% of the total cellular DNA. Physical mapping using six restriction endonucleases confirmed the linearity ofthe genome and its unique ends (Grant and Chiang, 1980).

B. Gene Content The first mitochondrial genes were identified by DNA sequence analysis and heterologous hybridi-

Chapter 34

Mitochondrial Genetics

zation(Pratje et al., 1984; Boer et al., 1985). In 1993, the mitochondrial genome sequence was completed (Vahrenholz et al., 1993) (Genbank Accession No U03843). It is a 15.8 kb linear DNA molecule(15758 bp, 45% GC) that contains thirteen genes. They encode subunit 1 of cytochrome c oxidase (complex IV), apocytochrome b (complex III), five subunits of NADH:ubiquinone oxidoreductase (complex I), a reverse transcriptase-like protein, three tRNAs and the rRNAs for the large (L) and small (S) ribosomal subunits (Fig. 1). The rRNA genes are discontinuous and split into four (S) and eight (L) modules, interspersed with one another, and with protein and tRNA genes. The small RNAs, S1 to S4 and L1 to L8, are thought to interact by way of extensive intermolecular pairing to form the conventional rRNA molecules (Boer and Gray, 1988b). The genes are located on both DNA strands: cob, nd4 and nd5 are transcribed from one strand, and the other genes are expressed from the opposite strand (Fig. 1). The non-coding sequences are very short in C. reinhardtii and the genes do not contain introns. The ends ofthe genome have been studied in detail by Ma et al. (1992a) and Vahrenholz et al. (1993). They are characterized by terminal inverted repeats of 531 or 532 bp in length. Moreover, both left and right ends exhibit long 3´ single-stranded extensions of identical sequence (39–41 nucleotides). The 86 outermost nucleotides of each end are identical to an internal 86-bp sequence, which is located at the 3´ end of the L2b gene, 674 bp from the right end ofthe

663 molecule (Vahrenholz et al., 1993) (Fig. 1). Chlamydomonas smithii, Boynton et al. (1987) have shown that the mitochondrial DNAs of the two interfertile species C. smithii and C. reinhardtii are co-linear with the exception of an insertion (ca. 1 kb) present in the cob gene of C. smithii DNA. Colleaux et al. (1990) have sequenced this insertion and shown that it corresponds to a group I intron of 1075 bp (the Cs cob. 1 or a intron). It possesses a 237 codon ORF in frame with the upstream exon of cob, whose product shows 36% amino acid identity with the ISceI endonuclease of S. cerevisiae. The intron putative ribozyme sequence shows 60% identity at the nucleotide level with the Neuropora crassa cob. 1 intron. The ORF product has endonucleolytic activity, specific for a sequence within the intron-free cob gene of C. reinhardtii (Ma et al., 1992b).

C. Replication The presence of two long but non-complementary single-stranded ends excludes the possibility of circularization or concatamerization ofthe molecule to form a replication intermediate, in contrast to what is found for the replication of some phages. The fact that the 5´ ends of the molecule can be efficiently labeled by T4 polynucleotide kinase (Grant and Chiang, 1980) also excludes the possibility that replication could proceed by a mechanism involving proteins linked covalently to these ends. Vahrenholz et al. (1993) have proposed two models for replication,

664 both of them based on the presence of the 86-bp internal repeat sequence (cfabove). One ofthe models postulates the intervention of a reverse transcriptase, which could be encoded by the ORF of the rtl gene.

D. Transcription and Processing Both strands of the mitochondrial DNA encode genetic information and two large transcription units have been postulated (Fig. 1). The leftward one includes the nd5, nd4 and cob genes whereas the rightward one comprises the cox1, nd2, nd6, nd1, rtl, rRNA and tRNA genes. The short intergenic region between the nd5 and cox1 genes might act as a bidirectional promoter but some controversy exists about this point, since transcript mapping studies have identified low levels of nd5 precursor RNA whose 5´ end overlaps the beginning ofthe cox1 gene (Gray and Boer, 1988). As shown by primer extension, S1 nuclease and RNase protection experiments, mature RNAs derive from nearly the entire genome and are generated by precise endonucleolytic cleavage of the long precursors. Pratje et al. (1984) and Boer and Gray (1986a) have identified a number of conserved sequence elements with potential hairpin structure within the 3´-end terminal regions of nd2, nd5 and cox1 genes. Their location close to mRNA termini suggests that these motives may play a role in the endonucleolytic cleavage of the primary transcripts. Moreover, in spacer regions associated with the rRNA coding segments and the tRNA genes, a set of short repeated sequences containing a palindromic consensus motif could also be involved in post-transcriptional mitochondrial RNA processing (Boer and Gray, 1991).

E. Translation The universal genetic code is used in mitochondria of C. reinhardtii (Kück and Neuhaus, 1986;Michaelis et al., 1990) but codon distribution is highly biased. Nine codons are not used at all and four codons occur exclusively in the rtl gene (Colleaux et al., 1990; Michaelis et al., 1990). This latter gene is probably the ORF of a ancient group II intron (Boer and Gray, 1988a). In spite of the absence of certain codons, a minimum of 23 tRNAs (assuming that separate are used) is needed initiator and elongator to translate the mitochondrial genetic information (Boer and Gray, 1988c). As only three mitochondrial

Claire Remacle and René F. Matagne tRNA genes have been detected (cf above), most of the tRNAs used in the organelle have to be imported, probably from the cytosol, as is the case for some tRNAs in higher plants (Maréchal-Drouard et al., 1993) and trypanosomes (Schneider, 1994). Compilation of the eight protein-coding gene sequences allowed the definition of a putative ribosome binding site (Colleaux et al., 1990). A consensus sequence (5´ AAAUUUAU 3´) complementary to the terminal sequence of the small subunit rRNA and located three to seven nucleotides upstream of the ATG initiation codon was identified.

III. Mitochondria and the Electron Transport Chain During the vegetative cell cycle of C. reinhardtii, mitochondria can undergo dramatic changes in number and shape, the number of organelle per cell varying from 50 small mitochondria to one giant, globular or highly branched mitochondrion (Blank et al., 1980; Ehara et al., 1995). The chondriome (i.e. the mitochondria of a cell collectively) can be viewed as a dynamic and flexible system which can probably fuse and dissociate in the cell. During zygote maturation, the volume of the chondriome decreases to about half the volume found in young zygotes and a few degenerated mitochondria are observed in every mature zygote (Brand and Arnold, 1987). This loss of mitochondrial mass could be correlated with the uniparental inheritance of the organelle genome (see below). As in higher plants and various fungi, the mitochondrial electron transport chain of C. rein­ hardtii is composed of two distinct pathways branching at the level ofthe ubiquinone pool (Fig. 2): the classical cyanide-sensitive pathway (or cytochrome pathway) and the alternative pathway (corresponding to the alternative oxidase), insensitive to cyanide but sensitive to salicylhydroxamic acid (SHAM) (Wiseman et al., 1977; Goyal and Tolbert, 1989; Matagne et al., 1989). The alternative oxidase of C. reinhardtii has been detected recently using a monoclonal antibody to the alternative oxidase from voodoo lily (Derzaph and Weger, 1996). Complex I, complex III, complex IV, the mitochondrial ATP synthase as well as a fraction displaying both NADH and NADPH:ubiquinone oxidoreductase activityhave been identified in C. reinhardtii membrane prepar-

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Mitochondrial Genetics

ations (Atteia, 1994). This NAD(P)H dehydrogenase could correspond to one of the two rotenoneinsensitive NAD(P)H dehydrogenases identified in the inner membrane ofplant mitochondria (reviewed by Siedow, 1995).

IV. Mutations Affecting the Mitochondrial Genome

A. Lethal Minute Colony Mutants When C. reinhardtii is grown heterotrophically (darkness + acetate) for several days in a medium containing the intercalating dyes acriflavine or ethidium bromide, the majority of the cell population is converted into mutant cells which display a delayed lethality: plated onto acetate agar medium and maintained in the dark, the mutant cells do not divide, whereas in the light they form minute colonies and growth ceases after 8 – 9 mitotic divisions (Alexander et al., 1974; Gillham et al., 1987; Matagne et al., 1989). The lethal minute phenotype is associated with marked defects in structure and function of mitochondria and to specific loss of mitochondrial DNA (Gillham et al., 1987). In the yeast Saccharomyces cerevisiae, mutants deprived of mitochondrial DNA are conditional lethals which survive when grown on a fermentable carbon source. In C. reinhardtii, the lethality probably results from the loss of some essential mitochondrial functions. Inactivation ofmitochondrial genes coding for subunits ofcomplex I, III or IV does not, however, prevent the survival of the cells (see below).

665

B. Obligate Photoautotrophic RespiratoryDeficient Mutants

In addition to lethal minute colony mutations, acriflavine and ethidium bromide induce a low percentage of viable mutants that grow in the light but do not divide under heterotrophic conditions (Matagne et al., 1989; Dorthu et al., 1992; Colin et al., 1995). Most of these obligate photoautotrophic mutants lack the cyanide-sensitive cytochrome pathway of respiration whereas the SHAMsensitive alternative pathway is still functional. Under mixotrophic conditions (light + acetate), their total consumed respiratory rate (5–10 nmoles cells) is 20–50% of that of wild-type cells, per and their growth rate is reduced. Under photoautotrophic conditions (light without acetate), their growth relative to wild-type is scarcely different and their respiratory rate is similar to that of the wildper cells). type (ca. 6 nmoles consumed The mutant colonies deprived of the cytochrome pathway ofrespiration can also be identified by an in vivo staining test, based on the reduction of 2,3,5triphenyltetrazolium chloride (TTC) to red formazan (Dorthu et al., 1992). The mitochondrial origin of the mutations has been demonstrated by genetic analysis: in crosses between wild-type and mutant cells, the meiotic progeny most often inherit the phenotype of the mating-type minus ( or paternal) parent, a transmission pattern which is typical of the mitochondrial genome. The mitochondrial mutations were named dum, for dark uniparental transmission by the minus parent (Matagne et al., 1989). Phenotypically, the dum mutants are identical to the

666 dark-dier nuclear mutants deprived of cytochrome c oxidase activity (Wiseman et al., 1977; Dorthu et al., 1992). Thirteen respiratory-deficient dum mutants have been partially or completely characterized at the biochemical and molecular levels (Table 1). Seven mutants (dum1, 2, 3, 4, 11, 14 and 16) have a terminal or subterminal deletion of the left end of the mitochondrial genome. The cob gene is partially or totally deleted and therefore, complex III activity is missing. Whereas wild-type cells contain a single type of mitochondrial genome, two or three types of mitochondrial DNA molecules are present in these mutants: deleted monomers and one or two types of dimers resulting from head-to-head fusions of monomers, with identical or different deletions (Dorthu et al., 1992; Randolph-Anderson et al., 1993; Colin et al., 1995). The absence of the left terminal inverted repeat region in the mutants could account for dimer formation. The dimer genomes may be more stable than the monomers because they have normal terminal inverted repeat structures at each end, derived from the right end of the monomer genome (Randolph-Anderson et al., 1993). The dimers would result from illegitimate recombinational events, by a mechanism which involves the joining of monomers at precisely defined sites (Dorthu et al., 1992). The seven deletion mutants also have in common the property of mitotically segregating two types of

Claire Remacle and René F. Matagne cells: viable cells (90–98%) and cells (2–10%) which divide eight to nine times under light to produce depigmented lethal minute colonies. These minute colonies are phenotypically identical to those produced after acriflavine or ethidium bromide treatment and deprived of mitochondrial DNA. The dum1 mutant has been studied in more detail (Randolph-Anderson et al., 1993). The mitochondrial genome of dum1 is unstable, with the extent of the deletion varying from 1.5 to 1.7 kb among single cell clones. PCR amplifications performed on DNA of minute colonies have shown that the deletion can extend to the nd4 gene. However, these results do not rule out the possibility that some molecules may have larger deletions or that part ofthe minute colonies may have lost all their mitochondrial DNA. Loss of complex III activity in the dum15 mutant results from a double base-pair substitution at codon 140 of the cob sequence, changing TCT (Ser) into TAC (Tyr) (Colin et al., 1995). This amino acid substitution occurs in the segment GQMSFWGAT (137–145) of the polypeptide which is highly conserved in bacterial and eukaryotic apocytochrome b proteins (di Rago et al., 1989). According to the eight transmembrane helix model for cytochrome b (Brasseur, 1988; di Rago and Colson, 1988; Colson, 1993), this segment is part of the protruding loop between helices III and IV and would be involved in electron transfer at the

Chapter 34

Mitochondrial Genetics

ubiquinone redox site located at the outer (positive) side of the mitochondrial inner membrane (di Rago et al., 1989). Despite the presence of a double bp substitution, the dum15 mutation reverts to restore the wild-type (reversion rate: phenotype. In two revertants analyzed (M. Dinant and C. Remacle, unpublished), the original cytochrome b sequence was restored by a single bp transversion replacing the mutant TAC codon by TCC, synonymous with TCT (Ser). The dum18 and dum19 mutants have both lost cytochrome c oxidase (complex IV) activity, as a consequence of a frameshift mutation in the cox1 sequence. Addition ofone T at codon 145 and deletion of one T at codon 152 have been identified in dum18 and dum19 respectively. Less than one third of the correct polypeptide sequence is thus present in either mutant strain. The leaky dum6 and dum20 mitochondrial mutants display reduced growth under heterotrophic conditions. The dum6 mutant was originally thought to bepartially deficient in complex IV activity (Dorthu et al., 1992), and thus to be mutated in cox1. However, other biochemical analyses have failed to confirm a significant defect in cytochrome c oxidase activity (Remacle et al., 1995) and no modification of the cox1 DNA sequence has been detected (M. Dinant and R. F. Matagne, unpublished). Therefore the position of the dum6 mutation site remains uncertain. The dum20 mutant was shown to lack the rotenonesensitive NADH:ubiquinone oxidoreductase (complex I) activity (M. Colin, unpublished), probably as a consequence of a mutation affecting one of the NADH dehydrogenase subunits encoded in the mitochondrial genome. A rotenone-insensitive NADH:ubiquinone oxidoreductase activity was detected in purified mitochondria from dum20 and wild-type cells, confirming the presence of this enzyme in C. reinhardtii mitochondria (see above). The newly isolated dum24 mutant does not grow in the dark and, in contrast to other mutants, produces small-sized colonies under photoautotrophic conditions (F. Duby and R. F. Matagne, unpublished). Southern blot analyses of the mitochondrial DNA have revealed a large deletion encompassing the cob and nd4 genes. The absence of both gene sequences was confirmed by PCR amplification. Thus, the simultaneous loss of cytochrome b and ND4 subunits is not lethal to the cell, contrary to the hypothesis formulated by Randolph-Anderson et al. (1993). In fungi, animals and higher plants, complex I is the

667 largest of the respiratory chain complexes and comprises more than 30 to 40 different subunits. In Neurospora crassa and probably other organisms, complex I has an L shape with a hydrophilic domain protruding into the matrix aqueous phase and a hydrophobic domain localized within the inner membrane and containing all the mitochondriaencoded subunits (for recent reviews, see Siedow, 1995; Whitehouse and Moore, 1995). Although little is known about the functions of most of the mitochondrial DNA-encoded subunits of complex I, the ND4 subunit appears to be important for the assembly of the hydrophobic arm. A mutant human cell line lacking the ND4 subunit fails to assemble the other mitochondrial-encoded subunits and lacks complex I activity (Hofhaus and Attardi, 1993). The absence of this enzyme activity does not, however, prevent the growth of the cell line.

C. Myxothiazol-Resistant Mutants Myxothiazol and mucidin (strobilurin A) are complex III inhibitors preventing electron transfer at the ubiquinone redox site located close to the outer side of the inner mitochondrial membrane (von Jagow and Link, 1986). In S. cerevisiae (di Rago et al., 1989) and in the photosynthetic bacterium Rhodobacter capsulatus (Daldal et al., 1989), mitochondrial mutants resistant to myxothiazol or mucidin have been mapped to specific loci in the cob gene. Spontaneous mutants displaying heterotrophic growth in the presence of myxothiazol or mucidin can be isolated easily from haploid cells of C. reinhardtii. The mutations, of nuclear origin, are thought to reduce drug permeability (Bennoun et al., 1991). Mitochondrial mutants resistant to these inhibitors have been selected as diploid colonies growing in the dark on acetate containing medium or supplemented with mucidin and mucidin myxothiazol (Bennoun et al., 1991, 1992). Mutants with same phenotype have also been recovered in our laboratory following a similar strategy (M. Colin and R. F. Matagne, unpublished). Bennoun et al. (1992) have reported that resistant mutants occur spontaneously and that after at a rate of approximately manganese treatment, the frequency increases to approximately In our experiments, mucidinresistant strains appear spontaneously at a rate of and manganese treatment does not modify

668 this frequency. It remains an open question whether manganese, which is mutagenic in yeast (Putrament et al., 1973), increases the mitochondrial mutation rate in C. reinhardtii. Sixteen mutants resistant to myxothiazol or mucidin have been characterized physically (Table 2). All mutations correspond to amino acid substitutions and are clustered in a short region of the apocytochrome b polypeptide. (MUD2 according to The mutation Bennoun’s nomenclature) is the most frequent. Mutations at positions 129 and 132 lie in a hydrophobic region of helix III while the mutation at position 137 lies in the extramembrane loop, between helix III and helix IV. Mutations at positions 129 and 137 have also been detected in yeast and/or in Rhodobacter (Daldal et al., 1989; di Rago et al., 1989). The mutated amino acid residues are probably involved in the binding of myxothiazol which interferes with the ubiquinone (Colson et al., 1990). redox site

V. Transmission of Mitochondrial Genes in Meiotic Zygotes In contrast to the chloroplast genome which is preferentially transmitted to the meiotic progeny from the mating-type plus or maternal parent, the mitochondrial DNA is most often inherited from the mating-type minus or paternal parent. This was first demonstrated in reciprocal crosses between C. reinhardtii and C. smithii by using the RFLP differentiating the two species (Boynton et al., 1987; Matagne et al., 1988). Point mutations affecting the mitochondrial genes follow the same rule. The percentage of meiotic products inheriting the mitochondrial marker of origin is most often null, or lower than 3% (Bennoun et al., 1991; Dorthu

Claire Remacle and René F. Matagne et al., 1992; Colin et al., 1995). Modifications of the organelle genome can, however, influence the mode of mitochondrial DNA transmission. In crosses of the wild-type with a mutant deleted in the cob gene, the transmission is almost strictly paternal when the whereas in the reciprocal wild-type parent is x mutant maternal cross (wild-type transmission occurs much more frequently (16–28% of the meiotic products inherit the wild-type genome origin; Matagne et al., 1989; Dorthu et al., of 1992). Thus, the uniparental paternal transmission of the mitochondrial genome can be strongly biased parent contains altered mitochondrial when the DNA molecules. Maternal chloroplast inheritance results from the within elimination of chloroplast DNA of the first hours following gametic fusion (Kuroiwa et al., 1982; Munaut et al., 1990). In contrast, and origins mitochondrial DNAs of both persist during the 24 hours which follow zygote formation (Beckers et al., 1991). In the course of zygote maturation in the dark, the amount of mitochondrial DNA progressively decreases whereas is retained. In the mature zygospores, the origin persist traces of mitochondrial DNA of even after several weeks but when the zygospores are DNA is transferred to light to induce meiosis, the no longer detected 24 hours after the transfer. When the zygotes are matured under continuous light on nitrogen-free agar plates, the maternal mitochondrial DNA is eliminated after five days. As the total mitochondrial mass diminishes during the maturation of the zygote (Brand and Arnold, 1987), it has been suggested that the uniparental transmission of the mitochondrial genome could result from the origin during the elimination of mitochondria of maturation process (Beckers et al., 1991). Sexual functions, including those responsible for the uniparental transmission of the organelle

Chapter 34

Mitochondrial Genetics

genomes, are under the control of the complex and (Chapter 6, Ambrust). Diploid gametes, which behave as minus in sexual agglutination, often transmit their chloroplast genes in or diploid gametes crosses with haploid is recessive (Matagne and Mathieu, 1983). Thus with respect to sex expression, but dominant to in the control of chloroplast gene transmission. Moreover, the ploidy of the gametes also influences the pattern of chloroplast gene transmission (Matagne and Mathieu, 1983). With respect to mitochondrial is dominant to and nuclear DNA transmission, ploidy has little, if any, effect on the transmission pattern in crosses of diploids with haploids (Matagne et al., 1988).

VI. Transmission of Mitochondrial Genes in Vegetative Zygotes and Mapping of Mitochondrial Mutations by Recombinational Analysis

A. Inheritance of Mitochondrial Genes in Vegetative Zygotes In C. reinhardtii, 1–10% of zygotes fail to undergo meiosis and divide mitotically to give rise to stable diploid progeny (Ebersold, 1967). Such zygotes are called mitotic or vegetative zygotes. The transmission of the mitochondrial genome in vegetative zygotes was first investigated by Boynton et al. (1987) by performing crosses between C. reinhardtii and C. smithii. The mitochondrial genomes of these two

669 interfertile algae are colinear with the exception of the intron located in the cob gene of C. smithii (see above). The C. smithii strain also possesses additional NheI, NcoI and HpaI restriction sites, located in cob, nd4 and cox1 genes, respectively (Fig. 3). A strain possessing an recombinant mitochondrial genome has also been isolated and used in crosses (Remacle and Matagne, 1993). The transmission pattern ofmitochondrial markers in crosses between strains which both possess or both lack the intron is different from that observed In crosses of the type in crosses of transmission is biparental with a strong bias for RFLP markers inherited from parent. Recombinant genomes are also the produced, providing evidence that mitochondria fuse in the vegetative zygotes. When a number of vegetative zygotes are heteroplasmic, the various types of mitochondrial DNA copies segregate during the successive mitotic divisions and after about 15 divisions, most cells are homoplasmic for all the markers (Remacle et al., 1990; Remacle and Matagne, 1993). A biparental transmission with a bias for markers (MUD2 or wild-type alleles) from the parent is also observed in crosses between strains (Bennoun et al., 1991; Remacle and Matagne, 1993). The mode of transmission is different in crosses of the type Almost all the diploid cells possess intron-containing molecules (especially when the but some of them inherit the paternal parent is RFLP markers from the intron-free strain (Boynton et al., 1987; Remacle et al., 1990; Remacle and Matagne, 1993). The conversion of into

670 molecules is frequently accompanied (94% of cases) by the co-conversion of the MUD2 marker located 27 bp from the insertion site of the intron. The conversion also influences the transmission of the more distant Nh, Nc and H markers in such a way that parent is the biased in crosses where the transmission of the markers inherited from the parent is no longer observed (Remacle and Matagne, 1993). The conversion ofintron-less molecules into introncontaining molecules and the co-conversion of the MUD2 marker is reminiscent of the transposition of intron of yeast, and other group I introns the (Dujon, 1989; Belfort, 1990). In several microorganisms and phages, and in the C. reinhardtii chloroplast, it has been shown that the endonuclease encoded by the ORF of the mobile group I intron recognizes a very long sequence (18–24 nucleotides) in the intron-free gene and induces a double-strand break at or near the insertion site of the intron. A copy of the intron is then inserted at the break, probably by a double-strand break repair mechanism. The intron conversion is associated with coconversion of flanking DNA sequences over a few hundred bp both upstream and downstream of the intron, the efficiency of co-conversion decreasing regularly with distance (Jacquier and Dujon, 1985; Zinn and Butow, 1985). In the present case, the unidirectional spreading of the intron to the intron-free molecules is probably related to the action of the endonuclease encoded by the intronic ORF (Ma et al., 1992b), but the exact sequence recognized by the endonuclease remains to be determined. More surprisingly, the conversion of molecules also influences the intron-free into transmission of the RFLP markers located at much longer distances (up to 5 kb) from the insertion site. This could result from a preferential transmission of the copies due to the elimination of a fraction of genomes, or to a cothe temporarily cleaved conversion extending for distances as long as several kilobases. Recently, Bussieres et al. (1996) have also reported that in crosses of C. eugametos with C. moewusii, co-transmission of chloroplast markers associated with a mobile intron takes place over distances longer than 3 kb. The transmission of mitochondrial markers has been analyzed also in diploids obtained by artificiallyinduced fusion between vegetative cells of the same or opposite mating-types (Remacle et al., 1990; Remacle and Matagne, 1993). As in vegetative

Claire Remacle and René F. Matagne zygotes obtained by sexual mating, recombination of mitochondrial markers and polarized transmission of the intron occur in the fusion products but the mating-type of the parental cells no longer influences the transmission pattern. It thus appears that the mating-type locus controls to variable degrees the transmission pattern of the mitochondrial genome. In meiotic zygotes, some gene functions determined by the MT locus trigger the differentiation of the diploid cell into a zygospore, but also the elimination of the mitochondrial DNA of origin. These functions are weakly expressed in vegetative zygotes, preventing zygospore formation and strongly reducing the elimination of the mitochondrial genome. The mating-type related functions are totally suppressed in the diploid cells produced by artificial fusion between vegetative cells. This mechanism controlling mitochondrial gene transmission fully parallels that responsible for chloroplast DNA transmission in meiotic zygotes, vegetative zygotes and artificial fusion products (Matagne, 1981; Chapter 6, Ambrust).

B. Mapping Mutations by Recombinational Analysis As mentioned above, recombination between mitochondrial RFLP markers occurs in vegetative diploids, but too few clones have been analyzed to determine with accuracy the frequency of these recombinational events. The mitochondrial dum mutants are good candidates for studying recombination in a large population of diploid colonies. In crosses between strains mutated in cob (dum11, dum15) and strains mutated in cox1 (dum18, dum19), 66–100% of the vegetative zygotes segregate respiratory-competent cells capable of growth in the cells dark (Remacle et al., 1995). As these represent half the recombinants (the double mutants not being detected), this indicates that the majority of vegetative zygotes transmit the mitochondrial genomes of both parents and that intermolecular recombinational events are frequent in these zygotes. The recombination frequencies have been calculated from the whole population of diploid progeny cells, when all of them have become homoplasmic (Remacle et al., 1995). In crosses between strains mutated in the cob gene and strains mutated in the cox1 gene, the frequency of recombination is 13.7% (± 3.2%) (Fig. 4). The corresponding physical distance between the

Chapter 34

Mitochondrial Genetics

mutations is 4.3 kb. One can thus estimate that the genetic distance for mitochondrial markers corresponds to 3.2% (± 0.7%) per kb. In crosses between strains carrying mutations separated by ca. 20 bp (dum18 × dum19, dum11 × dum15), a recombination frequency of 0.04% (± 0.02%) is found (Fig. 4). In that case, the genetic distance is 2% (± 1%) per kb, which is a value comparable to the one obtained for markers separated by 4.3 kb. The dum6 and dum20 mutants, although not yet characterized at the molecular level, have also been used for recombinational studies. Based on their positions on the linear genetic map (Fig. 4) and on the relation between recombination frequency and physical distance, these two mutations are predicted to be located in one of the nd genes (nd2 or nd6) to the right of cox1 (Fig. 1). This conclusion is in agreement with the observation that dum20 has lost complex I activity (Table 1). The recombination frequency per kb obtained for mitochondrial markers in C. reinhardtii is higher than the basal 1% per kb observed for chloroplast markers in the same organism (Harris et al., 1989). It is also somewhat higher than the mitochondrial recombinational frequency found in Schizosaccharo­ myces pombe (1.6% per kb, deduced from the data of Lückermann et al., 1979; Lang and Wolf, 1984; Weber and Wolf, 1988). In contrast, these frequencies are much lower than the values obtained in S. cerevisiae for mitochondrial markers (15–25% per kb, which corresponds to the maximal recombination rate for a multicopy organelle genome). Recombinational analysis thus constitutes an

671

excellent tool for mapping mitochondrial mutations that are less than 5–7 kb apart in C. reinhardtii.

