The Molecular Biology of Cyanobacteria

The Molecular Biology of Cyanobacteria

Advances in Photosynthesis VOLUME 1

Series Editor: GOVINDJEE Department of Plant Biology University of Illinois, Urbana, Illinois, U.S.A. Consulting Editors: Jan AMESZ, Leiden, The Netherlands James BARBER, London, United Kingdom Robert E. BLANKENSHIP, Tempe, Arizona, U.S.A. Norio MURATA, Nagoya, Japan William L. OGREN, Urbana, Illinois, U.S.A. Donald R. ORT, Urbana, Illinois, U.S.A.

Advances in Photosynthesis provides an up-to-date account of research on all aspects of photosynthesis, the most fundamental life process on earth. Photosynthesis is an area that requires, for its understanding, a multidisciplinary (biochemical, biophysical, molecular biological, and physiological) approach. Its content spans from physics to agronomy, from femtosecond reactions to those that require an entire season, from photophysics of reaction centers to the physiology of the whole plant, and from X-ray crystallography to field measurements. The aim of this series of publications is to present to beginning researchers, advanced graduate students and even specialists a comprehensive current picture of the advances in the various aspects of photosynthesis research. Each volume focusses on a specific area in depth.

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

The Molecular Biology of Cyanobacteria Edited by

Donald A. Bryant Department of Biochemistry and Molecular Biology The Pennsylvania State University, University Park Pennsylvania, U.S.A.

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

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Contents Preface

xiii

Color Plates

CP-1

1

1–25

Molecular Evolution and Taxonomy of the Cyanobacteria Annick Wilmotte Summary I. Introduction II. Guidelines to the Taxonomy of the Cyanobacteria III. Fossil Record of the Cyanobacteria IV. Results of Chemotaxonomic Studies V. Results of Macromolecular Methods VI. Conclusion Acknowledgments References

2

3

The Oceanic Cyanobacterial Picoplankton Noel G. Carr and Nicholas H. Mann

27–48

Summary I. Introduction II. Macromolecular composition III. Phycobiliproteins IV. Plasmids V. Phages VI. Transcription VII. Translation VIII. Nutrition IX. Adaptive Responses Acknowledgment References

27 28 30 31 33 33 35 36 37 43 44 45

Prochlorophytes: The ‘Other’ Cyanobacteria? Hans C. P. Matthijs, Georg W. M. van der Staay and Luuc R. Mur Summary I. Prochlorophytes and Chloroplast Ancestry II. Other Non-phylogenetically Directed Studies III. Concluding Remarks References

4

2 2 4 5 5 7 21 21 22

Molecular Biology of Cyanelles Wolfgang Löffelhardt and Hans J. Bohnert Summary I. Introduction II. Cyanelle Wall Biosynthesis and Structure III. Molecular Genetics IV. Protein Transport V. Phylogenetic Analyses VI. Conclusions Acknowledgments References v

49–64 49 50 55 62 62

65–89 66 66 68 69 79 81 82 84 84

5

Chloroplast Origins and Evolution Susan E. Douglas

91–118

Summary I. Introduction II. The Procaryotic Ancestry of Plastids and Their Subsequent Evolution III. Secondary Endosymbiosis in Plastid Evolution IV. Conclusions and Future Prospects Acknowledgments References

6

Supramolecular Membrane Organization Elisabeth Gantt

91 92 93 108 111 111 112

119–138

Summary I. Introduction II. Localization of Intrinsic Membrane Proteins and Enzymes III. Future Focus Acknowledgments References

7

Phycobilisome and Phycobiliprotein Structures Walter A. Sidler

119 120 121 134 135 135

139–216

Summary I. Introduction II. Phycobilisomes III. Phycobiliproteins Constituting the Phycobilisome Core IV. Phycobiliproteins Constituting the Rod Elements of PBS V. Linker Polypeptides, the Skeleton of the PBS VI. Organization of the Genes Encoding the Phycobilisome Elements Acknowledgments References

8

The Use of Cyanobacteria in the Study of the Structure and Function of Photosystem II Bridgette A. Barry, Renee J. Boerner and Julio C. de Paula

140 141 143 152 162 191 199 205 205

217–257

Summary I. Introduction II. A Comparison of the Biochemical Properties of Cyanobacterial and Higher Plant Photosystem II III. Site-Directed Mutagenesis Studies of the Donor Side of Photosystem II IV. Biophysical Studies of Cyanobacterial Photosystem II V. Concluding Remarks References

9

The Cytochrome Complex Toivo Kallas

218 218 219 231 239 247 247

259–317

Summary I. Introduction II. Role of the Cytochrome Complex in Cyanobacteria III. Relation to Quinol-Cytochrome c Oxidases in Chloroplasts, Mitochondria, and Other Bacteria IV. Polypeptides, Redox Centers, Substrate Binding Sites, and Subunit Topology V. Electron and Proton Transfer Pathways VI. Three-Dimensional Structure and Biogenesis vi

259 260 261 261 264 280 286

VII. Genetics and Mutational Analysis VIII. Unresolved Questions and Perspective Acknowledgments References

10

Photosystem I in Cyanobacteria John H. Golbeck

319–360

Summary I. Introduction II. Unifying Principles III. Architecture of Photosystem I IV. Integral Polypeptides V. Peripheral Polypeptides Acknowledgments References

11

289 303 304 304

The F-type ATPase in Cyanobacteria: Pivotal Point in the Evolution of a Universal Enzyme Wayne D. Frasch Summary I. Introduction II. Organization of Subunits III. Gene Organization IV. Mechanism of ATP Synthesis V. Characteristics of the Metal-Nucleotide Binding Sites VI. Location of the Metal-Nucleotide Binding Site on the subunit—the Catalytic Site VI. Location of the Noncatalytic Metal-Nucleotide Binding Site VII. Model of the Metal-Nucleotide Binding Sites of CF1 VIII. Regulation of Catalytic Activity Acknowledgments References

320 320 321 323 329 343 354 354

361–380 361 362 363 364 366 369 369 372 373 373 376 376

381–407 12 Soluble Electron Transfer Catalysts of Cyanobacteria Larry Z. Morand, R. Holland Cheng, David W. Krogmann and Kwok Ki Ho Summary I. Ferredoxin II. Flavodoxin Reductase (FNR) III. Ferredoxin IV. Plastocyanin V. Cytochrome VI. Low-Potential Cytochrome c VII. Hydrogenase References

382 382 387 389 392 394 396 398 402

409–435

13 Cyanobacterial Respiration G. Schmetterer Summary I. Introduction II. Primary Electron Donors to the Respiratory Electron Transport Chain III. Primary Oxidoreductases IV. Quinone Pool complex V. Cytochrome VI. Peripheral Intermediate Electron Carriers vii

409 410 412 415 420 422 423

VII. Terminal Oxidases VIII. Conclusion Acknowledgments References

14

426 428 429 429

The Biochemistry and Molecular Regulation of Carbon Dioxide Metabolism in Cyanobacteria F. Robert Tabita

437–467

Summary I. Introduction II. Pathways of Carbon Dioxide Metabolism III. Enzymes of Fixation: Structure, Function, and Regulation of Activity IV. Organization of Reductive Pentose Phosphate Cycle Genes V. Regulation of Expression of Reductive Pentose Phosphate Cycle Genes VI. Conclusion Acknowledgments References

15

Physiological and Molecular Studies on the Response of Cyanobacteria to Changes in the Ambient Inorganic Carbon Concentration Aaron Kaplan, Rakefet Schwarz, Judy Lieman-Hurwitz, Michal Ronen-Tarazi and Leonora Reinhold Summary I. Introduction II. Adaptation to Changing Ambient Concentration and Gene Expression III. Mechanism of Inorganic Carbon Uptake IV. Role of Carboxysomes V. Types of Concentration-Dependent Mutants and the Relevant Genomic Lesions VI. Concluding remarks Acknowledgments References

16

Assimilatory Nitrogen Metabolism and Its Regulation Enrique Flores and Antonia Herrero Summary I. Introduction II. Nitrogen Fixation III. Nitrate and Nitrite Assimilation IV. Assimilation of Organic Nitrogen V. Assimilation of Ammonium VI. Distribution of Assimilated Nitrogen VII. Global Nitrogen Control Acknowledgments References

17

Biosynthesis of Cyanobacterial Tetrapyrrole Pigments: Hemes, Chlorophylls, and Phycobilins Samuel l. Beale Summary I. Introduction II. Tetrapyrrole Precursor Biosynthesis viii

437 438 438 439 457 460 462 462 462

469–485

469 469 471 473 477 480 482 483 483

487–517 488 488 489 497 502 504 509 510 511 511

519–558 520 520 521

III. The Pathway from PBG to Uroporphyrinogen III IV. Steps Leading to Siroheme and Corrins V. Conversion of Uroporphyrinogen III to Protoporphyrin IX VI. The Fe Branch VII. The Mg Branch Note Added in Proof Acknowledgments References

18

Carotenoids in Cyanobacteria Joseph Hirschberg and Daniel Chamovitz

528 529 530 531 538 547 548 548

559–579

Summary I. Introduction II. Functions of Carotenoids III. Analytical Methods IV. Carotenoid Composition in Cyanobacteria V. Carotenoproteins: The Cellular Location of Carotenoids VI. Biosynthesis of Carotenoids VII. Molecular Characterization of Carotenoid Biosynthesis VIII. Inhibitors of Carotenoid Biosynthesis IX. Regulation of Carotenoid Biosynthesis and Accumulation References

19

559 560 560 562 564 565 567 571 572 574 575

581–611

Genetic Analysis of Cyanobacteria Teresa Thiel

Summary I. Introduction II. Gene Transfer III. Mutagenesis IV. Reporter Systems V. DNA Elements VI. Mapping VII. Expression of Foreign Genes in Cyanobacteria VIII. Developing a Genetic System: Practical Problems and Possible Solutions Acknowledgments References

20

The Transcription Apparatus and the Regulation of Transcription Initiation Stephanie E. Curtis and James A. Martin Summary I. Introduction II. Transcription in E.coli: Paradigms for Eubacteria III. Transcription in Cyanobacteria Acknowledgments References

21

The Responses of Cyanobacteria to Environmental Conditions: Light and Nutrients Arthur R. Grossman, Michael R. Schaefer, Gisela G. Chiang and Jackie L. Collier

582 582 582 592 596 598 599 601 604 606 606

613–639 614 614 614 619 635 635

641–675

641 641 643

Summary I. PBS Structure II. Chromatic Adaptation

ix

III. The Responses of Cyanobacteria to Nutrient Deficiency IV. Concluding Remarks Acknowledgments References

22

Short-term and Long-term Adaptation of the Photosynthetic Apparatus: Homeostatic Properties of Thylakoids Yoshihiko Fujita, Akio Murakami, Katsunori Aizawa and Kaori Ohki

654 668 668 668

677–692

Summary I. Introduction II. Short-term Adaptation: The State Transition III. Long-Term Adaptation: Regulation of PS I:PS II Stoichiometry IV. Relationship Between Short-Term and Long-Term Adaptation Acknowledgments References

23

Light-Responsive Gene Expression and the Biochemistry of the Photosystem II Reaction Center Susan S. Golden

677 678 679 683 689 690 690

693–714

Summary I. PS II: Agent and Target of Environmental Variation II. PS II Genes of Cyanobacteria III. Response of psbA Genes to Changes in Light Intensity IV. Response of psbD Genes to Changes in Light Intensity V. Functional Significance of Light-Responsive Regulation VI. Light Quality and psbA Expression VII. Light-Regulated Gene Expression and the Biochemistry of PS II Proteins VIII. Cyanobacteria as Models for Studying Photoinhibition Mechanisms in vivo IX. Future directions Acknowledgments References

24

Thioredoxins in Cyanobacteria: Structure and Redox Regulation of Enzyme Activity Florence K. Gleason Summary I. Introduction II. Structure of Cyanobacterial Thioredoxins III. Reduction of Thioredoxins IV. Functions of Thioredoxins in Cyanobacteria References

25

Iron Deprivation: Physiology and Gene Regulation Neil A. Straus Summary I. Introduction II. Iron in Photosynthetic Electron Transport III. Responses to Iron Deprivation IV. The Control of Gene Expression by Iron Acknowledgments References

x

693 694 694 699 704 705 706 706 708 710 710 711

715–729 715 716 720 722 723 726

731–750 731 732 732 734 743 745 747

26

The Cyanobacterial Heat-Shock Response and the Molecular Chaperones Robert Webb and Louis A. Sherman Summary I. Background II. Functional Aspects, Protein Folding and Localization III. Molecular Chaperones of the Cyanobacteria IV. Summary and Future Directions References

27

Heterocyst Metabolism and Development C. Peter Wolk, Anneliese Ernst and Jeff Elhai Summary I. What is a Heterocyst? II. Genetic Tools III. Metabolism of Mature Heterocysts IV. The Differentiation Process V. Pattern Formation and Perpetuation VI. Relationship of Diverse Differentiation Processes in Cyanobacteria Acknowledgments References

28

Differentiation of Hormogonia and Relationships with Other Biological Processes Nicole Tandeau de Marsac Summary I. Introduction II. Occurrence of Hormogonia Among Cyanobacteria III. Factors Modulating the Production of Hormogonia IV. Morphological, Ultrastructural, Biochemical and Genetic Changes During the Differentiation of Hormogonia V. Relationships of Hormogonium Differentiation with Other Biological Processes VI. Hormogonia and Symbiosis VII. Further Prospects Acknowledgments References

751–767 751 752 757 762 764 765

769–823 770 770 773 774 793 803 810 811 811

825–842 825 826 826 829 830 833 837 838 839 840

Organism Index

843

Gene and Gene Product Index

849

Subject Index

855

xi

Preface

More than twenty years ago, as a fledgling graduate student who was just starting to learn about these organisms that would become my primary research focus, the publication of Noel Carr and Brian Whitton’s The Biology of the Blue-Green Algae in 1973 was an event of great significance. Until the appearance of this treatise, there was no single volume available that presented a broad overview of the biology and biochemistry of these organisms. Nearly ten years later, I was privileged to be a contributing author to Carr and Whitton’s sequel volume The Biology of the Cyanobacteria. Although the intervening period had been marked by heated debates over the taxonomy and taxonomic position of the organisms, it was also a time when the comparative biochemistry of the group was intensively investigated. The Biology of the Cyanobacteria, published in 1982, appeared after the onset of the molecular biological revolution during the late 1970’s; however, only a few researchers (notably Bob Haselkorn and coworkers) had very actively begun to apply these techniques to cyanobacteria. An examination of The Biology of the Cyanobacteria will show that the discussion of the molecular biology of these organisms was confined to approximately two pages of text under a section entitled ‘New approaches.’ Even by the time that Peter Fay and Chase Van Baalen’s The Cyanobacteria was published in 1987, the discussion of the molecular genetics of Cyanobacteria had only expanded to represent a single chapter. Many of us still recall when talks on the molecular genetics of cyanobacteria were confined to the Friday afternoon session of meetings when almost no one cared to listen to another talk any longer—especially one filled with gene jargon! A primary objective in the development of The Molecular Biology of Cyanobacteria was to summarize more than a decade of progress in analyzing the taxonomy, biochemistry, physiology, and cellular differentiation and developmental biology of cyanobacteria by modern molecular methods and especially by molecular genetics. The title was not chosen because the book focuses on

some peculiar aspects of the genetics of these organisms but to pay respects to the two volumes of Carr of Whitton that played important roles in my own thinking about Cyanobacteria (and no doubt in the development of many others as well). Contributing authors were asked to describe not only what we know at present, but also to point out things we don’t know yet. I have attempted to assemble a book that would stimulate graduate students and other researchers in the same way that I was affected by the books mentioned above. It appears that cyanobacterial molecular biologists have indeed paid attention to the admonition of their erstwhile colleague, W. Ford Doolittle, to ‘study those things that cyanobacteria do well.’ During the past ten years or so, cyanobacteria have become the organisms of choice for detailed molecular analyses of oxygenic photosynthesis. Appropriately, about half of this book is devoted to descriptions of the major components of the cyanobacterial photosynthetic apparatus and their biosyntheses. The component light-harvesting and electron transport complexes are discussed in detail in terms of their function and assembly, and special emphasis has been given to structural aspects. In some cases it has been necessary to extrapolate structural and functional details from analyses of similar complexes from other photosynthetic bacteria and higher plant chloroplasts. Molecular biology has also had an important impact on the taxonomy of cyanobacteria and the origins of chloroplasts in algae and higher plants, and further advances in this area are expected in the future. A very important part of the book includes selected examples of the responses of cyanobacteria to environmental stresses and resulting cellular differentiation and development events. We know now that cyanobacteria do regulate gene expression (forgive me for bringing this up, Noel), but the things that they perceive to be important (light intensity, light wavelength, nutrient availability, etc.) are rarely the same things that an enteric bacterium views as important. As occurs in almost any book dealing with such a large and complex xiii

support over the years, and friendship contributed immeasurably to my own development and to this project: Alex Glazer, Roger Stanier, Germaine Cohen-Bazire, Nicole Tandeau de Marsac, Rod Clayton, Noel Carr, Philip Thornber, Peter Wolk, and especially John Golbeck, who suffered (mostly in silence) while listening to my incessant whining about this project. Noel and Nicole: in many ways, one could say that this book was born in the street cafes of Paris some fifteen years ago now. Fourthly, I would like to thank my students and postdoctorals for putting up with me during the production of the book—I realize that I have been a part-time mentor at times. Finally, but certainly not least of all, I want to thank my wife, Vicki Stirewalt, for allowing me the time and space required to complete this project. Thanks for taking care of many little things that allowed this project to move forward. I know that I owe you big for this one!

array of processes and organisms, some topics (e.g., ecological aspects) could not be covered. However, one can expect that molecular taxonomic methods and polymerase chain reaction technology will have made a significant impact in such aspects in the very near future. As with any project of this magnitude, there are many persons that I would like to thank. Firstly, and most of all, I would like to thank all of the contributing authors for putting up with my editorial idiosyncrasies and requests for changes and additions. Without the constant encouragement of many of you, it is doubtful that this project could ever be completed. Secondly, I would like to thank Larry Orr for his time, heroic effort and understanding in producing the page layout for the book. Larry and I endured a lot together as we learned how to go about producing a book, and we were the test case for the entire series. Thirdly, I would like to thank those whose training, continuing

–Donald A. Bryant

xiv

Color Plates

Color Plate 1. Diagram showing the major respiratory and photosynthetic electron transport components of cyanobacteria. For details, see Chapters 6–13.

CP-1 D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. CP1–CP10. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

Color Plate 2. ( A ) Trimeric structure of C-phycocyanin from Mastigocladus laminosus (Schirmer et al., 1985, 1987). The polypeptide backbones of the and subunits are shown in brown and green, respectively. The phycocyanobilin chromophores are shown in blue. (B) Trimeric structure of Porphyridium sordidum phycoerythrin (Fiener et al., 1992). The polypeptide backbones of the and subunits are shown in yellow and blue, respectively. The phycoerythrobilin chromophores are shown in red. (See Chapter 7, p. 172, Fig. 14.)

CP-2

Color Plate 3. The polypeptide backbones of B-PE subunit (A) and B-PE subunit (B) from Porphyridium sordidum. The helices are denoted according C-PC (Schirmer et al., 1985). The phycoerythrobilin chromophores are shown in red. The polypeptide back bones of B-PE subunit (C) and B-PE subunit (D) from Porphyridium sordidum (shown in red) are superimposed on the polypeptide backbones of C - P C subunit and subunit from Mastigocladus laminosus ( s h o w n in blue). Note that the structures of the two proteins only deviate significantly in the vicinities of the additional chromophore binding loops in the B-PE subunits. (Figures kindly provided by R. Fiener and R. Huber). (See Chapter 7, p. 187, Fig. 22.)

CP-3

Color Plate 4. Arrangement of structural elements in the PS I monomer pictured with the graphics program O52. The crystallographic 3fold axis is indicated by the symbol Transmembrane helices are shown as blue, and horizontal helices as white cylinders. (A) Crosssectional view through the membrane with the stroma ‘above’, the lumen ‘below’, and the connecting domain on the left. Head groups of the antenna chlorophylls are shown as green disks, electron carriers as yellow, and F x , FB and FA are indicated as red spheres. (B) View from the stroma toward the lumen, with the roughly 2-fold axis that relates helices a to h and a' to h' in subunits A and B indicated by the symbol •. Reproduced from Krauß et al. (1993) with permission. (See Chapter 10, P. 327, Fig. 4.)

CP-4

Color Plate 5. Upper: Stereo ribbon diagram of one of the energy-minimized NMR solution structures of PsaE from Synechococcus sp. strain PCC 7002. Lower: Two views of a space-filling model for PsaE from Synechococcus sp. strain PCC 7002. The two charged faces of the protein are shown. Amino acids shown in blue are positively charged (Arg, Lys, and His); amino acids shown in red are negatively charged (Glu and Asp); amino acids shown in yellow are polar (Ser, Thr, Tyr, Asn, Trp); and amino acids shown in white are non-polar (Ala, Ile, Val, Leu, Phe, Gly and Pro). PsaE contains no Gln, Met or Cys residues. Figures courtesy of Drs. J. Lecomte, C. Falzone, and D. A. Bryant. (See Chapter 10, p. 353, Fig. 8.)

CP-5

Color Plate 6. The structures of spinach ferredoxin:NADP+ oxidoreductase (lower left) and Anabaena sp. strain PCC 7120 ferredoxin. The iron-sulfur center of ferredoxin is likely to be drawn down into the cleft over the FAD cofactor. The numbered residues identify points of interaction described in the text. (See Chapter 12, p. 384, Fig. 1.)

CP-6

Color Plate 7. The structures of Synechococcus sp. PCC 6301 cytochrome (upper right). Synechococcus sp. PCC 6301 flavodoxin (upper left), and Enteromorpha prolifera plastocyanin (lower). The flavodoxin structure is presented in an orientation similar to that of ferredoxin in Fig. 1. The FMN would be drawn down into the cleft in FNR over the FAD cofactor. The numbered amino acid residues on the lower edge of the flavodoxin would interact with basic residues in the region 85 to 93 of FNR. The cytochrome structure has been rotated 180° from the conventional representation of c-type cytochromes to locate its ‘acid patch’ at residues 69 to 71 in the same position as the ‘acid patch’ at residues 42 to 44 in the plastocyanin molecule. (See Chapter 12, p. 389, Fig. 2.)

CP-7

Color Plate 8. Computer graphics generated models of the Synechococcus sp. strain PCC 6301 RubisCO (courtesy of J. Newman and S. Gutteridge). Top and bottom panels represent top and side views, respectively of the enzyme. The top view is down the prominent fourfold symmetry axis of the molecule and shows a considerably large solvent cavity. The A/B dimer (green/ochre) is shown at the top right section of the upper panel, with the small red sphere indicating the position of at the active site. The small subunits are shown in purple. The side view (bottom panel) is along one of the two-fold axes, with the small subunits (purple) at the top and bottom of the molecule. For simplicity, only the A/B (right) and G/H (left) pairs of large subunit dimers are shown (green/ochre). The small red spheres indicate the in the A/B dimers; the barrel domain of the A subunit is clearly seen, with the Mg atom at the A/B domain interfaces. For more details, see Newman and Gutteridge (1993). (See Chapter 14, p, 442, Fig. 2.)

CP-8

Color Plate 9. Structural model for a complex formed between dinitrogenase reductase (Fe-protein) and dinitrogenase (Mo-Fe protein) from Azotobacter vinelandii. The figure shows the backbone for only one dimer of the dinitrogenase molecule. The dinitrogenase subunit is shown in blue and the subunit is shown in red; the dinitrogenase reductase dimer is shown in light blue. The metal centers in the two proteins (see Fig. 3, Chapter 16) and an ADP molecule at the interface between the two Fe-protein are represented by models (yellow) surrounded by a dotted van der Waals surface. The pink regions surrounding the Ke-Mo-Cofactor at the upper left in the subunit show the locations of histidines that could participate in proton transfer to the bound substrate. The two purple residues at the right (one on the Fe-protein and one on the subunit) show residues that can be chemically crosslinked in the complex of the two proteins. This model was generated by graphic superposition of the crystal structures of the individual proteins (Georgiadis et al, 1992; Kim and Rees, 1993). Figure courtesy of Dr. Douglas C. Rees, California Institute of Technology; reprinted with permission. (See Chapter 16, p. 493, Fig. 2.)

CP-9

Color Plate 10. The quaternary structure of Salmonella typhimurium glutamine synthetase. The molecule is shown as line segments connecting the 468 sequential atoms for each of the six subunits of the top layer (Panel A) and for the six nearer subunits of the two layers (Panel B). Each active site is indicated by a pair of spherical Mn2+ ions (blue). Six central loops protrude into the central aqueous channel in Panel A. The maximum dimensions of the molecule, including side chains are 103 Å along the six-fold axis and 143 Å along one of the two-fold axes perpendicular to the six-fold axis. Figure courtesy of Dr. David Eisenberg, University of California, Los Angeles. (See Chapter 16, p. 507, Fig. 7.)

CP-10

Chapter 1 Molecular Evolution and Taxonomy of the Cyanobacteria Annick Wilmotte* Department of Biochemistry, University of Antwerp (UIA), Universiteitsplein 1, B-2610 Wilrijk, Belgium Summary I. Introduction A. Definitions B. The Botanical and Bacteriological Approaches to the Study of the Cyanobacteria C. Relationship Between Evolution and Taxonomy D. Denomination of Strains II. Guidelines to the Taxonomy of the Cyanobacteria A. The Botanical Approach B. The Bacteriological Approach III. Fossil Record of the Cyanobacteria IV. Results of Chemotaxonomic Studies A. LipidComposition B. Polyamines C. Carotenoids D. Biochemical Features V. Results of Macromolecular Methods A. Protein Electrophoresis and Isozyme Patterns B. Phycobiliprotein Patterns C. Immunological Studies D. Restriction Fragment Length Polymorphism (RFLP) 1. The Marine Synechococcus sp. Strains 2. The Symbiotic Cyanobacteria in Azolla 3. The Symbiotic Cyanobacteria in Cycads and Gunnera sp E. DNA Base Composition F. DNA Fingerprinting G. DNA-DNA Hybridizations H. Proteins and Protein-Coding Gene Sequence Analysis I. 16S rRNA Gene Sequence Analysis 1. Properties of 16S rRNA 2. Levels of Relationship Investigated 3. Sequence Determination Methods 4. Sequence Alignment and Data Analysis 5. Results of 16S rRNA Sequence Analysis of Cyanobacteria a. Branches A and B b. Branch C c. Branch D d. Branch E e. Branch F f. Branch G g. Branch H

2 2 2 3 3 4 4 4 4 5 5 5 7 7 7 7 7 8 8 8 8 9 9 9 10 10 11 12 12 12 12 12 14 17 17 18 19 19 20 20

* Present address: Laboratory of Genetics and Biotechnology, Vlaamse Instelling voor Technologisch Onderzoek, Boeretang 200, B-2400 Mol, Belgium D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 1–25. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

Annick Wilmotte

2

6. Comparison with the Current Evolutionary and Taxonomic Hypotheses 7. Possible Developments of the Use of the rRNA Cistrons to Study the Evolution and Taxonomy of the Cyanobacteria VI. Conclusion Acknowledgments References

20 21 21 21 22

Summary Molecular biology has provided new tools to decipher genetic information and can be used in attempts to reconstruct the evolution of organisms and improve their taxonomy. In the cyanobacteria, the use of molecular methods to study the genotypic relationships is underway, and initial results are promising. Different chemotaxonomic and macromolecular techniques are reviewed and their usefulness is evaluated. The most complete phylogenetic scheme of the cyanobacteria which is presently available is based on 16S rRNA sequence analysis. With this method, controversial taxonomic problems have been solved, such as the relationships among Pseudanabaena sp. strains or between the generaArthrospira and Spirulina. In other cases, additional 16S rRNA sequences are necessary to obtain a clear picture. In addition to the cultivated strains, molecular ecology studies have contributed to the determination of new 16S rRNA sequence types, that have been retrieved directly from natural populations. The corresponding morphologies are presently unknown but may be revealed by the use of labeled probes annealing to specific 16S rRNA regions. For taxonomic purposes, it is necessary to find morphological features and simple testing methods which are congruent with the genotypic groupings. This information may be used to evaluate and revise existing classifications. The first stage in the development of such a polyphasic taxonomy is now underway. I. Introduction In a review of the molecular evolution of cyanobacteria, Doolittle (1982) asked three questions. The first one concerned the phylogenetic position of the cyanobacteria within the procaryotes. The answer is that the cyanobacteria form one of the eleven major eubacterial phyla, as shown convincingly by the analysis of 16S rRNA sequences (Woese, 1987), 23S rRNA and protein sequences, such as the elongation factor Tu and the of ATP-synthase (Schleifer and Ludwig, 1989). The second question, the relationship between cyanobacteria and plastids, is treated by Susan Douglas in the Chapter 5 of this book. Thethird question, concerningthephylogenetic relationships among the cyanobacteria and the taxonomic implications of those relationships, is the subject of this chapter. Abbreviations: CCAP–Culture Collection of Algae and Protozoa; DAF – DNA amplification fingerprinting; G+C – guanine + cytosine; ITS – internal transcribed spacer; LPP – LyngbyaPlectonema-Phormidium; PCC – Pasteur Culture Collection; PCR – polymerase chain reaction; RFLP – restriction fragment length polymorphism; rRNA – ribosomal ribonucleic acid; SDS–sodium dodecyl sulfate; STRR–short tandemly repeated repetitive

Molecular information on phylogenetic relationships of organisms can be obtained by chemotaxonomic studies (fatty acids, quinones, carotenoids, etc.) and analyses of macromolecules (nucleic acids and proteins). The taxonomic utility of these molecules, or their shortcomings, is illustrated in this chapter. For students or readers not familiar with these topics, the actual status of evolutionary hypotheses and taxonomic schemes concerning the cyanobacteria is briefly presented, as well as the contribution of the botanical and bacteriological approaches to these topics and the information given by the fossil record.

A. Definitions ‘Systematics’ is defined as the comparative study of all the properties of organisms which can be used for .their taxonomy. ‘Taxonomy’includes ‘classification’, ‘Which is the arrangement of organisms in an orderly manner; ‘identification’ of new organisms; and ‘nomenclature, ’ that is concerned with the naming oforganisms and the rules governing the use ofthese names. Though they are sometimes considered as synonyms, taxonomy is only a part of systematics. The fundamental taxonomic unit is the ‘species,’ for

Chapter 1 Molecular Approach to Cyanobacterial Evolution and Taxonomy which many different definitions have been given and which is the topic of a comprehensive review by Castenholz (1992). ‘Phenotypic’ features are the outer manifestations of the genetic information of the organism, and are the result of complex interactions of molecules with each other and the environment; these interactions lead to recognizable differences in morphology, physiology, and biochemistry. ‘Genotypic’ characters include the genes and their direct products, the proteins. Phenotypes and genotypes are currently used to attempt to reconstruct the ‘phylogeny’ of organisms, the hierarchy resulting from the evolutionary processes. Hereby, only vertical evolution, the descent with modifications from an ancestor, is generally considered. Because genotypic characters, like DNA
DNA hybridizations or gene sequences, are considered to reflect well evolutionary processes, they are sometimes called ‘phylogenetic’ characters (Murray et al., 1990).

B. The Botanical and Bacteriological Approaches to the Study of the Cyanobacteria For a student with no prior knowledge of the cyanobacteria, the situation may seem very confusing. Several names are currently used for these organisms, including ‘cyanobacteria,’ ‘cyanophyceae,’ ‘cyano
phytes’ and ‘blue
green algae.’ The explanation is historical and a detailed review is given by Castenholz and Waterbury (1989). Until electron microscopy and biochemical analyses could show convincingly that the cyanobacteria were procaryotes, these organisms were generally considered as algae and were studied by botanists and phycologists. In nature, the cyanobacteria usually behave like algae. They possess chlorophyll a and perform oxygenic photosynthesis. Thus, the choice of one or another denomination is a matter of taste and reflects the interests, or more frequently the prejudices, of the authors. Botanists and bacteriologists have different backgrounds and experimental approaches. Dialogue between the two groups is necessary for mutual enrichment and progress.

C. Relationship Between Evolution and Taxonomy Before molecular approaches were developed, the relationship between evolution and taxonomy was considered differently in the botanical and bacter
iological worlds. For eucaryotic algae and higher

3

plants, complex morphological characters and fossil traces were generally present, and the idea that taxonomy should try to reflect the degree of phylogenetic relationships was well
established (Stuessy, 1990). Different evolutionary schemes based on the morphology of the cyanophyceae have been proposed by botanists and are reviewed by Desikachary (1973). Simple morphology was either assumed to be primitive (‘progressive evolution’) or derived by simplification from more complex forms (‘retrogressive evolution’). Schwabe (1960) even proposed a ‘reticular’ system where different morphological types evolved from one thermophilic ancestor with a complex morphology and were connected through intermediate taxa. On the bacteriological side, the shortcomings of morphological features were so evident that phylogenetic schemes were dismissed as pure speculation (Woese, 1987). Most bacterial classi
fications were based on phenotypic properties and had no ambitions to reflect evolutionary relationships. The suggestion that the structure of molecules contained information on the phylogeny of the organisms was made by Zuckerkandl and Pauling in 1965. The technical progress made in molecular biology has provided suitable tools to infer genotypic relationships and revolutionized the approach to the evolution and taxonomy of the living beings. Presently, the integrated use of phylogenetic and phenotypic characteristics, called ‘polyphasic’ taxonomy, is recommended by bacterial taxonomists (Murray et al., 1990). In cases where phylogenetic methods give different results than the phenotypic characters, these authors suggest that priority should provisionally be given to the latter and advise that thorough analysis of the phenotypes be performed to resolve the discrepancy. Thus, the ultimate goal of the modern bacterial taxonomy is to reflect the phylogenetic relationships to the greatest extent possible. This will to integrate phenotypic and genotypic characters is also clearly demonstrated on the botanical side by Anagnostidis and Komárek (1985, 1988, 1990), and Komárek and Anagnostidis (1986, 1989) in their modern approach of the classification of the cyanophytes. Unfortunately, the molecular data available for the cyanobacteria are still too fragmentary. Therefore, morphology, with its advantages and shortcomings (Wilmotte and Golubić, 1991), is still largely the basis used for the botanical or bacteriological taxonomy of cyano
bacteria.

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D. Denomination of Strains A major problem in the comparison and interpretation of taxonomic results is that the identification of strains may be incorrect. Komárek and Anagnostidis (1989) stated that the features of more than 50% of strains in the collections do not correspond to the diagnoses of the taxa to which they are assigned. Thus there is a real need for further characterization of the numerous cyanobacterial cultures available worldwide in laboratories and collections. In different taxonomic systems, the same taxon denominations (e.g., genus, order) have been used with different meanings. Therefore, when confusion is possible, the authors are mentioned by name. Finally, another problem is that the name of certain strains has changed. For example, the strain ‘Anabaena’ sp. strain PCC 7120 (Rippka et al., 1979) has been renamed Nostoc sp. strain PCC 7120 on the basis of DNA-DNA hybridization data (Lachance, 1981) and its hybridization patterns with highly repetitive (STRR) DNA sequences (Mazel et al., 1990). In such cases, the wrong name will be placed between ‘inverted commas’. II. Guidelines to the Taxonomy of the Cyanobacteria Excellent reviews of the historical development of the cyanobacterial taxonomy have been written by specialists including Desikachary (1973), Anagnostidis and Komárek (1985), Castenholz and Waterbury (1989) and Whitton (1992). However, a summary may be necessary to understand this chapter and this is given below.

Annick Wilmotte respectively. Since the last century, numerous new species have been described. In Geitler’s determination key published in 1932, about 1300 species and 145 genera were recognized. This Flora was devised for Germany, Austria and Switzerland but it was used all over the world and is still the basis of numerous taxonomic works. Among botanical taxonomists, there is a suspicion that too many species have been described over the years; many are based on a single character difference, such as the presence or absence of sheath or slight deviations in cell dimensions or forms (Anagnostidis and Komárek 1988,1990; Komárek and Anagnostidis 1986,1989). The problem of the morphological variability has prompted Drouet (1968) to revise the taxonomy profoundly. His basic idea was that there existed ecophenes, that were organisms sharing the same genotype but expressing distinct morphologies under the influence of environmental factors. He drastically reduced the number of species down to 62 by selecting certain morphological features which he believed to be invariant with the environment. Classical taxonomists were quite critical of this approach (Desikachary, 1973; Anagnostidis and Komárek, 1985). Later, DNA-DNA hybridizations showed that taxa placed by Drouet in the same species were genotypically different (Stam and Venema, 1975; Stam, 1980). Recently, a new and deeply reorganized taxonomic revision was published by Anagnostidis and Komárek (1985, 1988, 1990) and Komárek and Anagnostidis (1986, 1989). This revision is based on the definition of smaller, more coherent genera. The authors made an extensive review of the literature and tried to integrate all the biochemical, ultrastructural and molecular characters available with their considerable taxonomic experience.

A. The Botanical Approach B. The Bacteriological Approach Chronologically, the botanical approach was first to put its stamp on the cyanobacterial taxonomy. As for other algae, the classical taxonomy of the cyanobacteria is based on morphological features and their nomenclature is ruled by the Botanical Code. This means that each new species has to be described in Latin, and that its reference is a herbarium specimen. For the simple filamentous (Oscillatoriaceae) and the heterocystous species (Nostocaceae and Stigonemataceae), the starting points for valid publication of names are the monographs written by Gomont( 1892) and Bornet and Flahaut (1886–1888),

The classical botanical taxonomy was confronted in the seventies with a rather different approach: the bacteriological one. R. Y. Stanier and colleagues advocated that, since cyanobacteria were bacteria, their taxonomy should be treated accordingly and their nomenclature governed by the Bacteriological Code (Stanier etal., 1978). For example, the reference for each species would become a pure culture instead of a herbarium specimen. The possibility of having the same organism described under two different names in the Botanical and Bacteriological codes

Chapter 1 Molecular Approach to Cyanobacterial Evolution and Taxonomy would have created chaos. Mutual concessions and adaptations of the two Codes have ensured that species described validly under one of them were recognized under the other. Stanier and collaborators pioneered the use of physiological and genotypic characters determined with axenic cultures; characters employed included the pigment composition, fatty acid analysis, heterotrophic growth, nitrogenase activity, DNA base composition, and genome length (Kenyon et al. 1972, Herdman et al. 1979a,b; Rippka et al., 1979). Because the cyanobacteria are photoautotrophic organisms, the physiological studies did not furnish many useful taxonomic characters. The basis of the bacteriological taxonomy of the cyanobacteria was published by Rippka et al. (1979). This taxonomic system, which still relies largely on the morphology, allows the identification of the strains of the Pasteur Culture Collection at the generic level. The five major sections recognized by Rippka et al. (1979) coincide broadly with orders of other classifications, as illustrated in Table 1. A modified version of this system is given in the appropriate sections of Bergey’s Manual of Systematic Bacteriology written by Castenholz (1989a, b, c), Waterbury (1989) and Waterbury and Rippka (1989). This treatment is more global and includes taxonomic information on well-known taxa which are not in the Pasteur Culture Collection as well as ecological features. Rippka and Herdman (1992) have published a catalogue of the strains available in the Pasteur Culture Collection and prepared a taxonomic handbook. It is noteworthy that some of the revisions proposed by Anagnostidis and Komárek (1988) have been adopted. The previous paragraphs show that the taxonomy of the cyanobacteria is in constant evolution. Though it is confusing and not desirable, classification and nomenclature may vary because they reflect the current state of knowledge (Stackebrandt and Goodfellow, 1991). III. Fossil Record of the Cyanobacteria The great antiquity of the cyanobacteria is well documented, though the fossil record is fragmentary and biased towards the formations where preservation of the morphology was possible. The earliest unicellular and filamentous forms attributed to the cyanobacteria were found in sedimentary rocks formed 3500 million years ago. Endolithic forms

5

that reproduce with baeocytes were observed in rocks formed ca 1500 million years ago. Heterocystous forms, and forms with true branchings seem to have appeared more recently, after the Precambrian (Schopf and Walter, 1982; Schopf and Packer, 1987). As pointed out by Castenholtz (1992), however, the fossil cyanobacteria are morphologically strikingly similar to their actual present-day counterparts, which raises questions about the speed of evolutionary processes in the cyanobacterial lineage. IV. Results of Chemotaxonomic Studies Studies of cyanobacterial taxonomy by molecular methods, in the broad meaning of the word ‘molecular,’ should also include chemotaxonomic markers. Quite easy and rapid determination methods are generally available. However, data are relatively scarce and several studies are still preliminary. Problems of consistency and variations due to factors such as growth conditions have not always been systematically investigated (Holton, 1981). Moreover, since the molecules employed are typically synthesized through complex pathways, their presence or absence can have different causes. The chemotaxonomic markers presented below have been shown to be useful, or seem worthy of further study.

A. Lipid Composition The fatty acid composition of 66 cyanobacterial strains was studied by Kenyon and collaborators (Kenyon, 1972; Kenyon et al. 1972). A uniform fatty acid composition was observed for the groups Anabaena and Calothrix, but not for other taxonomic groupings. Not enough molecular data on the same strains are available for a comparative study. Sallal et al. (1990) have detected a highly polar, unknown glycolipid present only in the three heterocystous strains studied. They also showed that alcohol glycosides are not restricted to nitrogen-fixing strains. Caudales and collaborators (Caudales and Wells, 1992; Caudales et al. 1992) have determined the fatty acid compositions from free-living strains of the genera Nostoc and Anabaena sensu Rippka et al. (1979) and from symbionts of the water fern Azolla sp. Following their interpretation of the results, the symbionts are equally distant from both genera. Three marine, picoplanktonic Synechococcus strains had a similar fatty acid composition to the freshwater

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Annick Wilmotte

Chapter 1 Molecular Approach to Cyanobacterial Evolution and Taxonomy strain Synechocystis sp. strain PCC 6308 (Merritt et al., 1991). However, the 16S rRNA sequence analysis does not show a close genotypic relationship between these strains (J. B. Waterbury, personal communication to R. Rippka). Finally, the lipid analysis of Prochloron sp. indicated a closer relationship with the cyanobacteria than with the eucaryotic algae (Perry et al., 1978).

B. Polyamines The use of polyamines as chemotaxonomic marker in bacterial systematics has recently been reviewed (Hamana and Matsuzaki, 1992). The dominant polyamine in the cyanobacterial strains hitherto tested was either spermidine or sym-homospermidine. However, Hegewald and Kneifel (1983) observed that both types of polyamine were present in different strains assigned to the genera Oscillatoria, Phormidium, Calothrix and Chroococcus (using the botanical taxonomy). The authors warned that the taxonomic affiliation of some of their strains could be incorrect. Their data contradict the conclusion by Hamana et al. (1983) that higher concentrations of sym-homospermidine were present in nitrogen-fixing strains. In fact, this apparent relation was possibly due to the smaller sample of strains studied by the latter authors.

C. Carotenoids The results of previous analyses by chromatography of the carotenoid composition ofcyanobacteria were compiled by Hertzberg et al. (1971). They concluded that the variations in patterns were probably useful for species identification but not for determinations at a higher taxonomic level. Indeed, similar carotenoid patterns were observed for Phormidium ectocarpi strain PCC 7375 and ‘Phormidium persicinum’ strain CCAP 1462/5 (Healey, 1968), which have almost identical 16S rRNA sequences (Wilmotte et al., 1992; see Section V I.). However, the carotenoid content and composition was different in red or green isolates of the same species (Aakermann et al., 1992), suggesting the limited taxonomic utility of this character.

D. Biochemical Features Hall et al. (1982) examined the enzymology and regulatory patterns of aromatic amino acid pathway

7

in strains from the five sections defined by Rippka et al. (1979). The biochemical diversity allowed the distinction of subgroups within several of the genera including Synechococcus, Synechocystis, Anabaena, Nostoc, and Calothrix. This confirms the heterogeneity ofthese genera and suggests that biochemical diversity could provide useful taxonomic markers. However, these five genera were also represented by the greatest number of strains, and it is questionable whether a more detailed survey of other genera would have revealed similar heterogeneities. V. Results of Macromolecular Methods Macromolecules, nucleic acids and proteins, are copies or translations of the genetic information and thus may be the best tools to infer phylogenetic relationships (Murray et al., 1990). The 16S rRNA has given the most detailed hypothesis on the evolutionary relationships within the cyanobacteria and will be discussed in most detail. Macromolecules can be studied directly, by sequencing, or indirectly by electrophoresis, hybridization, or immunological methods.

A. Protein Electrophoresis and Isozyme Patterns In a review, Holton (1981) observed that few studies made use of isozyme patterns, though he believed that they were useful for taxonomic studies at the genus and species level. An interesting study by Klein et al. (1973) showed the taxonomic utility of esterase isozyme patterns. Shared bands allowed the recognition of four clusters of related strains among the thirteen Oscillatoriaceae strains tested. On the basis of their similarity, the strains Phormidium ectocarpi strain PCC 7375 and ‘Phormidium persicinum’ strain CCAP 1462/5 (F. T. Haxo and D. J. Chapman, personal communication) were assigned to the same species. This result was later confirmed by 16S rRNA sequence analysis (Wilmotte et al., 1992; see Section V I). Malate-dehydrogenase electrophoretic patterns were used to characterize eight cyanobacterial strains, and shared bands were observed within the genera Anabaena and Nostoc. On the contrary, the two unicellular strains, Synechococcus sp. strain PCC 6301 and Synechococcus elongatus strain CCAP 1497/1, had no bands in common (Schenk et al.,

8

1973). Stulp and Stam (1984) studied the electrophoretic patterns forfive enzymes in axenicAnabaena sp. strains. They showed considerable intra- and interspecific heterogeneity of enzymes in the strains tested. These patterns were used to confirm that two Anabaena sp. strains were in fact identical. Finally, zymograms for six enzymes were identical for twelve cyanobacterial symbionts isolated from five cultivated Zamia integrifolia (cycad) plants in one locality. However, there were differences in the band patterns obtained for the symbiont of a diiferent cycad species and for a Nostoc species isolated from the soil at the same site (Zimmerman and Rosen, 1992).

B. Phycobiliprotein Patterns Schenk and Kuhfittig (1983) have investigated the phycobiliprotein patterns in twenty-one cyanobacterial species by means of polyacrylamide discgel electrophoresis. A conspicuous heterogeneity was observed, even between strains assigned to the same species. Therefore, the authors concluded that this method was useful only for the identification of identical strains but not for establishment of classification schemes. During a thorough survey of the genus Pseudanabaena, Guglielmi and CohenBazire (1984b) determined the electrophoretic patterns of the phycobiliprotein subunits by SDSpolyacrylamide gel electrophoresis. They proposed to restrict the genus Pseudanabaena to the strains able of synthesizing four subunits of phycocyanin (e. g., strains PCC 6903 and PCC 7409) and excluded the strain PCC 7403 from this genus. This divergence is supported by other molecular markers (see Sections V G and V I). Finally, Bryant (1982) examined the entire Pasteur Culture Collection for strains capable of phycoerythrocyanin synthesis. Although it was recognized that the inability to synthesize this phycobiliprotein provides no useful taxonomic information, the ability to produce this protein is probably quite useful as an exclusionary character at the genus level for filamentous, heterocystous strains. It is also interesting to note that the ability to form phycoerythrocyanin is found in the members of the genus Chroococcidiopsis (Bryant, 1982).

C. Immunological Studies Immunological comparisons of proteins have been used for bacterial systematics (Schleifer and

Annick Wilmotte Stackebrandt, 1983), but have rarely been used for studies of the cyanobacteria. Ladha and Watanabe (1982) have observed a high degree of antigenic similarity among cyanobacterial symbionts from different species of Azolla but no cross-reactions between the symbionts and free-living cultures (see Section V D, 2). Zilinskas and Howell (1987) showed that the antigenic determinants of two rod linkers polypeptide were very conserved in nine strains belonging to sections I, III, IV and V, whereas for a third polypeptide, only strains of section IV crossreacted. Marine picoplanktonic Synechococcus sp. strains belonging to the same serogroup (polyclonal antibody) appeared to be genotypically quite diiferent (Wood and Townsend, 1990; see Section V D, 1). These authors suggested that monoclonal antisera directed against specific surface epitopes could improve the sensitivity of the method. Finally, immunological characterization by Bullerjahn et al. (1990) showed that the chlorophyll a/b-binding protein from Prochlorothrix hollandica was very similar to its counterpart in Prochloron sp., but not to light-harvesting chlorophyll a/b proteins of maize.

D. Restriction Fragment Length Polymorphism (RFLP) The RFLP technique is generally useful to identify and classify organisms at the population or the species level. It has been productively used to study marine Synechococcus sp. strains and cyanobacterial symbionts.

1. The Marine Synechococcus sp. Strains The taxonomy of small, planktonic, phycoerythrincontaining Synechococcus sp. strains is problematic. Wood and Townsend (1990) tested the genetic homogeneity of a group of eight strains that showed cross-reactivity to the antiserum directed against Synechococcus sp. strain WH 7803. The probes were derived from several heterologous genes of ‘Anabaena’ sp. strain PCC 7120 and from genes of Synechococcus sp. strain WH 7803. In the tree topology obtained, the eight strains were distributed into four different branchings. Moreover, the genetic distances between terminal taxa on different branches within the serogroup were as large as the distances found between members of the serogroup and two freshwater Synechococcus sp. strains that show no

Chapter 1 Molecular Approach to Cyanobacterial Evolution and Taxonomy cross-reactivity to the antiserum. This conclusion was confirmed by an analysis of the previously published results of Douglas and Carr (1988).

2. The Symbiotic Cyanobacteria in Azolla A filamentous, nitrogen-fixing symbiont lives in the leaf cavities of the water fern Azolla sp. and is generally identified as Anabaena azollae, although it is not clear whether one or more taxa are involved. The symbiont and its host are associated during all the life cycle. In total, the symbioses involving seven Azolla sp., belonging to the sections Euazolla and Rhizosperma, have been studied. The probes were derived from heterologous genes from ‘Anabaena’ sp. strain PCC 7120 (Franche and Cohen-Bazire, 1987; Meeks et al., 1988;Gebhardt and Nierzwicki-Bauer, 1991) or genomic and plasmid DNA clones of freshly isolated symbionts (Plazinski et al., 1990, 1991). The results have shown that (i) the leaf cavities of Azolla sp. probably harbored one major symbiont, accompanied by minor species; (ii) there is a good correlation between the genotypic relationships among the freshly isolated symbionts and the classification of their hosts. The division into the sections Euazolla and Rhizosperma was confirmed, except for the symbiont from Azolla nilotica which showed genotypic differences with the symbionts from both sections; (iii) Only the minor constituents of the symbiosis could be established as free-living cultures. Plazinski et al. (1990) found species-specific probes, of which two were in fact derived from plasmid sequences (Plazinski et al., 1991). In addition, three probes could discriminate among symbionts from the same Azolla sp. collected in different geographical areas. The free-living ‘Anabaena’ sp. strain PCC 7120 shared little similarity to the studied symbionts, but Nostoc cycas strain PCC 7422 seemed closely related to them. The taxonomic affiliation of the symbionts to the genera Anabaena or Nostoc was also investigated. Using as taxonomic marker the presence of two conserved restriction enzyme sites in the nif genes of four Nostoc sp. strains and their absence in ‘Anabaena’ sp. strain PCC 7120 and ‘Anabaena variabilis’ strain ATCC 29413 (PCC 7937), Meeks et al. (1988) proposed that the symbionts belonged to the genus Nostoc rather than Anabaena sensu Rippka et al. (1979). However, the delimitation of the two genera is not clear and the two ‘Anabaena’ strains

9

used as reference by the authors have later been assigned to Nostoc (see Section I D; Caudales and Wells, 1992). Thus, the data from Meeks et al. (1988) only illustrate the genotypic variability existing within the genus Nostoc but do not allow classification of the symbionts.

3. The Symbiotic Cyanobacteria in Cycads and Gunnera sp. The symbionts associated with cycads and angiosperms are classically identified as members of the genus Nostoc. Lindblad et al. (1989) used heterologous probes from ‘Anabaena’ sp. strain PCC 7120 to characterize the symbionts freshly isolated from five cycad species from Central America. The results suggest that most cycad species contained one dominant symbiotic strain associated with one or several minor symbionts. For one cycad species, three free-living cultures appeared to be identical to the freshly isolated symbiont. The possibility of obtaining the symbiotic organism of this cycad in the free-living state may be related to the type of association, which involves the colonization of each new plant by the symbiont. In the case ofangiosperm symbioses, twelve freeliving cultures derived from symbionts of eight Gunnera species growing in Sweden, New Zealand and U.S.A. were characterized by Zimmerman and Bergman (1990). Two methods were used: the RFLP analysis using heterologous probes from ‘Anabaena’ sp. strain PCC 7120 and the immunostaining of the protein profiles. Three Nostoc sp. isolates from Gunnera sp. plants cultivated in a greenhouse in Sweden were identical whereas a fourth Swedish plant grown outdoors harbored a different symbiont. Six different genotypes were observed among the other isolates. These results suggest that more than one Nostoc species is involved in the symbiosis with Gunnera sp. plants and that the selection of the symbiont depends probably on the presence of compatible species in the environment.

E. DNA Base Composition DNA base composition is one of the few molecular characters that has been determined for almost 200 cyanobacterial strains (Herdman et al. 1979a). It is a ‘one-way’ taxonomic marker, however. Large differences in DNA base composition indicate that the strains cannot be closely related, whereas similar

10

G+C percentages give no clue concerning genotypic relationships.

F. DNA Fingerprinting Three short tandemly repeated repetitive (STRR) sequences from Calothrix sp. strain PCC 7601 hybridized only with the digested genomic DNA of the heterocystous strains tested. Mazel et al. (1990) suggested that the banding patterns could be used to distinguish strains at the species and genus level. It is noteworthy that repetitive elements have also been used to generate species- and strain-specific fingerprints in eubacteria (Versalovic et al., 1991). The taxonomic value of two insertion elements from Calothrix sp. strain PCC 7601 appeared to be restricted to the identification of identical strains (Mazel et al., 1991). DNA amplification fingerprinting (DAF), a PCRbased method using short oligonucleotides for production of characteristic banding patterns, has been used by Eskew et al. (1993) to study the cyanobacterial symbionts of Azolla sp. ferns. The authors generated fingerprints that were unique to the symbionts from three different Azolla sp. and were able to show the maternal transmission of one symbiont. This method is very promising for strain identification.

G. DNA-DNA Hybridizations Wayne et al. (1987) proposed the use of DNA-DNA hybridizations as a criterion for the definition of eubacterial species. For members of the same species, at least 70% hybridization is required, and 20% is required for congeneric strains. The use of this method for the cyanobacteria was pioneered by Stam (Stam and Venema, 1975; Stam 1980), who used a filterhybridization technique. The genotypic relationships among twenty-nine filamentous strains which belonged to the LPP group sensu Rippka et al. (1979) were determined. The results showed the genotypic homogeneity of a group of nineteen strains around the freshwater strain Plectonema boryanum strain PCC 73110. Other freshwater and marine LPP strains showed conspicuous genotypic differences. Comparison of the results with morphological characters indicated that the presence of false branching, used to distinguish the genus Plectonema from Phormidium sensu Geitler (1932), was variable within one species and its taxonomic relevance was

Annick Wilmotte questionable. Lachance (1981) investigated by DNA-DNA hybridizations the genotypic relationships among the heterocystous genera of sections IV and V The genera Nodularia, Cylindrospermum, Chlorogloeopsis and Fischerella sensu Rippka et al. (1979) appeared to form tight genotypic groupings. On the other hand, the genera Anabaena, Nostoc and Calothrix contained strains or clusters of strains which showed little genotypic similarity. Guglielmi and Cohen-Bazire (1984b) used DNADNA hybridizations to elucidate the relationships of nine strains belonging to the genus Pseudanabaena and the LPP group sensu Rippka et al. (1979). The results supported the distinction of two subgroups containing either two or four phycocyanin subunits. They also show the genotypic divergence of strain Pseudanabaena sp. strain PCC 7403, which correlates well with differences in pigment composition (see Section V B) and 16S rRNA sequence analysis (see Section V I). Stulp and Stam (1984) performed DNA-DNA hybridization studies with twenty-one Anabaena sp. strains. Strains assigned to the same species sensu Geitler (1932) on the basis of morphology indeed showed very high hybridization percentages (about 100%). On the other hand, strains from different species showed intermediate hybridization values. Wilmotte and Stam (1984) demonstrated by DNADNA hybridizations that the strains PCC 7942 and PCC 7943 belong to the same species as Synechococcus sp. strain PCC 6301, which is often wrongly designated as ‘Anacystis nidulans’ (Komárek, 1970). This result was later confirmed by results of RFLP analysis for strains PCC 6301 and PCC 7942. The genomes of the two strains seem identical, except for a rearrangement (Golden et al., 1989; Wood and Townsend, 1990). DNA-DNA reassociation studies have also used to investigate the genotypic diversity of Prochloron sp. isolates from different didemnid species and locations. The fourteen Prochloron sp. isolates appeared to belong to a single species (Stam et al., 1985; Holton etal., 1990). A new method of DNA-DNA hybridization, based on optical renaturation rates, has shown that five strains of Microcystis sp. isolated in Europe and North America belong to the same species (M. Herdman, personal communication). Interestingly, the strain Microcystis sp. strain PCC 7005 had previously been assigned to the genus Synechocystis

Chapter 1 Molecular Approach to Cyanobacterial Evolution and Taxonomy because it does not produce toxins nor gas vesicles (Rippka et al., 1979).

H. Proteins and Protein-Coding Gene Sequence Analysis A number of cyanobacterial proteins and proteincoding genes have been sequenced (review by Tandeau de Marsac and Houmard, 1987; see gene index for this book), but generally this has been performed in the course of genetic studies and not with an evolutionary framework in mind. Therefore, molecular information is scattered over a few organisms and gives a poorly resolved tree topology. A few sequences of plastocyanin and cytochrome have been determined (see Chapter 12) but were analyzed only in relation to the origin of plastids (Aitken, 1988). Masui et al. (1988) published a tree topology based on twenty sequences from two types of ferredoxin (Fig. 1). However, the strain selection was heavily biased towards unicellular and heterocystous cyanobacteria. Following this tree, the

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heterocystous strains are divided into two lineages. This contradicts the results from the 16S rRNA analysis (see Section V I). However, in addition to the sample bias mentioned above, the relatively small number of characters (about 100), the presence of functional constraints on a number of positions, and the occurrence of several ferredoxin isoforms may obscure the evolutionary pattern (Meyer et al., 1986). The partial sequence of the nifH gene was compared between acultivated Trichodesmium sp. strainisolated from Japan and natural populations of Trichodesmium sp. strains from the Caribbean Sea. A sequence similarity of 98% was observed (Zehr et al., 1990). More sequences are needed to determine the usefulness of the nifH gene for taxonomic studies of Trichodesmium species. A potential complication in using the nifH gene is the presence of the chlL gene; this gene, that exhibits high sequence similarity to nifH, encodes a subunit of protochlorophyllide reductase (see Chapter 17). Finally, it should be mentioned that strains that do not express phenotypic characters such as heterocysts and gas vesicles may

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Annick Wilmotte

still possess the genes responsible for the formation of these features (Damerval et al., 1989; Rippka and Herdman, 1992).

I. 16S rRNA Gene Sequence Analysis The 16S rRNA has been shown to be an adequate macromolecule to study the evolution of the eubacteria. Therefore, its properties and use (Woese, 1987) will be presented in more detail.

2. Levels of Relationship Investigated For the eubacteria, the 16S rRNA sequence analysis is used to determine a broad range of relationships, from phylum to species. In contrast with the DNADNA hybridization method, no limits of sequence similarity have been given to delineate the taxonomic level of the relationships. A comparison of DNADNA hybridization percentages and 16S rRNA similarities gave contradictory results (Fox et al., 1992; Ward et al., 1992).

1. Properties of 16S rRNA

3. Sequence Determination Methods The 16S rRNAs are universal molecules. Their similar structure in all living beings suggests that they evolved very early from the same ancestor and changed relatively little since their origin. Their function in protein synthesis is essential for the cell life and the functional constraints on certain domains are expected to be stable. Though the secondary structure is highly conserved, the primary structure is a mosaic of evolutionarily conserved and variable regions. The use of more or less conserved regions allows the study of closely or distantly related organisms. The 16S rRNA is a long molecule, containing about 1500 nucleotides. This provides a combination of a large number of characters and allows statistical evaluation of the data. There is no evidence of lateral gene transfer for this character, although in theory, gene transfers involving rRNA genes could happen. In four cyanobacteria, from two to six copies of the rRNA cistrons have been found (Nichols et al., 1982). Golden et al. (1989) transformed Synechococcus sp. strain PCC 7942 with an inactivated rRNA operon from strain PCC 6301. The modified rRNA operon could replace one of its two endogenous counterparts without apparent deleterious effects. Stackebrandt et al. (1991) transferred a complete ribosomal operon of Proteus vulgaris into the chromosome of Escherichia coli, and observed that it constituted 5% of the rRNA. To check for the possibility of gene transfer, sequence data from one or two other genes should be compared with the results obtained by means of the 16S rRNA sequences. According to Woese (1987), the 16S rRNA is a good ‘molecular chronometer’—that is, it is a good molecule for measuring the overall rate of evolutionary change in a line of descent.

Partial sequencing using reverse transcriptase (Lane et al., 1985) and the classical cloning method (Tomioka and Sugiura, 1983) will probably be replaced by gene amplification with the polymerase chain reaction (PCR), followed by direct sequencing (Urbach et al., 1992) or sequencing after cloning of the PCR products (Wilmotte et al., 1993). Primers complementary to conserved regions (Giovannoni et al., 1988; Wilmotte et al., 1993) are used for sequencing using the dideoxynucleotide method. In addition to axenic cultures, cyanobacterial strains contaminated by other bacteria can also be used, after design of PCR primers allowing a selective amplification of the cyanobacterial sequence.

4. Sequence Alignment and Data Analysis The purpose of an alignment is to place those nucleotides which derive from the same ancestor at the same position–one under the other. The conserved regions are easily aligned and the variable parts are placed between them to maximize similarity. When there have been deletions or insertions, alignment can be difficult and knowledge of the secondary structure can give clues. Secondary structure models are constructed on the basis of comparative analysis and chemical and enzymatic techniques (Noller et al., 1987; De Rijk et al., 1992). As an example, the secondary structure model of Chlorogloeopsis sp. HTF strain PCC 7518 (Wilmotte et al., 1993) is given in Fig. 2. A constantly updated alignment containing all the eubacterial 16S rRNA sequences submitted to EMBL, with the secondary structure features, is available (De Rijk et al., 1992). Numerical data analysis is a very complex and controversial issue and a good review of the different methodologies was written by Swofford and Olsen

Chapter 1 Molecular Approach to Cyanobacterial Evolution and Taxonomy

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(1990). Distance methods and parsimony analysis are most often used to infer phylogenetic trees. A summary of their most important characteristics is given below. Distance methods belong to the phenetic techniques and are based on the calculation of dissimilarities for each pair of organisms. These dissimilarity values are corrected for multiple mutations and entered in a matrix. A tree topology, in which the distances between organisms in the tree will be as close as possible to the matrix values, is constructed by an algorithm. Several algorithms are available (Swofford and Olsen, 1990). The neighbor-joining method of Saitou and Nei (1987) has been shown to be very efficient by computer simulation (Saitou and Imanishi, 1989). Parsimony analysis is based on the cladistic approach. It considers that only the similarity due to the possession of new (derived) characters is phylogenetically informative. Thus, only the variable positions, where at least two organisms share the same derived nucleotide, are used in the calculation of the tree topology. Ancestral sequences are reconstructed at each node of the tree. The final tree topology is the one requiring the minimal number of evolutionary changes. A statistical evaluation of tree topologies can be carried out by bootstrap analysis (Felsenstein, 1985). In short, new sequences are generated by random sampling of the positions with replacement. For each resampled data set, a new tree is constructed. The bootstrap value for each node is the number of resampled trees in which the same organisms are clustered together. Only the grouping of strains is considered, not the topology and lengths of the branches that diverge after the node. Felsenstein (1985) considered that a bootstrap percentage of 95% was the limit to recognize a statistically supported grouping. It is important to note that molecular evolution is a new field of science, which is still developing (Swofford and Olsen, 1990). Therefore, the inferred tree topologies must be considered hypotheses.

5. Results of 16S rRNA Sequence Analysis of Cyanobacteria The first demonstration of the usefulness of 16S rRNA sequences was obtained by the oligonucleotide catalogue. However, with this method, a maximum of 45% of the information content of the molecule

Annick Wilmotte was used. The heavily biased composition of strains, including only unicellular and heterocystous cyanobacteria, is probably responsible for the conclusion that heterocystous cyanobacteria seemed to have arisen from within the high G+C Synechocystis sp. cluster (Doolittle, 1982). The first global evolutionary scheme of cyanobacteria comprising 29 partial 16S rRNA sequences (about 700 positions) was published by Giovannoni et al. in 1988. The distance tree obtained allowed the following conclusions: (i) the simple unicellular and filamentous cyanobacteria (sections I and III) were scattered in different lineages and were sometimes mixed together; (ii) sections II, IV and V seemed to correspond to coherent phylogenetic clusters. The internodal distances between the branches diverging at the base of the tree were very short. Giovannoni et al. (1988) suggested that the rise of the oxygen concentration in the Precambrian atmosphere allowed the colonization of new biotopes and probably led to extensive divergence of the cyanobacteria. Most of the branches were long and unbranched, reflecting the selection of strains generally representing different genera and not expected to be close relatives. 16S rRNA sequence analysis has also demonstrated that the ‘prochlorophytes’, Prochlorothrix hollandica (Turner et al., 1989), Prochloron sp. and Prochlorococcus marinus (Urbach et al., 1992), are not genotypically close relatives but belong to two or three lineages. Recently, an average 16S rRNA similarity of about 98% was observed among cultivated isolates of Prochlorococcus marinus collected from the Sargasso Sea, north Atlantic, equatorial Pacific and Mediterranean Sea (E. Urbach, personal communication). The tree topologies of Figs. 3 and 4 contain all available complete or nearly complete cyanobacterial 16S rRNA sequences (1993). The trees were constructed with a distance method, the neighborjoining method, followed by a bootstrap analysis (Fig. 3) or with a parsimony method (Fig. 4). The two trees are largely congruent. Groupings supported by the bootstrap analysis at a level higher than 50% in Fig. 3 are also observed in Fig. 4, except the grouping of Lyngbya sp. strain PCC 7419 and Arthrospira sp. strain PCC 8005 and the grouping of Chlorogloeopsis sp. HTF with the cluster of Fischerella sp. strain PCC 7414 and Chlorogloeopsis fritschii strain PCC 6718. It is not surprising that the distance and parsimony methods give different branch topologies for the basic nodes that are not supported by the

Chapter 1 Molecular Approach to Cyanobacterial Evolution and Taxonomy

15

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bootstrap analysis. The consistency index of the parsimony tree (Fig. 4) is very low, 0.28, and indicates a high number of multiple mutations that can obscure the evolutionary patterns. The consistency index ranges from 0 to 1, and the maximal value is obtained

Annick Wilmotte

when all the characters have changed only once in the tree. Observations concerning the branches, presented below, gives interesting information on the genotypic relationships among the cyanobacteria and their

Chapter 1 Molecular Approach to Cyanobacterial Evolution and Taxonomy congruence with phenotypic characters. 16S rRNA sequence analysis is very useful to determine whether phenotypic similarities are due to recent divergence from a common ancestor or to convergent evolution.

a. Branches A and B The trees of Figs. 3 and 4 include organisms of unknown morphology, from which the 16S rRNA sequences were retrieved directly from the field. The Sargasso Sea isolates, SAR6, SAR7, SAR 100, and SAR139 (Giovannoni et al., 1990; Britschgi and Giovannoni, 1991), are closely related to Prochlorococcus marinus and the marine picoplanktonic Synechococcus sp. strain WH 8103, which were isolated from the same sea (Waterbury and Rippka, 1989; Urbach et al., 1992; J. Waterbury, personal communication to R. Rippka). Other sequences were isolated from the Pacific Ocean by Schmidt et al. (1991), but they were too short to be used for tree construction in Figs. 3 and 4. They were highly similar to the sequences from SAR6, SAR7, and Synechococcus sp. strains WH 7805 and WH 8103, suggesting a wide distribution ofgenotypically close strains in the Pacific and Atlantic Oceans. In contrast, the sequence types of possible cyanobacterial affiliation retrieved from a hot spring cyanobacterial mat in Octopus Spring in Yellowstone National Park (Weller et al., 1991, 1992; Ward et al., 1992), have no close relatives among the cultivated strains. The sequences OS-VI-L-8 and OS-V-L-13 belong to the same lineage in both trees, with a bootstrap support of 90% in Fig. 3. Representatives of the sequence type A, which is closely related to OS-VI-L-8, were too short to be included in Figs. 3 and 4. The sequence OS-V-L-16 is loosely associated with two filamentous strains in branch E. The OSVI-L-4 sequence is problematic. Weller et al. (1991) remarked that its position in their distance trees was unstable, diverging between the cyanobacteria and the proteobacteria, but shifting into the latter phylum when the strains or the sequence regions used to build the trees were changed. For the 400 first nucleotides, the average 16S rRNA sequence similarities of this strain with sixteen cyanobacteria and Escherichia coli are 85% and 78.4%, respectively. It falls within the range of average cyanobacterial similarities of 83.8–89.9%. For the last 400 nucleotides, strain OS-VI-L-4 shows 68.9% and 70.3% respectively sequence similarity with the same cyanobacterial sequences and E. coli. It is clearly

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outside the average similarities within the cyanobacteria, which remain 83.7–88.7% (unpublished calculations). This could suggest that this sequence is a chimera but, in theory, this should not have been possible with the method used (cDNA cloning). Caution is needed in interpreting its position (Weller et al., 1991). The determination of 16S rRNA sequences in the framework of molecular ecology studies, reviewed by Ward et al. (1992), is a welcome source of new data. They allow the exploration of regions of the cyanobacterial diversity to which cultivated strains do not give access. If the original organisms can be detected in microscopic slides by the use of labeled probes annealing to specific 16S rRNA regions (DeLong et al., 1989; Ward et al., 1992), their morphological characterization can be very useful in taxonomy. Results of Ward et al. (1992) suggest that unicellular cyanobacteria that are morphologically similar may be phylogenetically very different. This raises the question whether it will always be possible to find phenotypic markers which are congruent with the genotypic affiliations, particularly in the case of the simple morphological types.

b. Branch C The strain Mef 6705 (Fig. 5a), originally identified as Oscillatoria redekei Van Goor, and renamed Limnothrix redekei Meffert (Meffert, 1989) shows about 97% sequence similarity with the 16S rRNA sequences of Pseudanabaena galeata (Fig. 5b) and Pseudanabaena sp. strain PCC 6903 (Giovannoni et al., 1988; Nelissen et al., 1992). The three strains share similar morphological features: cell diameter between 1 and gas vesicles at the cross-walls, thylakoids parallel to the longitudinal walls and intercellular trichome breakage (Guglielmi and Cohen-Bazire, 1984a; Meffert, 1989). Guglielmi and Cohen-Bazire (1984b) have excluded the strain PCC 7403 (Fig. 5c) from the genus Pseudanabaena on the basis of differences in pigment composition and low DNA-DNA hybridization percentages. In Figs. 3 and 4, strain PCC 7403 diverges at the base of the branch leading to the other Pseudanabaena sp. strains (Nelissen et al., 1992), and it shares about 92% similarity with the 16S rRNA sequences of these strains. This suggested that these four strains probably belong to the same genus, and the definition of Pseudanabaena sensu Guglielmi and Cohen-Bazire should be widened to include strain PCC 7403.