VII. Mitochondrial Transformation Genetic transformation is the only way to evaluate in vivo the consequences of DNA modifications made in vitro, and to analyze the phenotypic changes caused by the presence of novel genes in a genome. Mitochondrial transformation was achieved in C. reinhardtii by Randolph-Anderson et al. in 1993. Using the biolistic method, these workers have reported successful transformation of the dum1 deletion mutant to respiratory competence with partially purified mitochondrial DNA from C. reinhardtii or C. smithii. Transformants obtained with C. reinhardtii mitochondrial DNA were homoplasmic for the normal genome typical of this species. Homologous recombination rather than genome replacement was documented in the case of transformation with C. smithii mitochondrial DNA that has distinct RFLP markers. The transformation frequency was very low cell treated) compared to the frequency of chloroplast Boynton and Gillham, 1993) transformation Kindle, 1990) in or nuclear transformation the same organism. The rather low efficiency of mitochondrial transformation could be explained by the high number of mitochondria at certain stages of the cell cycle (see above) and their small size relative to the size of the microprojectiles. Moreover, to

672 obtain a homoplasmic mitochondrial transformant requires segregation of the transformed organelle as well as segregation of transformed genomes within that organelle. Reducing the copy number of mitochondrial genomes in order to favor the sorting out of the recombinant molecules might increase the frequency of transformation as has been done successfully for chloroplast transformation (Boynton and Gillham, 1993). Cotransformation with chloroplast or nuclear genes and initial selection for these markers might also facilitate the recovery of mitochondrial transformants. In yeast, most mitochondrial transformation experiments involve cotransformation with nuclear genes (Butow et al., 1996). Finally, the use of recipient strains other than dum1, and the use of drug resistance genes (for example the MUD2 mutation in the cob gene) as donor DNA could improve the efficiency of transformation.

Acknowledgments Work carried out in this laboratory was supported by grants from the Belgian FRFC (2.4520.93), Actions de recherches concertées (ARC 93-98/170) and Fonds spéciaux pour la recherche dans les Universités. We thank Dr. David Stern for critical reading of the manuscript.

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Claire Remacle and René F. Matagne Planta 150: 236–241 Boer PH and Gray MG (1986a) The URF 5 gene of Chlamydomonas reinhardtii mitochondria: DNA sequence and mode of transcription. EMBO J 5: 21–28 Boer PH and Gray MW (1986b) Nucleotide sequence of a protein coding region in Chlamydomonas reinhardtii mitochondrial DNA. Nucleic Acids Res 14: 7506–7507 Boer PH and Gray MW (1988a) Genes encoding a subunit of respiratory NADH dehydrogenase (ND1) and a reverse transcriptase-like protein (RTL) are linked to ribosomal RNA gene pieces in Chlamydomonas reinhardtii mitochondrial DNA. EMBO J 7: 3501–3508 Boer PH and Gray MW (1988b) Scrambled ribosomal RNA gene pieces in Chlamydomonas reinhardtii mitochondrial DNA. Cell 55: 399–411 Boer PH and Gray MW (1988c) Transfer RNA genes and the genetic code in Chlamydomonas reinhardtii mitochondria. Curr Genet 14: 583–590 Boer PH and Gray MW (1989) Nucleotide sequence of a region encoding subunit 6 of NADH dehydrogenase (ND6) and tRNATrp in Chlamydomonas reinhardtii mitochondrial DNA. Nucleic Acids Res 17: 3993 Boer PH and Gray MW (1991) Short dispersed repeats localized in spacer regions of Chlamydomonas reinhardtii mitochondrial DNA. Curr Genet 19: 309–312 Boer PH, Bonen L, Lee RW and Gray MG (1985) Genes for respiratory chain proteins and ribosomal RNAs are present on a 16-kilobase-pair DNA species from Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 82: 3340–3344 Boynton JE and Gillham NW (1993) Chloroplast transformation in Chlamydomonas. Methods Enzymol 217: 510–536 Boynton JE and Gillham NW(1996) Genetics and transformation of mitochondria in the green alga Chlamydomonas. Methods Enzymol 264: 279–296 Boynton JE, Harris EH, Burkhart BD, Lamerson PM and Gillham NW (1987) Transmission of mitochondrial and chloroplast genomes in crosses of Chlamydomonas. Proc Natl Acad Sci USA 84: 2392–2395 Brand N and Arnold CG (1987) Chondriome characteristics in mature zygotes of Chlamydomonas reinhardii. Endocyt C Res 4: 275–284 Brasseur R (1988) Calculation of the three-dimensional structure of Saccharomyces cerevisiae cytochrome b inserted in a lipid matrix. J Biol Chem 263: 12571–12575 Bussieres J, Lemieux C, Lee RW and Turmel M (1996) Optional elements in the chloroplasts DNAs of Chlamydomonas eugametos and C. moewusii: Unidirectional gene conversion and co-conversion of adjacent markers in high-viability crosses. Curr Genet 30: 356–365 Butow RA, Henke RM, Moran JV, Belcher SM and Perlman PS (1996) Transformation of Saccharomyces cerevisiae mitochondria using the biolistic gun. Methods Enzymol 264: 265–278 Colin M, Dorthu MP, Duby F, Remacle C, Dinant M, Wolwertz MR, Duyckaerts C, Sluse F and Matagne RF (1995) Mutations affecting the mitochondrial genes encoding the cytochrome oxidase subunit I and the apocytochrome b of Chlamydomonas reinhardtii. Mol Gen Genet 249: 179–184 Colleaux L, Michel-Wolwertz MR, Matagne RF and Dujon B (1990) The apocytochrome b gene of Chlamydomonas smithii contains a mobile intron related to both Saccharomyces and

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673 Hofhaus G and Attardi G (1993) Lack of assembly of mitochondrial DNA-encoded submits of respiratory NADH dehydrogenase and loss of enzyme activity in a human cell mutant lacking the mitochondrial ND4 gene. EMBO J 12: 3043–3048 Jacquier A and Dujon B (1985) An intron-encoded protein is active in a gene conversion process that spreads an intron into a mitochondrial gene. Cell 41: 383–394 Kindle KA (1990) High-frequency nuclear transformation of Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 87: 1228–1232 Kück U and Neuhaus H (1986) Universal genetic code evidenced in mitochondria of Chlamydomonas reinhardii. Appl Microbiol Biotechnol 23: 462–469 Kuroiwa T, Kawano S, Nishibayashi S and Sato C (1982) Epifluorescent microscopic evidence for maternal inheritance of chloroplast DNA. Nature 298: 481–483 Lang BF and Wolf K (1984) The mitochondrial genome of the fission yeast Schizosaccharomyces pombe. 2. Location of genes by interspecific hybridization in strain and cloning of the genome in small fragments. Mol Gen Genet 196: 465–472 Lückermann G, Sipiczki M and Wolf K (1979) Transmission, segregation and recombination of mitochondrial genomes in zygotes clones and protoplast fusion clones of yeast. Mol Gen Genet 177: 185–187 Ma DP, Yang YW and Hasnain SE (1988) Nucleotide sequence of Chlamydomonas reinhardtii mitochondrial genes coding Nucleic for subunit 6 of NADH dehydrogenase and Acids Res 16: 11373 Ma DP, Yang YW and Hasnain SE (1989a) Nucleotide sequence of Chlamydomonas reinhardtii mitochondrial genes coding Nucleic Acids Res for and 17: 1256 Ma DP, Yang YW, King YT and Hasnain SE (1989b) Nucleotide sequence of cloned nad4 (urf4) gene from Chlamydomonas reinhardtii mitochondrial DNA. Gene 85: 363–370 Ma DP, Yang YW, King TY and Hasnain SE (1990) The mitochondrial apocytochrome b gene from Chlamydomonas reinhardtii. Plant Mol Biol 15: 357–359 Ma DP, King YT, Kim Y, Luckett WS Jr, Boyle JA and Chang YF (1992a) Amplification and characterization of an inverted repeat from Chlamydomonas reinhardtii mitochondrial genome. Gene 119: 253–257 Ma DP, King YT, Kim Y and Luckett WS Jr (1992b) The group I intron of the apocytochrome b gene from Chlamydomonas smithii encodes a site-specific endonuclease. Plant Mol Biol 18: 395–402 Maréchal-Drouard L, Weil JH and Dietrich A (1993) Transfer RNAs and transfer RNA genes in plants. Annu Rev Plant Physiol Plant Mol Biol 44: 13–32 Matagne RF (1981) Transmission of chloroplast alleles in somatic fusion products obtained from vegetative cells and/or ‘gametes’ of Chlamydomonas reinhardi. Curr Genet 3: 31–36 Matagne RF and Mathieu D (1983) Transmission of chloroplast genes in triploid and tetraploid zygospores of Chlamydomonas reinhardi: Roles of the mating-type, gene dosage and gametic chloroplast DNA content. Proc Natl Acad Sci USA 80: 4780– 4783 Matagne RF, Rongvaux D and Loppes R (1988) Transmission of mitochondrial DNA in crosses involving diploid gametes

674 homozygous or heterozygous for the mating-type in Chlamydomonas. Mol Gen Genet 214: 257–262 Matagne RF, Michel-Wolwertz MR, Munaut C, Duyckaerts C and Sluse F (1989) Induction and characterization of mitochondrial DNA mutants in Chlamydomonas reinhardtii. J Cell Biol 108: 1221–1226 Michaelis G, Vahrenholz C and Pratje E (1990) Mitochondrial DNA of Chlamydomonas reinhardtii: The gene for apocytochromc b and the complete functional map of the 15.8 kb DNA. Mol Gen Genet 223: 211–216 Munaut C, Dombrowicz D and Matagne RF (1990) Detection of chloroplast DNA by using fluorescent monoclonal antibromodeoxyuridine antibody and analysis of its fate during zygote formation in Chlamydomonas reinhardtii. Curr Genet 18: 259–263 Newton KJ (1995) Aberrant growth phenotypes associated with mitochondrial genome rearrangements in higher plants. In: Levings CS and Vasil IK (eds) The Molecular Biology of Plant Mitochondria, pp 585–596. Kluwer Academic Press, Dordrecht Pla M, Mathieu C, Paepe RD, Chétrit P and Vedel F (1995) Deletion of the last two exons of the mitochondrial nad7 gene results in lack of the NAD7 polypeptide in Nicotiana sylvestris CMS mutant. Mol Gen Genet 248: 79–88 Pratje E, Schnierer S and Dujon B (1984) Mitochondrial DNA of Chlamydomonas reinhardtii: The DNA sequence of a region showing homology with mammalian URF2. Curr Genet 9: 75– 82 Pratje E, Vahrenholz C, Buhler S and Michaelis G (1989) Mitochondrial DNA of Chlamydomonas reinhardtii: The ND4 gene encoding a subunit of NADH dehydrogenase. Curr Genet 16: 61–64 Putrament A, Baranowska H and Prazmo W (1973) Induction by manganese of mitochondrial antibiotic resistant mutations in yeast. Mol Gen Genet 126: 357–366 Randolph-Anderson BL, Boynton JE, Gillham NW, Harris EH, Johnson AM, Dorthu M-P and Matagne RF (1993) Further characterization of the respiratory deficient dum-1 mutation of Chlamydomonas reinhardtii and its use as a recipient for mitochondrial transformation. Mol Gen Genet 236: 235–244 Remacle C and Matagne RF (1993) Transmission, recombination and conversion of mitochondrial markers in relation to the mobility of a group I intron in Chlamydomonas. Curr Genet 23: 518–525

Claire Remacle and René F. Matagne Remacle C, Bovie C, Michel-Wolwertz M-R, Loppes R and Matagne RF (1990) Mitochondrial genome transmission in Chlamydomonas diploids obtained by sexual crosses or artificial fusions: role of the mating-type and of a one kb intron. Mol Gen Genet 223: 180–184 Remacle C, Colin M and Matagne RF (1995) Genetic mapping of mitochondrial markers by recombinational analysis in Chlamydomonas reinhardtii. Mol Gen Genet 249: 185–190 Ryan R, Grant D, Chiang KS and Swift H (1978) Isolation and characterization of mitochondrial DNA from Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 75: 3268–3272 Schneider A (1994) Import of RNA into mitochondria. Trends in Cell Biol 4: 282–286 Siedow JN (1995) Bioenergetics: The mitochondrial electron transport chain. In: Levings CS and Vasil IK (eds) The Molecular Biology of Plant Mitochondria, pp 281–312. Kluwer Academic Press, Dordrecht Vahrenholz C, Pratje E, Michaelis G and Dujon B (1985) Mitochondrial DNA of Chlamydomonas reinhardtii: Sequence and arrangement of URF5 and the gene for cytochrome oxidase subunit I. Mol Gen Genet 201: 213–224 Vahrenholz C, Riemen G, Pratje E, Dujon B and Michaelis G (1993) Mitochondrial DNA of Chlamydomonas reinhardtii: The structure of the ends ofthe linear 15.8-kb genome suggests mechanisms for DNA replication. Curr Genet 24: 241–247 von Jagow G, Link TA (1986) Use of specific inhibitors on complex. Methods Enzymol 126: 253–271 mitochondrial Weber S and Wolf K (1988) Two changes of the same nucleotide confer resistance to diuron and antimycin in the mitochondrial cytochrome b gene of Schizosaccharomyces pombe. FEBS Lett 237: 31–34 Whitehouse DG and Moore AL (1995) Regulation of oxidative phosphorylation in plant mitochondria. In: Levings CS and Vasil IK (eds) The Molecular Biology of Plant Mitochondria, pp 313–344. Kluwer Academic Publishers, Dordrecht Wiseman A, Gillham NW and Boynton JE (1977) Nuclear mutations affecting mitochondrial structure and function in Chlamydomonas. J Cell Biol 73: 56–77 Zinn A and Butow R (1985) Non reciprocal exchange between alleles of the yeast mitochondrial 21 S rRNA gene: kinetics and the involvement of a double stand-break. Cell 40: 887–895

Chapter 35 Chlororespiration, Sixteen Years Later Pierre Bennoun

Institut de Biologie Physico-Chimique, Centre National de la Recherche Scientifique,

UPR 9072, 13 rue Pierre et Marie Curie, 75005 Paris, France

Summary I. Introduction II. The Thylakoid Electrochemical Gradient Present in the Dark A. The Basis of the Model of ‘Chlororespiration’ B. The Need for a New ATP Driven Gradient Generator III. Reduction of Plastoquinone in the Dark A. The Paradoxical Status of Chlamydomonas B. Mitochondrial Control of the Redox State of Plastoquinone IV. Oxidation of Plastoquinol in the Dark A. The Ambiguity Raised by Mitochondrial-Chloroplast Interactions B. Plastoquinol Oxidation via a Mitochondrial Oxidase V. Conclusion Cautionary Note Acknowledgments References

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Summary The model of chlororespiration was proposed by Bennoun (1982) in order to take into account experimental data concerning the state of thylakoid membranes in darkness. This model postulated the presence, in the thylakoid membranes, of an electrogenic respiratory electron transport chain. As observed in cyanobacteria, plastoquinone would be reduced by an NAD(P)H dehydrogenase, and reoxidized at the expense of oxygen by an oxidase. Such a pathway would explain both the formation of a permanent thylakoid electrochemical gradient in the absence of ATP synthase, and the redox properties of plastoquinone observed in the dark. This model which stimulated many experiments must now be revised in the light of recent work. Particularly, the occurrence of mitochondrial-chloroplast interactions has to be taken into account: the mitochondrial electron transport chain controls strictly both the redox state of plastoquinone and the permanent electrochemical gradient. The possibility that a respiratory process generates the permanent electrochemical gradient is no longer consistent with the available data. This gradient rather results from the operation of a new ATPdependent electrogenic pump. With regard to the existence of a chloroplast NAD(P)H dehydrogenase, clear-cut physiological evidence exists for an electron pathway connecting stromal reductants to plastoquinone in Chlamydomonas reinhardtii, whereas molecular evidence for such a chloroplast enzyme is lacking. However, the presence of a proton pumping enzyme closely related to the bacterial Complex I has been demonstrated in higher plants, which shows a higher specificity for NADH than for NADPH. Finally, there is at present no molecular evidence for the existence of a chloroplast oxidase. Given the existence of chloroplast-mitochondrial

J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 675–683. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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interactions, plastoquinol oxidation might alternatively occur through the mitochondrial electron transport chain, via a mobile electron carrier. Studies on the state of thylakoid membranes in darkness will have to evaluate the structure and function of the new ATP-dependent electrogenic pump present in the thylakoid membrane, to analyze thoroughly the bioenergetic interactions between chloroplast and mitochondria, and to elucidate the mode of oxidation of plastoquinone in the dark. Alternative models to that of chlororespiration are discussed in order to stimulate further experiments. Clearly, the state of the thylakoid membranes in darkness offers more than ever a vast field of investigations.

I. Introduction In the presence of ATP, an electrochemical gradient is observed in darkness across thylakoid membranes, presumably due to the functioning of the ATP synthase (Joliot and Joliot, 1980). The discovery that this permanent gradient was still present in the ATP synthase compels us to absence of question this mechanism (Bennoun, 1982). In the absence of ATP synthase, this gradient can either originate from a new ATP-dependent electrogenic pump, or from an electrogenic respiratory electron transport chain present in the thylakoid membranes. Bennoun (1982) was led to favor the second possibility in order to integrate in a single scheme another set of experimental data. The redox state of plastoquinone in the dark is partly reduced when observed in vivo under aerobic conditions. At the same time, following plastoquinone reduction by light through PS II, a high rate of reoxidation is observed in the dark. This suggests indeed that, in the dark, plastoquinone is reduced on one side, and reoxidized on the other side at a similar rate. The model of ‘Chlororespiration’ that Bennoun (1982) proposed thus consists of an electrogenic respiratory electron transport chain in the thylakoid membranes, including a pathway of reduction of plastoquinone through an NAD(P)H dehydrogenase, and a pathway of oxidation of plastoquinone at the expense of oxygen through an oxidase (Fig. 1). This pattern is comparable to that prevailing in photosynthetic prokaryotic organisms, in which photosynthesis and respiration occur in the same membranes. This chapter will not attempt to review all the studies devoted to this model in the last fifteen years, but rather to analyze the basis of this model, and to examine the recent experimental data that question this model. Abbreviations: ADP–adenosine di-phosphate; ATP–adenosine tri-phosphate; NAD – nicotinamide dinucleotide; PS I – Photosystem I; PS II – Photosystem II

II. The Thylakoid Electrochemical Gradient Present in the Dark

A. The Basis of the Model of ‘Chlororespiration’ Using delayed luminescence measurements, Joliot and Joliot (1980), showed the existence of a permanent electrochemical gradient across thylakoid membranes in dark-adapted algae. This phenomenon was attributed to the reverse functioning of the ATP synthase (Joliot and Joliot chloroplast 1980). However, Bennoun (1982) observed this permanent gradient in the Chlamydomonas FUD50 ATP synthase mutant lacking the chloroplast (as well as in any other mutant with this type of defect). A new ATP-dependent electrogenic pump could give rise to such a permanent gradient in the absence of ATP synthase. However, Bennoun (1982) preferred the alternative model of Chlororespiration, because it could also take into account the properties of the dark oxidation and reduction of plastoquinone. The problem raised by this permanent gradient was nevertheless central to this model in which proton translocation is coupled to electron transfer through a respiratory pathway associated with the thylakoid membrane (Fig. 1). In this model, the NAD(P)H dehydrogenase and possibly the oxidase as well might contribute to the formation ofthe permanent gradient. The electron flow in such a pathway would be regulated by the gradient itself, the concentrations of NAD(P)H and of oxygen. Consistent with this model, reduction of plastoquinone was observed in the presence of uncouplers (Bennoun 1983). This observation might result from uncoupling of the NAD(P)H dehydrogenase owing to the dissipation of the proton gradient, which would accelerate the input of electrons in the chlororespiratory chain. However, taking into account the now-recognized mitochondrial-chloroplast interactions (Section II.B), this effect might result as well from an indirect effect of inhibition of mitochondrial ATP synthesis.

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B. The Need for a New ATP Driven Gradient Generator More recently, Bennoun (1994) studied the effect of myxothiazol, an inhibitor of the mitochondrial cytochrome complex, on the redox state of plastoquinone in darkness and on the permanent electrochemical gradient. A fast decrease of the electrochemical gradient and a slow reduction of plastoquinone were observed following myxothiazol addition in darkness. The use of mitochondrial mutations in the cytochrome b gene conferring resistance to myxothiazol led to the conclusion that both effects were attributable exclusively to the inhibition of mitorespiration (Bennoun, 1994). These effects of myxothiazol were observed in the absence ATP synthase. If the permanent electroof chemical gradient was indeed generated by the chlororespiratory pathway, the reduction of plastoquinone would have increased the electron flow in the chlororespiratory chain, and thus increased the electrochemical gradient observed in the dark. As this gradient decreased under these conditions, Bennoun (1994) concluded that the permanent electrochemical gradient present in the absence of ATP synthase could not result from the coupling to the chlororespiratory chain. He rather proposed the presence of a new ATP-driven electrogenic pump translocating protons or other ions across thylakoid membranes (Fig. 2). In this model, the observed decrease of the permanent gradient upon specific inhibition of mitochondrial

respiration by myxothiazol indicates that, in one way or another, mitochondrial ATP is accessible to the chloroplast compartment as mentioned in Fig. 2. That chlororespiration was not responsible for the formation of the permanent electrochemical gradient was confirmed by the demonstration that, in Chlorella sorokiniana, this gradient was present under strict anaerobic conditions (Joliot and Joliot, 1994). This

678 observation was possible because fermentation processes are quite active in this species, and can provide ATP immediately when placed under anaerobiosis in the dark where mitochondrial ATP supply is lacking. Moreover, this gradient was present as well under strict anaerobic conditions in a Chlorella ATP synthase sorokiniana mutant deficient in (P. Bennoun, unpublished) and remains sensitive to submicromolar concentrations of tri-N-butyltin as in the wild-type strain (this compound blocks the proton channel of ATPases, see Joliot and Joliot, 1980). Unfortunately, the situation of Chlamydomonas under anaerobiosis in darkness is different: at least for short incubations, the gradient is very weak presumably because fermentation processes are less active, so that similar studies are not possible at present.

III. Reduction of Plastoquinone in the Dark

A. The Paradoxical Status of Chlamydomonas The plastoquinone pool observed in vivo is partly reduced in the dark, although a high rate of plastoquinone reoxidation is observed following reduction by Photosystem II (Bennoun, 1983). This simple observation suggests the existence of electron flow through plastoquinone occurring in darkness. Reducing equivalents can be supplied to the chloroplast by starch degradation, leading to the formation of reduced pyridine nucleotides (Gfeller and Gibbs, 1985). A partial reduction of plastoquinone is observed upon addition of acetate in the dark, indicating that reducing equivalents can be supplied to the chloroplast by this compound as well (Endo and Asada, 1996). Diner and Mauzerall (1973a,b) assumed the existence of an interaction between a reductant believed to be NADPH, plastoquinone, and oxygen, to explain the light saturation curve for oxygen evolution observed in Chlorella vulgaris and in Phormidium luridum. Plastoquinone reduction in the dark would proceed through a chloroplast NAD(P)H dehydrogenase in the model of ‘Chlororespiration’ proposed by Bennoun (1982). This prediction of the existence of a chloroplast NAD(P)H dehydrogenase has received large support in higher plants. Sequencing of higher plant plastid genomes revealed the ubiquitous presence (with the exception of some Pinus species) of open reading

Pierre Bennoun frames homologous to genes encoding subunits of mitochondrial Complex I (Meng et al., 1986; Ohyama et al., 1986; Ohyama et al., 1988; Sugiura, 1992). These ndh genes are transcribed (Matsubayashi et al., 1987) and the polypeptides products of some ndh genes have been found in thylakoid membranes (Nixon et al., 1989; Whelan et al., 1992; Berger et al., 1993; Guedeney et al., 1996; Kubicki et al., 1996; Martin et al., 1996). A protein complex of 670 kDa with NAD(P)H dehydrogenase activity and containing at least two ndh gene products was isolated from thylakoid membranes of potato chloroplasts (Guedeney et al., 1996). Cuello et al. (1995) and Sazanov et al. (1996) reported the existence of a chloroplast complex of 700 kDa showing a higher specificity for NADH than for NADPH. Comparison of bacterial, mitochondrial and chloroplast ndh genes revealed that the chloroplast NADH dehydrogenase complex is related to the proton-pumping complex of bacteria (Friedrich et al., 1995). This bacterial protein is simpler than the mitochondrial one that contains as many as forty polypeptides. In contrast to this situation, ndh genes could not be detected in the chloroplast genome of various Chlamydomonas species (Boudreau et al., 1994), and might be absent from other algal chloroplast genomes as well (Reith and Munholland, 1993). Moreover, the biochemical characterization of a thylakoid NADH dehydrogenase from Chlamydomonas is difficult because of the problems encountered in preparing large amounts of intact chloroplasts devoid of mitochondrial contamination, and of the observed co-purification of thylakoid and mitochondrial membranes (Atteia et al., 1992). This co-purification raises questions about the conclusions of previous work regarding the presence of this complex in thylakoid membranes (Godde and Trebst, 1980; Godde, 1982; Wu et al., 1989; Peltier and Schmidt, 1991). The situation in Chlamydomonas is therefore quite paradoxical: the chlororespiration modelwas deduced from observations made in this alga, but the predicted chloroplast ndh genes and the corresponding dehydrogenase were characterized only in higher plants. This uncomfortable situation raises several questions. Are ndh genes absent in the chloroplast genome ofChlamydomonas but present in the nuclear genome? This could create a problem for the import of their gene products into the chloroplast in light of their high hydrophobicity (Popot and de Vitry, 1990).

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Are other enzymatic systems involved in the dark reduction of plastoquinone? For instance, an alternative NADH dehydrogenase, different from Complex I, exists in yeast mitochondria. Also, the existence of a chloroplast succinate dehydrogenase has been reported by Willeford et al. (1989) in C. reinhardtii. However, this work was performed on a material that was not totally free from mitochondrial contamination. The reduction of plastoquinone in the dark can be observed in the absence of the main complexes involved in photosynthesis (PS I or PS II centers, ATP synthase, cytochrome ). However, Bennoun (1983) observed that plastoquinone reoxidation in the dark proceeds faster in mutants complex than in mutants lacking the cytochrome lacking the PS I reaction centers. This might indicate complex could be one of the entry points that the of electrons to the plastoquinone pool. In this respect, Lavergne (1983) reported the existence in Chlorella sorokiniana of an unknown redox component ‘G’, able to control the reduction of cytochrome Spectroscopic data showed that this component resembles a high-spin cytochrome c´ which could be involved in the dark reduction of the cytochrome complex (Joliot and Joliot, 1988).

B. Mitochondrial Control of the Redox State of Plastoquinone Rebeillé and Gans (1988) treated a Chlamydomonas mutant devoid of active ribulose 1,5-bisphosphate carboxylase oxygenase simultaneously by antimycin A (inhibiting cytochrome complex) and salicyl hydroxamic acid (inhibiting the alternative oxidase). This treatment resulted in a pronounced decrease in cellular ATP and a rise in NADPH concentration. This effect was accompanied by a reduction of plastoquinone and a transition from State I to State II (State transitions are treated in Chapter 30, Keren and Ohad). Gans and Rebeillé (1990) proposed that an inhibition of mitochondrial respiration, leading to a decrease of ATP level in the cell, would stimulate the glycolytic pathway in the chloroplast. This stimulation would in turn increase the NADPH level and reduce plastoquinone. Lastly, plastoquinone reduction would induce the State transition. Thus, this scheme proposes a reduction of plastoquinone caused by an interaction occurring between chloroplast and mitochondria. If true, any

conclusions drawn from the use of inhibitors of respiration raise questions because these inhibitors could act on both the mitochondrial and the chloroplast respiratory chains. For this reason, the claim by Ravenel and Peltier (1991) of a direct inhibition of chlororespiration by myxothiazol (inhibiting cytochrome complex) was questionable as well. The effect of myxothiazol was reexamined in detail by Bennoun (1994), who clearly demonstrated that a decrease of the permanent gradient across thylakoid membranes and a reduction of plastoquinone resulted from the specific inhibition of mitorespiration by this drug, thus establishing the occurrence of mitochondrial-chloroplast interactions. This was made possible by using a mitochondrial mutant resistant to myxothiazol due to a single mutation in the cytochrome b gene. A model for the reduction of plastoquinone incorporating these interactions and the new ATP-dependent gradient generator is depicted in Fig. 3.

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IV. Oxidation of Plastoquinol in the Dark

A. The Ambiguity Raised by MitochondrialChloroplast Interactions There is no doubt that plastoquinol oxidation in the dark requires the presence of oxygen (Joliot, 1965; Diner and Mauzerall, 1973; Bennoun, 1982; Peltier et al., 1987; Bennoun, 1994). On the other hand, the presence of the cytochrome complex, plastocyanin, Photosystem I or II reaction centers, and of ATP synthase is not essential for this process (Bennoun, 1983). One question to be answered is whether the oxygen consumption associated with plastoquinol oxidation occurs within the chloroplast, in the mitochondrion, or possibly in both compartments. The model of chlororespiration postulated the existence of a chloroplast oxidase connected with the plastoquinone pool and responsible for the oxidation of plastoquinol in the dark (Bennoun, 1982). Results by Peltier et al. (1987) on the sensitivity of a flash induced oxygen signal towards inhibitors of respiratory chains led these authors to favor this model. However, such conclusions cannot be drawn, unless the specificity of the inhibitors used for one compartment is demonstrated. As reported above (Section I), the inhibition of mitorespiration brings about indirect effects leading to the reduction of plastoquinone. Even if the hypothesis of the existence of a chloroplast oxidase is correct, mitorespiration rates will influence the redox state of plastoquinone. These mitochondrial-chloroplast interactions make it impossible to decide at present with certainty in is consumed during plastowhich compartment quinol oxidation in the dark.

B. Plastoquinol Oxidation via a Mitochondrial Oxidase An alternative model to that of plastoquinol oxidation through a chlororespiratory pathway involves mitochondrial-chloroplast interactions. In the presence of a chloroplast NAD(P)H dehydrogenase connected with the plastoquinone pool (Section III. A), reduced pyridine nucleotides could be exported by the chloroplast and reoxidized through the mitochondrial electron transport chain. Such interactions have been described in the light. The recycling of NADPH produced in the chloroplast by the mitochondrial electron transport chain and the import of mitochondrial ATP by the chloroplast were

shown to occur in a suppressed strain, that recovered photosynthetic competence, of the chloroplast mutant FUD50 lacking ATP synthase (Lemaire et al., 1988). However, this pathway was not operative in the wild-type strain. Peltier and Thibault (1988) proposed that plastoquinol oxidation in the light could occur through reverse electron flow from a chloroplast NADH dehydrogenase complex and reoxidation of NADH in the mitochondrion. Such an interaction between chloroplast and mitochondrial electron transport chains requires additional support, but would this pathway operate in the light, it would likely operate in the dark as well. If plastoquinone reduction in the dark occurs through a proton pumping NADH dehydrogenase, reverse electron flow should occur from plastoquinol at the expense of the electrochemical to gradient. This process could consume the permanent gradient built up by the new ATP-driven gradient generator postulated by Bennoun (1994) as mentioned in Section II. Following reverse electron flow, the reductant formed could be reoxidized at the expense of oxygen through the mitochondrial electron transport chain as it might occur in the light (see above). One observation that favors this alternative model is that quinone treatment inhibits both plastoquinone oxidation and reduction in darkness (Joliot and Joliot, 1985). Such a treatment is known to inhibit enzymes using nucleotides like ATP synthase and NADH dehydrogenase, but has no inhibitory effect on oxidases (Diner and Joliot, 1976; Joliot et al., 1989; Lavergne, 1989; Bulté and Wollman, 1990). In the same manner, plastoquinol oxidation in darkness is very rapid in vivo, yet drastically slowed down in vitro (in open cell preparations or in isolated chloroplasts), a condition in which the gradient is abolished. This dependence upon the integrity of the system would be consistent with a mechanism of oxidation involving the thylakoid electrochemical gradient. Lastly, contrary to the postulated NADH dehydrogenase complex whose presence was recognized at least in higher plants (Section III), no chloroplast oxidase could ever be characterized by biochemical or molecular approaches.

V. Conclusion The model of chlororespiration as proposed by Bennoun (1982) made three assumptions: the

Chapter 35

Chlororespiration

existence of a chloroplast NAD(P)H dehydrogenase connected to plastoquinone and involved in its dark reduction, the existence of a chloroplast oxidase connected to plastoquinone and involved in its dark oxidation, and an electrogenic electron transfer between these complexes responsible for the permanent thylakoid electrochemical gradient present ATP synthase. in the absence of It is now clear that the origin of the permanent electrochemical gradient does not rely on chlororespiration, but likely on the presence of a new ATPdriven gradient generator associated with thylakoid membranes. This opens new exciting perspectives of research, and the new technique of luminescence digital imaging described in this volume (Chapter 23, Bennoun and Béal) should be of great help to characterize this protein complex. With regard to the chloroplast NAD(P)H dehydrogenase, a way of entry of electrons in the plastoquinone pool definitely exists, as demonstrated by the changes in the redox state of this pool in the dark, induced for instance by acetate or upon inhibition of mitorespiration. This pathway is independent from the known complexes of the photosynthetic electron transport chain. In higher plants, as predicted by the model of chlororespiration, compelling evidence has been gathered for the presence of a chloroplast NADH dehydrogenase analogous to the proton pumping enzyme of bacteria (Section III.A). Such molecular evidence is lacking in Chlamydomonas, and the difficulties encountered would likely require some work to be performed on purified chloroplast or thylakoid membranes. Unfortunately, if Chlamydomonas is especially suitable for physiological investigations, such fractionations remain difficult in this alga. On the other hand, Chlamydomonas is an excellent model system for genetic studies, and progress in screening procedures which use fluorescence digital imaging (see Chapter 23, Bennoun and Béal) may allow one to select for mutants affected in the redox state of plastoquinone in darkness. This could eventually lead to the characterization of electron carriers involved in the dark reduction of plastoquinone. Lastly, molecular evidence for a chloroplast oxidase has not been provided for any organism. Due to mitochondrial-chloroplast interactions, it is impossible at present to decide whether, in the course of plastoquinol oxidation, oxygen is consumed in the chloroplast or within mitochondria. A set of data argue against the existence of this enzyme (Section

681 IV.B), especially the fact that, contrary to the prediction, the formation of the permanent thylakoid electrochemical gradient is independent of the process of chlororespiration (Section II.A). As this prediction was one of the bases for developing this model, it is wise to examine alternative schemes ofplastoquinone oxidation. In this respect, the combination of an NADH dehydrogenase and of an ATP-dependent electrogenic pump offers the basis of a mechanism for this reaction: a reverse electron flow from driven at the expense of the permanent to electrochemical gradient. Several arguments favor this hypothesis, especially the observation that plastoquinol oxidation strongly depends on the integrity of the biological material, as also does the establishment of a permanent gradient. Here again, the properties of Chlamydomonas are such that, in the future, a genetic approach to this problem might be more fruitful than a molecular one. In any case, the mechanism of the dark oxidation of plastoquinol remains at present an open question. Recent work has demonstrated the occurrence of mitochondrial-chloroplast interactions that need to be examined in more details. In particular a close control of both the permanent gradient and the redox state of plastoquinone by the mitochondrial electron transport chain was demonstrated thanks to the use of specific mitochondrial mutants (Bennoun, 1994). This conclusion, together with the evidence for a new ATP-driven gradient generator associated with thylakoid membranes (Bennoun, 1994), lead us to reconsider the model of chlororespiration. New models attempting to describe the properties of thylakoid membranes in darkness should integrate these two facts. Obviously, the questions raised by the observation of thylakoid membranes in darkness will require new experimental efforts. Whatever model prevails, it will imply the existence of new complexes in the thylakoid membranes, whose abundance is expected to be low, since major thylakoid membrane polypeptides have already been attributed to photosynthetic complexes. This will of course be an additional obstacle to their molecular characterization, and even once this characterization is achieved, the physiological role of these complexes will need to be clarified. For instance, what is the requirement for a permanent gradient across thylakoid membranes? Whether this gradient is of importance for the biogenesis of these membranes or for the activity of some of their complexes are open questions.

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Cautionary Note The existence of mitochondrial-chloroplast interactions requires that physiological studies conducted in vivo on thylakoid membranes be performed with strains that are fully competent for mitorespiration. However, most Chlamydomonas strains used nowadays all over the world do not meet this requirement. When analyzed genetically, they display many mutations slowing down heterotrophic growth, that should be removed by repeated crosses. Combinations of mutations leading to cell death in the dark and suppressor mutations are not uncommon either. A clone of about 2 mm should be recovered in a week at 25° on acetate plates in the dark. In the very same way, most Chlamydomonas strains commonly used are not fully competent for photosynthetic growth. Again repeated crosses should be performed and selected at each generation for growth on minimal medium at low light intensity (1000 to 2000 lux cool fluorescent light), a condition in which the primary processes of photosynthesis are limiting.

Acknowledgments The author is pleased to acknowledge R. Delosme and O. Vallon for critical reading of the manuscript. This work was supported by UPR 9072 of the Centre National de la Recherche Scientifique.

References Atteia A, de Vitry C, Pierre Y and Popot JL (1992) Identification of mitochondrial proteins in membrane preparations from Chlamydomonas reinhardtii. J Biol Chem 267: 226–234 Bennoun P (1982) Evidence for a respiratory chain in the chloroplast. Proc Natl Acad Sci USA 79: 4352–4356 Bennoun P (1983) Effects of mutations and of ionophore on chlororespiration in Chlamydomonas reinhardtii. FEBS Lett 156: 363–365 Bennoun P (1994) Chlororespiration revisited: Mitochondrialplastid interactions in Chlamydomonas. Biochim Biophys Acta 1186: 59–66 Bennoun P, Delosme M and Kück U (1991) Mitochondrial genetics of Chlamydomonas reinhardtii: Resistance mutations marking the cytochrome b gene. Genetics 127: 3356–343 Berger S, Ellersiek U, Westhoff P and Steinmüller K (1993) Studies on the expression of NDH-H, a subunit of the NAD(P)Hplastoquinone-oxidoreductase of higher-plant chloroplasts. Planta 190: 25–31 Bulté L and Wollman FA (1990) Stabilization of states I and II by p-benzoquinone treatment of intact cells of Chlamydomonas

Pierre Bennoun reinhardtii. Biochim Biophys Acta 1016: 253–258 Boudreau ER, Otis C and Turmel M (1994) Conserved gene clusters in the highly rearranged chloroplast genomes of Chlamydomonas moewusii and Chlamydomonas reinhardtii. Plant Mol Biol 24: 585–602 Cuello J, Quiles MJ, Albacete ME and Sabater B (1995) Properties of a large complex with NADH dehydrogenase activity from barley thylakoids. Plant Cell Physiol 36: 265–271 Diner B and Joliot P (1976) Effect of the transmembrane electric field on the photochemical and quenching properties of Photosystem II in vivo. Biochim Biophys Acta 423: 479–498 Diner B and Mauzerall D (1973a) Feedback controlling oxygen production in a cross-reaction between two photosystems in photosynthesis. Biochim Biophys Acta 305: 329–352 Diner B and Mauzerall D (1973b) The turnover of photosynthesis and redox properties of the pool of electron carriers between the photosystems. Biochim Biophys Acta 305: 353–363 Endo T and Asada K (1996) Dark induction of the non-photochemical quenching of chlorophyll fluorescence by acetate in Chlamydomonas reinhardtii. Plant Cell Physiol 37: 551–555 Friedrich T, Steinmüller K and Weiss H (1995) The protonpumping respiratory complex I of bacteria and mitochondria and its homologue in chloroplasts. FEBS Lett 367: 107–111 Gans P and Rebeillé F (1990) Control in the dark of the plastoquinone redox state by mitochondrial activity in Chlamydomonas reinhardtii. Biochim Biophys Acta 1015: 150–155 Gfeller RP and Gibbs M (1985) Fermentative metabolism of Chlamydomonas reinhardtii II. Role of plastoquinone. Plant Physiol 77: 509–511 Godde D (1982) Evidence for a membrane bound NADPHplastoquinone oxidoreductase in Chlamydomonas reinhardii. Arch Microbiol 131: 197–202 Godde D and Trebst A (1980) NADH as electron donor for the photosynthetic membrane of Chlamydomonas reinhardtii. Arch Microbiol 127: 245–252 Guedeney G, Corneille S, Cuiné S and Peltier G (1996) Evidence for an association of ndh B, ndh J gene products and ferredoxinNADP-reductase as components of a chloroplastic NAD(P)H dehydrogenase complex. FEBS Lett 378: 277–280 Joliot P (1965) Cinétiques des réactions liées à l’émission d’oxygène photosynthétique. Biochim Biophys Acta 102: 116–134 Joliot P and Joliot A (1980) Dependence of delayed luminescence upon Adenosine Triphosphatase activity in Chlorella. Plant Physiol 65: 691–696 Joliot P and Joliot A (1985) Slow electrogenic phase and intersystem electron transfer in algae. Biochim Biophys Acta 806: 398–409 Joliot P and Joliot A (1988) The low-potential electron-transfer chain in the cytochrome b/f complex. Biochim Biophys Acta 933: 319–333 Joliot P, Verméglio and Joliot A (1989) Evidence for supercomplexes between reaction centers, cytochrome and cytochrome complex in Rhodobacter sphaeroides whole cells. Biochim Biophys Acta 975: 336–345 Kubicki A, Funk E, Westhoff P and Steinmüller K (1996) Differential expression of plastome-encoded ndh genes in plant mesophyll and bundle-sheath chloroplasts of the Sorghum bicolor indicates that the complex I-homologous NAD(P)H-plastoquinone oxidoreductase is involved in cyclic

Chapter 35 Chlororespiration electron transport. Planta 199: 276–281 Lavergne J (1983) Membrane potential-dependent reduction of cytochrome b-6 in an algal mutant lacking Photosystem I centers. Biochim Biophys Acta 725: 25–33 Lavergne J (1989) Mitochondrial responses to intracellular pulses of photosynthetic oxygen. Proc Natl Acad Sci USA 86: 8768– 8772 Lemaire C, Wollman FA and Bennoun P (1988) Restoration of phototrophic growth in a mutant of Chlamydomonas reinhardtii in which the chloroplastic atpB gene of the ATP synthase has a deletion: An example of mitochondria-dependent photosynthesis. Proc Natl Acad Sci USA 85: 1344–1348 Martin M, Casano LM and Sabater B (1996) Identification of the product of ndhA gene as a thylakoid protein synthesized in response to photooxidative treatment. Plant Cell Physiol 37: 293–298 Matsubayashi T, Wakasugi T, Shinozaki K, Yamaguchi-Shinozaki K, Zaita N, Hidaka T, Meng BY, Ohto C, Tanaka M, Kato A, Maruyama T and Sugiura M (1987) Six chloroplast genes (ndhA-F) homologous to human mitochondrial genes encoding components of the respiratory chain NADH dehydrogenase are actively expressed: determination of the splice sites in ndhA and ndhB, pre-mRNAs. Mol Gen Genet 210: 385–393 Meng BY, Matsubayashi T, Wakasugi T, Shinozaki K, Sugiura M, Hirai A, Mikami T, Kishima Y and Kinoshita T (1986) Ubiquity of the genes for components of a NADH dehydrogenase in higher plant chloroplast genomes. Plant Sci 47: 181–184 Nixon PJ, Gounaris K, Coomber SA, Hunter CN, Dyer TA and Barber J (1989) psbG is not a Photosystem two gene but may be a ndh gene. J Biol Chem 264: 14129–14135 Ogawa T (1991) A gene homologous to the subunit-2 of NADH dehydrogenase is essential to inorganic carbon transport of Synechocystis PCC6803. Proc Natl Acad Sci USA 88: 4275– 4279 Ohyama K, Fukuzawa H, Kohchi T, Shirai H, Sano T, Sano S, Umesono K, Shiki Y, Takeuchi M, Chang Z, Aota S, Inokuchi H and Ozeki H (1986) Chloroplast gene organization deduced from complete sequence of liverwort Marchantia polymorpha chloroplast DNA. Nature 322: 572–574 Ohyama K, Kohchi T, Sano T and Yamada Y (1988) Newly

683 identified groups of genes in chloroplasts. TIBS 13: 19–22 Peltier G and Schmidt GW (1991) Chlororespiration: An adaptation to nitrogen-deficiency in Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 88: 4791–4795 Peltier G and Thibault P (1988) Oxygen-exchange studies in Chlamydomonas mutants deficient in photosynthetic electron transport: evidence for a Photosystem II-dependent oxygen uptake in vivo. Biochim Biophys Acta 936: 319–324 Peltier G, Ravenel J and Verméglio A (1987) Inhibition of a respiratory activity by short saturating flashes in Chlamy­ domonas: Evidence for a Chlororespiration. Biochim Biophys Acta 893: 83–90 Popot JL and de Vitry C (1990) On the microassembly ofintegral membrane proteins. Arch Rev Biophys Chem 19: 369–403 Ravenel J and Peltier G (1991) Inhibition of Chlororespiration by myxothiazol and antimycin A in Chlamydomonas reinhardtii. Photosynth Res 28: 141–148 Ravenel J and Peltier G (1992) Stimulation ofthe chlororespiratory electron flow by Photosystem II activity in Chlamydomonas reinhardtii. Biochim Biophys Acta 1101: 57–63 Rebeillé F and Gans P (1988) Interaction between chloroplasts and mitochondria in microalgae: role of glycolysis. Plant Physiol 88: 973–975 Sazanov LA, Burrows P and Nixon PJ (1996) Detection and characterization of a complex I-like NADH-specific dehydrogenase from pea thylakoids. Biochem Soc Trans 24: 739–743 Sugiura M (1992) The chloroplast genome. Plant Mol Biol 19: 149–168 Whelan J, Young S and Day DA (1992) Cloning of ndhK from soybean chloroplasts using antibodies raised to mitochondrial complex I. Plant Mol Biol 20: 887–895 Willeford KO, Gombos Z and Gibbs M (1989) Evidence for chloroplastic succinate dehydrogenase participating in the chloroplastic respiratory and photosynthetic electron transport chains in Chlamydomonas reinhardtii. Plant Physiol 90:1084– 1087 Wu M, Nie ZQ and Yang J (1989) The 18-kD protein that binds to the chloroplast DNA replicative origin is an iron-sulfur protein related to a subunit of NADH dehydrogenase. Plant Cell 1: 551–557

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Chapter 36 Perspectives Laurens J. Mets

Department of Molecular Genetics and Cell Biology,

University of Chicago, Chicago, IL 60637, U.S.A.

Jean-David Rochaix

Departments of Molecular Biology and Plant Biology, University of Geneva,

30 Quai E. Ansermet, CH-1211 Geneva 4, Switzerland

Summary I. Introduction II. The Niche of Chlamydomonas in Photosynthesis Research A. Natural Features B. Engineered Features 1. Chloroplast Genome Engineering 2. Mitochondrial Genome Engineering 3. Nuclear Genome Transformation, Analysis and Engineering 4. Advances in Colony Screening Methods C. Constraints that May Be Surmounted by Further Work D. Comparison with Cyanobacteria III. Forefront Problems in Photosynthesis and Organelle Research A. Structure-Function Analysis of Individual Complexes B. Integrative Aspects of Chloroplast Biogenesis 1. Organellar and Nuclear Genome Cooperation and Integration 2. Unusual Features of Organellar Gene Expression Systems Acknowledgments References

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Summary Studies using the unicellular green alga Chlamydomonas reinhardtii have earned a role in building our understanding of photosynthesis and chloroplast biogenesis through a combination of fortuitous natural characteristics and hard work by many investigators. The most critical contributor to this role is the development of tools for genetic and molecular genetic analysis. Photosynthetic processes span a wide range of temporally and mechanistically distinct phenomena that are intertwined with one another in ways that are both subtle and profound. Many of the biophysical, biochemical and physiologic techniques that can be exploited for investigating photosynthesis operate within restricted ranges of this temporal domain. Genetic analysis is an J.-D. Rochaix, M. Goldschmidt-Clermont and S. Merchant (eds): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, pp. 685–703. © 1998 Kluwer Academic Publishers. Printed in The Netherlands.

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investigative approach that extends across these domains and provides a basis for integrating the understanding that arises from many other types of studies. Studies exploiting the favorable characteristics of C. reinhardtii are continuing to prove their worth in the forefront areas of structure-function analysis of photosynthetic complexes, metabolic regulation and coordination of photosynthesis, and regulation of chloroplast biogenesis. Although they have already been uniquely valuable in the past, the range and precision of the tools for genetic analysis in this alga are currently being improved rapidly and dramatically. Research in photosynthesis will be a significant beneficiary.

I. Introduction Photosynthesis is a biologically catalyzed molecular process with massive global scale impact. Energy flux in living organisms is the very substance of life itself, and the vast majority of net biologically available energy that enters the global ecosystems arises in the catalyzed conservation of light energy to chemical form in the primary reactions of photosynthesis. The primary photochemical charge separation reactions take only picoseconds to occur, but on a geologic time scale, through a chain of intervening biochemical reactions and biological processes, they have inexorably led to a complete change in the composition of the earth’s atmosphere and have fueled organic evolution. The enzyme ribulose-1,5 -bisphosphate carboxylase/oxygenase (Rubisco) has fingered over at one time or another every single atom of carbon present in all living organisms and in all fossil fuel deposits. Every molecule of oxygen present in the modern atmosphere was generated in the photolytic oxidation of water catalyzed by Photosystem II (PS II), the oxygenic photosynthetic reaction center. Photosynthesis, as a process, figures prominently in the solution to many of the major Problems of Our Time. All of animal and human nutrition depend upon photosynthetically driven productivity in crops. Photosynthetic energy capture, both as actually practiced by living organisms and as a model for artificial systems, holds considerable promise as a source of renewable, sustainable energy for industrial and domestic needs, even in energy-intensive industrial societies. Since photosynthesis is the process that reduced concentrations to the 300 ppm level present in the modern pre-industrial earth’s atmosphere, it also holds promise as the solution to the threat of global warming caused by the recent man-made rise in atmospheric concentrations. Considering the net equation governing carbon balance in photosynthesis at minimum quantum requirement for Z-scheme photosynthesis,

it is clear that thermodynamic equilibrium subremoval from the atmosphere. stantially favors In the absence of coupled light energy, reoxidation of carbon,

is also thermodynamically favorable and constitutes a basic biological and industrial fueling reaction. Global carbon balance depends upon the relative net rates of these two countervailing reactions. Since net rates are governed by details of catalytic mechanisms of each step in the overall process, they cannot be understood without delving into the details of catalysis of each step. This requirement for understanding each step underlies the broad scope of research that is covered in this series. Evolutionary considerations tell us that in an efficient (selectively competitive) organism that depends upon a multistep process like photosynthesis for survival, no one step in the process can be limiting, and all must operate at balanced rates. Through the medium of evolutionary selection, the genetic, biochemical and physiologic conditions that govern the net rate ofcarbon fixation by Rubisco, for example, must be balanced with the genetic, biochemical, and photophysical mechanisms that govern excitation migration and trapping in photosynthetic antennae. An understanding of one process cannot necessarily be obtained without considering its implications for, and interactions with, the other. Why are the primary reactions of photosynthesis of high quantum efficiency rather than high energy efficiency? Why do oxygenic photosynthetic organisms exploit two photosystems and two quanta of light per electron to accomplish reactions for which one photosystem and one quantum would be thermodynamically

Chapter 36 Perspectives sufficient? The answers to these questions may not lie solely within the realm of photochemistry but may depend upon understanding natural selection operating on exigencies that constrain some other aspect of the photosynthetic process. Successful design of artificial photosynthetic systems will require understanding the natural reaction centers in this broader context. The individual steps in the photosynthetic process occur on a vast range of mechanistically distinct time domains, from the few tens of femtoseconds required for energy transfer of chlorophyll excited states from one molecule to another (the photophysical time domain) to the few tenths of a second required for a single turnover of Rubisco (the biochemical time domain) to a few hours required for chloroplast biogenesis (the molecular biological time domain) to the millennia involved in atmospheric changes (the evolutionary time domain) (Fig. 1). Understanding the process linkages that arise across this vast temporal landscape is not a trivial technical or conceptual matter. The scaling problem faced by photosynthesis research is actually multidimensional, encompassing time, space, and energy and combinations of these dimensions like rate, mass and power. Progress in research requires focus on particular steps in the overall process and typically yields models that are valid within a limited scope oftime. Each investigative technique yields information within only a limited temporal domain. Human attention naturally focuses in a particular, familiar technical and temporal domain and it takes conscious effort to remind ourselves that we must step back and consider that the critical insights necessary for answering our most engaging local mystery may arise from a different domain investigated with a different technique. Consider that in a growing cell, protein synthesis requires more energy than any other aspect of metabolism. For this reason alone, the distinction between post-translational and pre-translational coordination of gene expression (Chapters 9, Nickelsen; 10, Stern and Drager; 11, Herrin et al.; 12, Hauser, et al.), which may be necessary for adjusting the critical time frame of an adaptive response, could be critically related to starch metabolism (Chapter 29, Ball), cellular motility, nutrient acquisition and utilization (Chapters 28, Spalding; 32, Davies and Grossman; 33, Fernandez et al.), respiratory metabolism (Chapters 18, Redding and Peltier; 35, Bennoun), and to factors that determine the metabolic fate of absorbed light energy, such as regulation of non-photochemical quenching,

687 reaction center photoinhibition, and antenna state transitions (Chapter 30, Ohad). As we understand more and more about the mechanisms that operate within limited domains, expanding our understanding to the forefront problems of regulation and dynamic coordination, which also operate across these time domains, will require intentional cross-disciplinary and cross-dimensional dialog. While it is obvious in studies of photosynthesis that building understanding requires consideration of many time domains, the same fundamental chemical, physical, molecular and evolutionary processes constrain all biological reactions and functions. Photosynthesis differs only in the arbitrary technical fact that its rapid processes can be initiated synchronously and therefore discerned with very short flashes of light. Our feeling is that better understanding of all biological processes can be obtained by consciously forcing a broad temporal perspective. Photosynthesis serves as an instructive perspective model for studies of other biological processes. It is clear that no one area of photosynthesis research can proceed to a conclusion without encountering a fundamental need to understand other areas. This is the rationale that organizes the presentation of this volume and of this series. This volume focuses on studies of Chlamydomonas, which has come under the scrutiny of photosynthesis researchers principally because of the breadth and detail of genetic analysis that it will endure. Genetic analysis is an investigative tool that intrinsically spans across temporal boundaries and hence serves as a point ofcontact for integrating different fields of study. Our intent in this perspective chapter is to assess the unique current and future niche occupied by Chlamydomonas, in particular, in efforts to advance broader understanding of photosynthesis, with emphasis on the role of genetic analysis.