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The strain Gloeobacter violaceus strain PCC 7421 (Giovannoni et al., 1988) is loosely associated with this group. This unicellular organism, assigned to section I, has several unique features: it lacks thylakoids (Rippka et al., 1974) and has phycobilisomes of unusual structure which are attached to the inner surface of the cytoplasmic membrane and which are arranged as a cortical layer (Guglielmi et

Annick Wilmotte al, 1981). It would be interesting to determine the sequence of the other known Gloeobacter sp. strain PCC 8105.

c. Branch D A lineage, statistically supported at a level of 100%, contains marine, phycoerythrin-containing Phor-

Chapter 1 Molecular Approach to Cyanobacterial Evolution and Taxonomy midium species with a trichome diameter smaller than (Wilmotte, 1991). Three morphologically similar strains, isolated in Europe, USA, and Australia, have almost identical 16S rRNA sequences and were assigned to the same species, Phormidium ectocarpi Gomont. The strain Phormidium minutum Lindstedt (D5) has deeper constrictions at the cross
walls, and this is correlated with a sequence divergence of about 6% from the P. ectocarpi strains (Wilmotte et al., 1992). The positions of Prochlorothrix hollandica and two unicellular strains, Synechococcus sp. strain PCC 6301 and the thermophilic Synechococcus lividus strain CCCY7C
S, appear unstable (Tomioka and Sugiura, 1983;Giovannoni et al., 1988; Turner et al., 1989).

d. Branch E The grouping of Plectonema boryanum strain PCC 73110 and Oscillatoria amphigranulata strain CCC
NZ-concert-Oa (Giovannoni et al., 1988) is observed in 65.4% of the bootstrap trees (Fig. 3), and they are sister
groups in the parsimony tree (Fig. 4). These two strains share a similar trichome structure with slight constrictions at the cross
walls and a cell width of In Plectonema boryanum, a thin sheath is sometimes present (Stam and Holleman, 1975). This strain does not contain phycoerythrin nor phycoerythrocyanin (Bryant, 1982) and is able to fix nitrogen anaerobically (Rippka and Herdman, 1992). On the other hand, Oscillatoria amphigranulata strain CCC
NZ
concert
Oa (Fig. 5d) contains phycoerythrin and can perform chromatic adaptation. This thermophilic cyanobacterium, able to perform anoxygenic photosynthesis with sulfide as an electron donor (Garcia
Pichel and Castenholz, 1990), should probably be renamed (R. W. Castenholz, personal communication).

e. Branch F This complex branch contains exclusively simple filamentous strains, the cyanelle of Cyanophora paradoxa, and the chloroplast of the liverwort, Marchantia polymorpha. The strains Oscillatoria sp. strain PCC 7105 and Microcoleus sp. strain 10mfx have 16S rRNAs that are 98.8% similar in sequence (Giovannoni et al., 1988; Wilmotte et al., 1992) and belong to the same lineage in 100% of the bootstrap trees. However,

19

they probably have quite different morphologies. No information could be obtained from the scientist who cultivated Microcoleus sp. strain 10mfx, in spite of repeated requests. Following a short description by L. Richardson (NASA Ames), that was kindly transmitted by S. Giovannoni, this strain has a cell width of and a thin sheath. On the other hand, Oscillatoria sp. strain PCC 7105 has cells that are isodiametric to cylindrical (with a width 1.4 to 2.8 No sheath and no constrictions at the cross
walls are visible (Wilmotte, 1991). The cyanelle of Cyanophora paradoxa (V. L. Stirewalt and D. A. Bryant, personal communication) and the chloroplast of the liverwort (Ohyama et al., 1986) are clustered in 86% of the bootstrap trees. It is noteworthy that the DNA base compositions of their 16S rRNA genes, 53 and 55 GC% respectively, are similar to the values of their cyanobacterial counterparts. Thus, it is improbable that G+C biases can distort their relationships (see Lockhart et al., 1992). The position of cyanelles and plastids among the cyanobacteria is variable, depending on the organisms and sequence positions used (Douglas and Turner, 1991). There is a loose grouping of three filamentous strains, Oscillatoria sp. strain PCC 7515, Microcoleus sp. strain PCC 7420 and Oscillatoria limnetica (Solar Lake strain; Giovannoni et al., 1988; Nelissen et al., 1992) that have little morphological similarity (Wilmotte and Golubić, 1991) and should be further investigated. Arthrospira sp. strain PCC 8005 (Nelissen et al., 1992) is clustered with Lyngbya sp. strain PCC 7419 (Fig. 25, Rippka et al., 1979) and not with Spirulina sp. strain PCC 6313 (Giovannoni et al., 1988), although it has helical trichomes (Fig. 5e) like the latter strain (see Fig. 17, Rippka et al., 1979). The genera Arthrospira and Spirulina were fused in the same genus, Spirulina, by Geitler (1932) and Rippka et al. (1979). Later, Rippka and Herdman (1992) recognized the separation of the two genera, already proposed by Anagnostidis and Komárek (1988) and Castenholz (1989a). Ultrastructural studies (Gug
lielmi and Cohen
Bazire, 1982) supported this separation and indicated the presence of one circle of pores in the peptidoglycan near the cross
walls in Arthrospira sp. strain PCC 7345 and several Oscillatoria sp. strains sensu Rippka et al. (1979). In addition, Arthrospira sp. strain PCC 8005 often forms straight trichomes which look like Oscillatoria sp.(Fig. 5e). Thus, the grouping of Arthrospira sp.

20 strain PCC 8005 with Oscillatoriaceae strains (Figs. 3, 4; Nelissen et al., 1992) seems globally congruent with the phenotypic information. The strain Oscillatoria sp. strain PCC 6304 (Giovannoni et al., 1988) occupies an isolated position in the tree of Fig. 3 and is the only strain with a curved tip (Fig. 5f). Morphologically similar strains should be investigated (Wilmotte and Golubić, 1991). f. Branch G This heterogeneous branch contains unicellular and filamentous strains. The existence of a lineage containing the three baeocyteforming strains (section II) is not well supported. In only 35% of the bootstrap trees, Dermocarpa sp. PCC 7437 is clustered with Pleurocapsa sp. strain PCC 7321 and Myxosarcina sp. strain PCC 7312 (Giovannoni et al., 1988). Other genera of section II should be investigated. The positions of the other strains in branch G, simple unicellular and filamentous cyanobacteria (Giovan noni et al., 1988; Nelissen et al., 1992; Urbach et al., 1992), are unstable and their relationships are still uncertain. New sequences from closely related strains should provide useful information. The strain Chamaesiphon sp. strain PCC 7430 is unicellular and has the peculiar ability to reproduce by asymmetric fission or budding. This uniqueness is reflected in the isolated position of its 16S rRNA sequence (Giovannoni et al., 1988). To ascertain whether this type of division is a good phylogenetic marker, the 16S rRNA sequences of other Chamaesiphon sp. strains should be determined.

g. Branch H The heterocystous strains (Giovannoni et al., 1988; Ligon et al., 1991) are all situated in the same lineage, which is quite well supported by the bootstrap analysis (92.6%). However, within this group, the mutual interrelationships are less clear. The exception is the cluster of Anabaena sp. strain PCC 7122 and Nodularia sp. strain PCC 73104, supported at a level of 95.2%. The genotypic unity of the three strains from section V, or the order Stigonematales, is supported in only 53.6% of the bootstrap trees (Fig. 4). The strain Chlorogloeopsis sp. HTF (‘Mastigocladus sp. HTF’) strain PCC 7518 forms cell aggregates and short filaments (Castenholz, 1989c). Heterocysts are not observed in strain PCC 7518, though they were

Annick Wilmotte present in the original clone (Castenholz, 1969). However, nif genes have been observed in strain PCC 7518 (Rippka and Herdman, 1992). The 16S rRNA sequence analysis demonstrates that this strain belongs to the heterocystous cluster and has probably lost the capacity to differentiate heterocysts by mutation. In addition, it appears equally distant from Fischerella sp. strain PCC 7414 and from Chloro
gloeopsis fritschii strain PCC 6718. This result and DNADNA hybridization data (Rippka and Herdman, 1992) suggest that the three strains belong to different genera (Wilmotte et al., 1993).

6. Comparison with the Current Evolutionary and Taxonomic Hypotheses The only evolutionary hypothesis that may be supported by the trees of Figs. 3 and 4, is the rather late differentiation of the heterocystous species. The suggestion of Giovannoni et al. (1988), that the strains which do not form hormogonia, Anabaena sp. strain PCC 7122 and Nodularia sp. strain PCC 73104, were the last to appear is not confirmed in the tree of Fig. 3. Giovannoni et al. (1988) also suggested that the simple unicellular and filamentous types (sections I and III) had multiple evolutionary origins. This is in agreement with the tree topologies of Figs. 3 and 4, but none of the branches (C, D, and G) in which simple unicellular and filamentous strains are mixed, is convincingly supported by the bootstrap analysis (Fig. 3). However, Waterbury (personal commun ication to R. Rippka) observed a rather close relationship between the filamentous Phormidium sp. strain PCC 7375 and the unicellular Synechococcus sp. strain PCC 7335. The 16S rRNA sequences of Phormidium fragile strain PCC 7376 and Synecho
coccus sp. strain PCC 7002 exhibit very striking similarity (97%; J. B. Waterbury, personal com munication). Synechococcus sp. strain PCC 7002 is the reference strain of the Marine cluster C, to which Synechococcus sp. strain PCC 7335 has also been assigned (Waterbury and Rippka, 1989).It is noteworthy that both of these strains are marine and have quite similar genomic base compositions and cell diameters. Unfortunately, no other gene sequences have been determined for these pairs of strains in order to confirm their very close genotypic relationships. In agreement with the two preceding observations, it seems that the delimitation between unicellular and filamentous strains is not always sharp. For example, Synechococcus sp. strain PCC

Chapter 1 Molecular Approach to Cyanobacterial Evolution and Taxonomy 6301 forms also short chains of cells (Komárek, 1970). Moreover, filamentous mutants of unicellular strains are known (Ingram and Van Baalen, 1970) and a Pseudanabaena strain with short cells can become unicellular (Guglielmi and Cohen
Bazire, 1984a). Only a small genetic change may be sufficient to provoke or hinder the separation of the daughter cells after cell division (Wilmotte and Golubić, 1991). Conclusions on the congruences and differences of the 16S rRNA sequence data with different taxonomic systems are premature. Certain genotypic relationships have been recognized and other ones have been ignored, but too few taxonomic problems have been investigated to evaluate these classifications by comparison with what is presently known on the genotypic relationships of the cyanobacteria.

7. Possible Developments of the Use of the rRNA Cistrons to Study the Evolution and Taxonomy of the Cyanobacteria 16S rRNA gene sequences are probably too conserved to investigate intraspecies variability (Ward et al., 1992). The sequence of the internal transcribed spacer (ITS), situated between the 16S rRNA and the 23S rRNA genes, may be an adequate marker, however. When the reverse primer used for PCR is comple
mentary to the 5' end of the 23S, the ITS is also amplified (Wilmotte et al., 1993). In Synechococcus PCC 6301, the ITS is 545 nucleotides long and includes and (Tomioka and Sugiura, 1984). Species
specific probes inferred from the ITS sequence have been used for eubacterial identification (e.g. Rossau et al., 1992). The ITS sequence similarity for Pseudanabaena sp. strain PCC 7409 and Limnothrix redekei strain Mef 6705 is 88%, whereas their 16S rRNA genes share 99.7% sequence similarity. The ITS sequence of Pseudanabaena sp. strain PCC 7403 is longer and hence different from the two preceding ITS sequences, so that no meaningful alignment is possible (A. Wilmotte, unpublished data), except for the region ‘AAGAACCTTGAAAACTGCATAG’ corres
ponding to positions 438–459 of the ITS of Synechococcus sp. strain PCC 6301 (Tomioka and Sugiura, 1984). The high degree of conservation of this ITS region in ten cyanobacterial strains (A. Wilmotte, unpublished data) suggests that it has a function. In addition, the length variability of the ITS sequence may allow the detection of the presence of more than one cyanobacteria in a culture.

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The eubacterial 23 S rRNA is more variable than the 16S rRNA, and may be better suited to resolve close relationships (Ward et al., 1992). The only cyanobacterial 23S rRNA sequence available is that from Synechococcus sp. strain PCC 6301, and it is tentatively estimated to have 2869 positions (Kumano et al., 1983). Finally, for a rapid taxonomic survey, amplified portions of the rRNA cistrons may be digested by restriction enzymes. This kind of RFLP analysis has been used with success for eubacterial taxonomy (e.g., see Vaneechoutte et al., 1992).

VI. Conclusion This chapter has underlined the problems addressed by the evolutionary and taxonomic studies of cyanobacteria, and presented the promising tools offered by molecular techniques. However, the collection and analysis of molecular information are only now beginning and conclusions are still limited by the relatively small number of data available. It is hoped that more laboratories will become interested in ribosomal RNA sequences and other molecular techniques to determine cyanobacterial genotypic relationships. However, this molecular information should be integrated with other characteristics of the strains. This will form the basis for a polyphasic taxonomy that will not only be of practical use but will reflect as much as possible the evolutionary relationships of the strains.

Acknowledgments I would like to thank my colleagues for interesting discussions, B. Nelissen, G. Van der Auwera, R. De Baere and F. Haes for sequences, J
M. Neefs and P. De Rijk for sequence alignment, Y. Van de Peer for advice with the data analysis and Prof. R. De Wachter for financial support, useful discussions and interest. Many thanks are also due to V. L. Stirewalt and D. A. Bryant (Pennsylvania State University), R. Casten
holz (University of Oregon), S. Giovannoni (Oregon State University), M
E. Meffert (Limnological Institute, Plön), R. Rippka and M. Herdman (Pasteur Institute, Paris), E. Urbach (M.I.T.) and D. Ward (Montana State University) for giving strains, valuable information or unpublished results. Don Bryant improved this manuscript with many suggestions. I

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am grateful to S. Turner and N. Pace (Indiana University) for introducing me to the 16S rRNA analysis, as well as R. Rossau (Innogenetics) for teaching me PCR methods and for sharing his protocol. The National Fund for Scientific Research of Belgium (FNRS) is also thanked for financial support during six years. Finally, I would like to mention V. Demoulin (University of Liège) and W. Stam (University of Groningen) who awakened my interest in molecular evolution and taxonomy of the cyanobacteria. This interest was further stimulated by the numerous cyanophycologists, mostly IAC members, and cyanobacteriologists who shared with me their information, advice and enthusiasm. References Aakermann T, Skulberg OM and Liaaen-Jensen S (1992) Further studies on the carotenoids of blue-green algae (cyanobacteria) – a comparative investigation of strains from the genera Oscillatoria and Spirulina. Biochem System Ecol 20: 761– 769 Aitken A (1988) Protein sequences as taxonomic probes of cyanobacteria. Meth Enzymol 167: 145–154 Anagnostidis K and Komárek J (1985) Modern approach to the classification system of cyanophytes. 1–Introduction. Arch Hydrobiol Suppl 71, Algological Studies 38/39: 291–302 Anagnostidis K and Komárek J (1988) Modern approach to the classification system of cyanophytes. 3–Oscillatoriales. Arch Hydrobiol Suppl 80, Algological Studies 50/53: 327–472 Anagnostidis K and Komárek J (1990) Modern approach to the classification system of cyanophytes. 5–Stigonematales. Arch Hydrobiol Suppl 86, Algological Studies 59: 1–73 Bornet E and Flahaut C (1886–1888) Révision des Nostocacées hétérocystées. Ann Sci Nat, Bot 7, Sér 3: 323–381; 4: 343– 373; 5: 52–129; 7: 178–262 Britschgi TB and Giovannoni SJ (1991) Phylogenetic analysis of a natural marine bacterioplankton population by rRNA gene cloning and sequencing. Appl Environm Microbiol 57:1707– 1713 Brosius J, Dull TJ, Sleeter DD and Noller HF (1981) Gene organization and primary structure of a ribosomal RNA operon from Escherichia coli. J Mol Biol 148: 107–127 Bryant DA (1982) Phycoerythrocyanin and phycoerythrin: properties and occurrence in cyanobacteria. J Gen Microbiol 128: 835–844 Bullerjahn GS, Jensen TC, Sherman DM and Sherman LA (1990) Immunological characterization of the Prochlorothrix hollandica and Prochloron sp. chlorophyll a/b antenna proteins. FEMS Microbiol Lett 67: 99–106 Castenholz RW (1969) The thermophilic cyanophytes of Iceland and the upper temperature limit. J Phycol 5: 360–368 Castenholz RW (1989a) Subsection III, order Oscillatoriales. In: Staley JT, Bryant MP, Pfennig N and Holt JG (eds) Bergey’s Manual of Systematic Bacteriology, Vol 3, pp 1771–1780. Williams and Wilkins Co, Baltimore Castenholz RW (1989b) Subsection IV, order Nostocales. In:

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Annick Wilmotte and a colorimetric reverse hybridization assay for the detection of Bordetella pertussis. Molec Cell Probes 6: 281–289 Saitou N and Imanishi T (1989) Relative efficiencies of the FitchMargoliash, maximum-parsimony, maximum-likelihood, minimum-evolution, and neighbor-joining methods of phylogenetic tree construction in obtaining the correct tree. Mol Biol Evol 6:514–525 Saitou N and Nei M (1987) The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol 4: 406–425 Sallal AK, Nimer N A and Radwan SS (1990) Lipid and fatty acid composition of freshwater cyanobacteria. J Gen Microbiol 136: 2043–2048 Schenk HEA and Kuhfittig G (1983) Elektromorphe Phycochromoproteid-Muster: Eine Taxonomische Identifikationsmethode für Cyanobakterien-Species. Biochem System Ecol 11: 163–174 Schenk HEA, Hofer I and Metzner H (1973) Malat-Dehydrogenase Isoenzymbanden als potentielles chemotaxonomisches Kriterium für Cyanophyceen-Species. Biochem System 1: 179–184 Schleifer KH and Ludwig W (1989) Phylogenetic relationships among bacteria. In: Fernholm B, Bremer K and Jörnvall (eds) The Hierarchy of Life, pp 103–117. Elsevier Science Publishers, Amsterdam Schleifer KH and Stackebrandt E (1983) Molecular systematics of prokaryotes. An Rev Microbiol 37: 143–187 Schmidt TM, DeLong EF and Pace NR (1991) Analysis of a marine picoplankton community by 16S rRNA gene cloning and sequencing. J Bacteriol 173: 4371–4378 Schopf JW and Packer BM (1987) Early archean (3.3–billion to 3.5-billion-years-old) microfossils from Warsanoona group, Australia. Science 237: 70–73 Schopf JW and Walter MR (1982) Origin and early evolution of cyanobacteria: the geological evidence. In: Carr NG and Whitton BA (eds) The Biology of Cyanobacteria, pp 543–564. Blackwell Scientific, Oxford Schwabe GH (1960) Über den thermobionten Kosmopoliten Mastigocladus laminosus Cohn. Schweiz. Z Hydrol 22: 757– 792 Stackebrandt E and Goodfellow M (1991) Introduction. In: Stackebrandt E and Goodfellow M (eds) Nucleic Acid Techniques in Bacterial Systematics, pp xix–xxix. Wiley, Chichester Stackebrandt E, Niebel H and Söller R (1991) Expression of the chromosome-coded rRNA genes of Proteus vulgaris in Escherichia coli. J Gen Appl Microbiol 37: 141–146 Stam WT (1980) Relationships between a number of filamentous blue-green algal strains (Cyanophyceae) revealed by DNADNA hybridization. Arch Hydrobiol, Suppl 56, Algological Studies 25: 351–374 Stam WT and Holleman HC (1975) The influence of different salinities on growth and morphological variability of a number of Phormidium strains (Cyanophyceae) in culture. Acta Neerl Bot 24: 379–390 Stam WT and Venema G (1975) The use of DNA-DNA hybridization for determination of the relationship between some blue-green algae (cyanophyceae). Acta Bot Neerl 26: 327–342 Stam WT, Boele-Bos SA and Stulp BK (1985) Genotypic relationships between Prochloron samples from different

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Wayne LG, Brenner DJ, Colwell RR, Grimont PAD, Kandler O, Krichevsky MI, Moore LH, Moore WEC, Murray RGE, Stackebrandt E, Starr MP and Trüper HG (1987) Report of the ad hoc committee on reconciliation of approaches to bacterial systematics. Int J Syst Bacteriol 37: 463–464 Weller R, Walsh Weller J and Ward DM (1991). 16S rRNA sequences of uncultivated hot springs cyanobacterial mat inhabitants retrieved as randomly primed cDNA. Appl Environm Microbiol 57: 1146–1151 Weller R, Bateson, MM, Heimbuch BK, Kopczynski ED and Ward DM (1992) Uncultivated cyanobacteria, Chloroflexuslike inhabitants, and spirochete-like inhabitants of a hot spring microbial mat. Appl Environm Microbiol 58: 3964–3969 Whitton BA (1992) Diversity, ecology, and taxonomy of the cyanobacteria. In: Mann NH and Carr NG (eds) Photosynthetic Prokaryotes, pp 1–52. Plenum Press, New York Wilmotte A (1991) Taxonomic study of marine oscillatoriacean strains (cyanophyceae, cyanobacteria) with narrow trichomes. I. Morphological variability and autecological features. Arch Hydrobiol, Suppl 92, Algological Studies 64: 215–248 Wilmotte A and S (1991) Morphological and genetic criteria in the taxonomy of Cyanophyta/Cyanobacteria. Arch Hydrobiol, Suppl 92, Algological Studies 64: 1–24 Wilmotte A and Stam WT (1984) Genetic relationships among cyanobacterial strains originally designated as ‘Anacystis nidulans’ and some other Synechococcus strains. J Gen Microbiol 130: 2737–2740 Wilmotte A, Turner S, Van de Peer Y and Pace NR (1992) Taxonomic study of marine oscillatoriacean strains (cyanobacteria) with narrow trichomes. II. Nucleotide sequence analysis of the 16S ribosomal RNA. J Phycol 28: 828–838 Wilmotte A, Van Der Auwera G and De Wachter R (1993) Structure of the 16S ribosomal RNA of the thermophilic cyanobacterium Chlorogloeopsis HTF (‘Mastigocladus laminosus HTF’) strain PCC 7518, and phylogenetic analysis. FEBS Lett 317: 96–100 Woese CR (1987) Bacterial evolution. Microbiol Rev 51:221–271 Wood AM and Townsend D (1990) DNA polymorphism within the WH7803 serogroup of marine Synechococcus spp. (cyanobacteria). J Phycol 26: 576–585 Zehr JP, Limberger RJ, Okhi K and Fujita Y (1990) Antiserum to nitrogenase generated from an amplified DNA fragment from natural populations of Trichodesmium sp. Appl Environ Microbiol 56: 3527–3531 Zilinskas BA and Howell DA (1987) Immunological conservation of phycobilisome rod linker polypeptides. Plant Physiol 85: 322–326 Zimmerman WJ and Bergman B (1990) The Gunnera symbiosis: DNA restriction fragment length polymorphism and protein comparisons of Nostoc symbionts. Microb Ecol 19: 291–302 Zimmerman WJ and Rosen BH (1992) Cyanobiont diversity within and among cycads of one field site. Can J Microbiol 38: 1324–1328 Zuckerkandl E and Pauling L (1965) Molecules as documents of evolutionary history. J Theor Biol 8: 357–366

Chapter 2 The Oceanic Cyanobacterial Picoplankton Noel G. Carr and Nicholas H. Mann Department of Biological Sciences, University of Warwick, Coventry CV4 7AL, U.K. Summary I. Introduction A. The Oceanic Cyanobacteria B. Synechococcus (MC-A) C. Other Major Oceanic Cyanobacteria II. Macromolecular Composition A. DNA B. RNA and Protein Synthesis III. Phycobiliproteins IV. Plasmids V. Phages VI. Transcription VII. Translation A. Codon Usage VIII. Nutrition A. Nitrogen 1. Assimilation 2. Nitrogen Fixation B. Phosphorus C. Iron D. Carbon Assimilation IX. Adaptive Responses A. Highly Iterated Palindromic Sequences (HIP1) B. Protein Phosphorylation Acknowledgments References

27 28 28 28 30 30 30 31 31 33 33 35 36 36 37 37 37 39 40 42 43 43 43 44 44 45

Summary The initial interest in the phycoerythrin-containing picoplanktonic cyanobacteria, assigned to the genus Synechococcus, stemmed directly from a recognition of their considerable contribution to marine primary productivity, as well as to their widespread distribution in an environment hitherto characterized by its relative paucity of cyanobacteria. However, recent work has increasingly indicated that these organisms have features of their molecular biology which separate them from the well-characterized freshwater and halotolerant members of the genus and molecular phylogeny may indicate this separation to be deep. Much of the work described relates to molecular biological analyses of the mechanisms by which these organisms harvest light and acquire key nutrients in an environment which is highly variable with regard to the former and acutely oligotrophic with regard to the latter. Where appropriate, comparisons have been made to what is known ofthe molecular biology of nutrient acquisition by other ecologically significant oceanic cyanobacteria.

D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 27–48. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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I. Introduction

A. The Oceanic Cyanobacteria The limited diversity of cyanobacteria found in the open oceans is worth some consideration. It is in marked contrast to the wide range found among the freshwater plankton and represents only a tiny proportion of the ‘largest and most diverse group of prokaryotes’ to use the now well known phrase of Stanier and Cohen-Bazire (1977). The possibility that a saline environment may be the causative factor forthis restricted range can be excluded by considering inter-tidal and hyper-saline areas which have a rich and diverse cyanobacterial flora (Whitton and Potts, 1982). Relatively eutrophic ocean waters such as the Baltic Sea also have a reasonable range of cyanobacteria including strains of the genera Nodularia, Aphanizomenon and Anabaena (Geitler, 1932). Fogg (1982) has suggested that an explanation for the limited cyanobacterial component of open ocean water may lie in the turbulence of the sea which would interfere with the buoyancy regulation produced by gas vacuoles (Walsby, 1994) and other features and thereby prevent the development of blooms which are a characteristic feature of many successful freshwater species. An additional factor may lie in the nature of the ocean environment itself which, relative to freshwater, inter-tidal or terrestrial areas, is homogenous in its nutrient inputs and perhaps more importantly is acutely oligotrophic. Cyanobacteria are opportunistic phototrophs with a variety of strategies that permit them to adapt well to changes in theirphysical and inorganic environment (Whitton, 1992). If however that environment is rather uniform and some, perhaps all, of the potentially limiting nutrients (N, P, Fe) are in essentially equal short supply there would be little selective advantage in the ability to evolve mechanisms for the intermittent acquisition and storage of a particular nutrient such as to increase the growth rate and hence exploit particular niches. The concept of a single limiting nutrient at any one period of time may not apply in the Abbreviations: HIP – highly iterated palindromic sequence; ORF – open reading frame; PCB – phycocyanobilin; PEB – phycoerythrobilin; PUB – phycourobilin; SDS-PAGE – polyacrylarnide gel electrophoresis in the presence of sodium dodecylsulfate; Synechococcus (MC-A) – phycoerythrincontaining Synechococcus sp. strains of Marine-Cluster A as defined by Waterbury and Rippka (1989).

Noel G. Carr and Nicholas H. Mann open oceans that are under discussion; what there may be is plural limitation for several key nutrients. When enrichment occurs, naturally or of anthropogenic origin, different criteria apply and a wider range of cyanobacteria would establish themselves. There are three groups of cyanobacteria which are known to be present in distinct ocean provinces in numbers sufficient to make a measurable ecological contribution: the phycoerythrin-containing Synechococcus sp. strains, which are the subject of this chapter, the non-heterocystous, nitrogen-fixing Trichodesmium sp. and the heterocyst-containing Richelia intracellularis. There are, of course, many other reports ofcyanobacteria ofvarious genera from the different oceans (see Sournia, 1970; Fogg, 1982; Fogg, 1987) but none of these indicate a consistent, major input into their ecosystem. It should be emphasized that we are concerned here with the open oceans and not coastal areas or regions such as the Baltic that are strongly influenced by the surrounding land. To this restricted list should perhaps be added certain of the prochlorophyte species, as their apparently polyphyletic origin indicates a plural development along with some oceanic cyanobacteria to which they are phylogenetically related (Palenik and Haselkorn, 1992; Urbachetal., 1992) rather than their being a distinct separate phylogenetic branch (see Chapter 3). B. Synechococcus (MC-A) In 1979 two reports described the presence of a hitherto unrecognized group of cyanobacteria in the open sea (Johnson and Sieburth, 1979; Waterbury et al., 1979). These organisms were small, non-motile, non-nitrogen fixing unicells and were therefore assigned to the group Synechococcus. The taxonomy of this group is problematic and consequently we refer throughout this chapter to these phycoerythrincontaining marine Synechococcus sp. strains as Synechococcus (MC-A) in the sense proposed by Waterbury and Rippka (1989). A characteristic feature of these organisms is their possession of the accessory pigment phycoerythrin and indeed it was the fluorescence ofthis phycobiliprotein that led to their discovery: an orange fluorescence resulting from excitation at 540 nm was quite distinct from the red fluorescence due to chlorophyll a. The widespread distribution ofthese organisms, and the fact that sizefractionated productivity measurements showed that

Chapter 2 Oceanic Cyanobacterial Picoplankton they made a significant contribution to primary productivity, ensured continuing interest and they were soon recognized as major components of the ‘picoplankton’, the name given to organisms that passed through a filter, and which had been largely unrecognized by biological oceanographers (see Platt and Li, 1986). The perceived importance of the picoplankton was furthered by the idea that in the oceans there existed a ‘microbial loop’ through which a significant proportion ofprimary productivity was generated by organisms too small for effective grazing and was recycled by bacterial heterotrophy (Azam et al., 1983). In addition to the phycoerythrin-containing Synechococcus (MC-A) the picoplankton are known to comprise many other organisms; these include a range of small photosynthetic eucaryotes (see Thomsen, 1986) as well as green unicells of an apparently procaryotic nature. Some, perhaps most, of the latter could now be accounted for as prochlorophytes (see Chapter 3). Following their discovery there was immediate interest in examining the Synechococcus (MC-A) for two reasons. Firstly, there was theirratherwidespread distributionandvarying,but significant contribution to primary productivity–reports of between 10 and 30% were usual (Waterbury et al., 1986). Since it is generally thought that the oceans are responsible for about half of global productivity, it was evident that these were organisms of importance. What is less clearnow, with the largenumbers ofprochlorophytes being seen in the oceans by analytical flow cytometry (Chisholm et al., 1988; 1992; Vaulot and Partensky, 1990) is what proportion of picoplankton productivity can be assigned to its cyanobacterial component as most measurements of production were done with simple, size-fractionated populations. Secondly, the discovery oftheSynechococcus (MC-A) dramatically increased the number and type of cyanobacteria known to have an important role in the open oceans as distinct from coastal and littoral areas in which numerous andvariedcyanobacterialpopulations have been described (Whitton and Potts, 1982). Two distinct sub-populations of Synechococcus (MC-A) strains may be distinguished on the basis of the predominant chromophore associated with phycoerythrin (see Section III); the phycourobilin-rich strains are characteristic of the open oceans whereas those with a lower phycourobilin content are associated with shelf waters (Olson et al., 1990). A detailed description of the oceanic Synechococcus (MC-A) is given by Waterbury et al. (1986),

29 who providecomparisonsbetweenlaboratory cultures and natural populations with precise accounts ofthe source of the Woods Hole collection of these organisms. The Synechococcus (MC-A) are found extensively in the photic zone of all oceans with the exception ofthe Polar seas. Their numbers vary from one million per ml downwards and are of course variable with the seasons. Water temperatures below 5 °C seem to be inimical to growth. They thrive in conditions oflow photon flux (such as but will tolerate light intensities such as are found in the surface layers which can be of the order (Kana and Glibert, 1987a). With the exception of their pigmentation and rather smaller size these organisms are similar in appearance to the well-studied freshwater and coastal strains of the genus Synechococcus. Recently, populations of phycoerythrin-containing picoplankton have been reported in oligotrophic freshwater lakes (Hawley and Whitton, 1991). Some isolates of Synechococcus (MC-A) are motile by means of a mechanism as yet unknown (Waterbury et al., 1985). All are obligate photoautotrophs but, like many other cyanobacteria, can assimilate organic carbon material. All strains examined can use nitrate and ammonia as nitrogen source and several will utilize urea (see Glover, 1985; Waterbury et al., 1986; and Chapter 16). Recently, the ability of at least some of the strains in culture to assimilate amino acids has proved useful (Chadd, 1992) and may have important ecological implications (Paerl, 1991). Eighteen axenic strains have been shown not to fix nitrogen even under strictly anaerobic conditions and it is concluded that this property is unlikely to be present in the group. Searching for nif genes with DNA probes in these organisms has also been unsuccessful (J.P. Zehr, personal communication). A phycoerythrin-containing unicell (strain WH 8501) capable of aerobic nitrogen fixation has been isolated from tropical waters and assigned to the provisional assemblage Synechocystis (Waterbury and Rippka, 1989). In view of the requirement for vitamin shown by several strains of marine cyanobacteria by Van Baalen (1962), the absence of any such requirement in the Woods Hole cultures of the Synechococcus (MC-A) is of interest. The lack of cyanophycin was noted in the strains examined by Waterbury et al., (1986), and cyanophycin was experimentally proven to be absent by Newman et al., (1987) for one isolate. The relationship ofthe Synechococcus (MC-A) to other members of the order Chroococcales has been

Noel G. Carr and Nicholas H. Mann

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confused over the years by the casual identification by many workers of small unicellular cyanobacteria as ‘Synechococcus’ species. The position has been clarified by the publication ofWaterbury and Rippka (1989) in which formal descriptions and origins of many strains are classified as far as possible on bacteriological principles. The organisms which are the subject of this chapter are members of the Synechococcus group and fall into the Marine-Cluster A of Waterbury and Rippka (1989). The separation of Synechococcus (MC-A) from other Synechococcus species is supported by the RFLP analysis described below, and when 16S rRNA sequence analysis has been completed on a sufficient number of species, this division may turn out to be deeply branched. In order to measure and evaluate the diversity of the Synechococcus (MC-A) in natural populations, specific antisera were developed against Synechococcus sp. WH 7803 (Campbell et al., 1983; Campbell and Iturriaga, 1988) which cross-reacted with other species of the group. However, there are at least two distinct pigment types (see below) and other data indicate that the Synechococcus sp. strain WH 7803 serotype is heterogeneous (Wood 1985; Glover et al., 1986). The separation of the oceanic species which contained phycoerythrin into two groups by comparison ofanalysis ofrestriction fragment length polymorphisms and the clear distinction of both these from the freshwater Synechococcus sp. strain PCC 6301 was shown by Douglas and Carr (1988). Using a larger number of strains and incorporating existing data, Wood and Townsend (1990) demonstrated that Dollo Parsimony and cluster analysis of RFLP information showed that the group had at least four genetic lineages with differences as great as those that separated them from freshwater species. Wood and Townsend (1990) were able to conclude that natural assemblages of Synechococcus (MC-A) from a particular location were sometimes composed of genetically very distinct organisms.