II.The Niche of Chlamydomonas in Photosynthesis Research The feasibility of genetic analysis combined with a number of other fortuitous natural properties brought C. reinhardtii onto the stage of photosynthesis research (Chapter 2, Togasaki and Surzycki). The accumulated effort of numerous investigators, as represented in the chapters of this volume, has generated a number of additional tools for genetic manipulation and analysis of this species that justify

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Chapter 36 Perspectives the substantial recent rise in interest in its exploitation for an increasing range of studies in photosynthesis and organelle biology. In this section we lay out the natural and engineered dimensions and boundaries of this research niche, since they establish the locus of new research opportunities.

A. Natural Features Taxonomically, the green algae (Chlorophyta), including Chlamydomonas spp., Chlorella spp. and Scenedesmus spp., are more closely related to land plants than is any other algal group (Chapter 5, Nedelcu and Lee). They have proven sufficiently similar in terms of biochemistry and metabolic organization to serve as models for investigating and understanding at the cellular level many aspects of the photosynthetic processes that drive economic production in crops. When grown on chemically defined media, these unicellular algae are biochemically and physiologically homogeneous and behave reproducibly; properties that are difficult to obtain with preparations from multicellular plants. On the other hand, unicellularity restricts research focus to the cellular and sub-cellular level, since metabolic cooperation between cells does not exist. This restricted focus, however, makes clear the integrative aspects of cell autonomous photosynthesis (Chapters 2,Togasaki and Surzycki; 18, Redding and Peltier; 20, Timko; 28, Spalding; 29, Ball; 31, Merchant; 32, Davies and Grossman; 33, Fernandez et al.; 35, Bennoun) in a way that is more difficult to discern in studies of multicellular plants. The unicellular green algae grow rapidly, with doubling times as short as 2 h (some varieties of Chlorella) to 4 h (C. reinhardtii under favorable conditions) (Harris, 1989). Millions of genetically clonal colonies can be grown in days from single cells plated on an agar surface. This feature enables intensive selection or screening for phenotypes caused by rare mutations. Because the cells are haploid, mutant phenotypes are directly expressed following mutagenesis without the need for further genetic manipulation. While haploidy is a great advantage for studies of genes that control non-essential or dispensable characteristics of the cells, it greatly complicates the recovery of mutations in genes that are essential for cell growth and division. Fortunately for studies of photosynthesis, certain species of these algae (e.g. Chlorella vulgaris, Scenedesmus obliqus, and C. reinhardtii) can utilize a fixed carbon

689 compound to support heterotrophic growth even in the absence of light. This makes most aspects of photosynthetic function dispensable and readily amenable to mutational analysis. C. reinhardtii differs from the other species in that it cannot utilize glucose to support heterotrophic growth, but it can utilize acetate, which is assimilated through the glyoxylate cycle (Sager and Granick, 1953; Harris, 1989). C. reinhardtii actually grows more quickly in the light with acetate in the medium (‘mixotrophic’ conditions) than it does under strictly autotrophic as the only available carbon conditions with source and most photosynthesis mutants grow more slowly than wild type cells on acetate medium in the light. The resulting ‘small colony’ phenotype is sufficient for identifying many types of photosynthesis-deficient mutants. It should be noted that mutants deficient in cytochrome-dependent mitochondrial respiration also share this small colony phenotype under mixotrophic growth conditions (Matagne et al., 1989). Evidently photosynthesis stimulates respiratory acetate utilization (Chapter 35, Bennoun). While photosynthesis is dispensable under heterotrophic growth conditions, in C. rein­ hardtii, mitochondrial respiration is dispensable under photoautotrophic growth conditions (Matagne et al., 1989; Chapter 34, Remacle and Matagne). Thus, mutational analysis of mitochondrial as well as chloroplast function are feasible in the same organism. Unlike land plants, C. reinhardtii synthesizes chlorophyll in the dark at rates sufficient to support the biogenesis of a fully functional photosynthetic apparatus (Chapter 20, Timko). With a life cycle of just one day, the cells can (and do) generate a photosynthetic apparatus overnight that is ready to function as soon as there is light available in the morning. The discovery of the genetic and enzymatic basis for light-independent protochlorophyllide reduction in C. reinhardtii has led to intriguing insights into the evolution of photosynthetic organisms (Chapters 5, Nedelcu and Lee; 20,Timko), but here we wish to emphasize the often overlooked importance of this characteristic to the value of this species in the genetic dissection of photosynthetic functions. Without light-independent protochlorophyllide reduction, critical structure-function analysis of photosynthetic complexes by mutation would be impossible. Chlorophyll, at up to 30% of the mass, must be considered a structural component of the PS I (Chapter 17, Webber and Bingham), PS II (Chapters 15, Erickson; 16, Ruffle and Sayre), and

690 LHC (Chapter 19, Hoober et al.) complexes. Even the cytochrome complex contains a chlorophyll a molecule as a stoichiometric component (Chapter 24, Wollman; Huang et al., 1994). In the absence of chlorophyll, these complexes do not assemble. At the same time, the light energy absorbed by chlorophyll is potentially damaging if not handled carefully. Absorption of light by chlorophyll in the photosynthetic reaction centers generates the strongest and strongest reductant each a oxidant chlorophyll molecule, found in living cells. Every evidence indicates that photosynthetic cells rely upon an extraordinary array of regulatory mechanisms to balance the relative and absolute rates of production of these reactive species with the capacities and requirements of metabolism (Chapters 14, Olive and Wollman; 20, Timko; 26, Jacquot et al.; 29, Ball; 30, Ohad; 32, Davies and Grossman). These regulatory and protective mechanisms have been honed through eons of natural selection. It appears that most were in place prior to the emergence of land plants into an unfriendly, oxygen-containing atmosphere 500 million years ago, since most are found in green algae as well. Unraveling the intricacies of these interconnected regulatory mechanisms is one of the forefront challenges of photosynthesis research and an area in which we feel that genetic analysis in C. reinhardtii will play a crucial role. In the mean time, however, it is important to anticipate that any mutation that disrupts the balance for any reason, is likely to generate reactive and hence damaging photochemical intermediates in the light. This would make it difficult to distinguish the direct effects of a mutation from the photosensitized damage that it indirectly causes. Hence, it is an important property of C. reinhardtii that photosynthetically altered mutants can grow in the dark and that as they grow they synthesize the necessary chlorophyll to assemble the photosynthetic complexes for study. It is worth noting at this point that almost every mutation that affects some aspect of photosynthetic metabolism may well cause indirect photodynamic damage. For instance, Rubisco-deficient mutants are photosensitive (Chapter 27, Spreitzer). Growing them in the light not only causes damage, it also selects for a range of secondary mutations that suppress photosensitivity, but have nothing directly to do with Rubisco function (Spreitzer and Ogren, 1983). Again, accurate diagnosis of cause and effect in cells with genetically altered photosynthesis requires mini-

Laurens J. Mets and Jean-David Rochaix mizing light-induced stresses during growth of the study material. Among the green algae, Chlamydomonas rein­ hardtii stands out as the only species that can be grown heterotrophically that also has a readily controlled sexual life cycle (Chapter 1, Harris). Opposite mating type haploid cells can be induced to undergo gametogenesis and fuse to form a diploid zygote. After maturation, this zygote proceeds through meiosis, releasing the four genetically recombined haploid spores (the tetrad), which each grow into vegetative clones. This feature enables full genetic analysis of induced mutations. In the analysis of complete tetrads, an observed 2:2 segregation of a particular mutant phenotype gives confidence that it is caused by a single nuclear gene difference, rather than by a combination of differences in several genes. Through meiotic recombination, mutations can be genetically combined to test hypotheses about their interactions. Methods have been developed for selectively recovering mitotically dividing diploid cells, which are a minority of cells formed in the mating reaction (Ebersold, 1963, 1967). The availability of vegetative clones of heterozygous diploid cells enables testing dominance of mutations and complementation analysis for distinguishing among mutations with the same phenotype. In terms ofthe needs of genetic analysis, the one deficiency in methods for controlling the C. reinhardtii sexual life cycle is that the vegetative diploid cells cannot be induced to enter meiosis and yield haploid products again. In addition to its unique advantages among green algae for genetic analysis of nuclear gene mutations, Chlamydomonas is the only genus in which recombination of chloroplast genes occurs on a regular basis following sexual crosses (Chapters 6, Armbrust; 7, Sears). The prospect of conducting detailed genetic analysis of not only nuclear but also chloroplast gene mutations in a single organism attracted many researchers interested in photosynthesis and chloroplast molecular biology to work with Chlamydomonas. More recently, the pattern of transmission and recombination ofthe mitochondrial genome in crosses of C. reinhardtii has been established (Chapter 34, Remacle and Matagne), allowing full experimental control over combinations of nuclear and organellar genotypes. While organellar gene recombination can be exploited for creating new combinations of chloroplast or mitochondrial

Chapter 36

Perspectives

gene mutations, it has been largely supplanted for this purpose by the development of facile transformation methods (see below and Chapters 8, Goldschmidt-Clermont; 34, Remacle and Matagne). An important feature of the sexual life cycle of C. reinhardtii that has been underexploited in studies of organelle genetics is that newly mated cells are transiently heteroplasmic (carrying organellar genomes from both parents) just after mating. The two parental chloroplasts that enter the zygote remain apart for a few hours and then fuse into a single zygotic chloroplast that remains photosynthetically active for some time (Friedman et al., 1968; CavalierSmith, 1970, 1975, 1976). Bennoun et al. (1980) showed that both parental chloroplast genomes remain functional in gene expression for some time, in spite of the eventual degradation of the genome contributed parent in most zygotes. They showed that by the complementation between chloroplast gene mutations could be tested in this way, both prior to and after chloroplast fusion. Baldan et al. (1991) extended this observation and demonstrated that chloroplast and thylakoid membrane fusion could be exploited for initiating and studying the process of assembly of fully functional photosynthetic complexes during complementation of two different assembly-deficient strains (Chapter 14, Olive and Wollman). It is reasonable to expect that analysis of young zygotes will also allow complementation analysis of mitochondrial mutations. Biochemical and crystallographic studies of individual enzymes or photosynthetic complexes often require substantial amounts (kg) of starting material for a purification procedure. In this arena, algae do not often compete with crop plant material. Most labs are not equipped with fermentors large enough to grow more than a few grams of cells in a batch. Large scale commercial fermentors are not typically fitted with lights and become prohibitively expensive to use when doubling times are greater than 1–2 h (heterotrophically grown C. reinhardtii doubles in 12–18 h; many mutants grow even more slowly). Large scale outdoor algae farms are difficult to maintain axenic, and cultivation of C. reinhardtii is quite sensitive to contamination with a range of organisms from the environment, particularly when grown in acetate-supplemented media. Crop production for food is conducted on such a large scale that the economy and convenience of obtaining spinach from the market is hard to match with algal

691 cultures. However, there are instances in which the advantages of ‘genetically purified’ algal mutants will outweigh the disadvantages of growing the algae. Genetic purification can be used to eliminate through mutation and genetic combination those parts of the cell that are difficult or expensive to eliminate during a biochemical purification process. For instance, Pierre et al. (1995) were able to obtain highly purified comand active preparations of the cytochrome plex from C. reinhardtii by starting with mutants complex. Mutants deficient in the mitochondrial devoid of both PS I and PS II complexes have been exploited for studying single step excitation energy transfer from Chl b to Chl a in intact thylakoid membranes and even in intact cells (Eads et al., 1989; Du et al., 1994). In this case, the study could not be done without genetic purification, since the detergent treatments necessary for biochemically separating LHC complexes from PS I and PS II complexes in non-mutant organisms generate spectral changes in LHCII that complicate interpretation of the transfer kinetics (Eads et al., 1989). The opportunity for starting biochemical purifications with genetically purified biological material could be more generally exploited. One of the advantages of unicellularity is that it is often possible to feed cells exogenous metabolites in order to either modify metabolism or to support mutants with defects in metabolism. In an important recent development, it has proven possible to feed Chlamydomonas cells with exogenous lipids (Chapter 21, Trémolières). Certain lipids find their way from exogenous micelles into the thylakoid membranes, with the result that the lipid composition is modified. By constructing mixed micelles, it has been possible to recover mutants defective in the biosynthesis of specialized thylakoid membrane lipids, including sulfolipid, whose function has been enigmatic. Sulfolipid-deficient mutants of Chlamydomonas have much stronger deficiency in PS II activity (Sato et al., 1995) than would have been expected from the properties of comparable mutants in Rhodobacter sphaeroides (Benning et al., 1993) and Synechococcus sp. PCC7942 (Güler et al., 1996). The development of lipid feeding techniques and procedures for rescuing lipid-deficient mutants provides for the first time experimental tools necessary for studying the roles of specific lipids in the biogenesis and function of the various thylakoid membrane complexes. Such studies have been

692 impossible previously, since the lipid composition of the complexes is disturbed during detergent isolation. Although the route oflipid uptake is still unclear, it is intriguing to consider that lipid micelles might be able to deliver to the thylakoids other lipid-soluble components, such as quinones, carotenoids, phytol, and even chlorophyll.

B. Engineered Features In addition to its unique natural biological properties that have contributed to the value of C. reinhardtii in photosynthesis research, many important experimental tools have been engineered for use in this species through the hard and creative work of many investigators. Chief among these are methods for genetic transformation that were ushered in by the development of physical methods for introducing exogenous DNA into the cells (Boynton et al., 1988; Kindle, 1990).

1. Chloroplast Genome Engineering As reviewed in detail in Chapter 8 (GoldschmidtClermont), it is now possible to engineer the nucleotide sequence of the chloroplast genome virtually at will. Introduced DNA integrates via homologous recombination into the chloroplast DNA, causing replacement of endogenous sequences with introduced ones. Genes from other organisms have been engineered for expression in the chloroplast and exploited as selectable genetic markers for enhancing the recovery of specific transformants (Goldschmidt-Clermont, 1991). A system for removing the selectable marker following transplacement of an endogenous gene has also been demonstrated (Fischer et al., 1996), making it possible to construct multiply altered genomes by sequential transplacement. The sole remaining barrier to exploitation of this capability in photosynthesis research is that the complete sequence of the chloroplast genome of C. reinhardtii has not been fully determined. The experimental value of this sequence is so extraordinarily high, that we anticipate that it will be completed in the near future. There also exists an intriguing biological barrier to arbitrary alteration of the chloroplast genome in C. reinhardtii. It has not yet proven possible to recover mutations that abolish the function of any component of the chloroplast gene expression system, although nuclear gene mutants with diminished levels

Laurens J. Mets and Jean-David Rochaix of (though not absent) chloroplast ribosomes have been characterized in some detail (Harris et al., 1994). The identification of a chloroplast tRNA suppressor mutation that could only be maintained by heteroplasmic complementation with the wildtype version of the gene (Zhang and Spreitzer, 1990; Yu and Spreitzer, 1992) underscores the essential nature of chloroplast protein synthesis, even under growth conditions that do not require photosynthesis. It is possible that the chloroplast genome of C. reinhardtii carries a gene that is essential for intermediary metabolism ofcells growing on acetatesupplemented media as well as those known genes that are necessary for photoautotrophic metabolism. On the other hand, it is possible that the apparently essential role of the chloroplast gene expression system reflects the operation of a higher level checkpoint control that integrates chloroplast biogenesis with the overall cell cycle. The real answer awaits further experimentation. A technical approach that may help to determine the functions of apparently essential chloroplast genes is to develop an inducible/ repressible chloroplast gene expression system. This could be used to experimentally turn off the essential genes to determine how cellular metabolism is perturbed in the absence ofthe corresponding product. A naturally occurring chloroplast gene whose expression responds selectively to environmental cues has not been discovered, and it may be necessary to generate an artificial system using components from a characterized gene control system from another organism.

2. Mitochondrial Genome Engineering Experience with mitochondrial gene transformation in C. reinhardtii is relatively more recent than that with chloroplast transformation (Randolph-Anderson et al., 1993), but also indicates that exogenous DNA can be incorporated into the genome by homologous recombination (Chapter 34, Remacle and Matagne). There do not appear to be major technical barriers to the development of a full capability for engineering the mitochondrial genome, though transformation is rare and the requirements for engineering exogenous gene expression have not yet been worked out. The question arises whether some mitochondrial genome function might be essential for cell growth and division when the encoded respiratory activities are, themselves, not. In this case, the full 15.8 kb genome sequence from C. reinhardtii has been

Chapter 36

Perspectives

determined (Chapters 5, Nedelcu and Lee; 34, Remacle and Matagne), and includes genes for one subunit each of the cytochrome oxidase complex and the cytochrome complex and five subunits of the NADH dehydrogenase complex. The only other translatable sequence encodes a reverse transcriptaselike protein. Matagne’s group has isolated mutants deficient in each of the respiratory complexes (Chapter 34, Remacle and Matagne), and all are viable under phototrophic growth conditions, though the NADH dehydrogenase-deficient mutants are slow growing. Nevertheless, it is clear that some more extensive deletion mutations, including deficiency in the entire genome (Alexander et a.l., 1974; Gillham et al., 1987), are non-viable. Just as for the chloroplast genome, the essential feature of the mitochondrial genome has yet to be delimited and the rationale for its essentiality is still obscure.

3. Nuclear Genome Transformation, Analysis and Engineering Of all the unique tools that have been developed for genetic manipulation of Chlamydomonas, it is nuclear gene transformation that provides the broadest avenue for pushing our understanding of eukaryotic photosynthesis in new directions. The era of gene discovery for organellar genomes has now largely passed. What lies ahead is the equally important and challenging problem of investigating structurefunction relationships in their genes, whose identities are now known. However, numerous physiologic studies have defined regulatory and integrative processes whose mechanisms are not encoded in the organellar genomes. Previous screens for nonphotosynthetic mutants in C. reinhardtii have indicated the existence of hundreds of nuclear genes that are otherwise unknown. The proportion ofnuclear genetic information that is exclusively involved in photosynthesis or organelle biogenesis is at least 1.5 percent, based upon the proportion of random nuclear mutants that are non-photosynthetic (Krebtukova and Spreitzer, 1996). The analytic tools provided by nuclear transformation promise to enable detailed identification and then functional characterization of presently unknown genes with vital roles in photosynthesis. Among photosynthetic organisms, nuclear gene transformation methods per se are not unique to Chlamydomonas. They have been extensively developed for numerous land plants and have emerged as a prominent tool of commercial

693 crop variety development programs. However, for the reasons outlined above, none of these plants is as suitable for mutational analysis of photosynthesis as is C. reinhardtii. The status of nuclear genome transformation technology in C. reinhardtii is thoroughly reviewed by Kindle in Chapter 4. Here we wish to emphasize the salient features that are the underpinnings of its value for discovering gene function in photosynthesis. The most important experimental finding is that introduced DNA incorporates into the recipient cell genome principally by non-homologous recombination rather than by homologous recombination. While the details of this presumptively random insertion process have not been worked out, it is clear that many insertions occur within genes in ways that disrupt their function. In principle, the molecular ‘tag’ generated by the inserting DNA can be exploited for molecular cloning of the affected gene. While the results of actual insertional gene tagging experiments are a bit more complicated than this simple concept (Chapter 4, Kindle, and references therein), the approach is so important that we can expect rapid refinement based upon broad experience in many laboratories. A second vital experimental advance is that transformation frequencies are now high enough to support transformation analysis with clone libraries. This is important in two ways. First, a strain bearing a recessive mutation can be transformed with pools of cloned wild type DNA in order to identify the affected gene by complementation (Purton and Rochaix, 1994; Zhang et al., 1994). Experience with this practice is still at an early stage and strategies for distinguishing between suppression and complementation are laborious. Recently a high-efficiency method for transformation of C. reinhardtii by electroporation has been established in which the frequency of transformation is increased by two orders of magnitude as compared to the standard glass bead transformation method (Shimogawara et al., 1998). This new method appears to be very promising for gene rescue experiments. As pointed out by Kindle (Chapter 4), complementation cloning offers the possibility of stepping beyond one of the limitations of insertional mutagenesis. Intrinsically, insertional mutagenesis affects only the level of expression of the affected gene (usually to zero) rather than the quality of the encoded protein, as can be affected by chemical mutagenesis. Many regulatory genes may be essential and hence

694 unrecognizable by insertional mutagenesis, but detectable on the basis of altered phenotypes caused by amino acid substitutions. Hence, the spectrum of genes that can be characterized by chemical mutagenesis and complementation cloning is expected to be wider than the spectrum identifiable by insertional mutagenesis. On the other hand, complementation cloning is effective only for the recovery of genes defined by recessive alleles that have reversion/suppression frequencies below the transformation frequency and have counter-selectable (non-dividing under some condition) phenotypes. While these criteria are met by many strictly nonphotosynthetic mutants, they may not be met by mutants with more subtle physiologic alterations. As the transformation frequency becomes higher, this constraint is lessened. In fact, selection for the transformation event itself using a selectable marker on the cosmid library vector, e.g. Arg7 (Purton and Rochaix, 1994) or Nit1, followed by testing for complementation of the mutant allele may allow complementation cloning by screening methods. High transformation frequencies are also important to support studies of mutation libraries created in cloned genes in vitro. Once a null mutation in a gene has been isolated (e.g. by insertional mutagenesis), the wild type version of the gene can be mutagenized in vitro and then reintroduced to assess the effect of the mutation. While testing of specific structurefunction hypotheses may require making only one change in the gene in this type of experiment, in the exploratory stage of gene analysis, random in vitro mutagenesis followed by phenotypic selection or screening is likely to be extremely valuable. No experiments of this type have yet been reported for nuclear genes in C. reinhardtii, but they now seem feasible. Another hard-won advance in the capabilities for nuclear genome engineering in C. reinhardtii is the definition of modular gene expression control elements with sufficient precision to enable engineering of controlled expression of foreign genes (Chapter 4, Kindle). To date, these efforts have focused on the development of genes with selectable phenotypes for use as transformation markers and on reporter genes for use in analysis of gene expression controls. However, the ability to engineer expression of foreign genes, including enzymes that can be targeted to a particular cellular compartment (Chapter 13, Perret et al.) and whose expression can be either turned on or off, depending upon culture conditions

Laurens J. Mets and Jean-David Rochaix (Chapter 4, Kindle) introduces a very broadly important technique for studies of photosynthesis. With this technique, it is now possible to contemplate testing metabolic control systems by inserting foreign enzymes that would interfere with metabolite flow (see, for instance, Hallman and Sumper, 1996). The opportunities for exploiting engineered expression of foreign genes in C. reinhardtii appear quite open ended. A vital and fascinating biological phenomenon that remains to be understood in order to be predictably controlled is the appearance ofepigenetic silencing of the introduced genes during vegetative growth of the cells (Cerutti et al., 1997; Chapter 4, Kindle).

4. Advances in Colony Screening Methods There are many sensitive measurement techniques that are capable of characterizing subtle aspects of photosynthesis physiology and its regulation in intact unicellular organisms like Chlamydomonas (Chapter 22, Joliot et al.). The models generated from physiologic analysis can be tested, sustained or rejected, and extended by genetic and molecular genetic analysis, provided that the tools are available for generating and finding the appropriate mutant cells. Making mutants is not difficult, but finding them if they are not simply dead or alive under some condition is more of a challenge. Fortunately, many aspects of photosynthesis physiology affect, either directly or indirectly, dynamic changes in chlorophyll fluorescence yield in intact cells (for reviews of different aspects, see Dau, 1994; Govindjee, 1995; Schreiber et al., 1995). Recent advances in the sensitivity and temporal resolution of CCD detector video imaging allow time-resolved measurements of fluorescence yields from individual colonies of Chlamydomonas (Chapter 23, Bennoun and Béal). This brings dynamic analysis of photosynthesis physiology into the realm that makes it useful for identification of rare mutants among large numbers of non-mutant colonies. The approach has recently been applied to initiate a genetic analysis in Chlamydomonas of the regulation of non-photochemical quenching (Niyogi et al., 1997a,b) and state transitions (M. Fleischmann and J.-D. Rochaix, unpublished). The mutants obtained would have been extremely difficult to identify without this mass scale, subtle, non-invasive analytic technique. In addition to imaging of variations in steady state fluorescence from algal colonies, the technology is also available

Chapter 36 Perspectives for imaging colonies based directly upon chlorophyll fluorescence lifetime, either in the time domain or the frequency domain (Wang et al., 1992). Similar time or frequency gating techniques may also prove a valuable modification of colony imaging absorption spectroscopy (Arkin et al., 1990) for identifying colonies with altered time-dependent absorption changes.

C. Constraints that May Be Surmounted by Further Work Nuclear genes encoding subunits of the photosynthetic complexes are often identifiable by sequence before any mutants have been recovered. These genes are candidates for reverse genetics in which mutations are constructed in vitro and then used to replace the endogenous gene by homologous exchange. Preliminary attempts have been made to identify factors that control the ratio of homologous to nonhomologous integration in Chlamydomonas (Sodeinde and Kindle, 1993;Gumpel et al., 1994; Kindle and Sodeinde, 1994; Nelson and Lefebvre, 1995), but clear success in controlling insertion is still rare, even with strongly selectable phenotypes (Chapter 4, Kindle). As more and more genes are identified in plants by large scale sequencing projects, either from genomic DNA or from cDNA, the opportunity as well as the need for functional analysis through reverse genetics will continue to increase. Although precise nuclear gene replacements or disruptions have not been achieved in C. reinhardtii, it has been possible to recover deletions of specific nuclear genes in at least four cases: AtpC (Smart and Selman, 1991), PsaF (Farah et al., 1995), RbcS (Khrebtukova and Spreitzer, 1996) and Nit8 (Nelson and Lefebvre, 1995). These deletions are accompanied by other genomic rearrangements, and the molecular mechanism that generates them is still unknown. Nevertheless, they are excellent null mutations that can be used as recipients for modified versions of the wild type gene. The first step in such an analysis is to determine that transformation with the wild-type gene restores the wild-type phenotype. This has been achieved recently with the PsaF deletion mutant, and several site-directed mutations were analyzed (Hippler et al., 1998). The mutant gene copies were expressed and mutant PS I complexes could be isolated. Some of the mutations were shown to affect electron transfer from plastocyanin or cytochrome to PS I. Hence, reverse genetic analysis of nuclear

695 genes in C. reinhardtii is becoming more feasible. It is expected that a number of other nuclear genes involved in photosynthesis will soon be subjected to the same type of analysis. For genes defined by mutant alleles that do not fit the criteria for complementation cloning, an alternative approach is positional cloning. The general notion is that locating the mutation on a genetic map that is correlated with a physical DNA map of the same chromosome can yield a clone containing the gene. In Chlamydomonas, one cM genetic distance (1% recombination) corresponds, on average, to 50– 70 kb of DNA (Chapter 3, Silflow) with at least one dramatic expansion to the order of 1500 kb/cM in the vicinity of the mating type locus (Ferris and Goodenough, 1994) and a considerable contraction at the ARG7 locus to 6 kb/cM (Debuchy et al., 1989). At present, genetic maps of both phenotypic and molecular markers are still at relatively low resolution compared with the needs of positional cloning and a major effort at physical mapping has begun (Chapter 3, Silflow). However, results from both types of mapping effort are cumulative and we can hope that a combination of physical mapping of large genomic clones, like bacterial artificial chromosomes (BACs) and yeast artificial chromosomes (YACs) (Infante et al., 1995), and higher density genetic mapping of molecular markers will eventually build a correlated physical and genetic map of sufficient resolution to bring researchers close to a gene with minimal mapping effort. When starting from a small segment of DNA, the chances of success in complementation cloning increase considerably, even if positional cloning doesn’t narrow the location of the mutation to a single gene. A reliable method for diverting mitotically dividing diploid cells of Chlamydomonas through meiosis would extend the range of genetic analysis in this organism to include genes that are essential under all growth conditions. An approach of this type would provide, for instance, a valuable tool for clarifying what functions of the organelles are essential for cell division even when their major metabolic functions in energy metabolism are not essential. In the absence of a known environmental manipulation that could induce this transition in the life cycle, Dutcher attempted to exploit abortive zygote formation for this purpose (Dutcher, 1988). The results provided interesting insights into the regulation of meiosis, but the procedure itself has not come into broad use. A reinvestigation of this problem seems in order.

696 A major advance in capabilities for nuclear genome engineering and analysis would be obtained by the development of an autonomous replicon that could support the replication and transmission of introduced genes without the need for integrating into the recipient genome. Unfortunately, no natural infective, plasmid or episomal nuclear vectors are known in Chlamydomonas. On the other hand, experience with such vectors in other organisms, principally yeast, provide a blueprint for their engineered design and no technical barriers to their construction is apparent. Two strategies seem directly feasible. First, the only non-host protein that is necessary for the S-phase limited replication and maintenance of a high copy circular DNA molecule like the plasmid in yeast, is the FLP recombinase (Broach et al., 1982; Senecoff et al., 1985), though other plasmid genes influence the fidelity of transmission in yeast. Engineering expression of the FLP recombinase in Chlamy­ domonas now seems feasible, and it need not be carried on the vector. The other necessary components of such a vector are an origin of replication, an inverted repeat of the FLP recombinase recognition site, and a selectable marker. A vector carrying the latter two could be used for cloning an origin of replication from Chlamydomonas based upon the phenotype of stable maintenance of the vector as an independent replicon. A second strategy would involve construction of an artificial chromosome from Chlamydomonas components. The necessary parts are an origin of replication, telomeres (Hails et al., 1995), a selectable marker, and a mitotically functional centromere sequence. Only the latter is presently unavailable, but it is reasonable to expect a positional cloning strategy to succeed for this purpose, since tetrad analysis can be used to closely define the genetic position of centromeres.

D. Comparison with Cyanobacteria Many of the chapters in this volume focus on using Chlamydomonas for detailed structure-function analyses of individual enzymes and multiprotein complexes of the photosynthetic apparatus (Chapters 15, Erickson; 16, Ruffle and Sayre; 17, Webber and Bingham; 24, Wollman; 25, Strotmann et al.; 26, Jacquot et al.; 27, Spreitzer). With the exception of the function of LHC antennae and of the cytochrome complex (Chapter 24, Wollman), comparable structure-function analyses can be conducted in cyanobacteria, particularly in Synechocystis sp.

Laurens J. Mets and Jean-David Rochaix PCC6803, in which homologous gene replacement is facile (see The Molecular Biology of Cyano­ bacteria, Volume 1 in this series, D. A. Bryant, editor, 1994). Recently, the complete genome sequence of Synechocystis sp. PCC6803 has been determined (Tanaka et al., 1996), which provides a detailed guide useful for planning such studies. However, heterotrophic growth of this cyanobacterium is rather slow in comparison with C. reinhardtii. For chloroplast-encoded proteins, the molecular genetic tools necessary for structurefunction analyses are at least as good in Chlamydomonas as they are in cyanobacteria. However, for subunits of these complexes that are encoded in the nucleus in Chlamydomonas, the molecular genetic tools for structure-function analysis are still under development and not quite yet at the same level of productivity as those available in cyanobacteria. An exception is that genetic complementation analysis is a natural process for nuclear genes in Chlamydomonas, but artificial and not always reliable in cyanobacteria. Of course, the great strength of parallel studies in both Chlamydomonas and cyanobacteria is that the comparison provides the perspective of 1–2 billion years of evolutionary separation, with Chlamydomonas more closely related to land plants.

III. Forefront Problems in Photosynthesis and Organelle Research The main steps and components in the central pathways of photosynthesis have been known for some time. There are only a few persistent mysteries, such as the identification of the electron carrier(s) that mediate PS I-driven cyclic electron transport, the non-structural function of the cytochrome complex (Chapter 16, Ruffle and Sayre), and the role ofthe chloroplast NAD(P)H dehydrogenase complex. Attention is now focused on determining the detailed mechanisms of catalysis by each of the protein complexes and on the mechanisms that regulate and coordinate the overall process, from biogenesis to biochemical activity.

A. Structure-Function Analysis of Individual Complexes The establishment of an efficient chloroplast transformation system in C. reinhardtii together with the ability to perform precise chloroplast gene surgery

Chapter 36 Perspectives (Chapter 8, Goldschmidt-Clermont) has opened the door for an in-depth structure-function analysis of the photosynthetic complexes PS II, PS I, the complex, ATP synthase and Rubisco cytochrome (Chapters 15, Erickson; 16, Ruffle and Sayre; 17, Webber and Bingham; 24, Wollman; 25, Strotmann et al.; 27, Spreitzer) and also of the soluble electron carriers. Although this work is still at an early stage, new and important insights have been gained by combining molecular genetics, biochemical and biophysical approaches. The atomic structures have been determined for the reaction center from the purple bacterium Rhodopseudomonas viridis (2.3 Å resolution Deisenhofer et al., 1995), for the PS I reaction center from the cyanobacterium Synecho­ coccus elongatus (4.0 Å resolution, Schubert et al., 1997), for the beef mitochondrial cytochrome complex (2.9 Å resolution, Xia et al., 1997) and synthase (2.8 Å resolution, Abrahams et al., 1994), and for spinach Rubisco (1.6 Å resolution, Andersson, 1996). Structures have also been determined for plastocyanin (1.5 Å resolution from Chlamydomonas, Redinbo et al., 1993), ferredoxin (2.5 Å resolution from Spirulina platensis, Fukuyama (1.9 Å resolution et al., 1995) and cytochrome from Chlamydomonas, Kerfeld et al., 1995). Because of the conserved structure ofthese energy-transducing systems the structural data are directly applicable to Chlamydomonas and will greatly help in elucidating the structure-function relationship of these complexes in combination with site-directed mutagenesis and functional analysis. Recent studies on the PS I reaction center subunits PsaA and PsaB have allowed the identification of the amino acid side chain ligands to (Chapter 17, Webber and the primary donor Bingham; Redding et al., 1998). It is expected that the ligands of the primary electron acceptors and will soon be identified. Site-directed mutagenesis of psaC and PsaF from C. reinhardtii has also provided new insights into the acceptor and donor sides of PS I and its interactions with ferredoxin and plastocyanin/cytochrome c6, respectively (Fischer et al., 1998; Hippler et al., 1998). The recently established atomic structure of the soluble part of cytochrome f of turnip (Martinez et al., 1994) has greatly stimulated site-directed mutagenesis studies (Chapter 24, Wollman) and efforts are underway for determining the structure of the cytochrome complex from C. reinhardtii (Mosser et al., 1997). Rapid progress in the determination of atomic structures of photosynthetic complexes can be

697 expected in the near future. There is little doubt that the combination of atomic structure analysis with site-directed and region-directed mutagenesis in Chlamydomonas will be at the forefront in generating understanding of how these complexes function at the molecular level.

B. Integrative Aspects of Chloroplast Biogenesis 1. Organellar and Nuclear Genome Cooperation and Integration The analysis of numerous mutants of C. reinhardtii deficient in photosynthetic activity has revealed a complex interplay between the nuclear and chloroplast genetic systems. The salient features of these interactions are first the existence of a large number of nuclear loci required for the proper expression of chloroplast genes. Second, these loci encode factors that appear to be imported into the chloroplast where they act in a gene-specific manner. Third, these factors are required at various post-transcriptional steps including RNA stability (Chapter 9, Nickelsen), RNA processing (Chapter 10, Stern and Drager), RNA splicing (Chapter 11, Herrin et al.) and translation (Chapter 12, Hauser et al.). Recent work has identified the targets of some of these factors in the 5´- and 3´untranslated regions of chloroplast mRNAs. The genes of several of these factors have been cloned using genomic rescue of the mutants with cosmid libraries or gene tagging through transformation (Chapter 4, Kindle). Overall we still know very little of the details about how these proteins interact with their target RNAs or how they modulate chloroplast gene expression. It is clearly important to characterize these RNA-protein interactions further and to examine how they are influenced by different environmental and stress conditions. Several proteins have been found to bind to the 5´and 3´-untranslated regions of chloroplast mRNAs and recent studies suggest that proteins with molecular masses in the 46–47 kDa range play a role in chloroplast translation (Chapter 12, Hauser et al.). A multiprotein complex including a 47 and a 60 kDa protein has been proposed to specifically regulate translation of the psbA-mRNA in response to light. The light signal is mediated through the photosynthesis-generated redox potential (Danon and Mayfield, 1994). Recently, the genes of these two proteins have been cloned and found to encode a polyA-binding protein and disulfide isomerase,

698 respectively (Kim and Mayfield, 1997). Several of these RNA-binding proteins, including the 47 kDa protein, have been shown to be bound to low-density chloroplast membranes associated with thylakoid membranes, but with properties similar to those of the inner chloroplast envelope membrane (Zerges and Rochaix, 1998). These results suggest that the translation ofseveral chloroplast mRNAs ofthylakoid proteins may occur at either a subfraction of the envelope membrane or in a previously uncharacterized intra-chloroplast compartment. It is interesting that a similar association of translational activator proteins with the inner mitochondrial membrane has been found in Saccharomyces (for review, see Fox, 1996). It remains to be shown whether the membrane association of these proteins observed in Saccharomyces mitochondria and in Chlamydomonas chloroplasts reflects a compartmentalization of processing, translation and turnover of organellar RNAs. The interactions between the inner envelope membrane and the thylakoid membrane remain a fascinating area of research (Chapter 19, Hoober et al.; Chapter 21, Trémolières). Extensions of the inner envelope membrane and membrane vesicles in the stroma have been observed under particular conditions in C. reinhardtii (Chapter 19, Hoober et al.) and in higher plants (Morre et al., 1991) suggesting the possibility of vesicular trafficking between the inner envelope and the thylakoid membranes. In this respect it is particularly intriguing that in Arabidopsis a protein has been identified that appears to be involved in the biogenesis of thylakoid membranes and that belongs to the dynamin family (Park et al., 1998). Dynamins form a large group of GTP-binding proteins that are generally involved in trafficking of materials in eukaryotic cells and more specifically in vesicle formation during vacuolar protein sorting in Saccharomyces (Rothman et al., 1990). Since the steps of the chlorophyll biosynthesis pathway after Mg-protoporphyrin IX occur on the chloroplast envelope (Joyard et al., 1991), chlorophyll needs to be transported to the thylakoid by a mechanism that is still unknown. If synthesis of chlorophyll-binding chloroplast-encoded proteins occurred on the envelope membrane, chlorophyll could bind to them there and this could facilitate the coordination between these two processes. It is clearly important to characterize the low-density membrane system more thoroughly in Chlamydomonas. One problem is that known secretory pathways are usually too fast to detect proteins in intermediate stages. A

Laurens J. Mets and Jean-David Rochaix challenging problem will be to develop genetic screens for mutants affected in this process. Such an approach has been very rewarding in Saccharomyces. Although chloroplast transformation technology has provided important insights into chloroplast gene expression, we need to complement the in vivo studies with reliable in vitro assays for RNA stability, RNA processing and translation. The recent establishment of an in vitro plastid translation system with tobacco (Hirose and Sugiura, 1996) opens promising possibilities for a thorough analysis of chloroplast translation and its relation to RNA stability and processing. One basic problem in organellar biology is to understand how the different subunits of multiprotein complexes are assembled along with their necessary pigments and cofactors into functional membrane protein complexes. The challenge is to discover not only how they are put together, but also how the production of the various parts of each complex is coordinated. Biochemical and genetic studies with numerous mutants have shown that unassembled subunits are rapidly degraded (Chapters 12, Erickson; 17, Webber and Bingham; 24, Wollman). An additional regulatory mechanism has been shown to complex where operate for the cytochrome cytochrome f synthesis depends on its immediate interaction with other subunits of the complex (Chapter 24, Wollman). Similar mechanisms may function for the other photosynthetic complexes. Recently, the conserved chloroplast open reading frames ycf3 and ycf4 have been shown to be required for assembly of PS I in C. reinhardtii and, in the case of ycf3, also for tobacco (Boudreau et al., 1997; Ruf et al., 1997). A challenge is to understand how these two chloroplast-encoded factors act and to identify the corresponding assembly factors for the other photosynthetic complexes. The role of lipids in the assembly of some of the photosynthetic complexes appears to be important, but has not yet been explored thoroughly (Chapter 21, Trémolières). Little is known about the molecular mechanisms that control protein turn-over in the chloroplast, with the exception ofD1 protein turnover within PS II complexes under photoinhibitory conditions (Chapter 30, Keren and Ohad). Although the chloroplast clpP gene encodes a protein related to the ClpP subunit of the E. coli ATP-dependent protease, its function in the chloroplast has not yet been characterized, except that the protein appears to be essential for cell viability (Huang et al., 1994). Although it is indisputable that the nucleus is

Chapter 36 Perspectives heavily involved in chloroplast gene expression, it is also becoming apparent that the chloroplast can influence nuclear gene activity. Earlier studies with higher plants revealed that a functional plastid is required for the expression of nuclear genes encoding proteins ofthe photosynthetic apparatus. These genes are not expressed in plants containing defective plastids due either to the loss of carotenoids, which leads to photobleaching of the plastids in the light, or to plastid ribosome deficiency (Taylor, 1989). These experiments suggested the existence of a plastidderived signal that regulates in some way the transcription of these nuclear genes. Genetic analysis in Arabidopsis has identified several nuclear loci involved in this signaling pathway from the plastid to the nucleus (Susek et al., 1993). In Chlamydomonas the chlorophyll precursor Mg-protoporphyrin IX methylester and its immediate precursor appear to be involved in the regulation of accumulation of several mRNAs of nucleus-encoded chloroplast proteins although apparently in opposite ways. A first study suggested that this chlorophyll precursor(s) inhibits light induction of Cab and RbcS mRNAs (Johanningmeier et al., 1984; Johanningmeier, 1988). Recently it was shown that Mg-protoporphyrin IX acts as a signal in the light-induced expression of two nuclear heat-shock genes of C. reinhardtii (Kropat et al., 1997). This observation opens promising possibilities for identifying other components of the signal transduction chain from the chloroplast to the nucleus. The presence of both mitochondria and chloroplasts is a unique feature of plants and eukaryotic algae. It is well established that several biochemical pathways, e.g. photorespiration, use both the chloroplast and mitochondrial compartments. Recent work has shown that the reducing power generated by PS II can be shuttled to the mitochondrial compartment in the absence of PS I (Chapter 18, Redding and Peltier). The identification of the electron carriers involved in this pathway remains a challenging problem for which experimental analysis using Chlamydomonas offers unique advantages.

2. Unusual Features of Organellar Gene Expression Systems Besides their importance for understanding organellar biogenesis, the chloroplast and mitochondrial gene expression systems of C. reinhardtii have several unusual features that have interesting evolutionary implications (Chapters 5, Nedelcu and Lee; 11, Herrin

699 et al.). A striking example is provided by the chloroplast psaA gene, which comprises three exons that are widely separated on the chloroplast genome and that are transcribed separately. Assembly of the mature psaA mRNA requires two trans-splicing reactions (Kück et al., 1988; Choquet et al., 1989). Of particular interest is the tripartite structure of the first psaA intron in which the middle part encodes the trans-acting tscA RNA (Goldschmidt-Clermont et al., 1991). An intriguing question is whether this split intron structure reflects an intermediate stage in evolution between group II introns (in which the splicing is catalyzed in cis by the different intron domains) and eukaryotic introns (in which the splicing is catalyzed by trans-acting RNA-protein complexes, the snRNPs). It is still not known what type of proteins are associated with the tscA RNA and whether they are related to any of the eukaryotic snRNP proteins. Another unusual property of the organellar genetic systems of Chlamydomonas is the scrambled rRNA gene organization in the mitochondrial genome (Chapter 5, Nedelcu and Lee). It will be interesting to study how these mitochondrial ribosomes are assembled and how they function. The phylum Apicomplexa comprises a group of obligate intracellular parasites that often affect humans. It includes Plasmodium species, which cause malarial disease, Toxoplasma gondii, which causes toxoplasmosis, Eimeria species, which cause disease and economic loss in chickens and livestock, and Cryptosporidium species, which cause life-threatening diarrheal disease particularly in immunocompromised individuals. Recently, it has been found that these parasites harbor a plastid (Feagin, 1994; Wilson et al., 1994; Köhler et al., 1997) with functional ribosomes that are the target of some of the antibiotics that have therapeutic value (Beckers et al., 1995; Fichera et al., 1995). Some herbicides have already been shown to inhibit parasite growth (Stokkermans et al., 1996; Fichera and Roos, 1997) and many more should be considered as drug candidates for treating the diseases that these organisms cause. The relationship between Apicomplexan plastids and those of green algae is clearly supported by phylogenetic analysis of tufA genes (Köhler et al., 1997). The telomere sequences from nuclear chromosomes of the Apicomplexan Theileria parva are closely related to those of Chlamydomonas (Sohanpal et al., 1995). The mitochondrial genomes of these parasites contain scrambled rRNA genes like those of Chlamydomonas (Joseph et al., 1989; Vaidya et al., 1989). It seems clear that the level of

700 molecular genetic investigation that can be conducted in Chlamydomonas provides an important entrée for delving into metabolic pathways that may be shared between these parasites and algae, but absent from animals, and that hence may provide targets for chemotherapy. It is particularly intriguing to consider that the non-photosynthetic plastids of these parasites have been maintained through millennia ofevolution. Perhaps an understanding ofthe rather basic question of why Chlamydomonas plastid protein synthesis is essential, even in the absence ofphotosynthesis, may ultimately lead also to cures for some important diseases.

Acknowledgments LM is grateful for research grant support from the US National Science Foundation and from the US Department of Energy. JDR’s research was supported by grants from the Swiss National Fund and from the Human Frontier Science Program.

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Index

Symbols 460. See electrochemical gradient See Crp3:

222E 154–155, 157, 159

6.2z5 155

70S ribosomes 20

76-5EN 167

A A 330, 339

A´ 339

326, 329–330, 339, 690

326, 330–331, 341

AadA. See aminoglycoside-3´´ adenyl transferase

absorbance 464

absorption change 435, 440–442

absorption difference spectra

339

330

absorption spectrophotometry 434

absorption spectroscopy 440–443

ac-20 19–20

ac-206 15–16

ac-208 15

ac-21 17

ac-80 17

ac-u-g-23 97

ac115 206–207

ac20 186, 189

ac206 250, 603

ac208 228, 600, 601, 603

Ac29 99

Ac40 mutant 240

Acanthamoeba castellanii 78

acceptor side 333

PS I subunits 333

Acetabularia mediterranea 73

acetate

effect on gene expression 201

growth on 2

acetate auxotrophs 15

acetate flagellates 2

acetate-requiring mutants 523

chloroplast mutations 518

light sensitivity 518

temperature-conditional 518

acriflavine 665, 666

actinic flash 439, 440–441

actinic light 437–438, 441, 446

actinomycin D 189

action spectra 440

acylase

chloroplastic 422

cytosolic 422

5´ adenosyl-3´-phosphosulfate 615

adg1

Arabidopsis thaliana 561

ADP-glucose 554, 557, 560, 561

ADP-glucose pyrophosphorylase 552, 557–560, 625

affinity chromatography 202

affinity labeling 488

agglutinins 4

ALA 379, 389, 604. See also aminolevulinic acid

Alad 400

alanine: aminotransferase 533, 540

ALAS. See aminolevulinic acid synthase

algae

red 552

algal collections 8

alkaline phosphatase

as a reporter 47

alkylation damage 132

allantoate 643

allantoicase 644

allantoin 643

allantoinase 644

allophanate 643

allophanate lyase 645

alternative oxidase 541, 664

alternative pathway 541

amino acid

assimilation 642–643

auxotrophs 14

transporters 642

amino acid oxidase 620, 641

aminoglycoside-3´´ adenyl transferase 51, 471

as selectable marker 45, 47, 141, 143–144, 147, 166

reporter 142, 156, 202, 204, 207

cassette 487

acid regulation of synthesis 399–400 acid dehydratase 604. See also porphobilinogen synthase acid synthase 380

aminotransferase 642

ammonium 627

assimilation 639–642

transport 618, 629, 639–640

amplifier

lock-in 438

amylase 556

555–557, 559

555

amylopectin 550–551, 553, 560, 562–564

amyloplast 552, 554

amylose 550–551, 553, 557, 560, 562–563

Anabaena variabilis 341

Anabena 332

Anacystis 14, 15

anaerobiosis 559

antenna 435, 436, 440, 445

mobile 577

chla/b 247

chlorophylls 328

332–333

minor 242–243

Index

706 antenna complex (Continued) peripheral 234

antheraxanthin 571–572, 624

antibiotic resistance

genetic map 125 antibiotic resistance cassette 306, 310 antibiotic resistance markers 124 antimycin A 679 antisense RNA 170 aphVIII Streptomyces rimosus as selectable marker 45 Apicomplexa 699 apocytochrome b 663 appressed membrane 260, 263, 267–268, 272, 577 appressed regions 238 APS. See 5´ adenosyl-3´-phosphosulfate APS kinase 615 Arabidopsis 314, 366, 439, 618, 639, 646, 651, 698–699 hcf109 154, 173, 175

adg1 561

hcf5 173, 175

starch synthesis 561

arg2 36 Arg7 52, 54, 694 as selectable marker 43, 53–54, 57 locus 695 Ars. See arylsulfatase arsenate 645 arylsulfatase 616, 619, 627, 629 as a reporter 46, 48, 56–58

as reporter 48, 653

gene expression 46, 50

Ascobolus immersus 15 ascorbate 16 aspartate amino transferase 641 aspartate/malate shuttle 643 Aspergillus nidulans 616, 650 assembly 698 cofactor-protein 607

cytochrome

ccs loci 605

cytochrome

complex 472

metalloprotein 599 Photosystem II complex 234

photosynthetic complex 698 assimilation nitrate 645 nitrite 645 purines 643–645 urea 645 AST. See L-alanine aminotransferase atomic structures 697 ATP 266, 277, 434, 436, 439, 625 of 488 ATP sulfurylase 615 ATP synthase 234, 244–245, 442, 478–494, 676–678 conformational coupling 490 energy-linked binding change mechanism 490 mitochondrial 483 rotational coupling 491 three-site binding change hypothesis 489 ATP synthesis chemiosmotic basis 13

ATP-dependent electrogenic pump 676–677, 681 ATP-driven gradient generator 680 ATP/ADP ratio 434, 436 Atp/atp genes expression 483–484 coordination 484 diurnal cycle 484 light 483 organ-specific 483 plastid-derived signals 483 ATP:urea amidolyase 645 atpA 471 gene cluster 482 gene expression 153–156, 160, 171, 175, 205–206 ATPase 435 17, 487 latent 492 light-triggered 492 Mg-dependent 487 atpB 144 as selectable marker 141 gene cluster 482 gene expression 153, 155, 160, 168–170, 174–176, 204–205 AtpC 55, 695 as selectable marker 44 gene expression 47 atpE gene expression 171, 204 atpF spinach 142 atpH gene expression 171, 175 atrazine 302 autonomous replication 120 auxotrophs acetate 15 amino acid 14 5-azacytidine 103 azide sodium 224