C. Other Major Oceanic Cyanobacteria Richelia intracellularis is sometimes found freeliving but, as its name implies, is usually observed as an endophyte in species of the diatom Rhizolenia. It is easily seen in the light microscope and the cyanobacterium consists of a short filament with a terminal heterocyst (Mague et al., 1977). The number of filaments per diatom cell varies with the host

species andtheassociationwhich iswidelydistributed in temperate and tropical water is thought to make a significant contribution to the biologically available nitrogen (Fogg, 1982). Much more is known about Trichodesmium species, whose phycoerythrincontaining filaments are often observed as bundles in which many filaments are longitudinally aggregated together. It is this feature which separates them from the Oscillatoria sp., a distinction with which not all taxonomists are in agreement, but which is important oceanographically. It was considered that the assembly into bundles wouldpermit the development of physiologically specialized areas at the center of the bundle that would be relatively microaerobic and thus facilitate the operation of nitrogenase. Attempts were made to correlate the climatic and hydrographic conditions that allowed bundle formation with the measurement of nitrogen fixation (Carpenter, 1983; Fay, 1992). As an increasing proportion of nonheterocystous cyanobacteria were shown to be capable of nitrogen-fixation the solution to this question became less pressing. Molecular biology has now unequivocally shown Trichodesmium sp. to be nitrogen-fixing (see Section VIII B) and their considerable role in determining oceanicproductivity, particularly in tropical and semi-tropical areas, is being increasingly documented. Information on this genus will be considered, where appropriate, for comparison. II. Macromolecular composition

A. DNA The mean DNA base composition of a considerable number of strains of Synechococcus (MC-A) has been determined and the base ratios cover the range 54.9 to 62.4 mol % G+C (Waterbury et al., 1986). This broad span of base composition was taken to indicate that, though the organisms appear morphologically similar, they in fact exhibit considerable genetic heterogeneity. This taxonomic problem reflects the difficulty in the classification of small unicellular freshwater cyanobacteria. The base composition of DNA from Trichodesmium sp. is reported as 69 mol % A+T (Zehr et al., 1991a) Cuhel and Waterbury (1984) have reported a DNA content of 2.1 fg per cell for an axenic culture of Synechococcus sp. strain WH 7803. This value of

Chapter 2 Oceanic Cyanobacterial Picoplankton DNA per cell, if it were assumed to correspond to a single genome, would indicate a genome size of approximately basepairs. Such a genome size is only about 78% of the smallest genome size measured for freshwater cyanobacteria (Herdman et al., 1979) and only about 55% of the size of the genome of E. coli. It should be mentioned that this estimate of DNA content per Synechococcus sp. strain WH 7803 cell is based on an estimation of the phosphorus content of the cell and then on an estimate of the DNA contribution to total P content. Consequently, the genome size interpretation must be considered as, at best, a rough estimate and perhaps the best interpretation is that the smallest genome sizes of the freshwater and marine cyanobacteria are similar. In the case of freshwater cyanobacteria there is a widespread distribution of restriction endonucleases (see Houmard and Tandeau de Marsac, 1988) and a consequence of this is that DNA isolated from cyanobacteria is frequently resistant to cutting with a variety of restriction enzymes (van den Hondel et al., 1983; Herrero et al., 1984; Lambert and Carr, 1984). In addition it has been shown that the DNAs of three strains of cyanobacteria from the genera Anabaena, Plectonema and Synechococcus contain a high proportion of and 5-methylcytosine and that Dam-like and Dcm-like methylases are responsible, as well as the site-specific methylase counterparts of the restriction systems (Padhy et al., 1988). As yet there are no reports concerning the distribution of restriction enzymes in the marine cyanobacteria, and the authors have experienced no problem in restricting DNA from Synechococcus sp. strain WH 7803; however, evidence from studies with phages infecting Synechococcus (MC-A) suggest extensive DNA modification and/or the presence of restriction-modification systems (see Section IV). In the case of Trichodesmium sp., Zehr et al.(1991) describe the resistance of genomic DNA from strain NIBB 1067 to digestion by restriction endonucleases and report that as much as 15 mol % of the deoxyadenosine was modified at a position other than the usual position. In order for the adenine residues to be modified at this many positions, it is suggested that there must be several modification enzymes or at least one of the modifying enzymes must have a degenerate specificity. The extent of the modification also raises questions as to its significance for the ecology of the organism.

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B. RNA and Protein Synthesis Direct measurements of protein and RNA synthesis have been made in Synechococcus sp. strain WH 7803 using incorporation of and uracil (Kramer, 1990). Both compounds were readily transported, leucine being assimilated more rapidly and to a greater extent than uracil; short term experiments indicated that turnover in precursor pools was rapid compared to the generation time. The assumption that these pools form the immediate precursors of high molecular weight fractions was supported by inhibitor studies with chloramphenicol and rifampicin. In addition, use of the inhibitor DCMU suggested that transport of these substrates was linked to photosynthetic electron transport, but PSI-dependent phosphorylation was not the exclusive source of energy required to support transport. In keeping with these observations, incorporation of isotope into protein was dramatically reduced in the dark and incorporation into RNA was eliminated. Cuhel and Waterbury (1984) observed rapid uptake of nearly 20% of the isotope was incorporated, primarily into RNA, into cells within 30 hours, though non-trivial amounts were found in the protein fraction. thymidine was not significantly incorporated making this label inappropriate for measuring DNA synthesis. Kramer and Morris (1990) have shown that Synechococcus sp. strain WH 7803 responds to increases in irradiance by rapidly increasing the rates of macromolecule synthesis—the order being RNA, protein and DNA synthesis. Patterns of macromolecule synthesis following a decrease in irradiance indicated that the rate of protein synthesis was maintained despite a reduction in RNA synthesis. These studies tend to confirm the idea that, as in other procaryotes, the protein synthesizing system plays a central role in regulating growth. III. Phycobiliproteins A full description of phycobiliprotein structures and their arrangement into phycobilisomes will be found in Chapter 7 and the organization and transcription of phycoerythrin genes is discussed in Section VI. What is presented here is a brief account of some of the special features of phycobiliproteins associated with Synechococcus (MC-A). In addition to phycocyanin,

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allophycocyanin, and occasionally phycoerythrocyanin, the large amounts of phycoerythrin found in these organisms in their natural environment is consistent with their distribution through the photic zone and with them being organisms capable of maximum photosynthetic rates in low light (Kursar et al., 1981; Waterbury et al., 1986). The detailed biochemical analysis of their phycobiliprotein pigments, mainly by Glazer and colleagues, has shown adaptation and modification of phycobilisome components to maximize absorption of the wavelengths around 500 nm at which solar irradiance best penetrates through sea-water (see Ong and Glazer, 1988). Although considerable homology exists between the polypeptides of phycoerythrins there is marked variation in their bilin chromophore composition (de Lorimer et al., 1992a). The well-described, phycoerythrin-containing cyanobacteria from freshwater typically contain only phycoerythrobilin chromophores (PEB, absorbance maximum ~550 nm) while some ofthe Synechococcus (MC-A) have a major proportion of phycourobilin chromophores (PUB, absorbance maximum ~495 nm) previously known to be a component of the phycoerythrins of the eucaryotic red algae. In these Synechococcus (MC-A) the light energy absorbed by PUB is transferred to reaction centers via PEB (Ong and Glazer, 1991; Swanson et al., 1991). Furthermore, an unusual phycocyanin, termed R-phycocyanin II and carrying only one phycocyanobilin (PCB) and two PEB chromophores (Ong and Glazer, 1987), has been described in marine Synechococcus sp strains. The fine adjustment to environmental niche that is demonstrated by relative arrangements around a common structure of phycobilisomes from freshwater and marine cyanobacteria is understandable. Because such a large proportion of protein is invested in the construction of the light-harvesting antennae, the energetic cost to the organism is great— compare for example the rather modest investment that an antenna system based on chlorophyll b involves (Bryant, 1992). Thus, selection pressure to evolve an optimum protein-based light-harvesting apparatus will be great, and this is especially true in an environment whose productivity can be limited by energy and nitrogen supply. It is well established that phycocyanin in cyanobacteria can serve as a nitrogen reserve (see Allen, 1984) and the probability that this is the case for phycoerythrin in the Synechococcus (MC-A) is

Noel G. Carr and Nicholas H. Mann clearly relevant to their competition for nutrients with other phytoplankton. The relatively high ratios of phycoerythrin to phycocyanin formed under photon flux densities that permit near-maximum growth rate are consistent with this function (Wyman et al., 1985). Because these organisms will grow without photoinhibition at high photon flux well in excess of their optimum light requirement, phycoerythrin content can be markedly lowered, and reductions of up to twenty-fold with increasing photon-flux densities have been observed (Kana and Glibert, 1987a). Kursar et al. (1981) noted that a clone of the organism later designated Synechococcus sp. strain WH 7803 exhibited marked in vivo phycoerythrin fluorescence. Evidence that the phycoerythrin in Synechococcus sp. strain WH 7803 had novel features came from varying nitrate supply to turbidostats in which growth rate was, or was not, limited by photon-flux density (Wyman et al., 1985). A proportion of the light energy absorbed by phycoerythrin in fast-growing cultures under high photon-flux density was released as autofluorescence and was therefore uncoupled from transfer to reaction centers when nitrate was available in excess. This was confirmed by excitation with green light and the measurement of the induced delayed fluorescence, a parameter which measured excitation of PS II reaction centers and associated antenna chlorophylls. The delayed fluorescence observed was of similar intensity from N-limited and N-sufficient cultures, which had significantly different amounts of phycoerythrin (Wyman et al., 1985). These results indicated that this organism could accumulate phycoerythrin in excess of that amount needed for maximal photosynthesis and that at least some of this extra pigment autofluoresced its absorbed light energy. These experiments do not inform us as to the physical arrangements of the excess phycoerythrin and there is no reason to postulate that any non-phycobilisome form was present. The consequences of the ability of Synechococcus sp. strain WH 7803 to store nitrogen as phycoerythrin, a proportion ofwhich is energetically uncoupled, has nutritional implications. Nitrogen shift-down experiments showed that cultures carrying extra phycoerythrin could maintain growth in the absence of exogenous nitrogen for approximately a division time. The proportion of phycoerythrin declined during this period and all non-coupled pigment was lost (Wyman et al., 1985). Experiments using non-axenic cultures of this organism by Glibert et al. (1986), in

Chapter 2 Oceanic Cyanobacterial Picoplankton which there were methodological differences of design, did not demonstrate mobilization of phycoerythrin after nitrate starvation (also see Kana and Glibert, 1987b), although mobilization was observed in Synechococcus sp. strain WH 8018 (another strain belonging to Marine-Cluster A; T. M. Kana, personal communication). Picosecond time-resolved fluorescence spectroscopy of exponential cultures of Synechococcus sp. strain WH 7803 grown under non-light limiting conditions and in the presence of excess nitrate showed that phycoerythrin had a fast (100 ps) and a slower (1300 ps) decay component, the latter indicating that a significant fraction (of the order of 15%) of phycoerythrin was not transferring its excitation energy to the photosynthetic reaction center (Heathcote et al., 1992) and, given the methodology employed, this should be considered as a minimum value for that culture. In an examination of the timescales of the response of Synechococcus sp. strain WH 7803 to nitrogen-shift-up and shift-down experiments, rapid alteration of the transcription of the co-transcribed cpeA and cpeB genes was observed (M. Wyman and N. G. Carr, unpublished results). The specificity of this response was indicated by the much slower response of photosynthetic reaction center genes such as psbA. IV. Plasmids Since at least two isolates of the phycoerythrin containing marine Synechococcus (MC-A) can now be grown to single colonies on solid growth media, the potential exists to employ the diverse range of molecular genetic techniques available for analysis of their freshwater counterparts (see Chapter 19). This ability would be enhanced if plasmids capable of replicating in the Synechococcus (MC-A) strains were available. It may be possible that broad host range plasmids can be mobilized into marine species since the promiscuous IncQ plasmid pKT210 was successfully transferred by conjugation from E. coli to the freshwater species Synechocystis sp. strain PCC 6803 (Kreps et al., 1990); alternatively, vectors based on endogenous plasmids could be constructed. As yet nothing is known about the plasmid content of the open-ocean cyanobacterial species, though efforts in this laboratory to detect plasmids have been unsuccessful. Information is, however, available for halotolerant coastal species.

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Synechococcus sp. strain PCC 7002 (Agmenellum quadruplicatum strain PR-6) is a coastal species which is readily transformable and for which a variety ofvectorsbasedononeofthesixendogenous plasmids have been constructed (see Houmard and Tandeau de Marsac, 1988; and Chapter 19). One report (Matsunaga et al., 1990) describes thecharacterization offorty marine isolates from coastal areas ofJapan in terms of their plasmid content. Among the forty isolates, five could be demonstrated to contain 1-3 different plasmids. The plasmids ranged in size from 2.3 -16.8 kb and no phenotype could be ascribed to them. A 1.4 kb HindIII fragment from one of the plasmids (pSY11) of the isolate Synechococcus sp. strain NKBG 042902 was used to create a hybrid plasmid (pUSY02) with the E. coli plasmid vector pUC18. This hybrid plasmid could be re-introduced into a cured derivative ofits original host by a variety of transformation methods including electroporation, selection being made for ampicillin resistance. pUSY02 was also capable of replication in the freshwater species Synechococcus sp. strain PCC 7942. In contrast, however, theplasmid pSG 111, that is capable of replication in Synechococcus sp. strain PCC 7942, apparently could not replicate in the marine strain. It is worth noting that the marine isolate Synechococcus sp. strain NKBG 042902, although normally grown in the presence of 3% NaCl, was easily capable ofadapting to conditions of lower salinity and therefore must be considered as being quite distinct from the obligately halophilic Synechococcus (MC-A). Conjugative gene transfer has also been described for marine cyanobacteria (Sode et al., 1992). Transfer of the broad host range IncQ plasmid pKT230 from E. coli into Synechococcus sp. strain NKBG 15041C was detected at a comparatively high frequency and Southern blotting suggested that pKT230 was capable of autonomous replication in this strain. Transfer of Tn5 into Synechococcus sp. strain NKBG 15041C via the suicide vector pSUP1021 from the E. coli mobilization strain S17-1 was also detected, opening the possibility for random transposon mutagenesis. Again, the strains involved in this study were halotolerant and thus must be regarded as distinct from Synechococcus (MC-A). V. Phages Until the late 1980s it was conventionally thought

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that the concentration of bacteriophages in natural unpolluted waters was low. However, using a new method for the enumeration ofvirus particles, Bergh et al. (1989) reported concentrations ofup to viruses per milliliter in natural waters. Proctor and Fuhrman (1990) reported high viral abundance in the ocean and also counts of marine cyanobacteria (and bacteria) in the final irreversible stage of lytic infection. As many as 5% of cyanobacteria from diverse marine locations contained mature phage, and interpretation from culture data suggested that up to 70% ofprocaryotes could be infected. Suttle et al. (1990) provided evidence that infection of the phytoplanktonby viruses could, in addition to grazing and nutrient limitation, be a significant factor affecting primary productivity. Studies on the abundance and distribution of cyanophages in the marine environment have produced varying estimates. Suttle et al. (1993) report titers of cyanophages routinely occurring at concentrations in excess of with the highest abundances being detected closest to shore and in near-surface waters along a transect in the Gulf of Mexico. Cyanophages were detected over a wide range of water temperatures (12–30.5°C) and salinities (18–70ppt; Suttle and Chan, 1993), but the frequency of detection was markedly influenced by the strain of Synechococcus sp. used for screening; viruses infecting phycoerythrin-containing strains were particularly common. Cyanophage abundances are somewhat lower with titers often being in excess of phage though the pattern of higher abundances in inshore waters was confirmed (J. B. Waterbury, personal communication). The abundance of cyanophages in the marine environment is matched by their diversity. Waterbury and Valois (1993) have isolated over 50 Synechococcus sp. phages using dilution cultures. Electron microscopy revealed that the isolates included representatives from each of the three families of tailed phages; the vast majority were contractiletailed phages of the family Myoviridae. Within a single water sample collected in the Gulf Stream, eight morphologically distinguishable Synechococcus sp. phages were isolated, with five types being obtained following enrichment with a single host (Synechococcus sp. strain WH 8012). The phages that have been isolated so far have varying host ranges. Amongst phages infecting Synechococcus sp. strains of Marine-Cluster A, some could infect as many as 13 strains, whereas others will infect only

Noel G. Carr and Nicholas H. Mann one strain. One phage isolate could infect not only MarineCluster Astrains butalsostrainSynechococcus sp. strain WH 8101 from Marine-Cluster B (J. B. Waterbury and F. W. Valois, personal communication). Suttle and Chan (1993) were able to isolate marine cyanophages that included representatives from each of the genera proposed for freshwater cyanophages, namely Cyanopodovirus, Cyanostylovirus and Cyanomyovirus; only members of the latter genus could infect thephycoerythrin-containing strains. Wilson et al. (1993) reported the isolation by plaque purification of five strains of cyanophage capable of infecting and lysing the Synechococcus sp. strain WH 7803. The five cyanophage isolates were obtained from samples from distinct oceanic provinces: two from coastal waters off Plymouth Sound, one from Wood’s Hole harbor and two from coastal waters off Bermuda. Three distinct plaque morphologies were observed. Two cyanophage isolates (one from Plymouth Sound and one from Bermuda) gave large (> 3 mm) plaques, while two other isolates (again one from Plymouth Sound and one from Bermuda) gave small ~ 1 mm plaques. The fifth isolate (from Wood’s Hole harbor) gave an intermediate plaque morphology. Negative staining and electron microscopy revealed differences in morphology that correlated with plaque morphology and restriction analysis (see below). The ‘large’ plaque strains had large hexagonal heads, collars, and short tails with a pronounced sheath characteristic of the Myoviridae. The Wood’s Hole isolate had a similar sized head, longer tail with tail fibers, and a wider collar; it was also classified as a member ofthe Myoviridae. The ‘small’ plaque strains had isometric heads and long, rigid tails and consequently they were assigned to the family Styloviridae. None ofthe isolates could give a lytic infection with the halotolerant marine strain Synechococcus sp. strain PCC 7002 nor with the freshwater strains Synechococcus sp. strain PCC 7942 and Anabaena sp. strain PCC 7120. None ofthe isolates could infect the other marine isolate Synechococcus sp. strain WH 8103, but all could infect strains Synechococcus sp. strains WH 8012 and WH 8018. The effect of these cyanophages on Synechococcus sp. mortality is of central importance in assessing their contribution to what has been called ‘the viral loop.’ The marine cyanophages appear to be extremely sensitive to solar radiation suggesting that most cyanophage particles in surface waters should be

Chapter 2 Oceanic Cyanobacterial Picoplankton non-infective (Suttle et al., 1993). In addition, it appears that natural populations of Synechococcus sp. are resistant to their co-occurring phages (Waterbury and Valois, 1993). These observations seem at odds with the observed abundance of cyanophage, and to explain this paradox Suttle et al. (1993) have suggested that this apparent lack of infectivity is overcome by the host cell’s own DNA repair mechanisms. Laboratory studies by Wiggens and Alexander (1985) found that a minimum hostdensity threshold of approximately cells is required for productive host-phage interaction. The mechanism(s) of host resistance to infection, are not known but could arise from lysogeny (not so far observed), absence of the phage receptor (due to mutation or nutritional factors), prevalence of diverse restriction systems, and wide variations in mol % G+C content. Molecular biological studies on the marine cyanophages have so far been very restricted. DNA prepared from the five cyanophage strains studied by Wilson et al. (1993) yielded three distinct patterns of digestion with restriction endonucleases; the two isolates with the ‘large’ plaque morphology yielded identical restriction patterns, as did the two isolates with the ‘small’ plaque morphology. The Wood’s Hole isolate had a different restriction pattern from the other four isolates. All three Cyanomyovirus isolates had a genome size estimated from restriction analysis of the order of 80–85 kb, and the Stylovirus isolates were about 95-100 kb. Genomes sizes based on restriction analysis, that in some cases were larger than 100 kb have been calculated (C.A. Suttle, personal communication). After propagation of all five phages in the host Synechococcus sp. strain WH 7803 phage DNA was refractory to restriction digestion with several enzymes, including HindIII, MluI, XhoI, Sau3A, SmaI, and PvuII suggesting extensive modification of the phage DNA by Synechococcus sp. strain WH 7803. In contrast the enzymes BamHI, PstI, and ClaI were able to restrict the phage DNAs. A particularly interesting effect was seen with EcoRI: the enzyme would digest phage DNA from two of the three Cyanomyovirus strains and the Cyanostylovirus isolate propagated on Synechococcus sp. strain WH 7803, but would not digest the DNA from the third (Wood’s Hole) Cyanomyovirus isolate, suggesting that this particular phage may encode it own restriction-modification system. Southern blot experiments using the Wood’s

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Hole isolate as aprobe revealed significant similarities to the other two Cyanomyovirus strains, but only very limited cross-hybridization with the Cyanostylovirus isolates. SDS-PAGE analysis of viral polypeptides of representative phage isolates purified by CsCl density-gradient centrifugation showed almost identical polypeptide profiles for the Cyanomyovirus isolates and, as expected, marked differences for the Cyanostylovirus isolate (with the exception of the most abundant, presumably structural, protein which had a mass of about 45 kDa in all cases). No evidence has so far been obtained for lysogenic cyanophages infecting Synechococcus (MC-A). Thus, far greater efforts to study cyanophage mode(s) of replication and other aspects of the molecular biology of their life cycle will need to be made before their potential as genetic tools to be used with Synechococcus (MC-A) strains can be realized. In addition, analysis of interactions between phage and the receptors on the host cells may yield interesting insights into the effects of nutrient availability on phage infectivity. VI. Transcription Although several genes, particularly those for phycobiliproteins, have been cloned and sequenced from Synechococcus (MC-A) strains, there is virtually no information concerning the transcription of these genes and what information is available is largely based on inferences drawn by computer analysis of nucleotide sequence information. In only one case, that of the class-I phycoerythrin subunit (cpeBA) genes from Synechococcus sp. strain WH 7803 (Newman et al., 1993), has the transcription startpoint been identified. In this particular strain the arrangement of the two genes is typical of many cyanobacterial phycoerythrin genes, with cpeB being located upstream of cpeA: a similar pattern of organization is seen in Synechococcus sp. strain WH 8020 (Wilbanks and Glazer, 1993). The cpeBA operon of strain Synechococcus sp. strain WH 7803 is transcribed as a 1.3-kb dicistronic transcript, with the transcription startpoint being localized to 110/111 base pairs upstream of cpeB. No obvious promoter sequences conforming to that of the E. coli consensus can be detected upstream of the transcription startpoint(s). However, certain features that may be of transcriptional significance can be detected,

36

including a triple repeat of the pentanucleotide sequence 5'-CGGTT-3' within the 40 base pairs preceding the startpoint, and overlapping sequences some 200 bp upstream resembling integration host factor (IHF) binding sequences (Newman et al., 1993). Additional information can be obtained by comparing the upstream sequence from the cpeBA genes of Synechococcus sp. strain WH 7803 with an equivalent region from Synechococcus sp. strain WH 8020 (Wilbanks and Glazer, 1993). Such an alignment (Fig. 1) reveals two areas of sequence similarity. At a position centered on –10 from the startpoint there is the sequence 5'-TTANGTT-3' which may represent a component of the promoter and centered on position –60 is a highly conserved sequence of 21 bases with 19 identical residues between the two sequences. The significance of these sequences is yet to be established, though no indication, using gel retardation studies, of protein binding to a restriction fragment including these sequences was obtained with cell-free extracts from Synechococcus sp. strain WH 7803 (Newman et al., 1993). As regards mechanisms of transcription termination, information is entirely confined to structures inferred from analysis of sequence data. Given the size (1.3 kb) of the cpeBA transcript in Synechococcus sp. strain WH 7803, termination must occur very shortly after the cpeA gene. Indeed, there is a sequence immediately downstream from cpeA that conforms to the predicted structure of a rho-independent terminator. Similar structures can be detected immediately downstream from the cpeA gene of Synechococcus sp. strain WH 8020 and the pstS gene of Synechococcus sp. strain WH 7803.

Noel G. Carr and Nicholas H. Mann VII. Translation

A. Codon Usage Analysis of the nucleotide sequences of a large number of genes from many species has revealed that in the large majority of cases the various synonymous codons for a particular amino acid are not used with equal frequencies (see Ikemura, 1985). This is a reflection, at least in part, of directional mutation pressure which has led to variations in the mol % GC base composition within genomes. Although only a relatively small number of genes have been sequenced from the Synechococcus (MC-A) strains, it is already possible to obtain reasonably reliable data on the pattern(s) of codon usage. Eight genes from the lowPUB strain Synechococcus sp. strain WH 7803 have been completely sequenced (see Table 1), and these include the genes for the some of the phycobiliproteins as well as for other less abundant proteins. The overall mol % G+C composition of DNA from this strain is 61.3% (Waterbury et al., 1986) and in fact the base compositions of all eight genes fall within a few percent of this value (Table 1). In the high-PUB strain Synechococcus sp. strain WH 8103 which has a mol % G+C composition of 58.9% (Waterbury et al., 1986), the only available sequence information is for the mpeB and mpeA genes (encoding class-II phycoerythrin subunits; de Lorimer et al., 1992a); the mol % G+C values of 59.4 and 60.0 for these genes also fall close to the overall genomic base composition. A rather different pattern has been observed with the strain Synechococcus sp strain WH 8020 (Wilbanks and Glazer, 1993). A 14.9-kb

Chapter 2 Oceanic Cyanobacterial Picoplankton

region, containing both the mpe and cpe gene clusters as well as a number of other ORFs, has been completely sequenced and marked differences in mol % G+C compositions were observed. The cpeB and cpeA genes were 54 mol % G+C, the mpeB and mpeA genes were 48.5 mol % G+C and the ORFs between mpeC and cpeA were the lowest at 41.1 mol % G+C. The overall mol % G+C for the 15-kb region was 47.3% but the overall genomic base composition is not known; however, it would be surprising if it were outside the range 54.5 – 62.4 mol % G+C. There is as yet no explanation for these inter-strain differences. In the cases of both Escherichia coli and Saccharomyces cerevisiae there is a strong correlation between the degree of codon bias and the level of gene expression (Gouy and Gautier, 1982; Bennetzen and Hall, 1982). This would also appear tobe the case for Synechococcus sp. strain WH 8020 (Wilbanks and Glazer, 1993), as judged by differences in the frequencies with which the different nucleotides appear at the third position in codons. In the case of the phycobiliprotein genes the frequencies were C>T»A>G for while the pattern at for the other genes in the 15-kb region was T>A>C>G. In Synechococcus sp. strain WH 7803 a quite different nucleotide frequency is found at for the highly expressed biliprotein genes, namely C»T>G>>A. In this strain, however, there is no marked difference in the frequencies in genes which might be expected to be expressed at lower levels (e.g., woxA, pstS, phoB and phoR) with the frequenciesC>>G~T>A. Similarly in Synechococcus sp. strain WH 8103 a frequency of C>>T>G>A may be calculated for the published sequences for mpeB and mpeA (de Lorimer et al., 1992a). Thus, in the case of strain Synechococcus

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sp. WH 7803 a pattern of codon usage may be put together (Table 2) which applies equally well, apparently, to both highly expressed and less highly expressed genes. This codon bias successfully identifies coding regions using programs such as ‘codonpreference’ (Devereux et al., 1984) in nucleotide sequences from strain Synechococcus sp. strain WH 7803. Indeed, it can also be used to analyze nucleotide sequences from Synechococcus sp. strain WH 8103 and accurately identifies the coding regions for class II phycoerythrin subunit genes. Comparison of this codon usage table with one calculated for the freshwater strain Synechococcus sp. strain PCC 7942 (van der Plas, 1989) shows some similarities, but also several marked differences. For example, the most commonly used codons for serine, proline and lysine in strain Synechococcus sp. strain WH 7803 are TCC, CCC and AAG respectively, but in Synechococcus sp. strain PCC 7942 are AGC, CCG and AAA. In addition, TAG is by far the most commonly used termination codon in the freshwater strain, but it is TGA in the marine strain. Thus, there appear to be two distinct patterns of codon bias between freshwater and marine Synechococcus sp strains as well as between different Synechococcus (MC-A) strains; as more nucleotide sequence data become available for the marine strains, general patterns of codon bias should become discernible. VIII. Nutrition

A. Nitrogen 1. Assimilation All the Synechococcus (MC-A) strains tested by Waterbury et al. (1986) can use nitrate and ammonia

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as their sole nitrogen source and roughly half the strains tested could use urea as the sole nitrogen source. In addition, the major light-harvesting pigment phycoerythrin can function as a nitrogen reserve

Noel G. Carr and Nicholas H. Mann

material (see Section III). The assimilation of amino acids has also been demonstrated for Synechococcus sp. strain WH 7803 with (Kramer, 1990) and (Chadd, 1992) and for both this

Chapter 2 Oceanic Cyanobacterial Picoplankton strain and Synechococcus sp strain WH 8101 with a acid mixture (Paerl, 1991). methionine incorporation can be used effectively to label proteins in Synechococcus sp. strain WH 7803 and thereby facilitates the analysis ofthe response of this organism to changed environmental conditions in terms of the synthesis of novel proteins (Chadd, 1992). The uptake system for exogenous methionine does not become saturated until an external concentration exceeding 100 nM isreached. Although there are no reports ofamino acids serving as the sole nitrogen sources for Synechococcus (MC-A) strains, a stimulation of growth yield has been observed for Synechococcus sp. strain WH 7803 growing in ASW medium (Wyman et al., 1985) when lysine is added to the culture at a concentration of (N. G. Carr and N. H. Mann, unpublished results). In the freshwater strain Synechococcus sp. strain PCC 7942, ammonium acts as a repressor of proteins involved in nitrate assimilation (see Chapter 16). The gene ntcA, encoding a protein of the Crp and Fnr family of transcriptional activators, is required for full expression of ammonium repressible genes (Vega-Palas et al., 1992) including a 45-kDa nitratetransport protein, the gene for which (nrtA) has been sequenced (Omata, 1991). Glibert and Ray (1990) report that Synechococcus sp. strain WH 7803 will only take up nitrateafter is depleted, and that the nitrate uptake rate was only about 12% of that observed for This observation may be an example of phyletic inertia (Brand, 1986) or, alternatively, may reflect the chemical form of nitrogen to which the organisms are exposed in their natural environment. Southern blots ofchromosomal DNA from Synechococcus sp. strain WH 7803 have been probed with the ntcA and nrtA genes, and evenatverylow stringencies no specific cross-hybridizing bands were detected (D. J. Scanlan, personal communication). However, Synechococcus sp. strain WH 7803 does appear to exhibit an adaptive response to nitrogen limitation. SDS-PAGE analysis of extracts made from cells grown under nitrogen-limited conditions has revealed the synthesis ofseveral novel polypeptides including abundant species of 60 kDa and 16 kDa, the former being localized to the cell envelope (D. J. Scanlan, unpublished results).

2. Nitrogen Fixation None of the Synechococcus (MC-A) thus far examined

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can fix nitrogen (Waterbury et al., 1986), though it is interesting to note that a potential homologue of the gene hetA, that acts early in the process ofheterocyst differentiation (Holland and Wolk, 1990) has been partially characterized in Synechococcus sp. strain WH 7803 (G. Watson, personal communication). However, since the first description by Wyatt and Silvey (1969) there has been an increasing number of reports ofunicellular cyanobacteria which are known to fix nitrogen, under a variety of environmental conditions (see Gallon, 1992; Fay, 1992; and Chapter 16). Some of these are from coastal sources (Leon et al., 1986; Waterbury and Rippka, 1989), and recently two well-described unicellular, aerobic nitrogenfixers from a coastal area have been assigned to the Cyanothecegroup (Reddyetal., 1993). Theseworkers report extensive cyanophycin reserves in these organisms. It will be ofinterest to see ifthe nitrogenfixing tropical oceanic strain Synechocystis sp. strain WH 8501 (Waterbury and Rippka, 1989) also has this material, which has been considered both as a storage form and as a dynamic reservoir of nitrogen assimilation (Carr, 1988). There is no apparent reason why open-ocean, unicellular cyanobacteria should not have acquirednitrogen-fixing capacity exceptfor the euphotic zone’s permanent oxygenated state and the low surface area/volume ratio of the Synechococcus (MC-A), allied perhaps to the considerations discussed in Section I. Trichodesmium sp. is the only open-ocean cyanobacterium for which detailed information on nitrogen fixation exists (see Carpenter, 1983; 1992; Ohki et al., 1992b; Fay, 1992). In spite of the difficulties in maintaining cultures and of purification, the ability of this organism to fix nitrogen was established some years ago and this was later supported by the use of and acetylene reduction techniques. Because of the suggested connection between fixation and the aggregation of filaments into bundles, there has been speculation as to whether there existed ‘physiological specialization’ where inner areas of the bundles did not perform oxygenevolving photosynthesis thereby creating a relatively microaerobic zone. Polyclonal antibodies against dinitrogen reductase showed that this protein was present in all the cells ofa Trichodesmium sp. colony (Paerl et al., 1989), and it would thus appear that any microaerobic zone in such a colony of filaments would arise by self-shading in the course of the aggregation of filaments. Using Trichodesmium sp

40

strain NIBB 1067 in culture, Ohki and Fujita (1988) showed that acetylene reduction was maximal during exponential growth before bundle association of the filaments had developed. The nifH, nifD, and nifK. genes of the nif operon were detected in Trichodesmium sp. using DNA probes from Anabaena sp. strain PCC 7120 and were found to be contiguous (Zehr et al., 1991a). Although growth with nitrate, ammonium or urea prevents acetylene reduction, only urea repressed the synthesis of the Mo-Fe and Fe-proteins (Ohki et al., 1991). Both proteins were detected immunologically, but the Fe-protein had an apparent mass of 38 kDa in contrast to the 40-kDa which had been found in nitrogen-fixing cultures. The regulation of nitrogenase in relation to light-dark regimes has been examined in culture with the same organism (Ohki et al., 1992a) and in natural populations of Trichodesmium thiebautii (Capone et al., 1990). In each case the conclusion drawn was that nitrogenase activity was associated with the light period and that the enzyme was activated at the beginning of illumination. This was achieved both by de novo synthesis and activation and de-activation associated with change in molecular weight of the Fe-protein. The 38-kDa form appears to be the active protein which is converted to the 40 kDa form in natural populations at night and under artificially high oxygen concentrations, suggesting that it has a transient protective role (Zehr et al., 1993).

B. Phosphorus Many aspects of phosphorus acquisition by freshwater cyanobacteria are comparatively well documented (see Whitton, 1992), but as yet very little information is available on the mechanisms by which marine cyanobacteria fulfill this particular nutritional requirement. Studies with Synechococcus sp. strain WH 7803 have shown that it is unable to grow with organic sources of phosphate such as glycerol phosphate (K. M. Donald - personal communication) and therefore presumably lacks the inducible surface monoesterase activity commonly found in freshwater species (Healey, 1982). Under conditions in which inorganic phosphate (Pi) is severely limited Synechococcus sp. strain WH 7803 induces the synthesis of several novel polypeptides including particularly abundant species with apparent masses of 100 kDa and 32 kDa (Scanlan et al., 1993). The structural gene for the 32-kDa polypeptide was cloned using an oligonucleotide following N-terminal

Noel G. Carr and Nicholas H. Mann sequencing of the purified polypeptide. Nucleotide sequencing revealed an ORF potentially encoding a polypeptide of 326 amino acids (33.7 kDa); the deduced amino acid sequence exhibited 35% identity (51% similarity) with the periplasmic phosphatebinding protein (PstS) of E. coli. Although the Synechococcus sp. strain WH 7803 32-kDa polypeptide could not be localized to the periplasm, it did co-purify with the cell envelope. The aminotertninal 24 amino acids of the Synechococcus sp. protein appear to be proteolytically cleaved (between two alanine residues) during transport to the cell envelope. Thus, both on the basis of its inducibility by Pi limitation, similarity to PstS, and localization it seems highly likely that the Synechococcus sp. protein is indeed a homologue of PstS and fulfills a similar function, namely the sequestration of Pi within the periplasm (perhaps anchored or loosely attached to the cell wall) and its presentation to a high affinity Pi transport system in the cytoplasmic membrane. The PstS protein of E. coli is a component of the inducible high affinity Pst transport system for Pi (see Rao and Torriani, 1990); the expression of the Pst transport system is under the control of a two-component sensory system composed of a histidine protein kinase (PhoR) and a response regulator (PhoB). Recently, Aiba et al. (1993) have characterized the genes (sphS and sphR) coding for a histidine protein kinase and response regulator which are implicated in controlling the adaptive response of the freshwater strain Synechococcus sp. strain PCC 7942 to phosphate limitation. Given the similarity between the periplasmic Pi-binding protein from Synechococcus sp. strain WH 7803 and E. coli, and the occurrence of a phosphate-related two component sensory system in a freshwater cyanobacterium, it is worth asking the question as to whether the similarity in Synechococcus sp. strain WH 7803 extends to other components of the Pst system. Alignment of the E. coli and B. subtilis PhoB and PhoR response-regulator proteins revealed several regions that were highly conserved, and a degenerate 27-mer oligonucleotide probe was designed against one of these regions using the known pattern of codon usage in Synechococcus sp. strain WH 7803 (Table 2). This oligonucleotide was used to screen plasmid libraries of Synechococcus sp. strain WH 7803 DNA and a positive clone was obtained that after sequencing revealed two ORFs which encoded polypeptides highly related to the phosphate system histidine protein kinases and response regulators (G. M. Watson, personal

Chapter 2 Oceanic Cyanobacterial Picoplankton communication). The putative PhoB and PhoR homologues are 76.6% and 56.2% similar to SphR and SphS of Synechococcus sp PCC 7942, respectively. An alignment of the putative response regulator from Synechococcus sp. strain WH 7803 with those of Synechococcus sp. strain PCC 7942, E. coli and B.