B b-heme 607. See also heme b

355 BAC. See bacterial artificial chromosome Bacillus subtilis 607 back electron flow 580–582 bacterial artificial chromosome (BAC) 37, 695 bacterial photosynthetic reaction center H subunit 294 L subunit 294 M subunit 294 bacteriopheophytin 294–295, 351 bacteriorhodopsin 479 barley 397 barrel

loop 6 519

basal bodies 3, 7, 235

basal body DNA 7

BBY

Photosystem II particle 290

bchB Rb. capsulatus 391, 393

Index bchD

Rhodobacter spp. 388

bchE

Rhodobacter spp. 390

bchH

Rhodobacter spp. 388

bchI

Rhodobacter spp. 388

bchJ

Rb. capsulatus 391

bchL

Rb. capsulatus 391, 393

bchM

Rhodobacter spp. 389, 390

bchN

Rb. capsulatus 391, 393

bchX

Rb. capsulatus 394

bchY

Rb. capsulatus 394

bchZ

Rb. capsulatus 394

bcy1 622

benzoquinone 435

betaine lipids 417

BF25 245, 313

bicarbonate 276, 303, 620

bilins 379

biogenesis 234, 248–250

cofactor 472

cytochrome complex 472

Photosystem II 257–259

biolistic method 671

biolistic transformation. See transformation: particle bombardment

biosynthesis

chlorophyll 259

cofactor

molybdenum 647

molybdopterin 646

biparental chloroplast gene transmission 94–110, 123

biparental zygospore clone analysis 124

ble 31

Streptoalloteichus hindustanus as selectable marker 45, 47, 51

bleomycin 45

blue light signal

gametogenesis 98

branching enzyme 555, 557, 563–564

brc1 388

bromacil 302

5-bromo-4-chloro-3-indolyl sulfate 627

brs1 388

bsbI

gene expression 175

buoyant density gradients

CsCl 116

C C-terminal proteolytic processing 302

C4 photosynthesis 14

pathway 381–383, 389

CA. See carbonic anhydrase

707 -dependent AT Pase 17, 487

Ca1 48, 54, 628

gene expression 46

CA1 locus 531–532, 534

Ca1/Ca2 gene expression 57

Ca2 48

gene expression 46

release 583

nuclease 103

Cab genes, 589

Dunaliella tertiolecta 213

CabII genes 29

CabII-1

gene expression 46–47, 50

CabII-1 promoter CabII 30

Cah1 620

Cah2 620

Cah3 54

calcium 311–312

signal transduction. See signal transduction: calcium

calmodulin 404

Calvin cycle 351, 358, 538, 552, 573

cAMP-dependent protein kinase 622

carbon

assimilation 13

availability

acclimation 530–544

fixation 237

inorganic 618, 629

transport 531

isotope disequilibrium 531

transport

inorganic 629

carbon dioxide 620, 627.

acquisition 530–544

concentrating mechanism 521, 530–544, 628

fixation 19, 21, 349, 350–360, 516

reduction cycle 625

carbonic anhydrase 48, 534, 621, 627–628

genes 29

periplasmic 534

carbonyl cyanide-m-chlorophenylhydrazone 350

carbonyl cyanide-p-trifluoromethoxyphenylhydrazone 350

carboxysome 536

carotenes 242

572, 586

258, 264, 572, 586, 589, 590

de-epoxidation 571

404

desaturase 404, 405

carotenoid 401–406, 624

402

402

404

biosynthesis 401–406 regulation 406

cryptoxanthin 402

eyespot 3

lutein 402

luteoxanthin 402

lycopene 404

Index

708 carotenoid (Continued)

neoxanthin 402 neurosporene 404 phytofluene 404 triplet state 442 violaxanthin 402 xanthophyll 405 zeaxanthin 402

catalase 256

cbn1 397 cbn1-113 370 CBP2 Saccharomyces cerevisiae 188 CbR 590 ccb strains 467, 607 CCCP. See carbonyl cyanide-m-chlorophenylhydrazone CCM. See

mechanism CCS1 (c-type cytochrome synthesis) 467, 605–606 Ccs1 as selectable marker 44 CCS2 467, 605–606 CCS3 467, 605–606 CCS4 467, 605–606 ccsA 388, 398 cell architecture 3 cycle 48, 118, 152–153, 543, 622, 629 starch synthesis 558 division 622 cycle 540, 541–542 morphology 235 size 3 toxicity 646 cell wall 3 glycoprotein 235 cemA (ycf10) gene expression 171 proton channel 491

244, 457, 477–494, 676–678

activation 493

assembly 484–485

atpA cluster 482

atpB cluster 482

deactivation 493

evolution 482

gene arrangement 482

high resolution electron microscopy 481

kinetic properties 487–488

missense mutations 485

molecular genetics 482–487

molecular weight 478

mutants 485–487

nucleotide binding sites 479

proton conductivity 493

proton slip 493

regulation 492–493

signal sequences 484–485

site-directed mutations 485–487

stalk 481

structure 478–482

model 480

three-dimensional 481–482

disulfide bridge 492

subunits 479

thiol modulation 492–493

tightly-bound nucleotides 493

transit sequences 484–485

two-dimensional crystals 481

translocation 223

ch42

Arabidopsis 388

chaperones

chloroplast 485

chaperonin 258

genes 517

charge recombination 454, 582

Charophyceae 552 chill-sensitive plants 579 chimeric gene 471 chimeric protein 268, 373 Chl. See chlorophyll See singlet chlorophyll See triplet chlorophyll Chl a. See chlorophyll a Chl a/b and Chl/carotenoid ratio 240 Chl b. See chlorophyll b Chl-proteins 589 CHL1 soybean 388 Chlamydomonas spp. C. culleus 6 C. eugametos 2, 68, 97, 108, 127-128, 130, 417, 621 C. gelatinosa 70, 80, 192 C. humicola 128 C. limicola 341, 338 C. moewusii 2, 69, 72–73, 97, 120, 127, 130 C. monoica 2, 101, 108, 131, 562–563 C. mundana 604 C. pallidostigmatica 128 C. pitschmannii 69, 71, 77, 79 C. smithii 6, 70–71, 604, 663, 668 C. zebra 192 Chlamydomonas Genetics Center 7 chlamyrhodopsin 590 chlB 364, 393 gene expression 401 chlD Synechocystis PCC 6803 388 chlH Synechocystis PCC 6803 388 chlI Synechocystis PCC 6803 388 chlL 132, 364, 393 gene expression 401 chlM cyanobacteria 389 chlN 364, 393 gene expression 171, 401 chloramphenicol 160 chloramphenicol transacetylase 373 chlorate resistance 55 chlorate-resistant mutants 645 Chlorella 14–15, 79, 435–436

Index Chlorella ellipsoidea 77, 80, 85, 600, 629 Chlorella pyrenoidosa 70, 536, 589 starch 553 starch synthesis 556 Chlorella sorokiniana 243–443, 456–457, 465, 677–679 Chlorella vulgaris 83, 530, 678, 689 heterotrophic growth 689 starch synthesis 556 chlorina-type phenotype 365 Chlorobium 358 Chlorobium limicola 325, 336–337 Chloroflexus 358 Chlorogonium elongatum 71 Chlorophyceae 416–417, 552 chlorophyll 256–257, 259, 264, 267, 288, 364, 379, 461 accessory 326 antenna 298, 328 biosynthesis 259, 379–383, 388–397, 406 regulation 398–401, 406 cofactor 461 dimer 296 excited states 687 fluorescence 437, 452, 455 lifetime imaging 695 fluorescence decay 298 geranylgeranyl Chlides 368 light harvesting a/b binding protein 308 molecule 461 orientation 17 transition moment 296 RC I 396 reaction center 298 red 332 special pair 295, 296 synthesis 343, 364, 518 regulation 398–401 triplet 582 chlorophyll a 14, 242, 367, 589 biosynthesis 388 chlorophyll a´ 396 chlorophyll a/b proteins 243 chlorophyll b 242, 365, 367, 395 biosynthesis 396–397 chlorophyll b-less mutants 397 chlorophyll synthetase 394 chlorophyllide 365, 391 chlorophyllide reductase Rb. capsulatus 394 chlorophyll Z 297–298 chloroplast 236 ATP synthase 477–494 chaperones 485 complementation 691 development 257, 342, 364 regulation of 200 envelope 618 proteins 221 fusion 247 morphology 236–238 NAD(P)H dehydrogenase 675, 678, 681 nucleoids 105

709 oxidase 675, 680–681 reporters 141–142 selectable markers 140–141 supramolecular organization 234–251 transcription 153 transformation 140–148, 521 transportation 139 tRNA suppressors 518 synthetase 198 ultrastructure 368 chloroplast DNA 17, 96, 116–123 degradation 98, 100–103, 1 1 9 deletion 97 density gradient centrifugation 97 destruction 98 fragmented coding regions 83 genetic map 123 heteroplasmicity 518 inheritance 95–98, 668, 669 methylation 96–97, 119, 132 mutagenesis 521 mutations 131, 518 protection 100–103 recombination 123–130 repair 130–133 replication 116–123 origins 120 restriction map 120 RFLP 97 stain 96 DAPI 97 synthesis 98 chloroplast gene evolution 82–85 inheritance 131 light regulation 153 order 79 recombination 124 size 69–78 translational regulation 197–214 transmission biparental 123 chloroplast genome 100 engineering 692 evolution 63–87 organization 79–82 chloroplast ribosome 124 ribosomal proteins 96, 198 ribosomal RNA 85 chloroplast thylakoid membranes 211–212 chloroplast transit peptide 654 chloroplastic acylase 422 chloroplastic transcription 20 chlororespiration 559, 575, 675–682 ChlZ 306 chromate 629 chromogenic 628, 629 reagent 628 chromosomal rearrangement 99 chromosomes 7 cia4 628 cia5 628

710 CIA5 531

circadian control 152–153, 168

circadian rhythm 4, 542, 543

cis-elements 199

cis-vaccenate 416

cladistic analyses 65

Clark electrode 439

classical genetic methods 516

cleavage

proteolytic 646

cloning

complementation 693–694

nuclear genes 51–54, 158–159

positional 695

Clostridium pasteurianum 336, 338

Clp protease (clpP) 71

clpP 71, 83, 145, 466, 698

CMS. See cytoplasmic male sterility

co-transcription. See RNA: polycistronic

See carbon dioxide specificity factor 517

cob 664, 666

coding region

fragmented 64

intron-containing 63, 82

mitochondrial rRNA

fragmented 84

scrambled 84

rRNA 85

scrambled 64

Codium 80

Codium fragile 70, 73

codon

bias 30–31

initiation 467

start 33

stop 33

usage

mitochondrial 664

cofactor 467, 472, 598, 651

binding 608

molybdopterin 646

collections algal 8

colony screening 694–695

complementation 133

analysis 690

chloroplast 691

cloning 693, 694

functional 37

in vivo 646

intragenic 646

nucleus 690

confocal scanning laser microscopy 368

contractile vacuoles 3

copper 386, 460, 599, 605, 641

deficiency 600–601, 603

sensor 603–604

transport 605

affinity 604

transporter 602

uptake 604

copper-responsive elements 603

Index

coprogen oxidase, See coproporphyrinogen III oxidase

coproporphyrinogen III oxidase 386, 598, 604

copy correction 127, 133

corrinoids 379, 385

Corynebacterium glutamicum 639

cosmid library 54

cotransformation 50, 145

cotranslational integration. See membrane: insertion: cotranslational

coupling factor

F-54 17

cox1 664

COX3 (yeast mitochondria)

translational regulation 209

COXI. See cytochrome oxidase subunit 1 amino acid

CP26 366, 575

CP27 314

CP29 314, 575

CP43 289, 292, 306, 576, 578, 584–585, 589

folding model 307

CP47 289, 292, 306, 311, 585, 589

folding model 307

CP54 223, 224, 229

cpDNA 116. See chloroplast DNA

See D1 protein:

CPOX 400. See coproporphyrinogen III oxidase

Cpx1 386, 602

CRlpcr-1 44

cross-linking 333, 482

plastocyanin 335

crp1 173, 175

Crp3

mutant 175

crp3 175

crs 1

Zea mays 192

crs2

Zea mays 192

Cry1

as selectable marker 44, 47, 54, 55

cryptic gene 642

cryptogamic plants 417

cryptopleurine 44

Cryptosporidium 699

crystal structure 469

Cu. See copper

cucumber 397

cultures

synchronously grown 19

cupric reductase 604–605

CuREs. See copper-responsive elements

cw15 arg7A 369

cyanobacteria 422, 472, 530, 696

fatty acid synthesis 422

Cyanophora 83

Cyanophora paradoxa 314, 483

Cyc6 602

gene expression 46, 50, 57–58

promoter 30

cyclic electron transport 334, 351, 354, 359, 436, 441, 444, 576,

578, 589, 696

cyclic photophosphorylation 17

cycloheximide 485

cysteine 626, 629

Index Cyt. See cytochrome CYT18 Neurospora crassa 188 cylochrome 600, 602 assembly 603, 605, 607 deficiency 606 cylochrome b 443, 667 cytochrome 256, 270, 292, 297, 304–306, 311, 582, 586–587, 696 cytochrome 461, 464–465, 469 absorbance Em 464 cytochrome complex 243–245, 250, 260, 349–360, 433, 436– 437, 442–443, 455, 459–474, 538, 539, 574–577, 580, 584, 588, 679–680, 690–691, 696 accumulation 470 assembly 2 1 1 , 462, 470–473 control by epistatic synthesis 471 cofactor chlorophyll 461 quinone 464 crystallization 462–463 degradation 472 dimerization 462 PetL 463 electron transfer 464 mutants 461, 463 purification 461 reconstitution 463 subunit composition 461, 466 trans-membrane organization 461 cytochrome complex 677, 679, 697 cytochrome c 294 oxidase 662–663, 667 cytochrome c´ 465 cytochrome 273 cytochrome 20 cytochrome 222, 335, 387, 460, 598–599, 602–605, 697 structure 460 translocation 229 cytochrome f 14–17, 248, 460–462, 464–465, 471, 605 absorbance Em 464 assembly mutants 469 biosynthesis 468 mutants 469 regulation 472 interactions with plastocyanin 464 leader peptide 467 positive patch 464 precursor 469 processing 467 soluble 462 structure 469 translocation 224, 226–227, 468 cytochrome oxidase 601, 605 cytochrome oxidase subunit 1 amino acid (COXI) 65 cytoplasmic male sterility 662

D D. See electron donor: D D-loops 121. See also displacement loops

711 Dl protein 249, 289, 292, 295, 3 1 1 , 468, 577, 579, 580–581, 583– 584, 586, 588 292 cleavage 583–585 degradation 583–584 folding model 293 radiolabeling 266 translation 266 elongation 266 regulation 266 synthesis 588 D2 protein 249, 289, 292–293, 295, 311, 576, 578, 580–581, 583– 586 folding model 294 D503sup1,2 206 DA 628 damage oxidative 401, 580 DAPI. See DNA Fluorochrome, 4, 6-diamindio-2-phenylindole dark reactions 624 DBMIB. See 2,5-dibromo-3-methyl-6-isopropyl-p-benzoquinone DCCD. See dicyclohexyl carbodiimide DCIP 16, 21 DCMU 17, 21, 125, 292, 302, 350, 354, 357, 437, 439, 453, 584 resistance marker 141 de-epoxidation 571 deamination 642 debranching enzyme 563, 564 deep-etching technique 245 degradation chloroplast DNA 119 cytochrome complex 472 photosynthetic complexes 607 delayed fluorescence 439, 451. See also luminescence delayed luminescence 452, 452–57 digital imaging 451–452, 457 density gradient centrifugation CsC1 96, 97 desaturases 416 envelope 422 desaturation fatty acid 270 destacking 234 detecting light 440–441, 446–447 detergent 461, 463 Hecameg 461 octyl glucoside 461 dextrin 555 dextrinase 556 DGDG 419 DGTS 419 Dhc 29 diacylglycerol-trimethyl-homoserine 415–417 diaphorase 644, 646–647 2,5-dibromo-3-methyl-6-isopropyl-p-benzoquinone (DBMIB) 350, 354, 357, 463–465, 576 dicyclohexylcarbodiimide (DCCD) 491 differential scanning calorimetry 553, 562 digital imaging 451–457 dihydroxyacetone phosphate 558, 625 dimer 462 pyrimidine 130 thymine 130, 131

712 dimethylallyl pyrophosphate 402

2,5-diphenylcarbazide 351

diploid cells 690, 695

PEG fused 100

vegetative 100

dipyrromethane cofactor 385

displacement loops 120

disulfide bridge 492, 502

disulfide isomerase 697

diuron 302. See DCMU

3,8-divinyl chlorophyllide 390

3,8-divinyl protochlorophyllide 390

DNA

basal body 7

chloroplast 17

damage 109

excision 96

integration

transformation 49–50

methylation 51

recombination 96

repair 103, 109, 130–133

excision 105

recombination 105

repeated 63, 73

stability

transformation 49

See chloroplast DNA

DNA fluorochrome,

4, 6-diamindio-2-phenylindole 97–98, 118

DNA polymerase 122, 123

DNP-INT 463

domains

transmembrane-spanning 650

dominance

minus 101

donor side 335

PS II 580

double beam spectrophotometer 441, 446–447

doubling times 689

dtpB gene expression 200–202

dum 665, See also mutations: mitochondrial

Dunaliella bardawil 624

Dunaliella marina

starch synthesis 557

Dunaliella salina 585, 590, 608, 624

Dunaliella tertiolecta 213, 538, 623

Dunaliella tertiolectra 623

dynamin 698

dynein 3, 512

genes 29

dypyrromcthane cofactor 385

dysprosium ions 298

E early light-induced protein (ELIP) 589–590

EF particle density 239

Eimeria 699

electrochemical gradient 358, 451–452, 455, 457, 465, 680

permanent 455–457, 676–677, 681

thylakoid 453

Index electrochemical proton gradient 434–435, 439, 477, 478

permanent 435–436, 442

electrochromic probes 435, 442

electrochromic shift 442

electrodes

stationary 440

electron donor

D 299

Z 299

electron microscopy 98, 235, 249, 260

electron nuclear double resonance (ENDOR) 328–329, 300, 339

electron paramagnetic resonance (EPR) 296, 328, 330–331, 341,

445

power saturation 300

pulsed relaxation 300

spectroscopy 277

electron spin echo envelope modulation (ESEEM) 341

electron spin polarized (ESP) EPR 331

electron transfer 13–15, 434, 436, 440–441, 465, 518

back 580–581

reverse 357, 680

cytochrome

complex 464

kinetics 465

plastocyanin to

335

electron transfer reactions 436

electron transport 13

cyclic 334, 351, 359, 444, 696

electron transport chain 440

mitochondrial 664

respiratory 357

electrophoresis pulsed field gel 36

electroporation 49. See also transformation: electroporation

electrostriction 444

BLIP. See early light-induced protein

elongation factors 198

chloroplast 198

Em 464, 602–603

cytochrome

464

cytochrome f 464

emetine 44

endonuclease 128, 157, 159–160, 165, 172, 174

DNA 670

site-specific 185

homing 128, 129

SceI 663

endonucleolytic cleavage 664

endoplasmic reticulum 236

ENDOR. See electron nuclear double resonance

endosymbiotic origin 65

energy dissipation

non-radiative 571

energy storage 443

energy transduction 367

energy transfer 333, 445, 577

fluorescence 366, 492

singlet 366

energy-linked binding change mechanism 490

enhancement effect 350

envelope 364, 368

membrane

chloroplast 419

inner 236

Index lipid biosynthesis 423

outer 236

epigenetic silencing 694

epistatic effect 648

EPR. See electron paramagnetic resonance

ery-u-1 102

erythromycin 125

resistance 96, 102

resistance marker 141

Escherichia coli 332–333, 607, 647, 651

recA 142

ESEEM. See electron spin echo envelope modulation

ESP. See electron spin polarized EPR

ethidium bromide 665, 666

ethoxyzolamide 532

ethylmethanesulfonate 131

etioplast

development 257

Euglena 14–15

Euglena gracilis 83, 268

RNA polymerase 167

evolution 2

482

chloroplast genes 82–85

chloroplast genome 63–87

fossil evidence 86

genome size 75

mitochondrial genes 82–85

mitochondrial genome 63–87

modes and tempos 63–64

nucleotide substitution rate 64

photohydrogen 17

Photosystem I 358

Photosystem II 358

synonymous nucleotide substitution

evolutionary lineages 65

EXAFS. See extended X-ray absorption and fine structure

spectroscopy

excitation

flash 444

excitation energy 333, 339

transfer 17, 258, 332, 691

diffusion limited model 332

trap limited model 332, 333, 340

exciton transfer 436

excitonic coupling 314

exo-carboxypeptidase 587

exonuclease 155, 159, 165, 172, 174, 176

exoplasmic layer 238

extended X-ray absorption and fine structure spectroscopy

(EXAFS) 301

extrachromosomal amplicon 122

extragenic nuclear mutation 108

eyespot 3, 236, 590

Ezy1 29, 105

Ezy2 108

F F-54 coupling factor 17

F-60 17

713 composition 483

genes 483

mitochondrial

three-dimensional structure 485

synthase 697

F14 mutant 240

F15 206, 208

F16 155, 175

F34 160, 206–208, 239, 250

F34suI 206–207

F34su3 239

F35 160, 206

F54 205–206

F64 206–208

F8 355

326, 331, 337

photoreduction

low temperature 332

338

FAD 641

farnesylpyrophosphate synthase 404

fatty acid 417–422

composition 419

desaturation 270

export 423

import 423

synthesis 422

polyunsaturated 425

synthetase 422

326, 331, 332, 337

FCCP 350. See carbonyl cyanide-p-trifluoromethoxyphenylhyrazone Fd 354. See ferredoxin FdUr. See 5fluorodeoxyuridine FdUrd. See 5fluorodeoxyuridine 378, 387

fermentation 559

ferredoxin 213, 333, 349–360, 492, 502–508, 511, 598, 697

ferredoxin gene. See Frx1

ferredoxin NADP oxidoreductase 506

ferredoxin-GOGAT 640

ferredoxin-NADP oxidoreductase 507–508

ferredoxin-NADP reductase (FNR) 354, 552

ferredoxin: NADP+ reductase 351

ferredoxin:plastoquinone oxidoreductase 351

ferredoxin-thioredoxin reductase (FTR) 492, 502, 506–508

ferrochelatase 397, 400

fertilizer 616–617, 638

flagellar configuration 65, 84

flagellar movement 590

flagellar roots 3

flash

detection 438, 441, 446

excitation 440, 444

spectrophotometer 443, 446–441

spectrophotometry 441

spectroscopy 461, 464

flavodoxin 333, 598

flip-flop recombination 127

FLP recombinase 696

fluorescence 367, 433, 436–438, 451, 694

chlorophyll 452, 455, 695

Index

714 fluorescence (Continued) delayed 451 emission spectra 571 low temperature 439 energy transfer 366, 492 Fm 573–574 imaging 452–457, 694 induction 437, 452, 573 kinetic measurements 437 kinetics 437, 453–454

lifetime imaging 695

measurements 446, 574

77K 574 quenching 571, 574 spectroscopy 340 steady state 573 yield 436–438 kinetic analysis 436–438 fluorescence microscopy 97, 103 5-fluorodooxyuridine 106, 121, 131–132, 144, 518 (FdUrd) 5-fluorouracil 131 FMN 641 FNR. See terredoxin NADP oxidoreductase; ferredoxin: reductase synthase 453, 455 folding model PsaB 340 foreign genes expression 37 fossil evidence 86 Fourier transform infrared spectroscopy (FTIR) 296, 301 FPP synthase. See farnesylpyrophosphate synthase fractionation studies 240 fragmented coding regions 64 chloroplast 83 mitochondrial 83 freeze-fracture 234–236, 238, 240–241, 243–244, 246, 250, 463, 577 fructose 1,6-bisphosphate 625 fructose-1,6-biphosphate 558 fructose-1,6-bisphosphatase 503 fruclose-6-phosphate 558 Frx1 506 frxC. See chlL FTIR. See Fourier transform infrared spectroscopy FTR. See ferredoxin-thioredoxin reductase FUD34 206–207 FUD39 228, 313 FUD44 228, 311 FUD50 676, 680 FUD6 173 FUD7 250 Fus1 27, 29, 31, 33, 99 fusion 234, 248 gamete 99 Fv (Fv = Fm-Fo). See variable fluorescence parameter Fv/Fm 574, 580 Fv/Fo 574 326, 331–332, 336–338

G

gamete

pseudo-plus 99

fusion 99

gametogenesis 4, 98, 131, 472

blue light signal 98

nitrogen starvation 98

GDH. See glutamate dehydrogenase

GE2.10 154, 155

gel mobility shifts 202

gene 29–30

29–30

CabII 29

carbonic anhydrase 29

chimeric 471

cluster 80

content 63, 70

conversion 124, 127–129, 133

dynein 29

evolution

chloroplast 82–85

mitochondrial 82–85

expression. See gene expression histone 29, 33 inactivation. See gene targetting inheritance chloroplast 131 light-harvesting complex protein 29

Nit1 30 order 63 chloroplast 79 mitochondrial 79 organization rRNA, scrambled 699 RbcS2 29, 30 rearrangement 63 mechanism 81 recombination chloroplast 124 rescue 693 rRNA 28

5S 28 sequence ribosomal RNA 64 small subunit ribosomal RNA 68 silencing 48, 51 size chloroplast 69–78 mitochondrial 69–78 tagging. See insertional mutagenesis targeting 54–56, 147 transfer 78 gene expression

Ars 46, 50 atpA atpB AtpC atpE atpH

153–156, 160, 171, 175, 205–206 153, 155, 160, 168–170, 174–176, 204–205 47 171, 204 171, 175

Ca1 46 Ca1/Ca2 57

Ca2 46 gabaculine 383, 602, 604 galactolipids 415, 419, 422–423

CabII-1 46–47, 50

Index cemA 171 chlB 401 chlL 401 chlN 171, 401 Cyc6 46, 50, 58 dtpB 200–202 foreign 37 metal responsive 600 Nit1 46–47, 50, 58 nuclear control 467 Oee1 50 Pcy1 50 pet genes 467 petA 155, 159, 171, 173, 175, 205 petB 154–155 petD 155, 157, 160, 172–173, 175, 204–206 PetE (Pcy1) 47 PetE 50 Por1 401 psaA 54, 153, 160, 171, 184, 191, 208 psaB 153, 155, 160, 169, 176, 206, 208 psbA 141, 153, 156–157, 160, 186, 189, 200–203, 205–208, 211, 249 barley 211 tobacco 203 psbB 54, 154–157, 159, 171, 211 psbC 155, 160, 206–208, 211, 249 psbD 153–160, 171, 175, 206–208, 211 psbI 171, 175 psbM 171 psbT 171 rbcL 153–156, 160, 168–170, 176, 200–202, 212 RbcS2 47 rps4 204 rps7 171, 204 rrn16 168–169 rrnL 153, 186, 189 Rsp3 58 transformation 50 trnE1 167, 169 tscA 171 TubA 50 TubA1 57 TubB2 46–47, 50, 56–58 tufA 153, 168, 200 ycf10 175 ycf9 171 zygote-specific 100, 109 genetic analysis 6–7, 607, 687, 690 genetic code 664 genetic drift 124 genetic map 6, 26, 36 antibiotic resistance 125 chloroplast DNA 123 genetic purification 691 genetics mitochondrial 662–672 reverse 139–148, 354 genome chloroplast 79 database 608 evolution 64–87 mitochondria 79, 662