41

subtilis is shown in Fig. 2. Thus, the similarity with regard to a high affinity Pi transport system between the oceanic cyanobacteria and E. coli increases, and it must be supposed that Synechococcus sp. strain WH 7803 controls the inducibility of its high affinity

Noel G. Carr and Nicholas H. Mann

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Pi transport system via a two-component sensory system and presumably also has the cytoplasmic membrane components typical of a ‘shock sensitive’ transport system. To what extent arethese componentsofan inducible high affinity transport system for Pi common to cyanobacteria? A probe containing the 5' two-thirds of the Synechococcus sp. strain WH 7803 pstS gene was used to probe Southern blots of restriction enzyme digests of DNA from a range of both freshwater and marine cyanobacteria and homology, even at low stringency (~45% mismatching ), was confined to the phycoerythrin-containing marine Synechococcus sp. strains, with no detectable signal being obtained with freshwater strains, the halotolerant strain Synechococcus sp. strain PCC 7002, or to an abundant marine prochlorophyte, Prochlorococcus marinus. The marine Synechococcus sp. strains WH 8008, WH 8018 and WH 7803, suggested to be coastal or marine-shelf species (Olson et al., 1990), and belonging to different RFLP groups (Douglas and Carr, 1988) showed hybridizing fragments of identical size when probed with the Synechococcus sp. strain WH 7803 pstS gene. In contrast, Synechococcus sp. strain WH 8103, a high PUB-containing species characteristic of the open ocean, showed a quite different RFLP pattern using the pstS probe. Similar results were obtained with the antibody raised against the Synechococcus sp. strain WH 7803 PstS protein which, in immunoblots, cross-reacted with proteins from Synechococcus sp. strain WH 8103 and other phycoerythrin-containing marine strains, but not with halotolerant and freshwater strains (Scanlan et al., 1993). The lack of hybridization between the pstS probe and genomic DNA from freshwater strains may be explained by low levels of homology, since PhoB from Synechococcus sp. strain WH 7803 and SphS from Synechococcus sp. strain PCC 7942 are only 59.3% identical at the amino acid level and the genes exhibit 44% mismatch. Thus it would seem that the phycoerythrincontaining marine cyanobacteria may possess a system for the acquisition of Pi under limiting conditions that is similar, though somewhat distantly related, to that employed by the freshwater species. The availability of immunological and nucleic acid probes for the PstS protein and gene open the possibility ofusing molecular biological approaches for the characterization of the nutrient status with respect to Pi of natural assemblages of these oceanic strains.

C. Iron Iron is required by all cyanobacteria for a range of important physiological processes (see Chapter 25). Recently, Martin and his coworkers (Martin et al., 1990) have drawn attention to the possible role of iron in limiting the productivity of phytoplankton in certain ocean provinces: a suggestion that has provoked much discussion and some opposition (see Chisholm and Morel, 1991). The response of many procaryotic organisms to limited availability is to synthesize one or more high-affinity ferric iron chelators (siderophores) and associated components of a transport pathway. It is certainly the case that many species offreshwater cyanobacteria synthesize siderophores inresponse to restricted ironavailability (see Whitton, 1992) and siderophore production has been demonstrated for one halotolerant marine species Synechococcus sp. strain PCC 7002 (Kerry et al., 1988). A gene, irpA, encoding a protein involved in iron acquisition has been cloned and sequenced from the freshwater strain Synechococcus sp. strain PCC 7942 (Reddy et al., 1988). In addition to acquiring iron under conditions of restricted availability, cyanobacteria may also be capable of storing iron during times of relative abundance. The presence of a putative iron storage protein in freshwater cyanobacteria was established by Evans et al. (1977), and more recently bacterioferritin was purified and characterized from the freshwater strain Synechocystis sp PCC 6803 (Laulhère et al., 1992). The presence of a form of bacterioferritin in Synechococcus sp strain WH 7803 has been identified by Mössbauer spectroscopy (Mann et al., 1993). It has been shown that Synechococcus sp. strain WH 7803 is capable of growth under conditions of restricted iron availability following a lag phase of approximately three generation times, indicating that it can synthesize a high-affinity iron acquisition system. During this lag certain novel polypeptides (110 kDa, 96 kDa and 36 kDa) were synthesized, although no production of siderophores could be detected (Chadd, 1992). No close similarity to the iron acquisition system of freshwater strains is suggested, since no homologue of the irpA gene could be detected by Southern hybridization (H. E. Chadd, personal communication). A 36-kDa polypeptide synthesized in response to Fe-limitation was localized to the cell envelope. Antibodies have been raised against this protein, and immunoblotting detected a related polypeptide of similar size in the

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Chapter 2 Oceanic Cyanobacterial Picoplankton halotolerant strain Synechococcus sp. strain PCC 7002. In addition to induction ofiron acquisition systems, cyanobacteria exhibit a variety of other adaptive responses to iron limitation, including the replacement of iron-containing proteins with non-iron-containing functional homologs (see Chapter 25). Thus, ferredoxin containing an Fe-S center is replaced with flavodoxin as the terminal electron acceptor in photosynthesis (Hutber et al., 1977). Leonhardt and Straus (1992) have characterized an iron-stress operon involved in photosynthetic electron transport in the halotolerant marine cyanobacterium Synechococcus sp. strain PCC 7002. The iron-stress induced genes isiA and isiB have been cloned and sequenced and encode a PS II chlorophyll-binding protein related to PsbC (CP43; see Chapter 8) and flavodoxin, respectively. Using the cloned isiB gene as a probe, attempts to detect a flavodoxingene in arepresentative of the phycoerythrin-containing marine strains (Synechococcus sp. strain WH 7803) failed (D. J. Scanlan, personal communication). In laboratory cultures the marine nitrogen-fixing species Trichodesmium sp. strain NIBB 1067 has been shown to be capable of growth under conditions of restricted iron availability. These conditions cause effects on both the rates of photosynthesis and nitrogen fixation (Rueter et al., 1990). Natural samples of Triochodesmium sp. collected off Barbados responded to iron additions in ways that suggested that it may be iron-limited in its natural environment.

D. Carbon Assimilation The majority of cyanobacteria are photoautotrophs, though many species are capable of a degree of photoheterotrophy and a few exhibit facultative chemoheterotrophy (Rippka et al., 1979). The Synechococcus (MC-A) strains tested by Waterbury et al. (1986) were all obligate photoautotrophs, although several strains have been shown to incorporate exogenous organic molecules. Synechococcus sp. strain WH 7803 will incorporate acetate and adenine, but not glucose or thymidine (Cuhel and Waterbury, 1984) and both this strain and Synechococcus sp. strain WH 8101 will take up amino acids (see Section II B). Many cyanobacteria exhibit a bicarbonateconcentrating process which has been studied in freshwater species for some time (see Miller, 1990, and Chapter 15). Attention has focused recently on a

mechanism first suggested by Reinhold et al. (1987) that proposes that bicarbonate is concentrated in the cytoplasm and delivered into the carboxysome where carbonic anhydrase releases carbon dioxide which is fixed by Rubisco. Three strains of Synechococcus (MC-A) exhibited only slight, non-inducible bicarbonate concentration (Karagouni et al., 1990), although they do have carboxysomes (Waterbury et al., 1986). The explanation for this presumably is that an adequate supply of bicarbonate into the cell of a relatively slow growing organism with a large surface to volume ratio is achieved by the 1.8 mM external concentration present in sea water. It is ofinterest to note that inorganic carbon limitation has recently been observed for marine diatoms growing under otherwise optimal conditions (Riebisell et al., 1993). However, these organisms have a much smaller surface to volume ratio than the cyanobacterial picoplankton and assimilate and not bicarbonate. IX. Adaptive Responses

A. Highly Iterated Palindromic Sequences (HIP1) Recently the investigation of the mechanism by which cyanobacteria develop protection from heavy metal contamination has yielded some most interesting results. Metallothionein is a protein associated with protection from Cd and Zn and is conservatively and widely distributed in plants and microorganisms. A procaryotic metallothionein gene (smtA) was isolated and characterized from the freshwater cyanobacterium Synechococcus sp. strain PCC 7942 (Huckle et al., 1983). The smt locus contains, in addition to the smtA gene, the divergently transcribed smtB gene that encodes a metal-dependent represser of smtA (Morby et al., 1993). This is the first robust description in a cyanobacterium of a regulatory gene capable of acting in a trans configuration, and it is noteworthy that the gene is not involved in intermediary metabolism. Organisms exposed to heavy metals show an increased transcription of smtA and, most interestingly, cultures that had acquired cadmium resistance as a result of exposure to increasing concentrations over many weeks exhibited amplification of the gene (Gupta et al., 1992). Furthermore, such isolates exhibited deletion of 353 bp of the regulatory smtB gene and the boundaries of the region which had been deleted were traversed by identical octameric palindromes

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(5' GCGATCGC 3'; Gupta et al., 1993). Analysis of nucleotide sequences held in the ‘GenBank’ database (combined version of the GenBank/EMBL/DDBJ databases) revealed that this octameric highly iterated palindrome (HIP1) occurs on average every 664 bp within the genome of freshwater strains of Synechococcus sp. and also with a lower, but still relatively very high, frequency within the genomes of other cyanobacterial species, suggesting a widespread role for HIP1 in genome plasticity and cellular adaptation in these organisms (Gupta et al., 1993). In this respect the cyanobacteria were different from any other bacterial genera searched in the data base with exception ofhalobacteria and Thermus sp. in which other HIPs were identified (A. P. Morby, personal communication). Notable exceptions to this widespread distribution of HIP1 amongst the cyanobacteria are the Synechococcus (MC-A). Within 10-kb sample of nucleotide sequence from Synechococcus sp. strain WH 7803 from our own work, HIP1 was found only once and no occurrences were detected at all (A. P Morby, personal communication) in 14.9 kb of contiguous nucleotide sequence from Synechococcus sp. strain WH 8020 that had been deposited by Wilbanks and Glazer (1993). Furthermore, there was no evidence for any alternative highly iterated palindromes in these strains. The HIP1 sequence has also been found to be absent from a halotolerant Calothrix sp. (N. J. Robinson, personal communication). In light ofthe suggestion that the presence of HIP1 facilitates the adaptation of cyanobacteria to metal resistance, its absence from the Synechococcus (MC-A) is intriguing. The environment of these organisms, the open ocean, is extremely stable with respect to metal ion fluctuations and indeed to many other environmental variables that may challenge microbial growth, such as salinity variation, anthropogenic and natural organic molecules input for example. The volume of the ocean relative to such inputs acts as a massive homoeostatic mechanism. The argument may be advanced that these openocean organisms, have not evolved (or maintained) the HIP1 sequences because they are not exposed to the environmental fluctuations that characterize soil, freshwater and littoral locations and therefore, there is no advantage in their having a molecular strategy that allows them an adaptive response comparable to that in cyanobacteria found in other, less constant environments. This would have the implication that the nutritional fluctuations that these organism are

Noel G. Carr and Nicholas H. Mann exposed to — light, N, P, Fe, the list is quite limited (see Brand, 1986) — effect alterations in gene expression by means other than HIP1-facilitated rearrangements.

B. Protein Phosphorylation The occurrence of protein phosphorylation in freshwater cyanobacteria is well documented, particularly with respect to state transitions (for review see Allen, 1992), and has been implicated in adapting cells to changes in inorganic carbon availability, light-dark transitions and presence of exogenous organic carbon sources (Bloye et al., 1992). Thus far there are no published reports on the significance of protein phosphorylation in any ofthe marine strains, though patterns of protein phosphorylation in cellfree extracts of Synechococcus sp. strain WH 7803 have been observed that are reminiscent of those observed with freshwater species (N.J. Silman personal communication). In addition, Wilbanks and Glazer (1993) in the course ofsequencing a region of DNA from Synechococcus sp. strain WH 8020 encoding both phycobiliprotein and phycobilisome rod component genes characterized an incomplete ORF which potentially encodes a polypeptide with striking homology to the low-molecular-weight acid phosphatase from liver. They speculate that the protein product encoded by this ORF may be a candidate for a phosphoprotein phosphatase involved in mediating the State 2 to State 1 transition. In addition to the monoester phosphorylation of amino acids, phosphorylation of histidine and aspartate residues are involved in signal transduction through two component sensory systems (see Stock et al., 1989). The putative PhoR and PhoB homologues detected in Synechococcus sp. strain WH 7803 would be expected to exert control over the expression of Piregulated genes with the PhoR histidine protein kinase autophosphorylating at a histidine residue and activating the PhoB response regulator by transfer of the phosphate group to an aspartate residue on PhoR. Acknowledgments We thank our co-workers Dave Scanlan, Helen Chadd, Greg Watson, Nigel Silman, Julie Newman, William Wilson, Kirsten Donald and also Ian Joint and Michael Wyman from the Plymouth Marine Laboratory for discussions and access to unpublished work. Like many workers with the marine cyanobacterial

Chapter 2 Oceanic Cyanobacterial Picoplankton picoplankton we are indebted to John Waterbury for the gift of strains and valuable advice. We dedicate this chapter to the memory of Ian Morris who introduced us to oceanographic microbiology and to much else besides. References Aiba H, Nagaya M and Mizuno T (1993) Sensor and regulator proteins from the cyanobacterium Synechococcus species PCC7942 that belong to the bacterial signal-transduction protein families: Implication in the adaptive response to phosphate limitation. Mol Microbiol 8: 81–91 Allen JF (1992) Protein phosphorylation in regulation of photosynthesis. Biochim Biophys Acta 1098: 275–335 Allen MM (1984) Cyanobacterial cell inclusions. Ann Rev Microbiol 38: 1–25 Azam F, Fenchel T, Field JC, Gray JS, Meyer-Reil LS and Thingstad F (1983) The ecological role of water-column microbes in the sea. Mar Ecol Prog Ser 10: 257–263 Bennetzen JL and Hall BD (1982) Codon selection in yeast. J Biol Chem 257: 3026–3031 Bergh Ø, Børsheim KY, Bratbak G and Heldal M (1989) High abundance viruses found in aquatic environments. Nature (London) 340: 467–468 Bloye SA, Silman NJ, Mann NH and Carr NG (1992) Bicarbonate concentration by Synechocystis PCC6803: Modulation of protein phosphorylation and inorganic carbon transport by glucose. Plant Physiol 99: 601–606 Brand LE (1986) Nutrition and culture of autotrophic ultraplankton and picoplankton. In: Platt T and Li WKW (eds) Photosynthetic Picoplankton. Can Bull Fish Aquat Sci 214: 205–233 Bryant DA (1992) Puzzles of chloroplast ancestry. Curr Biol 2: 240–242 Campbell L and Iturriaga R (1988) Identification of Synechococcus spp. in the Sargasso Sea by immunofluorescence and fluorescence excitation spectroscopy performed on individual cells. Limnol Oceanogr 33: 1196–1201 Campbell L, Carpenter EJ and Iacon VJ (1983) Identification and enumeration of marine chroococcoid cyanobacteria by immunofluorescence. Appl Environ Microbiol 46: 553–559 Capone DG, O’Neil JM, Zehr J and Carpenter EJ (1990) Basis for diel variation in nitrogenase activity in the marine planktonic cyanobacterium Trichodesmium thiebautii. Appl Environ Microbiol 56: 3532–3536 Carpenter EJ(1983) Physiology and ecology ofmarine Oscillatoria (Trichodesmium) Mar Biol Lett 4: 69–85 Carr NG (1988) Nitrogen reserves and dynamic reservoirs in cyanobacteria. In: Rogers LJ andGallon JR(eds) Biochemistry of the Algae and Cyanobacteria, pp 13–21. Oxford Science Publications, Oxford Chadd HE (1992) Biochemical and molecular approaches to the study of iron nutrition in the marine cyanobacterium Synechococcus WH 7803. PhD thesis, University of Warwick Chisholm SW and Morel FMM (1991) What controls phytoplankton production in nutrient-rich areas of the open sea? Limnol Oceanogr 36: 1507–1969 Chisholm SW, Olson RJ, Zettler ER, Goericke R, Waterbury JB

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Chapter 2 Oceanic Cyanobacterial Picoplankton activity in relation to the light-dark regime in the filamentous non-heterocystous cyanobacterium Trichodesmium sp. NIBB 1067. J Gen Microbiol 138: 2679–2685 Ohki K, Zehr JP and Fujita Y (1992b) Trichodesmium: Establishment of culture and characteristics of In: Carpenter EJ, Capone DG and Rueter JG (eds) Marine Pelagic Cyanobacteria: Trichodesmium and other Diazotrophs, pp 307–318. Kluwer, Dordrecht Olson RJ, Chisholm SW, Zettler ER and Armbrust EV (1990) Pigment, size and distribution of Synechococcus in the North Atlantic and Pacific oceans. Limnol Oceanogr 35: 45–58 Omata T (1991) Cloning and characterization of the nrtA gene that encodes a 45-kDa protein involved in nitrate transport in the cyanobacterium Synechococcus PCC 7942. Plant Cell Physiol 32: 151–157 Ong LJ and Glazer AN (1987) R-phycocyanin II, a new phycocyanin occurring in marine Synechococcus species: Identification of the terminal energy acceptor bilin in phycocyanins. J Biol Chem 262: 6323–6327 Ong LJ and Glazer AN (1988) Structural studies of phycobiliproteins in unicellular marine cyanobacteria. In: Stevens Jr SE and Bryant DA (eds) Light-Energy Transduction in Photosynthesis: Higher Plant and Bacterial Models, pp 102– 121. American Society of Plant Physiologists, Rockville Ong LJ and Glazer AN (1991) Phycoerythrins of marine unicellular cyanobacteria: I. Bilin types and locations and energy transfer pathways in Synechococcus spp. phycoerythrins. J Biol Chem 266: 9515–9527 Padhy RN, Hottat FG, Coene MM and Hoet PP (1988) Restriction analysis and quantitative estimation of methylated bases of filamentous and unicellular cyanobacterial DNAs. J Bacteriol 170: 1934–1939 Paerl HW (1991) Ecophysiological and trophic implications of light-stimulated amino acid utilization in marine picoplankton. Appl Environ Microbiol 57: 473–479 Paerl HW, Priscu JC and Brawner DL (1989) Immunochemical local isation of nitrogenase in marine Trichodesmium aggregates: Relationship to fixation potential. Appl Environ Microbiol 55: 2965–2975 Palenik B and Haselkorn R (1992) Multiple evolutionary origins of prochlorophytes, the chlorophyll b–containing prokaryotes. Nature (London) 355: 265–267 Platt T and Li WKW (eds) (1986) Photosynthetic Picoplankton. Can Bull Fish Aquat Sci Vol 214 Rao NN and Torriani A (1990) Molecular aspects of phosphate transport in Escherichia coli. Mol Microbiol 4: 1083–1090 Reddy KJ, Bullerjahn GS, Sherman DM and Sherman LA (1988) Cloning, nucleotide sequence, and mutagenesis of a gene involved in iron-deficient growth of the cyanobacterium Synechococcus sp. strain PCC7942. J Bacteriol 170:4466– 4476 Reddy KJ, Haskell JB, Sherman DM and Sherman LA (1993) Unicellular,aerobicnitrogen-fixingcyanobacteria ofthegenus Cyanothece. J Bacteriol 175: 1284–1292 Reinhold L, Zvinan M and Kaplan A (1987) Inorganic carbon fluxes and photosynthesis in cyanobacteria – a quantitative model. In: Biggins J (ed) Progress in Photosynthesis Research, Vol IV, pp 289–296. Martinus Nijhoff, Dordrecht Rippka R, Deruelles J, Waterbury JB, Herdman M and Stanier RY (1979) Generic assignments, strain histories and properties of pure cultures of cyanobacteria. J Gen Microbiol 111: 1–61

47 Rueter JG (1988) Iron stimulation of photosynthesis and nitrogen fixation in Anabaena 7120 and Trichodesmium (Cyanophyceae). J Phycol 24: 249–254 Rueter JG, Ohki, K and Fujita Y (1990) The effect of iron on photosynthesis and nitrogen fixation in cultures of Trichodesmium (Cyanophyceae). J Phycol 26: 30–35 Scanlan DJ, Mann NH and Carr NG (1993) The response of the picoplanktonic marine cyanobacterium Synechococcus sp. WH7803 to phosphate starvation involves a protein homologous to the periplasmic phosphate-binding protein of Escherichia coli. Mol Microbiol 10: 181–191 Sode K, Tatara M, Takeyama H, Burgess JG and Matsunaga T (1992) Conjugative gene transfer in marine cyanobacteria: Synechococcus sp., Synechocystis sp. and Pseudanabaena sp. Appl Microbiol Biotechnol 37: 369–373 Sournia A (1970) Les cyanophycées dans le plancton marin. Ann Biol 9: 63–76 Stanier RY and Cohen-Bazire G (1977) Phototrophic prokaryotes: The cyanobacteria. Ann Rev Microbiol 31: 225–274 Stock JB, Ninfa AJ and Stock AM (1989) Protein phosphorylation and regulation of adaptive responses in bacteria. Microbiol Rev 53:450–490 Suttle CA and Chan AM (1993) Marine cyanophages infecting oceanic and coastal strains of Synechococcus: Abundance, morphology, cross-infectivity and growth characteristics. Mar Ecol Prog Ser 92: 99–109 Suttle CA, Chan AM, and Cottrell MT (1990) Infection of phytoplankton by viruses and reduction of primary productivity. Nature (London) 347: 467–469 Suttle CA, Chan AM, Feng C and Garza DR (1993) Cyanophages and sunlight: A paradox. In: Guerrero R and Pedrós-Alió (eds) Trends in Microbial Ecology, pp 303–307. Spanish Society for Microbiology, Barcelona Swanson RV, Ong LJ, Wilbanks SM and Glazer AN (1991) Phycoerythrins of marine unicellular cyanobacteria: II characterization of phycobiliproteins with unusually high phycourobilin content. J Biol Chem 266: 9528–9534 Thomsen HA (1986) A survey of the smallest eucaryotic organisms of the marine phytoplankton. In: Platt T and Li WKW (eds) Photosynthetic Picoplankton. Can Bull Fish Aquat Sci 214: 121–158 Urbach E, Robertson DL and Chisholm SW (1992) Multiple evolutionary origins of prochlorophytes within the cyanobacterial radiation. Nature (London) 355: 276–270 Van Baalen C (1962) Studies on the marine blue-green algae. Bot Mar 4: 129–139 van den Hondel CAMJJ, van Leen RW, van Arkel GA, Duy vesteyn M and de Waard A (1983) Sequence-specific nucleases from the cyanobacterium Fremyella diplosiphon, and a peculiar resistance of its chromosomal DNA towards cleavage by other restriction enzymes. FEMS Microbiol Lett 16: 7–12 van der Plas J (1989) Gene analysis in the cyanobacterium Synechococcus sp. PCC 7942. PhD Thesis, University of Utrecht Vaulot D and Partensky F (1992) Cell cycle distributions of prochlorophytes in the north western Mediterranean Sea. DeepSea Res 39: 727–742 Vega-Palas MA, Flores E and Herrero A (1992) NtcA, a global nitrogen regulator from the cyanobacterium Synechococcus that belongs to the Crp family of bacterial regulators. Mol Microbiol 6: 1853–1859

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Chapter 3 Prochlorophytes: The ‘Other’ Cyanobacteria? Hans C.P. Matthijs1, Georg W.M. van der Staay2 and Luuc R. Mur1 1

E. C. Slater Institute, BioCentrum Amsterdam, Laboratorium voor Microbiologie, Nieuwe Achtergracht 127, 1018 WS Amsterdam, The Netherlands; 2 MCB, University of Colorado, Box 347, Boulder, CO 80309-0347, USA

Summary I. Prochlorophytes and Chloroplast Ancestry A. Phylogenetic Considerations and an Inventory of Prochlorophyte-Type Organisms B. An Inventory of Experiments Relating Prochlorothrix hollandica to Either Cyanobacteria or Chloroplasts 1. Relationship to Cyanobacteria a. Morphology b. Photosynthetic Apparatus c. Respiration d. Carotenoids and Lipids 2. Relationship to Chloroplasts C. Studies on Conserved Genes and Derived Phylogenetic Conclusions D. Oxyphotobacteria, a Unifying Name for Cyanobacteria and Oxychlorobacteria (Prochlorophytes) II. Other Non-phylogenetically Directed Studies A. Separation of PS I and PS II Plus Adhering ChI a and b Antenna Complexes B. Ultrastructural Studies 1. Ultrastructure: Freeze-Fracture Faces 2. Ultrastructure: Image Analysis of PS I Particles C. Phosphorylation of Membrane-Bound Polypeptides D. Effects of Irradiant Light Intensity on Prochlorophytes III. Concluding Remarks References

49 50 50 51 51 51 51 53 53 53 54 55 55 56 57 57 59 60 61 62 62

Summary The interest in Prochlorophytes (Oxychlorobacteria) was originally boosted by the endosymbiont theory on chloroplast evolution. The first organism of this type: Prochloron didemni, was described as a procaryote performing oxygenic photosynthesis and containing both Chls a andb. The combination ofthose two pigments was until then only characteristic of Chlorophyte chloroplasts. The prochlorophytes accordingly were portrayed to be the potential endosymbionts from which (green) chloroplasts originated. Before Prochloron sp., the cyanobacteria were the only procaryotes known to perform oxygenic photosynthesis. Because of this qualifying property, the origin of chloroplasts was linked with endosymbiontic cyanobacteria. Following the discovery of more prochlorophyte species (e.g., Prochlorothrix hollandica, Prochlorococcus marinus) research efforts in various fields demonstrated that the predicted relationship between the prochlorophytes and green chloroplasts if any at all existed, was not very direct. Moreover, sequence analysis of conserved genes in general supported a closer relationship between prochlorophytes and cyanobacteria rather than chloroplasts. This has even led some to consider the prochlorophytes as another type of cyanobacteria. Given the identification of a number of distinctive differences in the organization of the photosynthetic apparatus in these two types of procaryotes

D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 49–64. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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performing oxygenic photosynthesis it would be appropriate to support and extend the suggestion of Gibbons and Murray (1978) and Florenzano et al. (1986) to institute two orders—i.e., Cyanobacteria and Oxychlorobacteria—within the class of the Oxyphotobacteria . The characterization of the photosynthetic apparatus of Prochloron sp. and Prochlorothrix hollandica, has clarified that the Chl a/b antenna is not related to the LHC II antennae in green chloroplasts. The properties of the prochlorophyte antenna will be related to the ultrastructural appearance ofthe thylakoid membranes and the regulation of light-harvesting. I. Prochlorophytes and Chloroplast Ancestry

A. Phylogenetic Considerations and an Inventory of Prochlorophyte-Type Organisms Interest in Prochlorophytes was originally boosted by the endosymbiont theory ofchloroplast evolution. The first organism of this type: Prochloron didemni, was described as a procaryote performing oxygenic photosynthesis and containing both Chls a and b (Lewin and Withers 1975; Lewin 1976). The combination of those two pigments was until then only characteristic of Chlorophyte chloroplasts. The prochlorophytes accordingly were portrayed to be the potential endosymbionts from which (green) chloroplasts originated (Lewin, 1981). Before the discovery of Prochloron sp. and the introduction of the new order of Prochlorales in the class of the photobacteria (Gibbons and Murray, 1978), the cyanobacteria were the only identified procaryotes performing oxygenic photosynthesis. Because of this qualifying property, the origin of chloroplasts was linked with endosymbiotic cyanobacteria. Some doubts remained, however, especially those concerning the ultrastructural consequences of the bulky phycobilisome antennae which are located on the cytoplasmic, stromal-facing surface of the thylakoid membranes in cyanobacteria; phycobilisomes would hamper thylakoid appression, an ultrastructural feature characteristically found in green chloroplasts. The partitioning of the thylakoid membranes into granal and stromal areas coincides with the lateral separation of PS I and II. In this respect, differences between organisms with phycobilisomes and ones with membrane-embedded, Abbreviations: Chl – chlorophyll; HPLC – high-pressure liquid chromatography; LiDS – lithium dodecylsulfate; LHC II – lightharvesting complex I I ; OEC – oxygen evolving complex; PAGE–polyacrylamide gel electrophoresis; PSI–Photosystem I; PS II – Photosystem II; RuBisCo – ribulose-1,5-bisphosphate carboxylase/oxygenase; SDS – sodium dodecylsulfate; TEM – transmission electron microscopy; Zwittergent 14, tetradecyl-N – N-dimethyl-1,3-ammonio-1-propanesulfonate

light-harvesting Chl a- and b-containing antennae may comprise more than just light harvesting properties (Barber, 1986, 1990). Evidently, this important difference between chloroplasts and cyanobacteria presents a dilemma which was potentially solved in 1975 by Lewin and Withers with the discovery of a green symbiont in marine didemnids: Prochloron didemni. This promised to become an important discovery–a missing link in the origin of chloroplasts. Establishment of the procaryotic nature of Prochloron sp., its performance of oxygenic photosynthesis, the presence of Chls a and b, and the ultrastructural appearance of the thylakoid membranes supported the exciting phylogenetic position of the organism (Lewin 1984). Prochloron sp. thus appeared to fill a gap in the evolution of chloroplasts. Renewed speculations on the progenitors of chloroplasts developed (Florenzano et al., 1986). Given the diversity of (photosynthetic) plastids, different origins for the various types of chloroplasts have been suggested to be possible (Raven, 1970; Douglas and Turner, 1991; Bryant 1992; see Chapter 5). The ‘color’ of the chloroplasts has been used to define the group name of the eucaryotes in which particular chloroplasts are present (e.g., Chlorophyta, Rhodophyta, Chromophyta). In this way, cyanobacteria have been indicated as potential ancestors of rhodophyte chloroplasts. The progenitors of green chloroplasts in these terms are likely to be found amongst the Prochlorales. The procaryotic progenitors of the Chromophyta presumably remain to be discovered. The limited availability of Prochloron sp. (in vitro culture attempts were not successful) and complications in the extraction of physiologically active Prochloron sp. cells from the host didemnids soon hampered experimental progress (Lewin and Cheng, 1989). It took eleven more years before a second prochlorophyte-type species was discovered during the course of a lake restoration program for which all phototrophic microorganisms present were separated

Chapter 3 Prochlorophytes (Oxychlorobacteria) into uni-algal cultures. One of these cultures was apple-green in color instead of the blue-green color typical for cyanobacteria. The same sorts of tests that were done earlier for Prochloron sp. confirmed this ‘Oscillatoria-type’ organism to be a prochlorophyte. In fluorescence microscopy, excitation of cyanobacteria at 625 nm usually gives bright orange-red fluorescence; however, with a prochlorophyte the fluorescence is faint. Although not always easy to judge when natural samples are used, the fluorescence microscopy method is quite straightforward for initial screening. Prochlorothrix hollandica is a free-living prochlorophyte that can be cultured in the laboratory (Burger-Wiersma, et al. 1986). Soon after, a third prochlorophyte, Prochlorococcus marinus, was found in the euphotic zone ofthe Atlantic ocean (Chisholm et al., 1988, 1992). The pigment composition of the latter organism is quite different from the first two ones; very interestingly, Prochlorococcus marinus does not contain Chl a proper but a similar pigment with a slightly different retention in reversed-phase HPLC, denoted as Chl (tentatively identified as divinyl-Chl a). Similarly Chl b is present in a slightly modified form, Chl and in addition Chl c has been found (Goericke and Repeta, 1992). Several new habitats of these marine organisms have recently been identified. Prochlorococcus marinus can also be grown in the laboratory (Chisholm et al. 1992), and it has been questioned whether these oxygenic procaryotes are actually prochlorophytes (Urbach et al. 1992). Several other new types of prochlorophytes have been discovered in fresh-water habitats, and these do contain ‘normal’ Chl a and b pigments (O. M. Skulberg and T. Burger-Wiersma, personal communication).

B. An Inventory of Experiments Relating Prochlorothrix hollandica to Either Cyanobacteria or Chloroplasts Prochlorophytes at first glance are likely to encompass properties of both cyanobacteria and green chloroplasts. The basic characteristics of prochlorophyte taxonomy, cytology and ecological physiology have already been reviewed elsewhere (Lewin and Cheng, 1989; Burger-Wiersma and Matthijs, 1990; Mur and Burger-Wiersma, 1992). The results of earlier studies and recent progress (here illustrated for Prochlorothrix hollandica), are summarized in Table 1. The gene sequence comparison data of Table 1 will be discussed below in Section I C.

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1. Relationship to Cyanobacteria a. Morphology Detailed data about the ultrastructural appearance of the cell wall and the type of the peptidoglycans present (of the their crosslinkage index and the thickness of the peptidoglycan layer clearly demonstrates a similarity between the cell walls of cyanobacteria proper and the ones of prochlorophytes. In addition, a lipopolysaccharide fraction that contained the O-methylatedsugar 3 -O-methyl-xylose (a chemical structure typically found in cyanobacteria and other phototrophic bacteria), was isolated from Prochlorothrix hollandica,. The usual appearance of RuBisCO in carboxysomes of cyanobacteria (see Chapter 15) was also noticed in Prochloron didemni as well as Prochlorothrix hollandica. Gas vesicles, as commonly found in other phototrophic microorganisms are present in Prochlorothrix hollandica. In all other respects the ultrastructural appearance of the prochlorophytes is typically procaryotic (SchulzBaldes and Lewin, 1976; Burger-Wiersma et al., 1986; Chisholm et al., 1988).

b. Photosynthetic Apparatus The thylakoid membranes are located in the border areas of the cytoplasm and are more or less parallel to the cell membrane. The thylakoid membranes appear to be locally appressed, and in this way the prochlorophytes deviate from cyanobacteria. This is due to the absence of phycobilisome structures and the presence of Chl a- and b-containing antennae. Regulation of light harvesting via the latter antennae involves (de)phosphorylation in cases of unbalanced excitation of the photosystems (see Section II C). At the carboxyl-terminus, the D1 polypeptide of green chloroplasts is seven amino acids shorter than its analog in cyanobacteria. Sequence analysis of the psbA gene of Prochlorothrix hollandica suggested that the D1 polypeptide in this organism mimics the chloroplast one rather than the cyanobacterial one (Morden and Golden 1989a). These authors later amended this result by critically reviewing the calculation procedure involved (1989b). The oxygenevolving complex of Prochlorothrix hollandica contains only the 33-kDa polypeptide; the 17- and 23-kDa components, that are present in eucaryotes, are lacking in Prochlorothrix hollandica just as in cyanobacteria (Van der Staay et al., 1992a; Mor et al.,

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1993). After fractionation of hydrophobic and hydrophilic membrane-associated proteins with Triton X-114, the 33-kDa protein was found in the hydrophobic detergent phase, in contrast to results obtained with Chlamydomonas reinhardtii. The more hydrophobic nature of the polypeptide is reflected in its tighter association with the thylakoid membrane. Washing with 0.8 M Tris at pH 8 or with 2 M NaCl, methods usually applied in the extraction of the oxygen-evolving complex, did not detach the oxygenevolving complex from Prochlorothrix hollandica (Mor et al., 1993). This was confirmed with PS II-

enriched membrane preparations (G.W.M. van der Staay, unpublished results). PS I structurally resembles its analog in cyanobacteria. An electron microscopy study of isolated PS I particles followed by image analysis revealed the typical trimeric PS I structure observed in cyanobacteria (van der Staay et al., 1993).

c. Respiration Chlororespiration in green chloroplasts proceeds via a NAD(P)H dehydrogenase and plastoquinol

Chapter 3 Prochlorophytes (Oxychlorobacteria) autooxidation (Peltier and Schmidt, 1991). Dark respiration in cyanobacteria involves a full respiratory chain, inclusive of a cytochrome terminal oxidase (see Chapter 13). Such a full respiratory chain was also found in Prochlorothrix hollandica. The terminal cytochrome c oxidase (of the cytochrome type) was largely present in the cytoplasmic membrane fraction (Peschek et al., 1989). Separation of the thylakoid and cell membranes can be achieved via the sucrose gradient separation techniques described for cyanobacteria (Omata and Murata, 1988).

d. Carotenoids and Lipids HPLC analysis of the bright-orange cell membranes showed mainly zeaxanthin and chlorophyll was nearly absent. The carotenoid complement is similar to that usually encountered in cyanobacteria, and the presence of a carotenoid-binding protein in the cell membrane points to a blue-light-shielding protective mechanism. Analysis of lipids indicated a strong similarity between Prochloron sp., Prochlorothrix hollandica and cyanobacteria (Volkman et al., 1988; Murata and Sato, 1983; Gombos and Murata, 1991). In addition to this, novel fatty acids were detected in Prochlorothrix hollandica: a hexadec-4-enoic acid (Volkmanetal., 1988) and 16:1 (4) and 16:2 (positions not determined; Gombos and Murata, 1991).