715 organization chloroplast 79–82 mitochondria 79–82 size evolution 75 variation 70 genomic library 36 geotactism 564 geotropism 564 geranylgeraniol 395 geranylgeranyl Chlides chlorophyll 368 geranylgeranyl pyrophosphate synthase 403 geranylpyrophosphate synthase 404 GGPP synthase. See geranylgeranyl pyrophosphate synthase gidA. See chlN Ginko biloba 393 glucose 558 glucose-6-phosphate dehydrogenase 557 reporter 142 GluRS. See glutamyl-tRNA synthetase glutamate dehydrogenase 641 glutamate semi-aldehyde aminotransferase 604 glutamate synthase 640 glutamate- 1-semialdehyde aminotransferase (GSA-AT) 381, 383, 399–400 glutamine synthetase 512, 533, 541, 640, 641 glutamyl-tRNA. See glutamyl-tRNA reductase 381, 382 glutamyl-tRNA synthetase 381, 398 GluTR 399. See glutamyl-tRNA reductase glyceraldehyde-3-phosphate 558, 625 glycerate 539 glycerol-3-phosphate dehydrogenase 559 glycerolipid 415–429 biosynthetic origin 419 metabolism 424 polar 418 glycine decarboxylase 535 glycogen 550, 560, 564 cyanobacterial 554, 563 glycolate 538–539, 541 glycolate dehydrogenase 533, 538, 541, 543 glycolate oxidase 538 glycolysis 557–559 glycoprotein cell wall 235 glyoxylate 538 glyoxylate cycle 689 GOGAT. See glutamate synthase Golgi apparatus 236 GPP synthase. See geranylpyrophosphate synthase grana membrane 247 grana stacks 428 granal lamellae 260 grc1 388 green algae phylogeny 65 green seaweed 552 greening 248, 365–366, 368, 400 group I introns 82, 663 transpostion 670 group II intron 83, 664, 699

Index

716 growth

heterotrophic 261, 690

pholoautotrophic 261, 262, 273, 350–360, 355

GS. See glutamine synthetase

GS/GOGAT cycle 639

GSA. See glutamate-1-semialdehyde

Gsa 400

GSA-AT. See glutamate-1-semialdehyde aminotransferase

Gulliver. See transposable elements

GUS

reporter 142. See also uidA: reporter

gyrase 122, 123

H H subunit bacterial reaction center 294

H + ATP synlhase 552

H+-ATPase 532

H+-translocation ATPase 477

evolution 354

Haematococcus pluvialis starch 553

hairpin structure 664

half-life of protein 471, 601

Hansenula anomala 652

hcf106 224

hcf109 154, 173, 175

hcf5 173, 175

heat emission 571

Hecameg 461, 463

helical slacks 260

603

Helicobacter pylori 607

Heliobacillus mobilis 325, 336–338, 341

Heliobacteria 325, 358

hem 604

hem A

E. coli 382

heme 379, 600, 604–605

attachment 605

binding 469

biosynthesis 397–398

lyase 398

non-plastidic 398

plane 465

transporter 606

heme a 398

heme b 398, 464–465, 607, 641. See also b-heme

assembly 467

ligands 469

443, 469, 607

443, 469, 607

heme c 607

assembly 467–468

ligand 465, 469

hemG E. coli 387

hemY

B. subtilis 387

herbicide, 584

atrazine 302

bromacil 302

diuron 302

metribuzin 302

photobleaching 387

resistant mutants 302

heteroduplex 127

heteroplasmic 146, 166

transformants 144–145

heteroplasmicity

chloroplast 518

heterotrophic growth 261, 690, 696

Chlorella vulgaris 689

Scenedesmus obliqus 689

hexokinase 557, 558

hexose-phosphate translocator 557, 558

high affinity nitrate transport 649

high light stress 305

histone genes 29, 33

HMG-CoA 404. See 3-hydroxy-3-methylglutaryl- CoA

HMG-CoA reductase 402, 406

homing endonuclease 128–129

homing intron 128

homologous recombination 49, 54–56, 81, 126–127, 132, 648,

692

homoplasmic 100, 146, 670

homoplasmic transformants 144–145

homothallic 101

Hordeum vulgare 364, 618

Hsp70 promoter 30

hydrogen

photoevolution 359

hydrogenase 17, 354

3-hydroxy-3-methylglutaryl-CoA (HMG-CoA) 402

1-hydroxymethylbilane 384

hypermethylation 96

I

ICR-191 131

ida4 31

imaging

delayed luminescence 451–457

digital 452

fluorescence 452–457, 694

immunocytochemical 240

labeling 250

localization 241, 652

immunocytochemistry 262, 272, 540

immunoelectron microscopy 243, 369, 370

imp1 99

imp11 mutant 99

import

chloroplast 220–222, 271

in organello synthesis 265

in vitro

synthesis 265

complementation 646

inclusion bodies 485

inheritance 93–110

biparental 9 4 – 1 1 0

chloroplast DNA 95, 95–98

maternal 94–110

Mendel’s laws 94

organelle 94–110

Index paternal 94–110

preferential

polarity 128

uniparental 4, 93, 94–110

bidirectional 102

initiation codon 467

imitation factors 198

chloroplast 198

inner envelope membrane 236, 698

inorganic carbon 618, 620–621, 627

transport 531, 618, 629

plasma membrane 618

insertion sequences 83

insertional mutagenesis 7, 37, 51–54, 560–561, 648, 693

inside-out vesicles 246

intergenic regions 70

intermolecular recombination 82

internal transcribed spacers 85

intrachloroplastic pathway 422

intragenic complementation 646

intragenic suppression 519

intramembrane particles 235

intramolecular recombination 81, 82, 118

intron 58, 70, 128, 600

containing coding regions 63

cyclization 184

group I 82, 184–189

group II 77, 83, 190–193, 699

trans-spliced 82

homing 78, 128–130

insertion 77

mobility 129, 185, 190

nuclear 31–33

number 63, 71

transposition 77, 128

inverted repeat (IR) 75, 77, 126–127

ionophore CR 454

IPP. See pyrophosphate

IR. See inverted repeat

iron 599, 604. See also

metabolism 608

transferrin 608

iron-sulfur center 505, 599

iso1 99

isoamylase 556, 563

Isochrysis galbana 623

pyrophosphate (IPP) 402

isopentenyl pyrophosphate 402

isoprene units 401–402

K kanamycin 44

Kautsky effect 573

571

kinase

protein 213

kinesins 3

kinetic complexity 662

kinetic measurements

fluorescence 437, 453

Klebsiella nidulans 616

717

L L subunit 295

bacterial reaction center 294

L-alanine aminotransferase 641–642

L-amino acid oxidase (LAO) 642

L-aspartate aminotransferase 642

L-ethionine 103

L-methionine-DL-sulfoximine 644

laboratory strains 4–6

land plants 64

LAO. See L-amino acid oxidase

Larix kaempferi 393

laser optoacoustic spectroscopy 444

lauryl maltoside 292

Lemna minor 621

Lemna perpusila 576

lesions

UV-induced 119

LHC 696

Lhcb genes 366

Lhca2 336

Lhca3 336

LHCI. See light-harvesting complex I

LHCII. See light-harvesting chlorophyll a/b-protein complex

LHCII complex. See light-harvesting chlorophyll a/b-protein

complex LHCP. See light-harvesting chlorophyll a/b-binding polypeptide library cosmid 54

genomic 36

life cycle 3

ligands 469

light

actinic 446

detection 446

modulated 440, 444

pipe 447

reactions 622

receptors

blue light 343

blue-UVA 343

phytochrome 343

regulation chloroplast genes 153, 176, 189, 200, 206, 212

sensitivity 262, 518

stress 578–590

light-energy distribution 247

light-harvesting chlorophyll a/b-protein complex (LHCII) 242,

248, 364–373, 428, 571–572, 574, 623, 691

apoproteins 372

assembly 364–373

degradation 580

immunolocalization 368

kinase 576, 579

model 366

membrane vesicle intermediate 364, 373

soluble intermediate 364, 372

N-terminal segment 577

phosphorylation 576–577, 580

protein import 372

reconstitution 366

trimer 428, 578

718 light-harvesting chlorophyll a/b-binding polypeptide (LHCP) 396

translocation 223–224, 229

genes 29

light-harvesting complex I (LHCI) 242, 332–333, 336–367

light-harvesting complex II. See light-harvesting chlorophyll a/b-

protein complex

light-sensitivity suppressors 518

light-triggered ATPase 492

lincomycin 160

linkage group 6

uni1 7

lipids 416–429

biosyntheyic origin

eukaryotic 419

prokaryotic 419

composition

fatty acid 419–420

manipulation 426

mutant 426

distribution 421

export 422

metabolic pathway

CDP-diacylglycerol 422

chloroplast envelopes 423

chloroplastic 423

desaturases 422–423

endoplasmic reticulum 422

extra-chloroplastic 423

fatty acid synthesis 422

glycerolipids 424

poly-unsaturated species 422

prokaryotic pathway 423

thylakoid membranes 423

micelles 691

signaling 416

thylakoid 228

liposomes 416, 425, 479

lock-in amplifier 438, 440

loroxanthin 366

low affinity NT 651

low-density membrane system 698

Lpcr-1. See Por-1

LRP. See promoter: chloroplast: light-regulated

lumen 436, 602–603

proteins 606

lumenal processing peptidase 467

luminescence 435, 438–439, 455, 457

delayed 452–457

kinetics 439

lutein 258, 366, 624

lyase

cytochrome c/heme 606

lycopene cyclase 405

M M subunit 295

bacterial reaction center 294

m-phenanthroline 396

155, 159, 175

155

154–155

ma1 618, 628

Index ma2 618, 628

macronutrients 614–630

maltase 556

maltotriose 556

manganese 17, 312

deficiency 17

manganese water oxidation complex 257, 265, 271, 274, 299–302,

570, 586

photoactivation 301

MAP. See mitogen activated protein

map

distances 126

genetic 6, 36

molecular 36

physical 36

mapping. See mitochondrial gene: mapping

Marchantia 81, 86

marker

molecular 36

recycling

transformation 145

selectable nuclear 43–45

selection

paternal 125

mass spectrometry 534

mat-3 131

mat3 106

maternal bias 124

maternal inheritance 94–110

mating type 669

change 100

minus dominance 101

mating type locus 695

chromosomal rearrangements 99

R domain 99

mating-type 668

locus 29, 31, 98–101

specific genes 99

MC9 370

MCA1 467

MCB1 467

MCD1 467

MCG1 467

Mehler reaction 276, 440

meiotic recombination 690

membrane

appressed (stacked) 234, 237, 243, 250, 260, 263, 267–268,

272, 463, 577

attachment site 123

chloroplast envelope 419

domains 234, 246

endoplasmic reticulum 419, 422

envelope 368, 422

outer 236

freeze-fractured 240

grana 247

hemi- 238

inner envelope 698

insertion 159

co-translational 159, 222

mitochondrial 419

integration. See membrane: insertion

non-appressed (unstacked) 234, 241–242, 244, 250, 260, 263,

267–268, 272, 463, 577

Index plasma 419

potential 435, 439

electrochromic shift 442

stroma 247

thylakoid 237–238, 419

vesicle intermediate 373

membrane-bound polypeptides 649

Mendel’s laws

inheritance 94

metabolic pathway

lipid

CDP-diacylglycerol 422

chloroplastic 423

desaturases 422

endoplasmic reticulum 422

extra-chloroplastic 423

fatty acid 422

glycerolipids 424

poly-unsaturated species 422

metabolic signals 655

metal

acquisition 598

cofactor 598

deficiency

copper 600

deficiency/availability

copper 599

iron 599

transport 598, 602, 608

metalloproteins

assembly 598–609,

methionine 629

methionine-D,L-sulfoximine 642

methylammonium 628–629, 648

-resistant mutants 639

sulphanilamide 645

methylation 96, 102

chloroplast DNA 119, 132

DNA 51

protein 507

methyltransferase 132

metribuzin 302

resistance marker 141

mevalonate kinase 402

mevalonic acid 402

Mg chelatase. See IX chelatase

Mg-PME cyclase. See IX monomethyl ester

Mg-protoporphyrin IX 698, 699

Mg-protoporphyrin IX methylester 699

387 . See divinyl protochlorophyllide 378 IX 388 IX chelatase 388, 400 IX methyltransferase 389

IX monomethyl ester oxidative cyclase 388– 390

MGDG 419

micelles 691

microalgae 530

micronutrient 605

microphone 444

microscopy

confocal scanning laser 368

719 electron 98, 260

fluorescence 97, 103

high resolution electron 481

immunoelectron 369–370

Mid 29, 31, 33, 99

midpoint potential

295 339 mismatch repair 127, 132–133 missense

mutations 485

rbcL 518

mitochondria 3, 357, 607

beef-heart 462

distribution 541

fragmented coding regions 83

monophyletic origin 65–69

polyphyletic origin 65–69

mitochondrial

carrier protein superfamily 532, 540

-chloroplast interactions 675–676, 680–681

genetics 662–672

genome

organization 79–82 size/copy number 662

mutations 662, 667

transformation 671

translation 664

membrane 419

rRNA coding regions

fragmented 84

scrambled 84

mitochondrial DNA

circular mapping 79

inheritance 668

terminal inverted repeats 663

mitochondrial genome

engineering 692

evolution 63, 64–87

mapping 670

mutations 665–668

recombination 669, 671

replication 663

size 69–78

transcription 664

mitogen activated protein 622

mitorespiration 680

mitotic offspring 94

mitotic segregation 95

MMS 132

MoCo. See molybdopterin cofactor

modulated light 440, 444

beam 438

molecular map 36

molecular marker 36

molybdate processing 646

molybdenum cofactor 608, 647

molybdopterin

biosynthesis 646

molybdopterin cofactor (MoCo) 641, 646–647

biosynthesis 647

carrier protein 647

mutants 647

oxidation 647

Index

720 monophyletic origin mitochondria 65–69 plastids 65–69 3-monovinyl protochlorophyllide 390 morphology cell 235 chloroplast 236–238 mRNA abundance 603, 606 MSX. See methionine-D,L-sulfoximine MT locus. See mating-type locus mtl1 102 mucidin 667–668 mussels 94 mutagenesis 298 directed 521 general 646 insertional 37, 51–54, 523, 560–561, 648, 693 site-directed 147, 265, 270, 273–274, 299, 311, 464 mutagens 132 mutants acetate-requiring 518, 523 assembly cytochromes 607

assimilation 645

ccb strains 607

chlorate-resistant 645

chlorophyll b-less 397

cofactor 646

binding 608

cryptic 646

complex 463

cytochrome

desaturases 427

hem 604

heme attachment

ccs 605

light sensitivity 518

suppressors 518

lipid biosynthesis 427, 429

lipid composition 427, 428, 429

polyunsaturated fatty acids 428

methylammonium-resistant 639

mitochondrial 667

NiR-deficient 654

nitrate reductase 646

nitrite reductase 654

nuclear 428

photosynthesis-deficient 427, 429, 518

PS I-deflcient 354

PSII-deficient 250, 429

random 522

RbcS2 523

regulatory 646, 648

resistance 645

screening 451, 452

random 518

self-sterile 101

site-directed 265, 273, 465

temperature-conditional 523

thylakoid membrane 429

transporter/transport 649

yellow (y) 4

zygote maturation 101

mutation

chloroplast DNA 131

frame-shift 270

length 75

missense 519

mitochondrial 662, 665

dum mutants 665–668

minute colony mutations 665–668

obligate photoautotrophic mutants 665–668

respiratory-deficient 665–668

mitochondrial genome 665–668

nonsense 519

second-site 519

site-directed 470

imitators 133 MVA. See mevalonic acid Mycobacterium leprae 607 Mycobacterium tuberculosis 607 myxothiazol 667–679

N density transfer 117, 120 Nacl 206, 207 Nac2 158, 206, 207, nac2-26 54, 154, 155, 156, 157, 158, 173, 175 NAD(P)H dehydrogenase 357, 665, 676, 678, 681, 696 chloroplast 675, 678, 681 nad5-nad4 80 NADH dehydrogenase 662, 680–681 NADH-GOGAT 640–641 NADH:ubiquinone oxidoreductase 663–664 photoreduction 335 NADP-malate dehydrogenase 503, 512 NADPH-plastoquinone reductase 435–436 NADPH-plastoquinone-oxidoreductase 575 NADPH-thioredoxin reductase 502, 508 NADPH: protochlorophyllide oxidoreductase. See protochlorophyllide reduction: light-dependent NADPH:ubiquinone oxidoreductase 664 nalidixic acid 123 naphthylphosphate 628, 629 napthylsulfate 46, 58, 629 Nar2 650 ncc1 154, 155, 156, 175 NCS. See 4-nitrocatecholsulfate nd1 664 nd2 664 nd4 664 nd5 664 nd6 664 ndh genes 678 ndhF 334 neoxanthin 258, 366 Neurospora crassa 606, 616, 647, 652, 663 RNA splicing 188 neurosporene. See carotenoid: neurosporene Nic7 99 as selectable marker 44 Nicotiana 651 Nicotiana plumbaginifolia 646 Nicotiana sylvestris 662 starch synthesis 561

Index Nicotiana tabacum 653

nigericin 223, 436

nitA

Volvox carteri 31

Nit1 (Nia1) 27, 30, 48, 652, 694

as selectable marker 43, 44, 51, 53–54

gene expression 46–47, 50, 58

Nit2 27, 44, 628

Nit8, (Nar2 ) 27, 55, 628, 695

as selectable marker 44

nitA

Volvox carteri 58

nitrate 620, 627, 638

assimilation 645

reductase 4, 487, 627–628, 641

gene 652

mutants 646

regulation 651

Volvox carteri 652

transport 649–651

high affinity 649

mutants 649

transporter 628

nitrite 620

assimilation 645

reductase 385, 627, 641, 654

immunological studies 654

mutants 654

redox center 654

transport 649–651

turnover 649

4-nitrocatecholsulfate 619

nitrogen 620–621, 625–627, 655

assimilation 638–655

regulation 638–655

starvation 131, 553

effect on ribosomes 202

gametogenesis 98

nitrogenase 394

nitroso-guanidine 132

nitroso-methyl-urea 132

NMR 338, 445, 481, See also nuclear magnetic resonance

N,N´-o-phenylenebismaleimide 482

non-heme Fe 303

non-invasive screening 18

non-photochemical quenching 571, 624, 687, 694

non-radiative dissipation 571

non-reciprocal cpDNA recombination 128 See also recombination:

chloroplast DNA

noncyclic photophosphorylation 17

nonsense

rbcL 519

norflurazon 401, 590

novobiocin 122, 123, 169

NPQ. See non-photochemical quenching

nptII 43

as selectable marker 44

Nrt2 650

NTR. See NADPH-thioredoxin reductase

nuclear

genes 517

cloning 51–54

promoters 30, 56–58

721 nuclear genome

engineering 693

organization 25–37

transformation 693

nuclear replication 117

nuclear transformation 41, 42–58, 56–58

nuclease 101, 108

103

endo 103

exo 103

zygote-specific 95

nuclease C 103

nucleoids 96–98, 103, 105, 118–119, 121

chloroplast 105

nucleotide analogs 488

nucleotide substitution

rate 64

null mutations 695

O o-phenanthroline 396

O-acetylserine 626

See singlet oxygen

evolution 622, 623. See also oxygen evolution

See triplet oxygen

obligate photoautotrophic mutants. See mutations: mitochondrial

octyl-glucopyranoside 461

octylglucoside 479

Odontella sinensis 314, 483

OEC 33. See OEE1

translocation 229, 223–224

OEC17. See OEE3

translocation 223

OEC23 228, 588. See also OEE2

translocation 223

OEE proteins 588

OEE1 33, 222, 228, 249, 270, 292, 308, 311–312

Oee1 as selectable marker 44

gene expression 50

OEE2 249, 312–322, 314

OEE3 249, 270.

olive

Antirrhinum majus 388

operon 80, 607

optical parametric oscillator 441

ORF 1995 145

ORF 472 145

organelle inheritance 94–110

organelle warfare 110

orthophosphate

regulation of ADP-glucose pyrophosphorylase 555, 557, 560

Oryza 86

Oscillatoria chalybea 654

Oscillatoria limnetica 359

osmotic pressure 464

oxidase 436

alternative 541, 664

chloroplast 675

damage 580

pentose phosphate pathway 652

oxidation

plastoquinone 680

722 oxidoreductase 606

2-oxoglutarate transaminase 642

oxygen

consumption 277, 440

emission 439–440

evolution 272, 301, 434–436, 439–444

gush 440

measurements 439–440

singlet 580, 582

oxygen-evolving complex

polypeptides

assembly 270

P P5/D1/D2 subcomplex 249

P6 subunit 249

581, 582

289, 295, 300, 580, 582

582, 690

midpoint potential 295

583

580, 581

572

581, 582

17, 326, 328–329, 338, 341, 579, 589

chromophore 396

midpoint potential 339

338

absorption difference spectra 339

p-benzoquinone 577

P-loop 488

p-nitrophenyl sulfate 619

palindromic sequences

GC-rich 81

palmitoylation 264

PAM. See pulsed amplitude-modulated measurements

Pandorina morum 69, 79

PAPS. See 3´-phosphoadensine-5´-phosphosulfate

paromomycin 45

particle bombardment. See transformation: particle bombardment

transformation 55

particle gun 140. See also transformation: particle bombardment

Pasteur effect 435

paternal inheritance 94–110

paternal marker selection 125

PBG 385. See porphobilinogen

PBGD 400. See porphobilinogen deaminase

PBGS 384, 400

PC. See plastocyanin

PC1 392

pc1 44, 401

Pchlide oxidoreductase 364

PCR. See photosynthetic carbon reduction

Pcy1 601, 602

gene expression 50

pD1 586, 588

pD1 protein 587

pd2 628

pd24 628

pd3 628

Index pde2 622

PDS 406. See phytoene desaturase PE 419

pea thylakoid translocation 223

Pediastrum boryanum 600

PEG fused diploids 100

pentose phosphate cycle 354

pentose phosphate pathway 557–559, 652

PEP. See phosphoenol pyruvate

PEP carboxylase 626

PEP phosphatase 626

pepper 405

periplasm

arylsulfatase 619

phosphatase 619, 620

carbonic anhydrase 534, 620

periplasmic space 619

petA 466, 471

gene expression 155, 159, 171, 173, 175, 205

petB 466

gene expression 154–155

petC 466

petD 466

gene expression 155, 157, 160, 171–173, 175, 204–206

PetE (Pcy1)

gene expression 47, 50

PetG 470

petG 461, 466

PetL 463, 465, 470

petL 461, 466

PetM 470

petM 461, 466

PF-1 565

PF-18 565

PF-2 565

PF-7 565

PF14 58

pf14 31, 47

PGA 625. See 3-phosphoglycerate

PGP1 531

571, 583

phemediphan 303

pheophytin 256–257, 264, 292, 298–299, 350, 396

Qx transition 299

phleomycin 45

Phormidium luridum 678

phosphatase 619, 627, 629

periplasmic alkaline 620

neutral 620

phosphate 617

esters 617, 619

translocator 558

transport 618, 629

phosphatidylcholine 425

phosphatidylethanolamine 415

phosphatidyiglycerol 415

phosphatidylinositol 415–416, 419

phosphinothricin 642

3´-phosphoadensine-5´-phosphosulfate 615

phosphoenolpyruvate 558, 626

phosphofructokinase 559

Index phosphoglucoisomerase 557 phosphoglucomutase 554, 557, 561 3-phosphoglycerate 552, 557–558, 560, 625 phosphatase 531, 539, 541, 543 phospholipids 621 phosphomevalonate kinase 402 phosphorescence emission anisotropy 491 phosphoribulokinase 17 phosphorus 617, 619, 621, 623, 625–627 phosphorylase 555–557, 559 phosphorylation 213, 247, 260–261, 488, 491, 573, 575, 625 LHCII 577, 580

protein 213

PS II

core proteins 585

reversible 247, 250

photoacoustics 351, 574, 577 measurements 443–445 spectrometer 444 spectroscopy 444 techniques 434 photoactivation 256–257, 270, 272–277, 301, 313 photoautotrophic growth 261–262, 273, 349–360, 355 photobaric effect 444 photobleaching 355, 699 herbicide 387 photochemical energy conversion quantum yield 443 photochemical energy storage 443 photodamage 264, 583, 690 photoevolution hydrogen 359 photohydrogen evolution 17 photoinactivation 578, 581 irreversible 579 PS II 586 photoinhibition 260, 263, 272–273, 276, 306, 308, 570, 578–579, 687

acceptor-side 580

donor-side 582

photolyase 130 photomorphogenesis 406 photooxidation 106, 273, 275 photooxidative damage 387, 401 photophosphorylation 13–15, 17, 477–478, 488, 491, 536 cyclic 17 noncyclic 17 photoreactivation 130 photoreduction 354 335 photorespiration 530, 535, 538–539, 643 photosensitivity 262, 354 photosynthetic complex 607 assembly 698

degradation 607

Photosystem I (PS I) 14, 349–360, 433, 436, 575, 584, 697 antenna 439 assembly 343 biogenesis 341–344 centers 234 closure 579 core 439 core proteins phosphorylation 585

723 cyclic electron flow 441, 589

deficient mutants 354–357, 454

donor side 580

evolution 358

immunocytochemical localization 241

photoinactivation 579

protein degradation 579

reaction center 292

reconstitution 338, 341

structure

model 327

structure and function 324–344

subunits

acceptor side 333

donor side 335

function 333

trimer 326 Photosystem II (PS II) 14, 287–315, 349–360, 433, 435–36, 452, 457, 518, 678 activity fluorescence mutations 426

assembly 249, 255–277

biogenesis 257–259

centers 234

complex

assembly 211, 234

core particles 292, 307

cyclic electron flow 580

evolution 358

functional analysis 287–315

herbicide binding 580

model structure

ribbon representation 290

monomerization 585

mutants 250, 454

oxygen evolving

core particle 307

particle

BBY 290

particles 239, 290–292

core 292

photoinactivation 586

protein turnover 579

protease 586

quinone acceptor 436

reaction center 307, 452

reassembly 587

repair cycle 587

phototaxis 3, 236, 590 photothermal effect 443 phycobilin 379 phycoerythrin 14 phylloquinone 330, 331, 341 phylogenetic tree 64, 68 phylogeny 64 of green algae 65 physical distance of a recombination unit 36 physical map 36 phytales 617 phytochrome 399 phytochromobilin 379 phytoene 404 phytoene desaturase 404, 590