2. Relationship to Chloroplasts The prominent divergence between prochlorophytes and cyanobacteria with respect to the photosynthetic apparatus (see Table 1), must be interpreted with some care. The mere absence of phycobilisomes and the presence of Chl a- and b- containing antennae, may spontaneously give rise to a number of the listed differences (Walsby, 1986). The membrane-intrinsic Chl a and b antennae may facilitate thylakoidmembrane appression and their hydrophobic nature could contribute to the formation of so-called grana stacks. One interesting consequence of the differences in thylakoid membrane structure is that in chloroplasts grana stack formation gives rise to the spatial separation of PS II in the grana lamellae and PS I in the stroma lamellae (Anderson and Andersson, 1988). Stacking and lateral heterogeneity has been demonstrated for thylakoid membranes in Prochloron sp. (Giddings et al., 1980), Prochlorothrix hollandica (Miller et al., 1988; Golecki and Jürgens, 1989; Van der Staay, 1992a) and Prochlorococcus marinus

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(Chisholm et al., 1988). Considerations on phylogenetic relationship may also require some of these secondary consequences to be taken into account. It can be concluded from Table 1 that despite the presence of Chl b, Prochlorothrix hollandica does not appear to be particularly closely related to the chloroplasts from higher plants or green algae. On the contrary, available data aiming at a comparison of Chl a/b antennae from green chloroplasts and prochlorophytes, i.e. immunological analysis via immunoblotting with anti-LHC II immunoglobulins from various sources (Hiller and Larkum, 1985; Bullerjahn et al., 1987) gave clear evidence for substantial differences among the Chl a/b binding polypeptides. A comparison of the structure of the LHC complexes of spinach and Prochlorothrix hollandica via circular dichroism analysis showed that the characteristic trimeric association of LHC II monomers present in chloroplast LHC II is absent in the prochlorophyte (Matthijs et al., 1988). It may thus be concluded that, instead of having contributed a clear proto-chloroplast organism fitting within the framework of the endosymbiotic theory, the discovery of Prochlorothrix hollandica has, rather, broadened the general view on evolutionary events. Support for the idea that the prochlorophytes have evolved independently comes from the observation that the Chl a/b-binding antennae of Prochlorothrix hollandica and Prochloron sp. are immunologically related (Bullerjahn et al., 1990). The Chl a/b antenna polypeptide from Prochlorothrix hollandica very interestingly crossreacted with an antibody raised against a 34-kDa Chl a-containing protein complex from the cyanobacterium Synechocystis sp. (Bullerjahn et al., 1987). This may point to an evolutionary divergence of an ancestral Chl-protein complex. It is plausible to assume that in the evolution of prochlorophytes this Chl-protein complex has acquired the ability to incorporate Chl b. Interestingly, considerable differences in the Chl a to b ratio of the antennae from Prochlorothrix (between 2.5 and 4; Bullerjahn et al., 1987; Matthijs et al., 1988; Van der Staay et al., 1992a,b; Post et al., 1992) and Prochloron sp. (about 1.5; Hiller and Larkum, 1985) exist. This supports a theory of independent evolution of the capacity to bind Chl b. One gene of great interest to be searched for would be the one encoding an enzyme that is specifically involved in the final step of Chl b synthesis. Such an enzyme has to our knowledge not yet been identified. The ability to synthesize Chl b and to incorporate Chl b into an antenna complex

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apparently may have developed several times (Miller and Jacob, 1989; Palenik and Haselkorn, 1992; Urbach et al., 1992; however, see Bryant, 1992). The questions as to whether these developments are really completely independent or whether gene transfers by natural transformation are involved (Palenik and Haselkorn, 1992; Bryant, 1992), or whether all Chl bcontaining organisms in fact had a single ancestor (Bryant, 1992), remain open. For Prochlorococcus marinus, a particularly recent origin of the pigment phenotype has been suggested (Urbach et al., 1992). The branching of a prochlorophyte from the cyanobacterial line requires at least three steps: the capacity to synthesize Chl b, the incorporation of Chl b in the Chl-binding antennae and the loss of phycobilisomes. The order of these steps remains unknown and can only be a subject of speculations. Phycobilisome proteins are absent in prochlorophytes (Lewin et al., 1976; Burger-Wiersma et al., 1986). With gene probes (generously donated by J.Houmard, Institut Pasteur), no positive evidence could be obtained for the presence of remnant phycobilisome genes in a series of relatively low-stringency Southern blot hybridization experiments with restriction fragments of chromosomal DNA from Prochlorothrix hollandica (H. C. P. Matthijs, unpublished results). A phycobilisome-less cyanobacterium probably would obtain an evolutionary advantage by the development of a Chl a/b-antenna system. Alternatively, a cyanobacterium already containing a Chl a/b antenna might have lost its phycobilisomes which might no longer be required for efficient photosynthesis in some environmental niches. Organisms representative of intermediates in these possible evolutionary scenarios may still be present in nature. Finding these species may not be easy, (it took surprisingly long to discover Prochlorothrix hollandica or Prochlorococcus marinus), considering that these species are dominant in their habitat and lack phycobilisomes! These speculations reveal an interesting (and provoking) question: ‘Do all cyanobacteria really contain Chl a only, or do cyanobacteria with some Chl b exist?’ (Van der Staay, 1992b). With the usually applied spectrophotometric methods, there are two obvious reasons why trace amounts of Chl b, if present at all, would probably remain unnoticed given the dominant presence of Chl a. The reaction center of PS I contains some 80–100 molecules of Chl a only, and PS II contains about 47 Chl a. In addition the PS I to PS II ratio may be significantly

greaterthan 3:l (Burger-Wiersma and Post, 1989; H. C. P. Matthijs , unpublished results). Even if Chl b were present in an antenna complex such as the ironstress-induced one of about 34 kDa that is identified to reside near to PS II in cyanobacteria (Sherman et al., 1987; Dekker et al., 1988; and Riethman and Sherman, 1988; see Chapter 25), it would clearly be vastly outnumbered by Chl a. This particular polypeptide is of potential interest because of its immunological relatedness to the Chl a/b-binding polypeptides of Prochlorothrix hollandica (Bullerjahn et al., 1987). Some of the techniques needed to enable studies on the possible presence of some Chl b in cyanobacteria, i.e. preparative separation ofChlprotein complexes prior to HPLC and spectroscopic analysis, have been developed for the prochlorophytes and will be reviewed below in Section II A. When cyanobacterial extracts, including extracts from ironstressed cells, were analyzed for the presence of Chl b by an HPLC technique, only minimal amounts of material with a retention time equal to that of Chl b were detected (van der Staay et al., 1992a).

C. Studies on Conserved Genes and Derived Phylogenetic Conclusions. Another potentially powerful tool to assess phylogenetic relationships is sequence comparison of conserved genes (see Table 1). Studies of the phylogenetic positions of the three prochlorophytes Prochloron sp., Prochlorothrix hollandica and Prochlorococcus marinus through 16S RNA sequence analysis and subsequent phylogenetic tree formation with the distance matrix method (Turner et al., 1989; Urbach et al., 1992; also see Chapters 1 and 5) has excluded that green chloroplasts and these prochlorophytes are constituents of a monophyletic group. Sequence analysis of several genes of Prochlorothrix hollandica, such as the rpoC1 gene (Palenik and Haselkorn, 1992) and the rbcL and rbcS genes (Morden and Golden, 1991), as well as the organization of the psbB, psbH, petB and petD genes (Greer and Golden, 1991, 1992) all point to a placement in the cyanobacterial domain of robust phylogenetic trees. The same conclusion was arrived at from a study of the Prochloron didemni atpBE genes (Lockhart et al., 1992). One exception, from which a deviation of cyanobacteria and prochlorophytes lineages could be deduced is the analysis of the psbA genes from Prochlorothrix hollandica (Morden and Golden, 1989a). Although it has been

Chapter 3 Prochlorophytes (Oxychlorobacteria) questioned whether the data treatment gives unequivocal results (Morden and Golden, 1989b), the psbA-encoded D1 protein was suggested to have a carboxyl-terminal deletion of seven amino acids, which is common for D1 proteins in chloroplasts but not in cyanobacteria (Morden and Golden, 1989a). Details of these studies and the derived phylogenetic trees are discussed elsewhere in this volume (see Chapter 5). It can be deduced from those data that the prochlorophytes and the cyanobacteria clearly group together within a common radiation (Urbach et al., 1992, Bryant, 1992). Within this so-called cyanobacterial radiation the three prochlorophyte species are quite distant from one another. This predicts the existence of more prochlorophyte species–a factual reality given recent reports on the presence of new species in several new natural habitats (O. M. Skulberg and T. Burger-Wiersma, personal communication).

D. Oxyphotobacteria, a Unifying Name for Cyanobacteria and Oxychlorobacteria (Prochlorophytes) Although biochemical and physiological data generally support the evidence for a close relatedness between cyanobacteria and prochlorophytes, some very distinct differences between these organisms have been demonstrated as well (Lewin and Cheng, 1989; Burger-Wiersma and Matthijs, 1990). Murray (1989) in a treatise on ‘the Higher Taxa, A Place for Everything...?’ suggested class status for the name ‘ oxyphotobacteria’ in order to accommodate newly recognized lower taxonomic categories, i.e. the cyanobacteria and the Prochlorales, a suggestion which permits a distinction in the deep phylogenetic clefts nowadays established. This prompted BurgerWiersma et al. (1989) to introduce the name ‘oxychlorobacteria’ for the prochlorophytes. In addition, the use of the name ‘prochlorophytes’ should in this respect be regarded as outdated for the very same reasons by which usage of the name blue-green algae for cyanobacteria has become obsolete (BurgerWiersma et al., 1989). In conclusion, the question mark in the title of this chapter expresses the opinion that it should at least be critically discussed whether the prochlorophytes (oxychlorobacteria) are to be referred to as another type of cyanobacteria. At present, the authors of this chapter support Murray’s idea of instituting the name oxyphotobacteria (oxygen-evolving procaryotes) at class status and to incorporate both the cyanobacteria

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(the former blue-green algae) and the oxychlorobacteria (prochlorophytes) into this class. II. Other Non-phylogenetically Directed Studies The scope of this contribution on the prochlorophytes extends beyond mere phylogenetic relationships. (Eco)physiological properties of the prochlorophytes, as studied with the tools of biochemistry and biophysics, should also be described. Limitation to just these techniques mostly stems from the limited number of studies in other fields and by other methods. For example, the application of molecular biological techniques has been limited, and no DNA transfer system has yet become available for any of the prochlorophytes. A basic requirement, the availability of axenic strains, has been independently achieved for Prochlorothrix hollandica (R. A. Lewin and R. Rippka, personal communication). Both of these axenic strains originate from the same isolate of Prochlorothrix hollandica from lake Loosdrecht, the Netherlands (Burger-Wiersma et al., 1986). The growth rate of the axenic strain (Pasteur Culture Collection strain 9006) is somewhat lower than that of the original uni-algal but non-axenic isolate (18 versus 40 h, unpublished results). To start cultures from small inocula of resting cells can be a tedious job. Low light intensity mild aeration and room temperature usually render success. Growth of the axenic strain on plates is only possible on very soft agar (0.5 to 0.7%) (S. S. Golden, personal communication). Attempts to grow Prochlorothrix hollandica heterotrophically have failed thus far. Otherwise, growth of Prochlorothrix hollandica does not require anything other than the media and light normally needed for culture of cyanobacteria (BurgerWiersma and Matthijs, 1990; Mur and BurgerWiersma, 1992). The appearance of the culture depends very much on the light intensity. With an incident light intensity below cultures appear apple-green; higher light intensity produces yellowish, though active, cultures. The cells of Prochlorothrix hollandica can be broken easily with a French press. Photosynthetically active membranes can be obtained by the same centrifugation steps routinely used in studies with cyanobacteria (van der Staay, 1992a, b). Prominent questions posed about Prochloron sp. and Prochlorothrix hollandica concern the locali-

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zation and functional properties of Chl b. Questions about the (eco)physiological properties of these new organisms concern the regulation of light energy distribution and the acceptability of various light intensities for growth. Different experimental approaches have been used to resolve the function and localization of the Chl a- and b-containing antennae in Prochlorothrix hollandica. The procedures involved are: 1) separation of PS I and PS II plus adhering Chl a and b antenna complexes with a variety of techniques; 2) ultrastructural studies; 3) state-transition-type experiments in which the fluorescence emission from PS II at 680 nm or the relative phosphorylation of antenna polypeptides are monitored ; and 4) effects of changes in ambient culture light intensity and on the presence of Chl bbinding protein complexes.

A. Separation of PS I and PS II Plus Adhering Chl a and b Antenna Complexes Solubilization of isolated thylakoid membranes from Prochloron sp. and Prochlorothrix hollandica with different detergents (SDS, and Zwittergent 14) enabled separation of several Chl protein complexes on non-denaturing ‘green’ gels (Hiller and Larkum, 1985; Schuster et al., 1985; Bullerjahn et al., 1987; van der Staay, 1992; van der Staay et al., 1992a,b). In early experiments various Chl-protein complexes, obtained with SDS or dodecylmaltoside solubilization, were resolved by non-denaturing PAGE. Analyses of complexes by fluorescence excitation and emission spectroscopy at 77 K revealed a connection between PS I and a substantial part of the Chl b comprised in Chl a- and b-containing Chl-protein complexes (Schuster et al. 1985, Hiller and Larkum 1985, Bullerjahn et al. 1987). In these experiments non-denaturing electrophoresis was used as the sole separation technique. (Note: In general the membrane-associated polypeptides in Prochlorothrix hollandica seem to be more hydrophobic than their counterparts in chloroplasts or cyanobacteria. This might explain the somewhat poorer resolution on denaturing gels of Prochlorothrix hollandica thylakoids in comparison to those of spinach or Anabaena sp.). Recent studies have revealed that in thylakoid membranes of Prochlorothrix hollandica the rate of electrophoretic migration of the Chl a/b complexes is equal to that of the PS I reaction centers (van der Staay, 1992). This could give rise to co-

isolation of PS I and Chl a/b antennae. Additional separation techniques were used to further address this problem (van der Staay et al., 1992a). With sucrose-gradient centrifugation as the primary separation method and by testing a series of different detergents, solubilization of isolated thylakoid membranes from Prochlorothrix hollandica with Zwittergent 14 permitted the separation of PS I and PS II into two discrete bands. The Chl a/b-binding antennae were found in the same fraction as PS II. SDS-PAGE and immunoblot analyses with specific antibodies against PS I and PS II core complexes clearly demonstrated the separation of both photosystems. The compositions of the PS I and PS II-plus-antenna fractions were analyzed by nondenaturing and denaturing electrophoresis. On ‘green’ gels PS I separated into two bands and a faint band comprised of free pigment. Even by illumination with ultraviolet light, a sensitive method to detect green bands of PS II or antennae, fluorescence could only be seen in the PS I fraction from the free pigment. The PS II fraction was resolved into three bands and the free-pigment zone. UV illumination induced bright fluorescence of all bands. The polypeptide composition of the various green bands was analyzed by denaturing SDS-PAGE. The PS I fraction yielded a dimer at 60 kDa and smaller PS Iassociated apoproteins with masses of 16.5, 14.4, 11 and 9 kDa. In one of the two PS I bands the 16.5 kDa component is missing (van der Staay et al., 1992a). This polypeptide, which is apparently loosely attached to the PS I reaction center, is presumably the psaD gene product which has been demonstrated to be an extrinsic polypeptide in spinach (Tjus and Andersson, 1991) and for Synechococcus sp. strain PCC 6301 (Li et al., 1991). In the three green bands derived from the PS II-enriched fraction of the sucrose gradient, at least three polypeptides could be attributed to the Chl a/b-binding antenna: a major band at 28 to 30 kDa and two weaker bands at 32 and 34 kDa. The molecular masses of the latter two apoproteins are similar to those of the D1 and D2 polypeptides. A fourth band at 36 kDa may be an antenna apoprotein as well (van der Staay et al., 1992a). Post et al. (1992) reported 30, 33 and 34 kDa polypeptides to be part of Prochlorothrix hollandica Chl a/b complexes. The pigment composition of the green bands was analyzed by HPLC. PS I-derived bands contained Chl a and only trace amounts of Chl b and zeaxanthin were present. The PS II-derived bands all contained Chl a and b, as well as appreciable amounts

Chapter 3 Prochlorophytes (Oxychlorobacteria) of zeaxanthin and also some The band which was mostly enriched in antenna had a Chl a to b ratio of approximately 3.2. The absorbance spectra (Fig. 1) show the differences between the thylakoid membranes and the PS I and PS II fractions at room temperature. PS II showed Chl b and carotenoids (mostly zeaxanthin) as a peak at 465 nm and a shoulder at 495 nm. The Chl peak in the red was centered at 671 nm. The spectrum of the PS I band was red-shifted, and the peak in the red was centered at 680 nm. A shoulder at 468 nm and a peak at 497 nm indicated the presence of in the PS I band. Low-temperature fluorescence emission and excitation spectroscopy indicated that was functionally linked to the PS I reaction center (excitation at 470 and more so at 505 nm, proved effective for 715 nm emission). Moreover, by comparing fluorescence emission spectra after excitation at different wavelength values, a functional connection between Chl b and the reaction center of PS II was indicated (van der Staay et al., 1992a). Interpretation of the experimental data which gave rise to the different views on the functional localization of the Chl a/b complexes–i.e., association with either PS I (Hiller and Larkum, 1985; Schuster et al., 1985; Bullerjahn et al., 1987) or PS II (van der Staay 1992; van der Staay et al., 1992 a,b)–needs further consideration. The conclusions cannot exclude other possible explanations. For example, the fluorescence excitation spectroscopy study of PS I-enriched fractions at 77 K which yielded a peak around 470 nm and which has been related to absorption via Chl b present in an antenna near to PS I (Bullerjahn et al., 1987) may be the result of co-purification of the Chl a/b complexes and PS I or may stem from absorption at this wavelength (van der Staay et al., 1992a). Alternatively, the use of Zwittergent 14 may give rise to disconnection of Chl a/b complexes and the PS I reaction center. In general, risks are involved in the use of detergents: on the one hand disconnection of associations existing in vivo can occur and on the other hand the creation of new assemblies between originally unlinked complexes may occur in vitro. These considerations should also be taken into account when fractionation studies are used to arrive at conclusions concerning the assembly of functional complexes in vivo. A conclusion may be that other techniques are required to arrive at definite conclusions on the functional localization of the Chl a- and b-containing antennae in the vicinity of either PS I or PS II. To this

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end, separation was achieved without any detergent through mild mechanical fractionation of the thylakoid membranes, or with digitonin very much like in the techniques applied for preparation of grana and stroma lamellae from green chloroplast thylakoid membranes (van der Staay, 1992). In this way grana-type, oxygenevolving particles which predominantly contained PS II and Chl a and b antennae were prepared. The stroma-lamellae preparations contained PS I but were lacking Chl a and b antennae. These observations support a model in which the Chl a and b antennae are functional as LHC II-like complexes, which also agrees with observations on the ultrastructural appearance of the thylakoid membranes of Prochloron sp. and Prochlorothrix hollandica.

B. Ultrastructural Studies 1. Ultrastructure: Freeze-Fracture Faces In all three types of prochlorophytes described to date, the thylakoid membranes show some appression (Giddings and Staehelin, 1980; Burger-Wiersma et al., 1986; Chisholm et al., 1988, 1992; see Fig. 2). In chloroplasts, stacking is accompanied by lateral heterogeneity. PS II and its antenna complexes are localized in the appressed membranes (grana), while PS I and the coupling factor are located in the unstacked membranes (stroma thylakoids). In all organisms exhibiting lateral heterogeneity, four fracture faces can be distinguished (see Fig. 2 and

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Chapter 3 Prochlorophytes (Oxychlorobacteria) Table 2). Similarly, lateral heterogeneity has been reported for the prochlorophytes Prochloron sp. and Prochlorothrix hollandica. In chloroplasts, particles of different sizes on the four fracture faces have been identified as components of the photosynthetic apparatus (Staehelin, 1986; Simpson, 1990; Olive and Vallon, 1991). In Table 2 the particle sizes on the different fracture faces of the thylakoid membranes of various organisms obtained after freeze etching are given; the nomenclature is according to Staehelin (1986). The protoplasmic fracture faces reveal several complexes of similar size, which hampers identification of these particles. In chloroplasts, the complex of PS I and its antenna LHC I could be attributed to the large particles (>10 nm, see Table 2) in the fracture face (Staehelin, 1986). In cyanobacteria, two properties give rise to a different membrane appearance. Firstly, stacking is sterically constrained by the bulky phycobilisomes. Secondly, cyanobacteria do not have any specific PS I-associated antenna equivalent to LHC I (Golbeck 1987; Sherman et al., 1987; see Chapter 10). Therefore, only two types of fracture faces can be identified. Due to the absence of a LHC I-type complex in cyanobacteria, the fracture facecontains only particles which are smaller than 10 nm (Golecki and Drews, 1982; Staehelin, 1986; Mörschel and Schatz, 1987). Freeze-fracture electron microscopy thus offers a powerful tool to assess the positioning of the photosystems and adjoining Chl-protein complexes. In Prochlorothrix hollandica and Prochloron sp., the fracture face contains only particle sizes smaller than 10 nm as in cyanobacteria. A report describing larger-size particles in Prochlorothrix hollandica (Golecki and

59 Jürgens, 1989) is probably due to a variation in the method of size determination, because all dimensions are larger than in the other reports.

2. Ultrastructure: Image Analysis of PS I Particles Apart from the presence or absence of a LHC I antenna, another typical difference between chloroplast and cyanobacterial PS I has been used to characterize further the status of PS I in Prochlorothrix hollandica (van der Staay et al., 1993). In order to conserve any anticipated multicomplex structure, PS I particles from Prochlorothrix hollandica were isolated from thylakoid membranes on sucrose gradients in the presence of The particles were characterized by electron microscopy and image reconstruction techniques. The fastest migrating, major green band in the gradient contained trimeric PS I particles with features similar to PS I from cyanobacteria. Ellipsoidal particles, characteristically found for PS I centers with LHC I (E. Boekema, personal communication) were absent. This result supports the view that PS I is not structurally linked with a Chl a/b -binding antenna. The results indicate that a trimeric PS I aggregates are broadly distributed among oxyphotobacteria. In contrast to these results, Post et al. (1993) concluded from energy distribution studies that the Chl a/b antenna may associate with PS I in some cases (i.e., under conditions of PS II overexcitation).

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C. Phosphorylation of Membrane-Bound Polypeptides In chloroplasts from higher plants and green algae, phosphorylation of membrane-bound polypeptides has been shown to play an important regulatory role in photosynthesis (for a review, see Allen, 1992). In particular the PS II-associated antenna, LHC II, is rapidly phosphorylated under conditions that reduce the plastoquinone pool. The phosphorylated LHC II detaches from PS II and migrates into the stromal thylakoids. LHC II is steadily dephosphorylated by a phosphatase; this allows LHC II to return to its original position near PS II. In this way the size of the antenna near PS II varies through phosphorylationinduced changes. A condition in which the antenna size is decreased through phosphorylation is called state 2 (relative overexcitation of PS II). The opposite situation in which PS I activity is relatively too high for the present PS II capacity (state I), requires an increase in size of the antenna near PS II which is established through dephosphorylation of LHC II. These so called state transitions, are seen as responses to prevent the overexcitation of PS II and still maximize photosynthetic electron transport by maintaining a balanced energy distribution between PS II and PS I. State transitions can be detected by monitoring the fluorescence, since a decreased antenna size results in a decreased variable fluorescence emission from PS II. Fluorescence changes in PS I/PS II transitions have been shown to occur in Prochlorothrix hollandica (Burger-Wiersma and Post, 1989). In PS II light, maximal fluorescence decreases by nearly 50% in 5 to 10 minutes; adding PS I light enhances fluorescence by some 30% in a few minutes. All changes were blocked in the presence of DCMU. These fluorescence properties very much resemble those observed for green chloroplasts, in which a direct relationship between fluorescence decrease and LHC II phosphorylation has been proven (Allen, 1992). Phosphorylation as a regulatory mechanism of light energy distribution thus may also be applicable in prochlorophytes. In Prochloron sp. a polypeptide of about 34 kDa has been demonstrated to become phosphorylated in experiments in which isolated thylakoid membranes were incubated with ATP under various conditions (Schuster et al., 1984). The phosphorylated polypeptide was shown to comigrate with the antenna-containing green band on non-denaturing gels, which suggested that this

polypeptide represents a component of the Chl a/bcontaining antenna. Interestingly, in contrast to observations in chloroplasts, the phosphorylation appeared to be independent of the light intensity and to happen in darkness as well. This result, and the observation that fluorescence measurements on whole cells showed no appreciable changes in fluorescence when conditions which otherwise would have induced state transitions in chloroplasts were used, gave rise to the hypothesis that the antenna in Prochloron sp. is continuously phosphorylated. The Chl a/b antenna, with the 34-kDa polypeptide being permanently phosphorylated, wouldwith reference to the working mechanism of the higher plant model be continuously shifted nearer to the PS I reaction center. This apparent lack of a regulatory response was explained by the need of Prochloron sp. cells to adapt to the light conditions in their habitat at several meters below the water surface where relatively little PS I light is available (Schuster et al., 1984). Prochlorothrix hollandica was isolated from a shallow lake, in which continuous mixing of the water column occurs such that this organism is normally subjected to constant changes in light intensity. Therefore, one could imagine Prochlorothrix hollandica to use protein phosphorylation as a regulatory mechanism for balanced light-harvesting more so than Prochloron sp. Up to now, two studies on the phosphorylation of membrane-bound proteins in Prochlorothrix hollandica have been published. Using isolated membranes to which reductants or oxidants were added, van der Staay et al. (1989) reported major phosphorylated polypeptides with masses of 37, 29 and 26 kDa. The rates of phosphorylation of these polypeptides was different; phosphorylation of the 29- and 37-kDa polypeptides proceeded linearly for 20 minutes and continued for at least 1 hour. The phosphorylation increase of the 26-kDa protein lasted for a much shorter time. Mgion specificity of the kinase and redox-state dependence of phosphorylation activity were documented. Reducing conditions markedly favored the phosphorylation of the 29-kDa polypeptide and to a lesser extent of the 37-kDa one. As demonstrated for LHC II (Bennett, 1980), the phosphorylated amino acids could be removed with trypsin. Phosphorylation in the presence or absence of the phosphatase inhibitor NaF revealed a marked (two-fold faster) stimulatory effect on the rate of phosphorylation of the 29-kDa polypeptide when NaF was present. In contrast to this expected result, the rate of phosphorylation of the 37-

Chapter 3 Prochlorophytes (Oxychlorobacteria) kDa polypeptide was reduced in the presence of NaF. The required adjustability of phosphorylation through redox and phosphatase control favor the 29-kDa polypeptide for having a regulatory role in light harvesting. The phosphorylated polypeptides were recovered in the green bands of a non-denaturing polyacrylamide gel. Subsequent SDS-PAGE analysis of the green bands showed the 37-kDa polypeptide to be present in all five chlorophyll-protein complexes, whereas the 29-, 26- and minor 16-kDa bands were only found in the fastest migrating green complex. If one assumes a faster migration of the phosphorylated bands, the phosphorylated complex of 29-kDa can be matched with the antenna polypeptide of about 30kDa (van der Staay, 1992). In addition to the 34-kDa band, Schuster et al. (1984) also identified some phosphorylated proteins in the 27- to 30-kDa range that were part of the fastest-running complex on green gels. This complex was identified as a chlorophyll a/b-protein complex (Bullerjahn et al., 1987). The experiments of van der Staay et al. (1989) did not reveal a clear physiological role. Phosphorylation was strongest in darkness, but in the light the degree of phosphorylation did not appear to depend on the irradiant light intensity; moreover, using PS Iand PS II-specific light, state-dependent changes in phosphorylation could not be demonstrated. Post et al. (1992) used both intact cells and isolated membranes from Prochlorothrix hollandica and observed phosphorylated polypeptides with apparent masses of 35, 25, 23 and 14 kDa. It seems likely that the differences in apparent molecular masses in the two studies reflect differences in the electrophoretic conditions such that the 37- and 35-, the 29- and 25-, the 26- and 23 -, and 16- versus 14-kDa polypeptides respectively, are equivalent. The kinase was activated in vivo with PS II (650 nm) light but was inactive in PS I (710 nm) light. Using these light conditions, fluorescence changes in whole cells of Prochlorothrix hollandica could be related to state transitions (Post and Burger-Wiersma, 1989; Post et al., 1992). Light as well as the reductant duroquinol stimulated kinase activity in vitro (Post et al., 1992). In agreement with van der Staay et al. (1989), it was demonstrated that NaF stimulated phosphorylation of the 25- and 23kDa polypeptides, and gave slight inhibition for the 35-kDa one. In contrast to van der Staay et al (1989), Post et al. (1992) judged the 25- and 23-kDa polypeptides not to be present in a green band on nondenaturing gels. Otherwise, the 35-kDa band was present in all green bands. From the latter results and

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earlier observations that a substantial part of the Chl a/b antenna purifies in parallel with PS I (Bullerjahn et al., 1987), a working mechanism for light energy distribution in Prochlorothrix hollandica was derived in which a phosphorylated antenna moves away from PS I and thereby may enhance PS II activity (Post et al., 1992). These mostly biochemically derived data indicate a mechanism for state transitions to be present in Prochlorothrix hollandica which differs from the one in chloroplasts. In the latter the Chl a/b antenna moves away from PS II. The mechanism proposed by Post et al. (1992) does not explain the stimulating effects of 650 nm (PS II) light and reducing conditions on kinase activity. Before a final opinion on the precise localization of the Chl a/b antenna and on the role which phosphorylation plays in the mechanism of regulation of light energy harvesting can be given, satisfactory separation techniques of thylakoid membranes of prochlorophytes are needed (see Section II A). A mechanism for regulation of PS II light-harvesting via a mobile antenna near PS II needs further investigation, since Prochlorothrix hollandica, like cyanobacteria, contains a PS I to PS II ratio of three or more (depending on the light intensity; H. C. P. Matthijs, unpublished results). Furthermore, Prochlorothrix hollandica appears to be more sensitive to photoinhibition than the green alga Chlamydomonas reinhardtii (Mor et al., 1992a). Protection against photoinhibition would be possible via state transitions in the classical chloroplast way by increasing or decreasing the lateral separation of PS I and PS II. The fluorescence data presented by Burger-Wiersma and Post (1989) support such a regulatory system. The observations of enhanced phosphorylation under reducing conditions (van der Staay et al., 1989; Post et al., 1992), as well as their enhancement in PS II light, also favor a classic state transition mechanism in Prochlorothrix hollandica.

D. Effects of Irradiant Light Intensity on Prochlorophytes A decrease of the ratio of chlorophyll a to accessory pigments is usually observed upon transfer of phototrophs from high to low photon-flux densities due to the increased production of light-harvesting pigments relative to reaction centers. However, the prochlorophytes have properties which may generate different results. Firstly, the Chl a/b antennae in at least Prochlorothrix hollandica have been shown to be relatively poor in Chl b content with respect to Chl

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a (see Section II A). Furthermore, just as in cyanobacteria, low-light conditions favor synthesis of PS I reaction centers more than PS II (see Chapter 22). These different consequences of low-light adaptation may be the reason that different responses of the Chl a/b ratio in Prochloron sp. have been reported (for a review see Burger-Wiersma and Matthijs, 1990). More evidence for the specific behavior towards a shortage of light comes from the actual estimation of reaction center and antenna content. In Prochlorothrix hollandica low-light conditions favor relative induction of both Chl a/b antenna and PS I reaction centers (H. C. P. Matthijs, unpublished results). Under these conditions TEM micrographs display more thylakoid membranes per cell and more areas of appressed membranes. The Chl a to b ratio at is about 10; however, in cultures this ratio approaches 18. Non-denaturing gel electrophoresis of thylakoids from these differently grown cells showed that in high-light-grown cells, the complexes CP 2, CP 3 and CP 5 (nomenclature according to Bullerjahn et al., 1987) are markedly reduced. The high-lightgrown cells have a yellow appearance and after separation the cell membrane has a bright orange color. Photoacclimation of Prochlorococcus sp. strains also demonstrated adaptational differences, that were stated to result from either changes in the number of LHC complexes per PS II or from the Chl b-binding capacity of the apoproteins that constitute LHC (Partensky et al., 1993). The latter possibility is very interesting, and in fact the possible incorporation of either Chl a or b into the apoproteins of LHC is still not completely resolved for the various Chl-protein complexes in green chloroplasts. The prochlorophytes with a wide range of Chl a to b ratios in the putative LHC-like assemblies are of interest with regard to the question of Chl a or b incorporation. III. Concluding Remarks The collected research results in Table 1 clearly demonstrate that the prochlorophytes share many properties with the cyanobacteria. The main discriminating factor is presently linked with the mere presence of Chl b in these organisms. Although the amount of Chl b is small (between 5 and 15 Chl a molecules per Chl b) relative to Chl a, Chl b nevertheless exerts a major effect on the organization and possibly the regulation of the photosynthetic

apparatus. The precise assembly of thylakoid membranes with regard to the association of the Chl a/b antennae with either PS I or PS II awaits further detailed experiments. It would also be of great importance to clone and sequence the genes for the polypeptides of the Chl a/b complexes and to compare these to the sequences of the LHC polypeptides of green chloroplasts. Further biochemical and physiological studies may help to answer the ecophysiological questions on why the prochlorophytes are so successful in some of the habitats in which they were discovered. Discovery of a wider range of prochlorophyte type organisms may help to define further the position of the prochlorophytes with regard to the cyanobacteria. Instead of ‘prochlorophytes’ it is suggested that a more appropriate name for these organisms is the ‘oxychlorobacteria.’ References Allen JF (1992) Protein phosphorylation in regulation of photosynthesis. Biochim Biophys Acta 1098: 275–335 Anderson J M and Andersson B (1988) The dynamic photosynthetic membrane and regulation of solar energy conversion. Trends Biochem Sci 13: 351–355 Barber J (1986) New organism for elucidating the origin of higher plant chloroplasts. Trends Biochem Sci 11: 234 Barber J (1990) The fluid-mosaic nature of the thylakoid membrane. In: Baltscheffsky M (ed) Current Research in Photosynthesis, Vol. II, pp 715–724. Kluwer, Dordrecht Bennett J (1980) Chloroplast phosphoproteins. Evidence for a thylakoid-bound phosphoprotein phosphatase. Eur J Biochem. 104: 85-89 Berhow MA and McFadden BA (1983) Ribulose 1,5-bisphosphate carboxylase and phosphoribulokinase in Prochloron. Planta 158: 281-287 Bryant DA (1992) Puzzles of chloroplast ancestry. Curr Biol 2: 240-242 Bullerjahn GS, Matthijs HCP, Mur LR and Sherman LA (1987) Chl-protein complexes of the thylakoid membrane from Prochlorothrix hollandica, a prokaryote containing Chl b. Eur J Biochem 168: 295–300 Bullerjahn GS, Jensen TC, Sherman DM and Sherman LA (1990) Immunological characterization of the Prochlorothrix hollandica and Prochloron sp. Chl a/b antenna proteins. FEMS Microbiol Lett 67: 99–109 Burger-Wiersma T and Matthijs HCP (1990) The Biology of the Prochlorales. In: Codd GA, Dijkhuizen L and Tabita FR (eds) Advances in Autotrophic Microbiology and One-Carbon Metabolism, Vol l, pp 1-24. Kluwer, Dordrecht Burger-Wiersma T and Mur LR (1989) Genus ‘Prochlorothrix’ In: Staley JT(ed) Bergey’s Manual of Systematic Bacteriology, Vol 3, pp 1805-1806. Williams and Wilkins, Baltimore Burger-Wiersma T and Post AF (1989) Functional analysis of the photosynthetic apparatus of Prochlorothrix hollandica (Prochlorales), a Chl-b containing procaryote. Plant Physiol

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Chapter 4 Molecular Biology of Cyanelles Wolfgang Löffelhardt Institut für Biochemie und Molekulare Zellbiologie der Universität Wien und Ludwig-Boltzmann-Forschungsstelle für Biochemie, A-1030 Wien, Austria

Hans J. Bohnert Department of Biochemistry, Department of Molecular and Cellular Biology and Department of Plant Sciences, The University of Arizona, BioSciences West, Tucson, Arizona 85721, USA Summary I. Introduction II. Cyanelle Wall Biosynthesis and Structure III. Molecular Genetics A. Genome Structures of Plastids B. Genes of the Translation Apparatus I. rRNA Genes 2. tRNA Genes 3. Genes for Ribosomal Proteins C. Genes for Components of the Photosynthetic Apparatus 1. Photosystem I 2. Photosystem II Complex 3. Cytochrome 4. Other Components of the Electron Transport Chain 5. Phycobilisomes 6. ATP Synthase 7. Rubisco D. Novel Genes in Cyanelle DNA 1. NAD Biosynthesis 2. Isoprenoid Pathway 3. Other Genes E. Characteristics of Cyanelle Genes IV. Protein Transport A. Import of Proteins B. Routing within Cyanelles C. Protein Translocation Machinery V. Phylogenetic Analyses VI. Conclusions Acknowledgments References

D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 65–89. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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Summary In a number of systematically unrelated eucaryotes, plastid-like organelles, termed cyanelles, are found which resemble present day cyanobacteria in morphology, biochemical organization of their photosynthetic apparatus, and in the presence of a peptidoglycan wall. The existence of cyanelles in species of different systematic position indicates repeated invasions of heterotrophically living cells by cyanobacteria-like organisms in the evolutionary past. Only one of these cyanelle-bearing algae has been studied to some extent. Investigations into the genome and gene structure of the cyanelle of the unicellular eucaryotic alga Cyanophora paradoxa are reviewed. The cyanelle genome of approximately 130,000 bp includes more genes than the chromosomes of typical higher plant chloroplasts, mainly because only very few introns are found in cyanelle genes. Genes are tightly packed in operons that are similar in structure to bacterial operons. Up until now, with perhaps 85% of the cyanelle genome sequenced, genes that are typically found in chloroplasts are found in cyanelles as well. A remarkable exception is the apparent absence of ndh gene homologs. Other cyanelle genes, absent from chloroplasts, encode proteins functioning in isoprenoid biosynthesis, chlorophyll biosynthesis, other metabolic processes, and protein transport and protein folding. Generally, more of the genes encoding components of multi-protein complexes (ribosomes, photosystems, ATP synthase, etc.) are retained by cyanelles. In their gene complement cyanelles resemble chromophytic and rhodophytic algal plastids, while in their gene organization parallels exist to cyanobacteria as well as to higher plant plastids.

I. Introduction Among the photosynthetic eucaryotes, those that contain chloroplasts – the green algae, mosses, ferns, and higher plants – are but one group of plastidcontaining organisms. Different forms of nuclear and plastid genome organization can be found in a number of other algal groups. As more information about the genetic organization of these plastids in different phyla (e.g. chromophyta and rhodophyta) becomes known, better educated guesses can be made about plastid evolution, and one can begin to understand the many-faceted metabolic interactions and regulatory connections between the nucleocytoplasmatic and plastid compartments. Comparative molecular biology of plastids of different algae will greatly enhance our understanding of the evolution of photosynthesis. The view of plastid acquisition by an endosymbiotic event, involving an ancestral photosynthetic procaryote and a heterotrophic eucaryote, is now widely accepted (Margulis, 1981; Gray, 1989). Questions that are still to be answered are: (i) how often did such an endosymbiotic event occur; (ii) were different types of procaryotic ancestors involved; (iii) what was the Abbreviations: b – base; bp – basepair; FNR – ferredoxin oxidoreductase; IR – inverted repeat; LSC – large single-copy; LSU – large subunit; ORF – open reading frame; OEM – Zouter envelope membrane; PBP – penicillin-binding protein(s); PBS – phycobilisomes; rbs – ribosome binding site; SSC – small single-copy; SSU – small subunit

nature of the evolutionary pressure governing the reduction of the endosymbiont’s genome; and (iv) how did the control systems for the genetic and metabolic integration of the former endosymbiont into the cell evolve. This review will discuss a group of organisms that contain a special type of plastids, cyanelles, that have cyanobacterial-type pigment composition and phycobilisomes. What makes these plastids, and the organisms within which they are found, unique is that cyanelles, like cyanobacteria, are surrounded by a peptidoglycan wall. Additionally, cyanelles resemble cyanobacteria in overall morphology and in the presence of carboxysome-like structures. Such associations between a eucaryote and a cyanobacterium are sometimes called ‘endocyanomes’. Equally remarkable is that cyanelles have been detected in different eucaryotic cells which are obviously unrelated evolutionarily and systematically. For example, an amoeboid ‘host’, Paulinella chromatophora, containing cyanelles has been described (Kies, 1984, 1992), while other cyanelles were detected in host cells that may represent either red algal, diatom, dinoflagellate or cryptomonad forms of organization (Kies, 1992). Table 1 lists some cyanelle-bearing organisms. Most of these organisms, and a number of others about which we have only anecdotal knowledge, are included in one taxonomic group (Glaucocystophyceae) mostly by virtue of the peculiar plastid type with which they are endowed, supported by morphological characters

Chapter 4 Molecular Biology of Cyanelles

(Kies, 1992). Remarkable in a historical sense is that cyanelles were at one time considered cyanobacterial endosymbionts of eucaryotic hosts (Pascher, 1929), requiring classification by family and species names (Hall and Claus, 1963). This view became untenable once the limited genome size of the cyanelle found in Cyanophora paradoxa was recognized (Herdman and Stanier, 1977). It also appears reasonable, although it has never been proven, to assume a genetic interdependence for other ‘endocyanomes.’ One exception appears to be Geosiphon pyriforme, in which the associated cyanobacteria have been shown to enter the host cell but are also viable outside the host cell (Mollenhauer, 1992). That cyanelles are found in systematically divergent eucaryotes in itself can be taken as a strong argument in favor of multiple endosymbiotic events through which cyanobacterial invaders colonized heterotrophic cells resulting in their conversion into photosynthetically competent organisms. This topic has been summarized recently (Valentin et al., 1992b) and those authors indeed suggest several independent endosymbiotic transfection events based in part on the information that has been obtained from the study

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of cyanelles. There are, in fact, several arguments that have to be considered in discussing polyphyletic or monophyletic plastid origins and singular or multiple primary endosymbiotic events (see Chapter 5). At present phylogenetic analyses seem to rule out prochlorophytes as direct ancestors of the chloroplasts (chlorophyll b-lineage) which is a drawback of the polyphyletic model (Turner et al., 1989; Palenik and Haselkorn, 1992). There may be multiple events of endosymbiosis which left traces that are difficult to interpret within the different algal phyla which are much less well studied than the chlorophyte – green plant lineage. The case to make this point are cyanelles, although it is unfortunate that only cyanelles of Cyanophora paradoxa have been studied in some detail (Schenk, 1992). It could be that the cyanelles from the amoeba Paulinella chromatophora and those from Glaucocystis nostochinearum are of different origin and that these cyanelles and those from Cyanophora paradoxa represent separate endosymbiotic events. The underlying assumption that prompted our work on the Cyanophora paradoxa cyanelle was that similar plastid forms in eucaryotic cells of different

68 evolutionary history might represent paradigms for the study of plastid acquisition (including ancient and possibly even relatively recent events; Margulis, 1981; Gray, 1989). It would be preferable over continued debate on this topic, if one were able to generate sufficient comparative data, or better yet to generate experimental proof, to settle the controversy. It must also be envisioned that cyanelles may be only marginally successful remnants ofendosymbiotic events and may represent an evolutionary dead-end. The historical term cyanelle, while it may not be completely satisfactory, has over the last several years come to describe precisely a unique type of plastid structure, genome organization, andmetabolic interaction. Maintaining the term cyanelle seems desirable, since these organelles have preserved more cyanobacterial features than, for example, rhodoplasts. The type specimen is Cyanophora paradoxa (Korschikoff, 1924). Two isolates of Cyanophora sp. are known which differ in the organization of their cyanelle DNAs (Löffelhardt et al., 1983; Breiteneder et al., 1988; see below). Reflecting the difficulties insystematic, evolutionary andmolecular descriptions of cyanelle-bearing organisms, various other names for these organelles have been proposed to incorporate new views or recent findings. We suggest that the established name be maintained. It is to be expected that in-depth analyses of cyanelle genomes of the less well studied endocyanomes or plastid genomes of other algal classes may uncover even more exotic forms of plastid organization. Additional schemes would have to be established to distinguish an increasingly greater number of cyanelle types. This review will point out the importance of Cyanophora paradoxa cyanelles for our comprehension of plastid evolution. It appears that cyanelles, while definitely having originated from an endosymbiotic event, represent a different mode for the establishment of plastids than the events that led to extant chloroplasts as they are found in green algae, mosses, ferns and higher plants. Instead, cyanelles appear either to be related to the plastids from red algae, brown algae and cryptomonads, or to result from separate endosymbiotic events that nonetheless resulted in similar genome structure and gene content. In making this statement one has to realize that there are also striking parallels to chloroplast gene organization in some operons. There is no single criterion that can be employed to assess relationship.

Wolfgang Löffelhardt and Hans J. Bohnert It is premature to speculate more on this topic since plastid gene organizations of members of diverse algal phyla have not yet been studied as extensively as in cyanelles. This may change in the near future with data on the organization of a large portion of a rhodoplast genome forthcoming (Reith and Munholland, 1993). Present results certainly appear to support a common origin of all plastid types. This review focuses on recent results concerning cyanelle genome organization. We include a progress report on the sequencing of the Cyanophora paradoxa cyanelle genome that is underway (H. J. Bohnert and D. A. Bryant, unpublished). For additional information on cyanelle evolution and systematics, growth and division, physiology and biochemistry (including molecular biology), the reader should consult previously published reviews (Trench, 1979, 1982a, 1982b; Kies, 1979, 1984, 1992; Reisser, 1984; Coleman, 1985; Wasmann et al., 1987; Schenk, 1990, 1992; Bohnert and Löffelhardt, 1992).

II. Cyanelle Wall Biosynthesis and Structure Probably the most intriguing feature of cyanelles is that, although residing in the ‘host’ cytoplasm, they are surrounded by a cell wall. The presence of peptidoglycan has been demonstrated through biochemical analysis in two species which contain cyanelles, C. paradoxa (Aitken and Stanier, 1979) and Glaucocystis nostochinearum (Scott et al., 1984). All components of an murein, namely N-acetyl glucosamine, N-acetyl muramic acid, L-alanine, D-alanine, D-glutamate and m-diaminopimelic acid, have been detected in the proper ratios. This explains the lethal effect of antibiotics on these obligatorily autotrophic organisms (Berenguer et al., 1987). In addition, Kies (1988) reported that growth in the presence of penicillin resulted in the loss of C. paradoxa cyanelles to the medium, whereas the cyanelles from G. nostochinearum and Gloeochaete wittrockiana appeared to be degraded within the cells. Kies suggested that the cyanelle outer envelope membrane (OEM) might originate from a host cell vacuole that upon damage of the cyanelle might turn into a lysosome. Onlythe integrity of the cyanelle wall prevented digestion of the organelle. This, in turn, would have to be interpreted to mean that integration of cyanelles into the Cyanophora paradoxa cell represented a precarious and unstable equilibrium. However, such a view as a

Chapter 4 Molecular Biology of Cyanelles predator/invader relationship does not easily explain that the majority of cyanelle genes are located in the nuclear DNA, which must mean that the association persisted over long periods of time. An alternative view is that the cyanelle OEM is derived from the outer membrane of the ancestral cyanobacterial invader and that its observed instability (Giddings et al., 1983) might be due to the lack of lipoprotein (Höltje and Schwarz, 1985) connecting it to the murein layer. The necessary presence of receptors for the import of approximately 800 nucleus-encoded cyanelle polypeptides would be difficult to reconcile with a vacuolar nature of the cyanelle OEM. A final decision between these two possibilities must await analysis of the lipid and protein composition of isolated cyanelle OEM. In addition, understanding the routing of proteins into the cyanelle should provide important necessary data, probably more important than sequence comparisons, on the evolution of Cyanophora paradoxa cyanelles. Berenguer et al. (1987) demonstrated the presence of seven penicillin-binding proteins (PBPs), ranging in size from 110 to 30 kDa, in the cyanelle envelope. Their distribution and binding characteristics to various antibiotics suggest that they should be similar to the well characterized PBPs from E.coli (Höltje and Schwarz, 1985), although immunological relatedness might be low. Heterologous hybridizations of PBP gene probes from E. coli to cyanelle DNA proved unsuccessful which might mean that the majority of the PBPs might be encoded in the nuclear genome. Several cDNA clones isolated with the aid of antisera directed against affinity-enriched cyanelle PBPs have been obtained and are being analyzed at present (M. Kraus, personal communication). Evidence for the biosynthesis of the soluble precursor UDP-N-acetylmuramyl pentapeptide in the cyanelle stroma through a pathway analogous to that in E. coli was obtained recently (Plaimauer et al., 1991). Results obtained in this study suggested that the enzymes acting on already polymerized peptidoglyan (such as DD- and LD-carboxypeptidase and endopeptidase) are located in the cyanelle periplasm. This necessitates the existence of a carrier lipid, undecaprenyl phosphate, in the cyanelle inner envelope membrane for the translocation of the soluble precursor to the periplasm, thus creating the membrane-bound activated substrate for the PBPs (Höltje and Schwarz, 1985). Considering the structure of a peptidoglycancontaining cell envelope, results from E. coli showed

69 that the peptidoglycan in the periplasmic space is sandwiched between an outer and an inner membrane, and that the PBPs reside at the outer surface of the inner membrane (Park, 1987). An intriguing question is how the targeting of the PBPs and the other periplasmic enzymes involved in cyanelle wall metabolism is achieved. Are they transported directly across the OEM, like mitochondrial cytochrome c, assuming that the corresponding genes are nuclear? Or, is import of cytoplasmic precursors into the cyanelle stroma followed by re-export across the IEM as in the case of mitochondrial cytochromes and (Segui-Real et al., 1992)? Or are some enzymes exported from the cyanelle, meaning that part of the corresponding genes are cyanelle-located? Considerable progress has been made in understanding the structure of the cyanelle wall. The 16 major muropeptides obtained after muramidase cleavage of purified peptidoglycan (Höltje and Schwarz, 1985) were isolated by preparative HPLC and subjected to amino acid analysis and molecular weight determination through plasma-desorption mass spectrometry (Pfanzagl et al., 1993). Five cyanelle muropeptides proved to be identical to monomers, dimers and a trimer known from E. coli, whereas 11 muropeptides where shown to be derived from the respective E. coli counterparts through substitution(s) at the D-glutamate residue(s). Very recently the substituent that forms an amide bond with the group of D-glutamate leading to a molecular weight increment of 112 was identified as N-acetyl putrescine (Pittenauer et al., 1993). Such a substitution is rare among procaryotic cell walls. A preliminary search for N-acetyl putrescine in the peptidoglycan from the cyanobacterium Synechococcus sp. was negative (U. J. Jürgens, personal communication). Unfortunately, data on the fine structure of cyanobacterial peptidoglycans are missing. A comparison of cell wall architecture in this diverse group with that of the unique organelle wall of C. paradoxa is certainly needed.

III. Molecular Genetics The focus on cyanelle gene organization, which will be reviewed below, should eventually be supplemented by including studies on the genetic system of the ‘host’. The two commonly studied strains of Cyanophora paradoxa can only be kept in liquid culture. This fact and the strictly photoautotrophic

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nature of C. paradoxa prevented establishment of a genetic system up to now. No information has yet become available about the genome size of C. paradoxa. Using fluorescence-assisted sorting of mithramycin- or DAPI-stained nuclei (DeRocher et al., 1990), the DNA contents per nucleus (genome sizes) of the two C. paradoxa strains have been measured. Both have indistinguishable DNA contents, equivalent to 0.27 ± 0.04 pg DNA per nucleus (K. Harkins and H. J. Bohnert, unpublished). This value might be the C or 2C, or even a higher C-value, since nothing is known about this aspect of the life cycle of Cyanophora paradoxa. To put this DNA amount into perspective: a C-value of approximately 0.2 pg per nucleus would be nearly identical to the corresponding value for Arabidopsis thaliana.

A. Genome Structures of Plastids A recent review on the structure of chloroplast DNAs is available (Sugiura, 1992). All our comparisons referring to chloroplast gene structure are based on this compilation of the chlorophyll-b plastid lineage. Land plant chloroplast DNAs are similar in structure and gene content. The reduction of the organelle genome by transfer of genes from plastid DNA to nuclear chromosomal DNA has been demonstrated to occur during evolution (Baldauf and Palmer, 1990). Knowledge about algal chloroplast DNAs is less developed. Apart from Euglena gracilis (Christopher and Hallick, 1989) and Chlamydomonas reinhardtii (Harris, 1989), scant information is available. Plastid DNAs from red algae, brown algae and cryptomonads that are being studied by several groups (Valentin et al., 1992a; Douglas, 1992b) indicate that the general structure is not significantly different in these phyla. A detailed map of the rhodoplast genome of the alga Porphyra purpurea has recently been published (Reith and Munholland, 1993). In the brown alga Pylaiella littoralis (Loiseaux-de Goër et al., 1988) two circular chromosomes have been obtained from the plastids. Differences in gene complement, relative to the chlorophyll-b lineage, are found in all these algal phyla. They will be discussed in the context of cyanelle gene organization and complement. Cyanelle DNA from C. paradoxa is of roughly the same size as chloroplast DNA, approximately 130 and 140 kbp, respectively, in the two strains known (Löffelhardt et al., 1983). The chromosome contains

Wolfgang Löffelhardt and Hans J. Bohnert an inverted repeat (IR) structure, as is present in many other plastid DNAs. The cyanelle IR is approximately 10.5 kbp in length. The two segments of this repeat separate the small single-copy region (approximately 17 kbp) from the large single-copy DNA region (approximately 90 and 100 kbp, respectively, for the two strains). The strain referred to in the vast majority of molecular genetic studies described here is the strain originally isolated by Pringsheim and deposited in the algal culture collections worldwide (LB 555 UTEX; strain 2980 of the Algal Culture Collection, Göttingen). The second strain (no. 1555) was isolated by Kies and is maintained both in his collection and by us. In spite of high overall sequence homology, the restriction pattern of cyanelle DNA from the Kies strain is different from that of the Pringsheim strain. However, 16 protein genes mapped to analogous positions on the cyanelle genome of both strains. Thus, differences in restriction fragment patterns were ascribed to numerous small insertions/deletions and, perhaps, inversions in intergenic regions (Breiteneder et al., 1988).

B. Genes of the Translation Apparatus I. rRNA Genes Cyanelle DNAs from both strains known contain the rRNA genes within the 10.5 kbp IR (Breiteneder et al., 1988). Each repeat includes one operon of rRNA genes which are arranged in the following order: 5'16S rRNA - trnI - trnA - 23S rRNA - 5S -3', as in chloroplasts (Sugiura, 1992). The rDNA unit occupies approximately half of the IR, 16S rDNA being located at the border facing the SSC region, a feature which is only paralleled in the IR of Chlamydomonas reinhardtii. Contrary to the situation in the genomes of higher plant chloroplasts, the tRNA genes in the cyanelle operon lack introns (Janssen et al., 1987). The rRNA genes are transcribed as one large primary transcript. We have sequenced the central part of this region mainly to learn about the 3 '-terminal end of the 16S-rRNA, that has a functional role in the recognition of start-sites for translation (Bonham-Smith and Bourque, 1989). The 3'-end of the cyanelle 16SrRN A shows a sequence complementary to ribosome binding sites (rbs) in bacterial mRNAs. Since sequences complementary to such ‘rbs’, which may base-pair with the 16S-rRNA, are found at the 5'-end

Chapter 4 Molecular Biology of Cyanelles of most cyanelle transcripts close to either ATG (methionine) or, in a few instances, to GTG (valine) start codons (see below), it is safe to assume that these sites are actually used, although a final proof has not been provided. In fact, cyanelle ‘rbs’ function as binding sites in E. coli (Michalowski et al., 1991a; R. Flachmann, personal communication). About 1 kbp of the 16S rRNA gene has been sequenced by Giovannoni et al. (1988) who, upon comparison with the corresponding sequences from other organisms, came to the conclusion that cyanelles are equidistant from extant cyanobacteria and chloroplasts. Analogous results were obtained when the 5'-terminal 512 bp of the cyanelle 23 S gene were compared to the corresponding sequences from cyanobacteria and plastids (Janssen et al., 1987). As was also observed for cyanobacteria, the cyanelle 23S rRNA shows ‘hidden nicking’ (Trench, 1982a) which causes the appearance of a 18S and a 14S RNA species on denaturing gels. These nicks appear to represent in vivo events (Marsh, 1979). 5S sequences were obtained at the RNA level and revealed distinct cyanobacterial signatures in secondary structure (Maxwell et al., 1986).

2. tRNA Genes The 4S RNA fraction obtained from isolated, sucrosegradient-purified cyanelles has been resolved by two-dimensional polyacrylamide gel electrophoresis yielding about 40 RNA species. Of these, 29 RNAs were identified as cyanelle tRNAs by aminoacylation using E. coli aminoacyl-tRNA synthetases (Kuntz et al., 1984). Single tRNA species were found for seven amino acids, two isoacceptors for six amino acids, three isoacceptors for two amino acids, and four isoacceptors for leucine. tRNAs for cysteine, glutamic acid, glutamine and glycine could not be identified by this method. Using the individual tRNAs as well as heterologous chloroplast probes in hybridizations to cyanelle DNA, the tRNA gene complement (26 genes) in cyanelles has been measured. Twenty-one tRNAs mapped to the LSC region, three to the SSC region and two to the IR (Kuntz et al., 1984). Since then, 19 tRNA genes have been characterized by sequencing, and three of these are located in the IR. The duplicated genes and were the first to be analyzed during sequencing of the rDNA spacer (Janssen et al., 1987). These tRNAs can be folded into the usual clover leaf structure and

71 require post-transcriptional addition of the 3'terminal-CCA-OH. The small size of the rDNA spacer (287 bp) and the very short distance (3 bp) between the spacer tRNAs are in excellent agreement with data obtained for plastids from chromophytes (Markowicz et al., 1988; Delaney and Cattolico, 1989), rhodophytes (Maid and Zetsche, 1991) and Cryptomonas sp. (Douglas and Durnford, 1990). An additional tRNA gene, trnC, which was not amenable to the previous detection technique, was identified in the IR (D. A. Bryant, personal communication). Two out of the three tRNA genes predicted from hybridization experiments for the SSC region were recently sequenced: and They are clustered upstream of psaC , but are not cotranscribed with this gene (Rhiel et al., 1992). In the LSC region 10 of the mapped loci revealed tRNA genes upon sequencing. In addition eight new genes were identified, among these the missing genes for glutamic acid and glycine. Thus, one can expect that, upon completion of the cyanelle genome sequence, a set of at least 32 genes, as required according to the Wobble Rule, will be found. Evrard et al. (1988) observed in the vicinity of the IR, adjacent to a gene for a leucine tRNA, with the anticodon UAA, that is split by a 232 bp class I intron, the only intron so far found in any cyanelle gene. Thus far it has not been possible to demonstrate a self-splicing capacity for this intron. It is interesting that this leucine tRNA also contains an intron in some cyanobacteria (Xu et al., 1990; Kuhsel et al., 1990). The presence of a cyanobacterial intron in this tRNA gene is compatible with the interpretation that at least some introns represent old features predating the procaryote-eucaryote transition. In the central part of the LSC region and were localized between petFI and rps18/rpl33 (Kuntz et al., 1988). They give rise to separate transcripts since ORF65 separating these tRNAs yields only a 400 bases RNA (Evrard et al., 1990c). Michalowski et al. (1991b) identified a gene yielding a transcript of approximately 100 bases between the genes for a putative prenyltransferase and nadA (Fig. 1). Recently, rps4 was found to be surrounded by four tRNA genes: trnG, trnM, trnT, and trnR (C. B. Michalowski and H. J. Bohnert, unpublished results). The list of sequenced genes is completed by trnP, another trnR, another trnL, and trnW (V. L. Stirewalt, J. Farley, M. B. Annarella and D. A. Bryant, personal communication).

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Wolfgang Löffelhardt and Hans J. Bohnert

Chapter 4 Molecular Biology of Cyanelles

3. Genes for Ribosomal Proteins During the initial work on cyanelle gene organization all groups utilized gene probes from identified chloroplast genes, resulting in the detection of genes that were common to both groups (for a review, see Wasmann et al., 1987). Using such probes from the higher plant chloroplast large ribosomal protein gene cluster (Zhou et al., 1989; Markmann-Mulisch and Subramanian, 1988), the presence of several genes was indicated in one region of the cyanelle chromosome (Löffelhardt et al., 1990). Evrard et al. (1990a,b) and Michalowski et al. (1990b) subsequently determined the structure of a portion of the cyanelle genome that contained a larger number of genes compared to higher plant plastids which are equivalent to the eubacterial S10 and spc operons (Lindahl and Zengel, 1986). In the case of one of these genes, rpl3, the product of this novel plastid gene could be detected in cyanelles and in transgenic E. coli through the use of heterologous antibodies directed against ribosomal protein L3 from E. coli (Evrard et al., 1990b). Some of these novel cyanelle genes have also been reported by Bryant and Stirewalt (1990). In cyanelles, the S10 and spc operons are fused and give rise to a primary transcript approximately 7,500 nucleotides in length which is successively cleaved into smaller transcripts, although the primary transcript is relatively stable. The str operon is located close to this fused S10-spc operon, being separated by approximately 2.7 kbp (Kraus et al., 1990). It is interesting that the gene rps10 (located at the 5'-terminus of the bacterial S10 operon) is separated from the cyanelle S10-spc operon (Fig. 1). It has been translocated to the 3'-end of the str operon and appears to be co-transcribed with tufA (Bryant et al., 1991;Neumann-Spallart et al., 1991). Formally, the location of the genes petFI, two tRNA genes, and the rpl33 and rps18 genes, which form a transcription unit as on higher plant plastid DNAs (Evrard et al., 1990a,b), may be considered as the result of a transposition event that separated rps10 from the rest of the genes of the S10-spc operon in cyanelles (Fig. 1). The genes rpl23 (a pseudo-gene in spinach plastid DNA; Thomas et al., 1988), infA (a pseudo-gene in tobacco plastid DNA; Sugiura, 1992), rpl36, and two genes derived from the procaryotic operon (rps11, rpoA; Lindahl and Zengel, 1986) that are found in the large ribosomal protein gene cluster of higher plant plastid DNAs are absent from the cyanelle gene cluster on DNA fragment BglII-5 (Fig. 1). Results from heterologous hybridizations

73 (Löffelhardt et al., 1991) and sequence analyses indicate that some of these genes reside on a different locus on cyanelle DNA, in the order 5'-rpl36-rps13rps11-rpoA-rps9-3' (V. L. Stirewalt and D. A. Bryant, personal communication). Of the additional genes, rps13 occupies the equivalent position as in the E. coli whereas rps9, also a nuclear gene in higher plants, forms a bicistronic operon with rpl13 in E. coli (Lindahl and Zengel, 1986). As in chloroplasts rps4, a component of the in E. coli but not in Bacillus subtilis (Boylan et al., 1989), occupies a solitary position in a different part of the cyanelle genome (C. B. Michalowski and H. J. Bohnert, unpublished). The same applies for rps14 (V. L. Stirewalt and D. A. Bryant, personal communication) which in E. coli is located in the spc operon. The cyanelle ribosomal protein gene complement includes, in addition, the clustered genes rpl20 and rpl35 (Bryant and Stirewalt, 1990) and the genes rpl1, rps2, rpl19 and rpl21 that are not linked (V. L. Stirewalt, M. B. Annarella and D. A. Bryant, personal communication). Table 2 lists the 12 genes that are absent from higher plant plastid genomes as well as the four genes that have been identified on higher plant plastomes but not yet on cyanelle DNA. Compared to higher plant chloroplasts, that encode 21 ribosomal proteins, cyanelles have retained a larger number of ribosomal protein genes – 30 have been identified thus far (Fig. 1) – within their genome. Additional genes might still be detected, with rpl24 being a candidate based on hybridization results (Löffelhardt et al., 1991). Together these genes increase the cyanelle ribosomal protein gene complement to more than 50% of an estimated total of 60 genes. Algal plastid genomes also might surpass the number of ribosomal protein genes present in higher plant chloroplasts as indicated by the additional gene rpl5 on the plastome of Euglena gracilis (Christopher and Hallick, 1989). This situation could be even more pronounced in chlorophyll b-less algae. The str operon of Cryptomonas sp. also contains rps10 at the 3'-end and in addition rpoA, rpl13 and rps9 upstream of rps12 (Douglas, 1991). In this case the bicistronic E. coli operon rpl13-rps9 has been fused together with an gene into the str-operon. Gene rpl13 has not been reported before for any plastid genome. A high number of novel ribosomal protein genes has recently been detected on the genome of the red alga Porphyra purpurea (Reith and Munholland, 1993). A portion of the large cluster of genes ranging from rpl2 to rpl6 has been sequenced.

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It contained all the genes found in cyanelles and, in addition, the genes rpl29 (E. coli S10 operon) and rpl24 (E. coli spc operon). Also, genes rps 1 and rps6, which had not been detected on plastid genomes previously, are present on the rhodoplast genome. With the exception of a partial sequence comprising two genes (rpl15 and sec Y, Nakai et al., 1992) data are missing concerning the organization of cyanobacterial S10, spc and that might deviate from what is found in E. coli. Notably, most cyanelle ribosomal protein genes give better identity scores upon comparison with B. subtilis or B. stearothermophilus than with E. coli (Michalowski et al., 1990b). The cyanobacterial origin of cyanelles is clearly shown in the comparison of the str operon sequences from plastids and cyanobacteria (Kraus et al., 1990). Elongation factor Tu is cyanelle-encoded as generally observed in all groups of algae with the exception of some charophytes (Baldauf and Palmer, 1990), whereas EF-G is the product of a nuclear gene (Löffelhardt et al, 1990).

C. Genes for Components of the Photosynthetic Apparatus Maps for cyanelle genes for photosynthetic functions based on heterologous hybridizations have been published (Bohnert et al., 1985; Lambert et al., 1985). All chloroplast gene probes used yielded positive results. In many cases the corresponding genes have been sequenced and some novel genes that are nucleus-encoded in higher plants have been identified.

Wolfgang Löffelhardt and Hans J. Bohnert

1. Photosystem I Higher plants contain approximately 17 subunits in the Photosystem I-light-harvesting complex I, five of which are chloroplast-encoded (Sugiura, 1992). Homologs to all chloroplast psa genes were detected on the cyanelle genome. The psaA and psaB genes encoding the reaction center heterodimer have been sequenced (V. L. Stirewalt and D. A. Bryant, personal communication) and give rise to a 6-kb mRNA (Löffelhardt et al., 1987). The psaC gene, encoding the 9 kDa protein harboring the two [4Fe-4S] centers and was recently identified in the SSC region of cyanelle DNA (Rhiel et al., 1992). Identity scores for this highly conserved protein are in the range of 90–95% compared to plastid and cyanobacterial counterparts. A gene with homology to psaF, specifying the plastocyanin-docking protein that is nucleus-encoded in higher plants and located on the lumenal side of the thylakoid membrane, was detected recently (V. L. Stirewalt and D. A. Bryant, personal communication). In cyanelles, the gene product of psaF is presumably synthesized as a pre-protein containing a leader sequence in analogy to its counterpart in the cyanobacterium, Synechocystis sp. strain PCC 6803 (Chitnis et al., 1991). Since there is no plastocyanin present in cyanelles, the PsaF protein must interact with cytochrome Sequence information was also obtained on the genes psaI and psaJ, the latter being immediately adjacent to psaF (V. L. Stirewalt and D. A. Bryant, personal communication). The psaI gene product was recently shown to be contained within the Photosystem I complex of Anabaena variabilis ATCC 29413 with 45% sequence

Chapter 4 Molecular Biology of Cyanelles identity relative to the cyanelle counterpart (Ikeuchi et al., 1991). Yet another gene that is not encoded in the tobacco or rice chloroplast genomes, but that is encoded in the plastid genome of Marchantia polymorpha, is the cyanelle gene psaM (V. L. Stirewalt and D. A. Bryant, personal communication).

2. Photosystem II Of the 12 known chloroplast genes for subunits of Photosystem II, 10 have been detected thus far on the cyanelle genome. Janssen et al. (1989) reported the sequence of the cyanelle psbA gene encoding the D1 protein of the Photosystem II reaction center. The cyanelle D1 protein shows higher overall homology with chloroplast than with cyanobacterial counterparts. However, in contrast to plastids from higher plants and chlorophyll b -containing algae, the cyanelle gene contains a specific region at its carboxy-terminal end, an insertion of seven codons, that is also found in cyanobacteria and in the plastids from chlorophyll b-less algae such as the Rhodophyceae (Maid and Zetsche, 1990), Phaeophyceae (Winhauer et al., 1991), and Xanthophyceae (Scherer et al., 1991). The genes psbB, encoding the larger PS II antenna polypeptide CP47, and psbN, are clustered together with an interspersed small ORF as found in chloroplasts. However, psbH, specifying the 10 kDa phosphoprotein, known to be adjacent to psbN in chloroplasts and cyanobacteria (Mayes and Barber, 1991) has been shifted to a position upstream of psbB on the cyanelle genome (V. L. Stirewalt and D. A. Bryant, personal communication). Likewise, the linkage observed in chloroplasts between psbB and petB/D is not found in cyanelles. The transcription unit psbD-psbC, that encodes the second antenna polypeptide CP43 and the reaction center protein D2 and that is found in both chloroplasts and cyanobacteria, is conserved in cyanelles (V. L. Stirewalt and D. A. Bryant, personal communication) giving rise to a 3,200-b mRNA (Löffelhardt et al., 1987). A short distance downstream and transcribed from the opposite strand, psbK was identified (Stirewalt and Bryant, 1989a); the predicted protein is highly similar in sequence to its homolog from Synechococcus sp. strain PCC 6301 (Fukuda et al., 1989). Cytochrome is encoded in chloroplasts and cyanobacteria by the twin genes psbE-psbF. In cyanelles this cluster contains in addition the genes psbL and psbJ, that are assumed to specify smaller intrinsic membrane proteins of PS II (Cantrell and

75 Bryant, 1988). This arrangement is similar to that in cyanobacteria, Euglena gracilis plastids (Cushman et al., 1988), and higher plants (Sugiura, 1992).

3. Cytochrome

Complex

As in chloroplasts, four components are encoded by the cyanelle genome and the Rieske-iron-sulfurprotein appears to be the product of a nuclear gene (J. Jakowitsch, personal commun.). Hybridization information is available for petB and petD encoding cytochrome and subunit IV, respectively (Bohnert et al., 1985). These genes are co-transcribed resulting in a 1,400-b mRNA, as in cyanobacteria (Kallas et al., 1988), whereas petA, encoding cytochrome f and located between the atpBE and psbDC transcription units, likely yields a bicistronic mRNA of 1,500 nucleotides together with the downstream psaM(V. L. Stirewalt and D. A. Bryant, personal communication; Löffelhardt et al., 1987). A leader sequence encounteredwithall cyanobacterial andplastidgenes investigated (Widger, 1991) is also present in cyanelle petA. Finally, petG, encoding subunit V, was characterized by sequencing (Stirewalt and Bryant, 1989b).

4. Other Components of the Electron Transport Chain Neumann-Spallart et al. (1990) and Bryant et al. (1991) recently reported the nucleotide sequence of gene petFI, encoding ferredoxin I. Its 450-b mRNA represents one of the few monocistronic transcripts in cyanelles. The amino-terminal amino acid sequence of the purified protein (Stefanovic et al., 1990) could be shown to correspond to the gene sequence. Evrard et al. (1990c) reported the presence of a reading frame for a 6.5 kDa hydrophobic protein, presumably an intrinsic protein of thylakoid membranes, in cyanelles that is also present in chloroplasts.