724 phytoene synthase 404

phytoglycogen 563

phytol 395

phytylation 394

piezoelectric transducer 444

Pinus sp. 678

Pioneer 1. See transposable elements

PK. See pyruvate kinase

plasma membrane 419, 618

plasmid rescue 51, 53

Plasmodium 699

plastids

development 259

monophyletic origin 65–69

polyphyletic origin 65–69

plastocyanin 14–15, 17, 20, 222, 263, 349–360, 460, 463, 575,

580, 584, 598–599, 602–604, 697

biosynthesis 600, 607

copper 460

cross-linking 335

degradation 605

diffusion 464

structure 460

translocation 223, 224, 229

plastoquinol 436, 581, 584

oxidation 680, 681

plastoquinol pool 581

plastoquinone 213, 256, 263–264, 349, 350–360, 440, 676,

678–679

reduction in dark 678–679

plastoquinone pool 435–136, 439, 571, 576, 580–581,584, 623,

678, 680

Platymonas subcordiformis 68, 73, 78, 79

pmp1 628

PMP1 531

poly-unsaturated fatty acids 425

polyA-binding protein 697

polyadenylation. See RNA: polyadenylation; RNA:

polyadenylation: chloroplast

polyadenylation signal 33

polymorphism

restriction fragment length 36, 97

polynucleotide phosphorylase 174

polyphenol oxidase 256

polyphosphates 621

polyphyletic origin 68

mitochondria 65–69

plastids 65–69

polysomes

membrane bound 210–211, 586

thylakoid-bound 469

FOR. See protochlorophyllide reduction: light-dependent

Por1 44, 391, 393

gene expression 401

porphobilinogen (PBG) 383, 385

porphobilinogen deaminase (PBGD) 384–385, 400

porphobilinogen synthase 383

Porphyra purpurea 314, 483

Porphyridium cruentum 14

porphyrins 378. See also

PPOX 387. See protoporphyrinogen oxidase

PPT. See phosphinothricin

PQ. See plastoquinone

Index pre-amylopectin 563

prenyltransferase 403

preuroporphyrinogen. See 1-hydroxymethylbilane

processing

exonucleolytic. See exonuclease

molybdate 646

RNA. See RNA: processing: chloroplast

processing protease 587

prolamellar bodies 364

promoter 466

chloroplast 157, 168–169, 171–172

light-regulated 168

nucleus encoded RNA polymerase 168

spinach 169

distal 466

light-induced 653

mitochondrial 664

nuclear 47–48, 56–58

genes 30

Propionigenium modestum 491

protease 472, 586

protein

chimeric 268

degradation 470, 598, 601

lumen 601

half-life 601

import 372

kinase

cAMP-dependent 622

MAP 622

translocation 379

chloroplast 220–222, 400–401

thylakoid 223–229

protein-chromophore complex 288

Proteobacteria 358

proteolipid 480

proteolylic cleavage 646

proteolytic degradation 212

proteolytic processing 268, 270

protochlorophyll 391

protochlorophyllide 391

photoconversion. See protochlorophyllide reduction: lightdependent

reductase 200. See also protochlorophyllide reduction

reduction 391

light-dependent 391–393, 392, 400

light-independent 391–393, 689

protoheme 379, 397. See also heme b

proton

electrochemical gradient 434, 464

gradient 358, 442, 676

motive force 492, 618

pump 465

translocation 491

uptake 439

protoplasmic layer 238

protoporphyrin IX 387, 397

protoporphyrinogen IX 379, 386

protoporphyrinogen oxidase (PPOX) 387

Prototheca wickerhamii 65, 68, 73, 79, 82

PS I. See Photosystem I

PS II. See Photosystem II

PS II-H 308

Index PS II-I protein 306 PS II-J protein 309 PS II-K protein 309 PS II-L protein 309 PS II-M protein 310 PS II-N protein 310 PS II-S protein 314 PS II-T protein 310 PS II-X 314 PsaA 328, 331–332, 336, 338, 341, 343 psaA 82–83, 147, 190–191, 325, 336, 341, 344, 354, 357, 699 gene expression 54, 153, 160, 171, 184, 191, 208 355 deletion mutants 357 mutants 355 PsaB 328, 331–332, 336, 338–339, 341, 343, 579 folding model 340 psaB 147, 325, 336, 344, 354, 357 gene expression 153, 155–156, 160, 169, 176, 206, 208 deletion mutants 357 mutants 355 PsaC 326, 332–333, 338, 341 psaC 341, 354–355 PsaD 326, 333, 341–342 PsaD 334–343 PsaE 333 PsaE 334 PsaF 221, 326, 342 PsaF 52, 55, 695 PsaG 336 PsaH 333, 336 PsaI 326, 335 PsaJ 326, 335 PsaK 335–336 PsaL 326 psaL 335 PsaN 335–336, 342 translocation 223 psbA 125, 127, 143, 147, 587 as selectable marker 141 barley gene expression 211 gene expression 141, 153, 157, 160, 186, 189, 200–203, 205– 208, 211, 249 spinach gene expression 156 tobacco gene expression 203 psbB 585 gene expression 54, 154–157, 159, 171, 211 psbC 585 gene expression 155, 160, 206–208, 211, 249 psbD 173, 191 gene expression 153–160, 171, 175, 206–208, 2 1 1 psbE 582 psbF 582 PsbH 578 psbH 308 psbI 292, 306, 309 gene expression 171 psbJ 309 psbK 309 psbL 310

725 psbM 310 gene expression 171 psbN 310 PsbO 311 PsbP 312, 313, 314 PsbQ gene 313 PsbS 313 psbT 310 gene expression 171 PsbW 314 PsbX 314 Pseudomonas C12B 616 Pseudomonas putida 619 Pseudomonas spp. 607 psoralen 123 PSY 406. See phytoene synthase Psy 404 pullulan 556 pullulanase 556, 563 pulse labeling protein 266, 468, 471, 605, 607 RNA 152, 170. See also RNA: pulse labeling pulse-chase experiment 584 pulsed amplitude-modulated measurements 573 pulsed field gel electrophoresis 36 pump ATP-dependent electrogenic 677, 681 purine 620, 643–645 assimilation 643–645 transport 644 pyrenoid 3, 98, 236–237, 518, 533, 536–538, 541, 552 starch sheath 541 pyridoxal-phosphate 383, 641 cofactor 383 pyrimidine dimers 130 pyruvate kinase 625

Q 289, 295, 302, 436, 573, 576 573–574, 579, 580, 582–583 582 579–581 289, 295, 302, 573–574, 580, 583–584, 623 non-reducing 623, 624 582 581 572–574. See site 464 See non-photochemical quenching Qo site 464, 465, 576 571, 574 572 quantum efficiency 581 quantum noise 437, 440, 441, 446 quantum requirement 686 quantum yield 443, 445 spectra 445 quenching non-photochemical 624, 687, 694 quinone 17, 263, 276, 436 binding site 460–461 affinity 465

726 quinone, binding site (Continued) 453

Qi 464

Qo 464

R r1 389

Raman spectroscopy 461

RAS 622

rbcL 69, 75, 167, 517

gene expression 153–156, 160, 168–170, 176, 200–202, 212

mutations 516

missense 518

nonsense 518–519

transcription 523

RbcS 30, 69, 51, 85, 517, 695

deletion mutant 523

gene family 523

genes 29

gene expression 47

RbcS1, RbcS2 as selectable marker 44

RCII core 578

rcl-u-1-10-6C 97

reaction center 325, 336, 697

bacterial

H subunit 294

L subunit 294

M subunit 294

chlorophyll 298

Photosystem II 452

type 1 358

type 2 358

types 288

rearrangement

gene 63

RecA 106, 132

recA E. coli 142

receptors 512

recombination 75, 690

chloroplast intramolecular 118

chloroplast DNA 118–119, 123–130

DNA 96

nip-flop 127

homologous 49, 54–56, 81, 126–127, 132, 142–143, 692

intermolecular 82

intramolecular 81–82

mitochondrial genome 669, 671

non-reciprocal 128

recombination hotspots 127

recombination unit

physical distance 36

recombination/repair 132

red algae 552

red drop phenomenon 13

red pigments 333

redox

balance 575–576

change transients 440

center 651, 654

cofactor 654

Index control 578

potential 643, 697

state 434, 574

regulation 504

667, 668 redox site reduced 581

regulatory mutants 648

repair

chloroplast DNA 130–133

cycle

PS II 579

mismatch 132–133

repeated DNA 63, 73

repeated sequences

dispersed 63

interspersed 26–27

simple 28

tandem 27–28

repeats

inverted 75

short direct 75

short dispersed 75

replication 75

autonomous 120

chloroplast DNA 116–123

bidirectional 121

origins 120, 122

dispersive 118, 119

mitochondrial genome 663

nuclear 117

semiconservative 117

replication complex 122

replication origins 43

reporter 46–47, 154–157, 202–204, 207

(uidA) 142

alkaline phosphatase 47

aminoglycoside-3´´ adenyl transferase (aadA) 142

arylsulfatase 46, 48, 56–58, 653

chloroplast genes 141–142

gene 603

resistance

arsenate 645

chlorate 645

erythromycin 96

methyl-ammonium 642

MSX 642

PPT 642

sulphanilamide 645

resistance marker

DCMU 141

erythromycin 141

metribuzin 141

spectinomycin 141

streptomycin 141

resonance Raman spectroscopy 299

respiration 440

respiratory-deficient mutants. See mutations: mitochondrial

restriction fragment length polymorphisms (RFLP) 36, 97, 668,

670

restriction map

chloroplast DNA 120

restriction-modification 102

reticulocyte lysate 485

Index reverse electron flow 357, 680 reverse genetics 139–148, 354, 695 reverse transcriptase 77 -like gene 72 -like protein 663 reversible phosphorylation 247, 250 RFLP. See restriction fragment length polymorphism (RFLP) rhizobia 607 rhizosphere 617 Rhodobacter capsulatus 391, 606, 667 Rhodobacter sp. 294, 607 Rhodobacter sphaeroides 331, 339 Rhodopseudomonas viridis 295, 297, 299, 302, 697 rhodopsin 3 rhodopsin-like light sensor 590 Rhodospirillum rubrum 482, 521 Rubisco 516 ribonuclease. See also exonuclease ribonucleoprotein complex 202, 205–209 ribosomal proteins chloroplast 198 ribosomal RNA gene organization scrambled 699 gene sequence 64 variable regions 85 ribosome deficient mutants chloroplast 208–209 pausing 259 ribosome binding site mitochondrial 664 ribosomes chloroplast 212 cytoplasmic 364 nitrogen starvation 202 ribozyme. See RNA: catalytic ribulose-l,5-bisphosphate carboxylase/oxygenase (Rubisco) 19, 96, 97, 237, 260, 515, 516–524, 573, 621, 686, 531–533, 535–539, 541, 543, 552, 604, 621, 687, 697 active site 517 barrel loop 6 519

activase 517, 522, 536, 552

catalytic efficiency 515, 519, 521–522

large subunit 517–519, 523

loop 6 522

nuclear mutants 523

rbcL 517

rbcL mutations 516

RbcS 517

Rhodospirillum rubrum 516

small subunit 517, 519, 523

specificity factor 517–521

stability 517–522

Synechococcus 516

thermostability 522, 523

uniparental inheritance 97

Rieske iron-sulfur protein 17, 461–463, 465, 472 targeting sequence 469 translocation 224 rifampicin 20 right-side-out vesicles 246

727 RNA affinity chromatography 202 3´ UTR. See RNA: untranslated region: 3´ antisense 170 binding proteins 158, 205–209, 698 capping 171. See also vaccinia virus guanylyl transferase catalytic 184, 184–189 degradation 151. See also exonuclease genes 28 initiation codons 205 inverted repeat. See RNA: secondary structure leader. See RNA: untranslated region: 5´ polyadenylation 171, 176, 199 chloroplast 158

polycistronic 154, 166

polymerase

chloroplast 166 nucleus-encoded 167

sigma factor 167

subunit C 71

processing 159, 173, 697

chloroplast 157–158, 170–177

endonucleolytic cleavage 664

in vitro 172

pulse labeling 152, 154, 156, 170 secondary structure 151, 160, 169–171, 173, 175–176, 185, 190, 192–193, 664

splicing 184–193, 697

stability 151, 172–173, 175–177, 697

translation 160–161

chloroplast 151–161 untranslated region

3´ 155–157, 170, 174, 176

5´ 156–159, 173, 200, 203–214

RNAse E 174 rotational coupling 491 rotenone 665 rpo 85 rpoB tobacco 166 rpoB1 145, 166–167 rpoB2 145, 166–167 rpoC2 71, 145, 166 rps12 95, 191 rps3 145 rps4 gene expression 204 rps7 gene expression 171, 204 rrn16 gene expression 168–169 rRNA. See ribosomal RNA rRNA genes 5S 28 rrnL 80, 185, 186 gene expression 153, 186, 189 rrnS 80, 185 Rs3 387 Rsp3 31, 47 gene expression 58 Rsp4 29 Rsp6 29 rt1 77, 664

Index

728 Rubisco [or] RuBP-carboxylase. See ribulose-1,5-bisphosphate carboxylase/oxygenase

S S-adenosyl-L-methionine 389 S-state 300 EPR multiline signal 306 582 S2,3 582 Sac1 622 sac1 622, 624, 627 sac2 627 sac3 627 Saccharomyces 698

Saccharomyces cerevisiae 483, 543, 606, 618, 622, 628, 639, 662, 667 RNA splicing 188 salicyl hydroxamic acid 679 salicylaldoxime 15 salicylhydroxamic acid 664 salvage/turnover/repair 119 SAM. See S-adenosyl-L-methionine SceI 663 Scenedesmus 14, 15, 366 Scenedesmus obliquus 14, 65, 70, 79, 83–84, 259, 268, 270, 530, 587, 600, 689 heterotrophic growth 689 screening colony 694–695

mutant 451–452 non-invasive 18

random 518 seaweed green 552 SecA 23–224, 228 second-site mutations 519 secondary structure RNA. See RNA: secondary structure SecY 224, 228 sedoheptulose-1,7-bisphosphatase 512 segregation 247 mitotic 95 vegetative 124 selectable markers 648, 692 aadA 45, 47, 141, 143–144, 147, 166 aphVIII from Streptomyces rimosus 45

Arg7 43, 53, 54, 57 atpB 141

AtpC 44 ble from Streptoalloteichus hindustanus 45, 47, 51 Ccs1 44

chloroplast 140–141 Cry1 44, 47, 54, 55 Nic7 44

Nit1, (Nia1) 43, 44,51, 53, 54 Nit8, (Nar2) 44 nptII 44 nuclear 43–45 Oee1 44 petA 44

psbA 141 RbcS1, RbcS2 44 Thi10 44 tscA 141 Selenastrum minutum 626 selenate 629

semiconservative replication 117 semiquinone 454 semiquinone cycle 465 sequences dispersed repeated 63 insertion 83 short direct repeats 75 short dispersed repeats 75 sex-limited gene 100 uniparental inheritance 98 sex-linked genes 99 uniparental inheritance 98 sexual cycle 4 sexual differentiation 99 Shine-Dalgarno sequence 198, 203, 467 sigma factor RNA polymerase 167 signal 299 299 signal hypothesis 220 signal recognition particle CP54 372 signal sequence 219 chlroplast targeting 220–222 mitochondrial targeting 221 thylakoid targeting 222–229 signal transduction 603 calcium 400 pathway 543–544 signal-to-noise ratio 436–437, 439–442, 445–446 signals metabolic 655 sim30 206 singlet chlorophyll 570 singlet energy transfer 366 singlet oxygen 580, 582 sink carbon 551 siroheme 379, 385 site-directed mutation 470 mutagenesis 147, 270, 274, 299, 464, 607 mutants 273 Skeletonema capstatum 623 slk1 622 small subunit 68 ribosomal RNA 68 gene sequence 68 mechanism 488 SNF3 628 snRNP 699 sodium azide 224 spacer DNA 63 spacers internal transcribed 85 special pair chlorophyll 295–296

Index spectinomycin 20, 51, 156, 487

spectinomycin resistance marker 141

spectrometer

photoacoustic 444

spectrophotometer

double beam 441, 446–447

flash 443, 446–447

spectrophotometry

flash

methods 441

spectroscopy

absorbance 464

absorbtion 602, 440–443

EPR 277

extended X-ray absorption and fine structure 301

flash 461, 464

fluorescence 340

laser optoacoustic 444

photoacoustic 444

resonance Raman 299, 461

X-ray absorbance near-edge 301

spectrum

absorption difference

330

spinach 169, 277

atpF 142

Spirodela oligorrhiza 259, 577, 583–584

Spirogyra 80

Spirogyra maxima 70

Spirulina platensis 697

splicing 184–193

light regulation 189

self- 184–189

spy200 393

spy6 393

SQDG 419

SQR. See sulfide:quinone oxidoreductase

sr1 mutation 95

sr2 mutation 95

SSU rRNA. See small subunit ribosomal RNA: gene sequence

STA-1 562–563

STA1 560

STA2 560–561

STA3 560, 562

STA4 560

STA5 560–561

STA6 560–561

STA7 560, 563

STA8 560, 563

starch 550–565

breakdown 559

floridean 552

granules 551

lyase 556

storage 552, 554

structure 550–554, 551

synthesis 237, 552, 554–565

synthase 555, 561

granule-bound 555, 556, 562

soluble 555, 556, 562, 564

transient 551, 554

start codon 33

state transition 234, 247, 435, 438–439, 444–445, 463–464, 466,

570, 572, 573–578, 679 687, 694

729 cycle 575

kinetics 577

process 577

state I 242–243, 574, 576–577, 679

state II 242, 574, 576–577, 624, 679

582

stationary phase 603

statolith 564

stigmatellin 463

stop codon 33

STOR 110. See also salvage/turnover/repair

Streptoalloteichus hindustanus 45

Streptomyces rimosus 45

streptomycin 95, 96, 125

streptomycin resistance marker 141

stress

high light 306

light 578–590

nutrient 614–630

stroma membrane 247

stromal peptidases 220

structure-function analysis 696

697

cytochrome

cytochrome 697

ferredoxin 697

697

plastocyanin 695, 697

PS I 695, 697

Stylosanthese hamata 618

sub1 397

sub2 397

sub8 397

sub9 397

succinate dehydrogenase 357, 679

sucrose 551

suF15 206, 208

sugary 1

Zea mays 563

sulfatase 616, 627

sulfate transport 617, 629

sulfide:quinone oxidoreductase 358

sulfite reductase 385

sulfolipid 415, 423, 616, 691

6-sulfoquinovosyl diacylglycerol 621

sulfonates 615

sulfoquinovosyldiacylglycerol 415

6-sulfoquinovosyl diacylglycerol 616

6-sulfoquinovose 616

sulfur 615–617, 619, 621–623, 625, 627

cycle 616

sup1 108

sup4B 206

supercomplex 464, 465

superoxide dismutase 600

suppressors

informational 519

intragenic 518

light-sensitivity 518

surface charge

screening 247

surface charge density 246

symporter 618

synchronously grown cultures 19

730 Synechococcus Rubisco 516 Synechococcus elongatus 326, 697 Synechococcus sp. PCC 6301 335 Synechococcus sp. PCC 7002 334–335 Synechococcus sp. PCC 7942 621 Synechococcus vulcanus 312 Synechocystis 332 Synechocystis sp. PCC 6803 268, 296, 299, 301–302, 304, 308– 309, 311, 314, 326, 333, 334, 336–338, 621, 696

synonymous substitution 85

synthesis. See also pulse-labeling

chloroplast DNA 98

in organello 265

in vitro 265

starch 237

thylakoid 364

T TAB1 208 tab1 206 TAC. See transcriptionally active chromosome tagging transposon 37. See transposable elements targeting gene 55, 56 targeting sequence cytochrome f 469 Rieske protein 469 TATA box 30 TBC1 207, 207–208 tbc1 206 TBC2 207 tbc2 206 tbc3 206, 207, 2 1 1 tbc3-rb1 206 TCA cycle 535 TCA1 467 Tcr1 27 Tcr2 27 Tcr3 27 telomeres 28 temperature-conditional mutants 523 tentoxin 479 tetrad analysis 690 tetrads 690 tetrapyrrole biosynthesis 378–379, 398–401, 406 regulation 398–401, 406 pathway 602, 604 tetratrico peptide repeat (TPR) 158 tha1 224 Thalassiosira pseudonana 623 Thalassiosira weisflogii 623 thermal dissipation 571 thermoluminescence 439 band 298, 300 Thi10 99 as selectable marker 44 thiamine 96 thiol modulation 492, 493 analog of ATP 488

Index thioredoxin 213, 492, 502–505, 508–512, 606, 640

thioredoxin reductase. See NADPH-thioredoxin reductase

thioredoxin-f 492

thioredoxin-m 492

thm24 155, 175

thylakoid

synthesis 364 thylakoid electrochemical gradient 453

light-driven component 453

permanent component 453

thylakoid insertion proteins (TIP) 226 thylakoid membrane 237–238, 453, 419, 675–676

appressed 234, 237, 240, 243, 246, 250, 428

biogenesis 428

grana stacks 236, 428

lateral mobility 234, 238, 247, 250, 577

LHCII 428

lipid biosynthesis 423

non-appressed 160, 238, 267, 268, 272, 436, 577

particle

density 241

EF 239

PFu 241

PSII-associated

tetramer-like structure 245

supramolecular organization 234–251 ultrastructure 234, 428 thylakoid-bound polysomes 469 thymidine kinase 117 thymidylate synthetase 131 thymine dimers 130, 131 time resolution 436, 437, 438, 444, 446 TIP. See also thylakoid insertion proteins tip 227 tobacco 166, 405 TOC1 26 TOC2 27 tomato (plants) 405, 621, 640 topoisomerase 123 Toxoplasma gondii 699 toxoplasmosis 699 TPR repeat. See tetratrico peptide repeat trans-acting factors 199 trans-splicing 354, 699 group II intron 82 transcript 5´ end 600, 605 abundance 466 degradation 467 exonucleolytic 467

maturation 467

monocistronic 466

polycistronic 466

stabilization 467

transcription 466–467, 523 activation 605 activators 603 chloroplast 20, 153, 166–171 DNA topology 169

mitochondrial genome 664

petA 466–467

petD 466–467

polycistronic unit 171, 172, 174, 175

Index regulation 603

run-on assay 152, 154, 170

termination 169–171

169–171

transcriptionally active chromosome 167

transducer

piezoelectric 444

transfer sequence

thylakoid 342

transformants heteroplasmic 144–145 homoplasmic 144–145 transformation 7

biolistic method 671

chloroplast 140–148, 521

co-transformation 50, 145

DNA integration 49–50, 142–148

DNA stability 49

electroporation 43, 46, 49

gene expression 50

glass bead 43, 45–46, 55, 140

marker recycling 145

mitochondrial 671

mutagenesis 523

nuclear 56–58, 625–653

particle bombardment 43, 45, 55, 140

Pcy1 601

silicon carbide whiskers 43, 46

transformation, biolistic. See transformation: particle bombard ment

transit peptide/ sequence 108, 342, 505, 510, 517, 558. See also

signal sequence

translation 20, 266, 467

chloroplast 197, 198–214

and RNA stability 160–161

D1 266

in vitro 259

mitochondrial 664

petA 467

petD 467

psaB mRNA 343

translation regulation

ADP in 266

cis elements 199

D1 266

eukaryotic vs. prokaryotic 199

light 697

redox potential 697

trans-acting factors 199

translocation 250, 469

translocation/import

post-translational 605

transmembrane organization 245–246

transmembrane-spanning domains 650

transmission of mitochondrial Genes. See mitochondrial DNA:

inheritance transport

ammonium 618, 629, 639–640, 648

chromate 629

chloroplast 139

copper 604

protein 602

cyclic 354

731 energy dependent 617

inorganic carbon 618

metal 602

nitrate 649–651

high affinity 649

nitrite 649–651

nitrogen 638–655

phosphate 618, 629

post-translation 600

purine 644

selenate 629

sulfate 617, 629

transporter

amino acid 642

bispecific 649

heme 606

low affinity

nitrate 651

metal 608

nitrate 628

proton-dependent 651

transposable elements 26, 37

Gulliver 6, 27

Pioneer I 27

TOC1 26

TOC2 27

transposition intron 77

transposon insertion 648

transposon tagging 27, 37

transposons. See transposable elements

tri-N-butyltin 678

triose phosphate 3-phosphoglycerate-phosphate translocate 373

triplet chlorophyll 582

triplet oxygen 570

triplet state

carotenoid 296, 442

tRNA suppressors

chloroplast 518

synthetase

chloroplast 198

382

chloroplast

tetrapyrrole synthesis 381–382

trnE1 gene expression 167, 169

trnI 191

Trx1 509

Trx2 509

trypsin 584

tscA 192, 344, 699

as selectable marker 141

gene expression 171

tub box 30

TubA gene expression 50

TubA1 gene expression 57

TubB2 47

gene expression 46, 47, 50, 56–58

genes 29–30

genes 20, 29

tufA gene expression 153, 168, 200

732 turnip 462 turnover 688 298–300 EPR signal 308 290–300, 302

U UDP-galactosyl transferase 422 UDP-glucose 557 uidA reporter 155, 172, 203–204 uncoupler 435–436, 443, 465 uniparental inheritance 4, 93–110 antibiotic resistance 97 bidirectional 102 doubly 94 Rubisco 97 Rubisco large subunit 97 sex-limited genes 98 sex-linked genes 98 unstacked membrane 234, 241, 244. See also thylakoid membrane: non-appressed regions 238, 240, 246, 250 unstacked membranes 234 urate 643 urate oxidase 605, 641, 644 urea 620 assimilation 645 compartmentation of 645 uricase 644 UROD. See uroporphyrinogen III decarboxylase uroporphyrinogen I I I 379, 384–386 uroporphyrinogen III decarboxylase 386 uroporphyrinogen I I I synthase 384–385 UROS. See uroporphyrinogen III synthase UV cross-linking 158, 202, 208 UV irradiation 103–106, 109, 126, 130 UV-induced lesions 119 UV-light 132 UV-mutagenesis 131 UV-sensitive factor 105 UV-sensitive mutants 131 UVB 583–584 uvsE1 105, 106, 1 3 1

V vaccinia virus guanylyl transferase 171 vacuole 369–370 contractile 3 vanadate 532 variable fluorescence parameter 574 vegetative diploid 100 vegetative segregation 124 vegetative zygotes 126, 669 transmission 669 vesicular trafficking 698 Vicia faba 648 8-vinyl reductase 390 violaxanthin 258, 366, 572 vitamin 379

Index vitamin 330

Volvocales 417

colonial 2

unicellular 2

Volvox carteri 31, 417

transformation 45, 652

nitA 58

W water oxidation 301, 311–312 water oxidizing complex 312 Wendy element 77, 82, 129

X X-irradiation 131 X-ray absorbance near-edge spectroscopy (XANES) 301 X-ray analysis 481 X-ray crystallography 583 X-ray diffraction starch 552, 553, 562 XANES. See X-ray absorbance near-edge spectroscopy Xantha-f barley 388, 400 Xantha-g barley 388 Xantha-h barley 388, 400 xanthine intracellular pool 644 xanthine dehydrogenase 641, 644 inactivation of 644 xanthophylls 242, 365, 405, 571, 624. See also carotenoid: xanthophyll cycle 401, 571 Xenopus 651 XPO4 629 XSO4 629. See 5-bromo-4-chloro-3-indolyl sulfate

Y y mutants 4, 393 y-1 248–249, 343, 364, 369, 393, 398, 589 y-y 389 y1-4 393 y1-a 393 y5 393 y7 393, 401 YAC. See yeast artificial chromosome YACs 695 ycf10 gene expression 175 ycf3 343, 698 ycf4 343, 698 ycf9 gene expression 171 yeast 2-and 3-hybrid systems 202 yeast artificial chromosome (YAC) 36, 695 yellow mutant. See y-1 582 582

Index

Z Z. See electron donor: Z

Z-scheme 133, 49, 350–360, 686

Zea mays 173, 175

hcf106 224

tha1 224

zeaxanthin 366, 571–572, 624. See also carotenoid:

zeaxanthin

zygospore 4, 7, 668

733 zygote

development 97–98, 103

exceptional 95, 104

formation 234, 248

lethality 106, 108, 110

maturation 98, 100–101

minus/minus 100

-specific gene 108

-specific nuclease 95

vegetative diploid 126