5. Phycobilisomes Cyanelles and red algal plastids appear to encode the chromophorylated polypeptide components of the phycobilisomes (PBS; see Chapter 7 for details), whereas some of the linker polypeptides are products of nuclear genes (Egelhoff and Grossman, 1983; Burnap and Trench, 1989). However, gene probes for two rod linkers from Calothrix sp. strain PCC 7601 (Tandeau de Marsac et al., 1988) gave distinct

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signals with cyanelle DNA but not with nuclear DNA from C. paradoxa (C. Neumann-Spallart, personal communication). This indicates that some linker genes might also reside on plastid DNAs of PBScontaining algae (see Valentin et al., 1992b). This has recently been confirmed by Apt and Grossman (1993b, 1993c) for the red alga Aglaothamnion neglectum, for which the phycocyanin-associated rod core linker CpcG and a putative allophycocyanin associated core linker protein ApcC have been identified on the plastid genome. Phycobiliprotein genes were identified on the cyanelle genome prior to the detection of their cyanobacterial counterparts (Lemaux and Grossman, 1984). Transcription units in the center of the SSC region and comprising the genes cpcA and cpcB for the and of phycocyanin, respectively, and the genes apcA and apcB, encoding the and subunits of allophycocyanin, respectively, have been sequenced (Bryant et al., 1985; Lemaux and Grossman, 1985). The apcE gene encoding the 97 kDa ‘anchor’ phycobiliprotein is located adjacent to and upstream from apcAB on the same strand (Bryant, 1988), but is transcribed separately. This matches the organization in cyanobacteria (Capuano et al., 1991). This large protein, that channels excitation energy transfer from the PBS to Photosystem II, contains one biliprotein domain and three conserved domains of 150 amino acids each that show homology to rod-linker polypeptides. Thus, an interaction of the 97 kDa polypeptide with three allophycocyanin hexamer equivalents in the PBS core was suggested (Bryant, 1988). A sixth gene, apcD, that encodes the terminal-acceptor protein allophycocyanin B, has recently been found in cyanelle DNA and is unlinked to the other biliprotein genes (Michalowski et al., 1990a). A homolog of the Synechococcus sp. strain PCC 7002 apcF gene, that encodes the allophycocyanin subunit denoted and that is associated with the ApcE anchor phycobiliprotein of the cores, has recently been identified upstream from the cpcB gene (V.L. Stirewalt and D.A. Bryant, unpublished results). A similar gene has been identified on the plastid genome of the red alga Aglaothamnion neglectum (Apt and Grossman, 1993c). Biliprotein genes have recently been detected on the plastomes of several rhodophytic algae and cryptomonads (Shivji et al., 1992; Valentin et al., 1992b; Douglas, 1992b;Roell and Morse, 1993; Apt and Grossman, 1993a, b, c; Reith and Munholland, 1993). Interestingly,the ofphycoerythrin

Wolfgang Löffelhardt and Hans J. Bohnert appears to be nucleus- or nucleomorph-encoded in a cryptomonad, Chroomonas sp. This alga contains a light-harvesting system consisting of soluble biliproteins located inside the thylakoid lumen (Jenkins et al., 1990).

6. ATP Synthase The bipartite set of genes observed in cyanobacteria (Cozens and Walker, 1987) and chloroplasts is also encountered on the cyanelle genome. The bicistronic transcription unit atpB-atpE is located adjacent to rbcL on the opposite strand (Wasmann, 1985; Löffelhardt et al., 1987; Lambert et al., 1985; V. L. Stirewalt and D. A. Bryant, personal communication), a feature typical for higher plant chloroplasts, but different from cyanobacteria. The second gene cluster is remarkable in being adjacent to the upstreamgenes 5'- rpoB - rpoC1 - rpoC2 - rps2 -3', a trait in genome organization that is only shared with chloroplasts. However, there is a difference in gene composition. While the basic gene order found in the cyanobacterial operon is preserved, when compared to the arrangement in higher plant chloroplasts, two additional genes, atpD and atpG, are present and gene atpI, that is present in chloroplasts, is absent from the cyanelle gene cluster (V. L. Stirewalt, M. B. Annarella and D. A. Bryant, personal communication). In the plastomes of the diatom Odontella sinensis (Pancic et al., 1992) and of red algae (Valentin et al., 1992a) atpD, atpG, and atpI are present in the respective operons that lack only atpC compared to the cyanobacterial gene cluster. Interestingly, the cluster in Porphyra purpurea (Reith and Munholland, 1993) appears to be the result of a fusion of the procaryotic rpoB operon, the S2 operon (containing the tsfgene encoding elongation factor Ts) and the cyanobacterial atp gene cluster with the order preserved: 5'-rpoB-rpoC1-rpoC2-rps2-tsf-atpI-atpHatpG-atpF-atpD-atpA-3', without any missing gene other than atpC. The Porphyra purpurea gene order and number could represent the gene arrangement established after a primary endosymbiotic event.

7. Rubisco The genes for both subunits of ribulose-1,5-bisphosphate carboxylase/ oxygenase (Rubisco) on cyanelle DNA have been characterized (Heinhorst and Shively, 1983; Wasmann, l985; Starnes et al., 1985; Valentin and Zetsche, 1990a). Cyanelle rbcS was the first

Chapter 4 Molecular Biology of Cyanelles reported plastid-encoded rbcS gene whereas SSU from higher plants, green algae and euglenoids are the products of nuclear genes. The two genes are cotranscribed, as in cyanobacteria (Starnes et al., 1985). Recently, an identical arrangement of the two subunit genes has been detected in the plastomes of a number of chlorophyll b-less algae including three brown algae (Boczar et al., 1989; Valentin and Zetsche, 1990b; Assali et al., 1991), two red algae (Valentin and Zetsche, 1990b, 1990c), a cryptophycean alga (Douglas and Durnford, 1989), and a diatom (Hwang and Tabita, 1989).

D. Novel Genes in Cyanelle DNA 1. NAD Biosynthesis Recently, we have characterized (Michalowski et al., 1991 a) an open reading frame that is located close to the S10-spc ribosomal protein gene cluster (Fig. 1). Comparisons with sequences in the data banks suggested that this ORF329 had homology with bacterial nadA genes encoding, quinolinate synthetase (E.C.4.6.1.3). The enzyme catalyzes the condensation of iminoaspartic acid with dihydroxyacetone phosphate to generate quinolinic acid. Cyanelle nadA has 33.7% and 38.9% identity with the functionally identified genes from E. coli and Salmonella typhimurium, respectively. If conservative exchanges are permitted, homology is in the range of 65 to 74%. No homology for this coding region is found in the completely sequenced chloroplast genomes. NAD biosynthesis is an essential function of all organisms, most ofwhich possess a salvage pathway for this compound (for a review, see Foster and Moat, 1980). A biosynthetic pathway for NAD starting from low molecular weight compounds of basic metabolism has up to now only been studied biochemically and genetically in microorganisms (Flachmann et al., 1988; Foster et al., 1990), although it is to be expected that plants possess this pathway. The detection of nadA in Cyanophora paradoxa cyanelles is the first genetic indication that this pathway is present in eukaryotic cells.

2. Isoprenoid Pathway Cyanelle DNA contains an open reading frame, ORF323 (Michalowski et al., 1991b), with homology to gene crtE, originally reported to encode prephytoene pyrophosphate dehydrogenase, from Rhodo-

77 bacter capsulatus (Armstrong et al., 1989). This ORF is transcribed in cyanelles and is, at least in vitro, translated into a peptide of the size expected for this sequence. Reinvestigation of the function of the crtE gene product, now also identified in higher plants where it is nucleus-encoded (Kuntz et al., 1992), indicated an earlier role in the pathway, i. e. the activity of a geranyl-geranyl pyrophosphate synthase. In a test system based on genes for carotenogenic enzymes from Erwinia uredovora (Misawa et al., 1990) and their expression in transgenic E. coli the cyanelle ‘crtE’ gene product did not show geranyl-geranyl pyrophosphate synthase activity (G. Sandmann, personal communication). Sequence comparisons (Löffelhardt et al., 1993) point to the function of a higher prenyl transferase (Table 3). The best identity score was obtained with a hexaprenyl pyrophosphate synthase from yeast mitochondria (Ashby and Edwards, 1990). Thus we assume that the cyanelle protein encoded by ORF323 catalyses consecutive additions either in cis or trans of isopentenyl pyrophosphate to farnesyl pyrophosphate (Sherman et al., 1989) leading to undecaprenyl pyrophosphate, or leading to the precursor of the nonaprenyl sidechain of plastoquinone, respectively. In the latter case this enzyme activity should be present in all plastids whereas in the former case it might be confined to the peptidoglycan-containing cyanelles. In accord with this interpretation we propose to rename the gene as preA (prenyl transferase) as shown in Fig. 1.

3. Other Genes In addition to the ‘extra’ genes absent from higher plant chloroplast genomes which have been discussed before, a survey of other novel genes that have recently been published for chlorophyll b-less plastids is given in Table 4. Cyanelles, rhodophyte and chromophyte plastids contain genes for at least three chaperonin-type proteins, dnaK (= hsp70), groEL and groES (V. L. Stirewalt, J. Farley, and D. A. Bryant, personal communication). Cyanelles and Porphyra purpurea rhodoplasts have a number of other genes in common encoding enzymes involved in the biosynthesis of amino acids, carotenoids, chlorophyll and fatty acids (see Reith and Munholland, 1993, for a detailed list; V. L. Stirewalt, J. Farley, M. B. Annarella and D. A. Bryant, personal communication; C. B. Michalowski, unpublished). The position of a number of these genes on the cyanelle

78

Wolfgang Löffelhardt and Hans J. Bohnert

Chapter 4 Molecular Biology of Cyanelles chromosome is included in Fig. 1; other genes identified but not shown include chlB, chlL, petK, chlN, hemA, clpP1, clpP2, hisH, trpG, and acpA. Also included in this figure are several open reading frames (ORF) which have not been identified by homology with functionally identified genes. Up until now, the number of gene markers on cyanelle DNA that have been either completely or partially sequenced exceeds 140 genes. Approximately 85% of the genome is sequenced. Preliminary sequence informationobtained from remaining portions of the cyanelle genome suggests a further increase in the number of novel genes(V. L. Stirewalt, J. Farley, M. B. Annarella and D. A. Bryant, personal communication; C. B. Michalowski, unpublished). Due to the absence of introns in protein genes, to the small intergenic distances, and to the small size of the IR segment, a gene number exceeding that of higher plant chloroplasts can be accommodated, although the cyanelle genome is slightly smaller than the typical chloroplast genome. A set of genes thus far absent from the cyanelle genome are the ndh genes encoding putative subunits of a NADH dehydrogenase complex. These are found on higherplant chloroplast DNAs, but neither the gene products themselves nor their function could be demonstrated in vivo(Sugiura, 1992). Ndh genes have not be detected on the plastid genomes of Euglena gracilis, which is nearly completely sequenced (Hallick et al., 1993) or of Porphyra purpurea (approximately 60% sequenced; M. Reith and J. Munholland, personal communication). The lack of these genes might be a general phenomenon for algal plastid genomes.

E. Characteristics of Cyanelle Genes Generalizations about cyanelle gene structure become approachable with the availability of the increasing number of sequences determined. In many cases, heterologous hybridizations using probes from either cyanobacterial or plastid genes are successful in finding the corresponding cyanelle gene (Löffelhardt et al., 1985; Lambert et al., 1985). However, the failure to obtain a positive hybridization result does not necessarily mean that a given gene is missing. Genes on cyanelle DNA have a characteristic bias for either A or T in the third codon position whenever this is possible. Genes are densely packed, and the intergenic regions are extremely A+T-rich. Putative control regions of gene expression show the following three features. Firstly, sequence elements which

79 closely resemble the well-known ‘–35’ and ‘–10’ promoter boxes of bacterial genes can always be found. In several genes, such elements have been identified in regions that may form a stem-loop structure. Stem-loop structures, the second conspicuous feature, are very often found close to the termination codons of identified genes. In cases where two genes are separated by only a few nucleotides, we have found instances where a stem-loop structure might be formed involving the protein initiation codon for the following gene. Stems range up to approximately 40 base pairs with loops of four to five nucleotides. Since such structures have been found ubiquitously at positions which separate operons (identified by transcripts analysis; Michalowski et al., 1990b), we consider them functional features. In several instances convergently transcribed genes or operons appear to share a single sequence capable of forming a stem-loop structure. The third obvious feature is the presence of ‘ribosomebinding sites’ with complementarity to the cyanelle 16S rRNA 3'-end at virtually every position that contains an open reading frame with homology to functionally identified bacterial genes. In the promoter region of one gene, nadA, a sequence with high similarity to bacterial cAMP receptor protein binding sites has been detected (Michalowski et al., 1991a).

IV. Protein Transport

A. Import of Proteins Like plastids, cyanelles have to import the majority of their proteins. This statement can be made safely, even when we consider that the dense spacing of genes and the near total absence of introns (as far as sequences are known) suggest that more functional genes will be encoded on the cyanelle DNA than on higher plant chloroplast DNAs. The first example for such an imported protein is oxidoreductase (FNR), as it is synthesized as a preprotein on cytosolic 80S ribosomes (Bayer et al., 1990). Recently, the sequence of the corresponding cDNA was determined (Jakowitsch et al., 1993). The mature protein, which is overall highly conserved with respect to other FNR proteins, lacks the Nterminal extension thought to be responsible for attachment to phycobilisomes in the enzyme from the cyanobacterium Synechococcus sp. strain PCC 7002 (Schluchter and Bryant, 1992) and behaves in

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this respect like the higher plant enzymes (Michalowski et al., 1989). The amino-terminal transit peptide sequence shows little if any homology with transit peptides of higher plant pre-FNR enzymes or with other amino-terminal extensions in general. This is not surprising since plastid pre-sequences are notoriously variable. However, overall characteristics, such as a lack of charge at the extreme amino terminus and presence of hydroxyl-amino acids, of the presequence suggest the function of a transit peptide in stroma targeting. Functionally, the protein import apparatus for cyanelles appears similar to that of chloroplasts. We have recently made progress in establishing an in vitro protein uptake system into cyanelles (C. Neumann-Spallart and J. Jakowitsch, personal communication). Isolated cyanelles, as well as isolated pea plastids, import and process in vitro synthesized FNR precursor peptides resulting in a product of slightly higher mobility. Such a system will be important in comparing chloroplast and cyanelle protein uptake and routing of cytoplasmically synthesized pre-proteins.

B. Routing within Cyanelles Chloroplast proteins that are destined to the thylakoid lumen are subject to an additional independent targeting or routing mechanism dependent on the presence of a amino-terminal leader sequence. This may either constitute the amino-terminal more hydrophobic part of composite transit sequences in the case of nuclear gene products or the targeting signal of chloroplast genes (Smeekens et al., 1991). This second organellar protein translocation machinery which is also invoked for the functionally equivalent targeting to the mitochondrial intermembrane space is assumed to originate from the ancestral procaryotic endosymbiont. The retention of procaryotic pre-protein translocases in plastid and mitochondrial membranes is postulated by the ‘conservative sorting’ hypothesis (Hartl and Neupert, 1990).

C. Protein Translocation Machinery In Fig. 1 we have included an open reading frame (ORF492; Flachmann et al., 1993) that is homologous to the E. coli secY gene (Akiyama and Ito, 1987), whose product is essential for growth and is considered a part of the protein export complex (for reviews, see Wickner et al., 1991; Bieker and Silhavy,

Wolfgang Löffelhardt and Hans J. Bohnert 1990). The cyanelle ORF492 is located at the 3'-end of the spc operon (Michalowski et al., 1990b). E. coli secY occupies the same position, although it is cotranscribed with the ribosomal protein genes in the spc operon. It appears that this ORF492, which is transcribed in the organelles as a monocistronic mRNA separate from the spc operon, encodes a SecY-like protein in cyanelles. This protein shows a sequence identity of 28.1% with the E. coli SecY counterpart and is functional in E. coli as shown by a complementation assay (Flachmann et al., 1993). Cyanelle SecY restored growth at 42 °C in transformed, thermosensitive secY mutants of E. coli (Shiba et al., 1984). A tentative model of the membrane topology of cyanelle SecY, based on computer predictions, on comparison with the E. coli protein, and on the analysis in bacteria of two secY-phoA fusion proteins (Flachmann et al., 1993) is given in Fig. 2. Very recently, secY genes have also been identified on the plastid genomes of Cryptomonas sp. (Douglas, 1992a) and the chromophytic alga Pavlova lutherii (Scaramuzzi et al., 1992a) by sequence similarity. In these cases the identity scores towards the cyanelle protein are in the range of 50%. Cyanelles and other chlorophyll b-less plastids might well encode additional subunits of the preprotein translocase as exemplified by the recent detection of secA on the plastid genome of a chromophyte (Scaramuzzi et al., 1992b) and a rhodophyte (Valentin, 1993). However, the nature of the cyanelle membrane(s) harboring SecY and the other components of the translocase remains to be demonstrated through the use of specific antibodies. Our working hypothesis assumes that cyanelles and, likely all plastid types, possess a thylakoid-bound, SecY-dependent protein transport system for the translocation of lumenal polypeptides. In the case of higher plant chloroplasts the respective genes must reside on the nuclear genome. In addition, the IEM in cyanelles may also contain SecY protein and the transport machinery. In contrast to the intermembrane space of the plastid envelope the periplasmic space of cyanelles is a defined compartment containing a number of identified proteins, including enzymes involved in wall synthesis (Plaimauer et al., 1991). The most likely candidates for proteins to leave the organelle stroma would be the pre-proteins of enzymes involved in the biosynthesis of the peptidoglycan wall and OM proteins.

Chapter 4 Molecular Biology of Cyanelles

V. Phylogenetic Analyses As a ‘bridge’ organism C. paradoxa has often been included in phylogenetic trees constructed from different traits using different computing programs (for additional information, see Chapter 5). Due to the problems still inherent to phylogenetic algorithms and the possible substitutional and constitutional bias encountered in the sequences to be analyzed (see Lockhart et al., 1992), the results should be considered not only with interest, but also with caution. At first, cyanelle RNA genes have been taken for comparisons. 16S rRNA-derived phylogenies show cyanelles and all kinds of plastids well within the cyanobacterial radiation (Giovannoni et al., 1988; Urbach et al., 1992; see Chapter 5) with a pronounced relation of cyanelles to plastids from Cryptomonas sp. and red algae (Douglas, 1992b). When 5S rRNA is the trait compared, all plastids group together with cyanelles coming closest to Porphyra sp. and Euglena gracilis plastids (Wolters et al., 1990). Based on D1 protein sequences Cyanophora paradoxa groups with the rhodophyte Cyanidium caldarium and the xanthophycean alga Bumilleriopsis filiformis between the cyanobacteria and prochlorophytes on one hand and the branch containing green algae, Euglena gracilis and higher plants on the other hand (Scherer et al., 1991). Ribosomal protein genes rpl33, rps18, rpl2, rps19, and rpl22 constituted the data set for trees that indicate a shorter evolutionary distance between cyanelles and higher plant plastids than between the latter and Euglena gracilis plastids (Evrard et al., 1990a). Trees constructed from the nucleotide

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sequences of three str operon genes (Kraus et al., 1990) were most conclusive for rps7 that is the least conserved gene; here cyanelles occupy an intermediate position between chloroplasts and cyanobacteria coming somewhat closer to the latter. With rps12 and especially tufA the order of branching of the algal chloroplasts became uncertain. A wealth of sequence information is available concerning ferredoxin I; in two trees obtained with different analysis programs, cyanelles group with cyanobacteria, the plastids from rhodophytic algae and Xanthophyceae being the next closest relatives (M. Kraus, unpublished; Lüttke, 1991). A recently published tree using sequences of the rpoC1 gene products (Palenik and Haselkorn, 1992) confirms the results of 16S rRNA data, i. e. that the cyanelle is the closest known relative to the ancestor of chloroplasts. When the LSU of Rubisco was used as a phylogenetic marker, cyanelles appeared to be separated from plastids of rhodophytes, chromophytes, and Cryptomonas sp. groupingwith the chlorophyll btype plastids (Valentin et al., 1992a). Rubisco SSUderived trees (Assali et al.,1991; Valentin et al., 1992a) show similar features: Cyanophora paradoxa groups with cyanobacteria (that also show no insertion —31 amino acids with rhodophytes/chromophytes and 12 amino acidswith chlorophytes, respectively— in the amino-terminal region of the SSU protein) and is separated from other plastid types, especially those from rhodophytes and Cryptomonas sp. This is in contrast to trees obtained from numerous other traits and indicates that Rubisco postulated to originate from lateral gene transferwith or bacteria

82 as donors (Assali et al., 1991) might not be the best choice as an evolutionary marker. Upon analysis of their trees several authors claim a monophyletic origin of plastids whereas others favor a polyphyletic origin, eventually separately for cyanelles (see Chapter 5 for additional information on this subject). In our opinion neither the dataset available nor the present state of the art in phylogenetic analysis allows a definitive answer to this important question. Nuclear gene sequences from C. paradoxa have become available only recently. The trees deduced from FNR sequences support the intermediate position of cyanelles between plant chloroplasts and, slightly closer, cyanobacteria (Jakowitsch et al., 1993). Very recently a trait pertinent to the ‘eukaryotic host’ became available: 18S rRNA data show a close relationship between C. paradoxa, Glaucocystis nostochinearum, and the cryptophycean algae Cryptomonas sp. and Pyrenomonas salina. They are sister groups, i. e. they share a common evolutionary history (D. Bhattacharya, personal communication). This is also supported by some of the plastid gene-derived trees mentioned above. Interestingly, due to its mitotic apparatus C. paradoxa was once classified as a cryptomonad (Pickett-Heaps, 1972). Phylogenetic analyses based on eubacterial (Tschauder et al., 1992), cyanobacterial (Nakai et al.,

Wolfgang Löffelhardt and Hans J. Bohnert 1992) and plastid SecY proteins using PAUP (version 3.0; D. Swofford, Illinois Natural History Survey, Champaign, Illinois) gave discouraging results: C. paradoxa and Synechococcus sp. strain PCC 7942 appeared to be unrelated. The structural constraints acting on this membrane protein seem to be different among different organisms (e.g., the size of the protein is quite variable); the G + C contents of the respective genomes are also quite variable. Thus, SecY is another example for a trait less suitable for phylogenetic analysis. Nonetheless, a dendrogram constructed from the aligned secY nucleotide sequences using the PileUp function of the GCG Sequence Analysis Software Package (Devereux et al., 1985) is shown in Fig. 3 that depicts the relationships between the secY genes from 12 organisms. An analysis based upon SecY protein sequences would group Cryptomonas sp. with Synechococcus sp. strain PCC 7942.

VI. Conclusions Our view regarding the position of C. paradoxa cyanelles in the context of plastid evolution is given in the Fig. 4 (compare to Fig. 9 of Chapter 5). This hypothetical scheme is not based on identities of

Chapter 4 Molecular Biology of Cyanelles

individual genes, or on identities of groups of genes, but it is rather based on genome, operon and gene organization features. The scheme owes much to the recent work on the plastid genome of Porphyra purpurea (Reith and Munholland, 1993) and to work on the genome organization of chlorophyll b-less plastids in general (Douglas, 1992b; see Chapter 5). Their data and the data resulting from the cyanelle sequencing project permit a generalizing view. According to this scheme, cyanelles would constitute a side line of plastid evolution branching off early from a semiautonomous endosymbiont, originating from a single, primary endosymbiotic event that may be ancestral to all plastid types. This may explain why Cyanophora paradoxa is relatively rarely encountered: The organism has been isolated only two times. Positioning cyanelles as shown appears justified regarding the many characteristics they share

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with rhodophytes and chromophytes. This location, however, does not exclude distinct parallels to the chlorophyte/ higher plant lineage. The descendence of chromophyte plastids from multiple secondary endosymbiotic events with rhodophyte-like invaders and heterotrophic host cells is now widely accepted (for reviews, see: Douglas, 1992b and Chapter 5). This view might also apply for euglenoid algae (Douglas, 1992b). Additional gene transfer events from the nucleus of the (eucaryotic) endosymbiont to the nucleus of the (secondary) ‘host cell’ is invoked. This led either to the complete loss of the endosymbiont nucleus in euglenoids, brown algae and diatoms, or to its retention as a nucleomorph in cryptomonads. In contrast to the plastids of the chlorophyll b-lineage, the other plastid types (Fig. 4) share an estimated total of 40 to 50 additional genes (Reith and Munholland, 1993; Valentin et al., 1992a;

84 V. L. Stirewalt, J. Farley, M. B. Annarella and D. A. Bryant, personal communication; C. B. Michalowski, unpublished results).

Acknowledgments We wish to thank our colleagues and collaborators, former students and postdoctoral fellows, who have worked on cyanelle gene structure over the last 12 years. During that time work has been supported by the Deutsche Forschungsgemeinschaft, EMBLHeidelberg, Fonds der Stadt Wien, Hochschuljubiläumsstiftung Wien, Austrian Research Council, National Science Foundation, and Arizona Agricultural Experiment Station, Tucson, Arizona. We thank Christine Michalowski and Marty Wojciechowski for establishing the PAUP-program for phylogenetic analyses. The cyanelle genome sequencing project is supported by a grant from USDANRI (Plant Genome) to HJB. and Donald A. Bryant (The Pennsylvania State University).

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Wolfgang Löffelhardt and Hans J. Bohnert Sugiura M (1992) The chloroplast genome. Plant Mol Biol 19: 149–168 Tandeau de Marsac N, Mazel D, Damerval T, Guglielmi G, Capuano V and Houmard J (1988) Photoregulation of gene expression in the filamentous cyanobacterium Calothrix sp. PCC 7601: Light harvesting complexes and cell differentiation. Photosynth Res 18:99–132 Thomas F, Massenet O, Dorne AM, Briat JF and Mache R (1988) Expression of the rpl23, rpl2 and rps19 genes in spinach chloroplasts. Nucl Acids Res 16: 203–209 Trench RK (1979) The cell biology of plant-animal symbiosis. Annu Rev Plant Physiol 30: 485–531 Trench RK (1982a) Physiology, biochemistry, and ultrastructure of cyanellae. In: Round FE and Chapman DJ (eds) Progress in Phycological Research, Vol 1, pp 257–288. Elsevier, Amsterdam Trench RK (1982b) Cyanelles. In: Schiff JA (ed) On the Origin of Chloroplasts, pp 55–76. Elsevier/ North Holland, New York Tschauder S, Driessen AJM and Freudl R (1992) Cloning and molecular characterization of the secY genes from Bacillus licheniformis and Staphylococcus carnosus: Comparative analysis of nine members of the SecY family. Mol Gen Genet 235: 147–152 Turner S, Burger-Wiersma T, Giovannoni SJ and Mur L (1989) The relationship of a prochlorophyte, Prochlorothrix hollandica, to green chloroplasts. Nature 337: 380–382 Urbach E, Robertson DL and Chisholm SW (1992) Multiple evolutionary origins of prochlorophytes within the cyanobacterial radiation. Nature 355: 267–270 Valentin K (1993) SecA is plastid-encoded in a red alga: Implications for the evolution of plastid genomes and the thylakoid protein import apparatus. Mol Gen Genet 236: 245– 250 Valentin K and Zetsche K (1990a) Nucleotide sequence of the gene for the large subunit of Rubisco from Cyanophora paradoxa – phylogenetic implications. Curr Genet 18: 199– 202 Valentin K and Zetsche K (1990b) Rubisco genes indicate a close phylogenetic relation between the plastids of chromophyta and rhodophyta. Plant Mol Biol 15: 575–584 Valentin K and Zetsche K (1990c) Structure of the Rubisco operon from the unicellular red alga Cyanidium caldarium: Evidence of a polyphyletic origin of the plastids. Mol Gen Genet 222: 425–430 Valentin K, Cattolico RA and Zetsche K (1992a) Phylogenetic origin of the plastids. In: Lewin R (ed) Origins of plastids, pp 195–222. Chapman & Hall, New York Valentin K, Maid U, Emich A and Zetsche K (1992b) Organization and expression of a phycobiliprotein gene cluster from the unicellular red alga Cyanidium caldarium. Plant Mol Biol 20: 267–276 Wang S and Liu XQ (1991) The plastid genome of Cryptomonas encodes an hsp70-like protein, a histone-like protein, and an acyl carrier protein. Proc Natl Acad Sci USA 88: 10783–10787 Wasmann CC (1985) The cyanelle and the cyanelle genome of Cyanophora paradoxa. Ph. D. thesis, Michigan State University, East Lansing, MI Wasmann CC, Löffelhardt W and Bohnert HJ (1987) Cyanelles: Organization and molecular biology. In: Fay P and Van Baalen C (eds) The Cyanobacteria, pp 303–324. Elsevier, Amsterdam Wickner W, Driessen AJM and Hartl F-U (1991) The enzy-

Chapter 4 Molecular Biology of Cyanelles mology of protein translocation across the Escherichia coli plasma membrane. Annu Rev Biochem 60: 101–124 Widger WR (1991) The cloning and characterization of Synechococcus sp. PCC 7002 petCA operon: Implications for the cytochrome c-553 binding domain of cytochrome f. Photosynth Res 30: 71–84 Winhauer T, Jaeger S, Valentin K and Zetsche K (1991) Structural similarities between psbA genes from red algae and brown algae. Curr Genet 20: 177–180 Wolters J, Erdmann VA and Stackebrandt E (1990) Current status

89 of the molecular phylogeny of plastids. In: Nardin P (ed) Endocytobiology IV, pp 545–552. INRA, Paris Xu MQ, Kathe SD, Goodrich-Blair H, Nierzwicki-Bauer SA and Shub DA (1990) Bacterial origin of a chloroplast intron: Conserved self-splicing group I introns incyanobacteria. Science 250: 1566–1570 Zhou DX, Quigley F, Massenet O and Mache R (1989) Cotranscription of the S10- and spc-like operons in spinach chloroplasts and identification of their gene products. Mol Gen Genet 216: 439–445

Chapter 5 Chloroplast Origins and Evolution Susan E . Douglas Institute for Marine Biosciences. National Research Council. 141 1 Oxford Street. Halifax. Nova Scotia B3H 321. Canada Summary .................................................................................................................................................................91 I. Introduction ...................................................................................................................................................... 92 II. The Procaryotic Ancestry of Plastids and Their Subsequent Evolution .......................................................... 93 A. Plastid Gene Content ......................................................................................................................... 93 B. Gene Clusters .................................................................................................................................... 95 1. Ribosomal RNA Operons .........................................................................................................96 2. Ribosomal Protein Operons...............................................................................................96 3. ATPase Operons ......................................................................................................................97 4 . Photosystem Operons ............................................................................................................100 5. Other ....................................................................................................................................... 100 C. Plastid Transcription ........................................................................................................................ 101 D. Plastid lntrons .................................................................................................................................. 102 E. Plastid Gene Sequences ........................................................................................................... 102 1. Ribosomal RNA ......................................................................................................................103 a. 5 s rRNA ........................................................................................................................103 b. SSU rRNA .....................................................................................................................103 c . LSU rRNA ................................................................................................................ 105 2. ATP Synthase Subunit Beta (atpf3) ........................................................................................ 105 105 3. Photosystem II Protein D l (psbA) .......................................................................................... 4 . Ribulose.1, 5.Bisphosphate Carboxylase (Rubisco) ............................................................... 105 5. Elongation Factor Tu (tufA) ................................................................................................... 107 6. Others .............................................................................................................................. 107 F. Nuclear Gene Sequences ............................................................................................................. 108 Ill. Secondary Endosymbiosis in Plastid Evolution ............................................................................................. 108 A. Electron Microscopic Studies ....................................................................................................... 108 B. Hybridization Studies .................................................................................................................... 109 C. Gene Sequences ............................................................................................................................. 109 1. Ribosomal RNA ...................................................................................................................... 109 a. 5 s rRNA .................................................................................................................... 109 b. SSU rRNA ................................................................................................................. 109 c . LSU rRNA .................................................................................................................. 109 2. GAPDH ................................................................................................................................... 110 IV. Conclusions and Future Prospects ................................................................................................................ 111 Acknowledgments ................................................................................................................................................. 111 References ............................................................................................................................................................ 112

Summary Plastids from extant plants exhibit considerable diversity in morphological and biochemical characters. Although most authors have agreed on xenogenous (endosymbiotic) rather than autogenous origins o f plastids (discussed b y Doolittle ( I 982) in 'The Biology o f Cyanobacteria'). details concerning the endosymbiotic events remain unresolved. D. A . Bryant (ed): The Molecular Biology of Cyariobacter-ia.pp . 91-1 18. O I994 Kluwer Academic Publishers . Printed in The Netherlands.

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In the eleven years since ‘The Biology of Cyanobacteria’, many data have accumulated that, while supporting the xenogenous origin of plastids, have revived the controversy over single (monophyletic) versus multiple (polyphyletic) origins. These arguments revolve around the number and nature of the primary endosymbiont(s) that gave rise to the first plastid-bearing eucaryotes. The question of whether secondary endosymbiotic events, originally hypothesized on the basis of electron microscopic evidence, were responsible for the formation of ‘complex’ plastids (those surrounded by more than two membranes) has now been investigated by molecular methods. The purpose of this chapter is to present recent evidence bearing on the probable nature of the procaryotic ancestor(s) involved in the primary endosymbiotic event(s), and on the secondary endosymbiotic events that gave rise to eucaryotes bearing complex plastids. Comparisons of gene content, gene arrangement, gene expression and gene sequences between extant cyanobacteria and plastids give important clues about the possible ancestors of plastids and of the subsequent transformation of a eubacterial genome into a plastid genome. In the last several years, four complete land plant plastid genomes have been sequenced, contributing vastly to our knowledge of plastid architecture and expression. In addition, a great deal of molecular data has been acquired on cyanelle and cyanobacterial genes and genomes. Increased emphasis has now been placed on the study of non-land plant plastid genomes and a number of rhodophyte, chromophyte, cryptophyte and euglenophyte plastid genomes have been extensively mapped and sequenced. These data are presented and the phylogenetic implications evaluated. I. Introduction On the basis of morphological and biochemical characteristics, plants have been assigned to three primary lineages (Table 1). Those plants possessing Chl a/phycobilin-containing plastids were assigned to the Rhodophyta (red algae), those possessing Chl a/b-containing plastids included the Chlorophyta (green algae) and Metaphyta (land plants), and those possessing Chl a/c-containing plastids were assigned to the Chromophyta (Christensen, 1964). However, algae exist that are hard to assign to one of these particular groups because of unique characteristics such as aberrant or supernumerary plastid membranes and/or unusual pigment complements. For example, some members of the Glaucophyta possess cyanelles surrounded by a residual murein sacculus (Wasmann et al., 1987; see chapter4). The Euglenophyta contains both aplastidial and plastidial (three-membraned) representatives, and mutants that have lost their plastids may be obtained, indicating that these organelles are dispensable in certain members of this group. The Dinophyta also contains aplastidial and plastidial (also three-membraned) members and some representatives contain endosymbiotic photosynthetic Abbreviations: CER – chloroplast endoplasmic reticulum; Chl – chlorophyll; GAPDH – glyceraldehyde-3-phosphate dehydrogenase; LSU – large subunit; rRNA – ribosomal ribonucleic acid; Rubisco – ribulose-1,5-bisphosphate carboxylase/oxygenase; SSU – small subunit

eucaryotes (see Dodge 1987, Schnepf 1993). Another anomalous situation is seen in the Cryptophyta, which possess Chl a/c-phycobilin-containing plastids that are surrounded by four membranes and contain a small, nucleus-like organelle called the nucleomorph (Greenwood, 1977) in a cytoplasmic space between the inner and outer plastid membrane pairs (Gillot and Gibbs, 1980). A similar situation is seen in the mastigamoeba Chlorarachnion sp. that possesses Chl a/b-containing plastids with a similar ultrastructure to those of the Cryptophyta (Ludwig and Gibbs, 1989). It was Schimper (1883) who first proposed that plant cells resulted from autonomous green cells and colorless hosts by endosymbiosis. Mereschowsky (1905) proposed cyanobacteria as the progenitors of plastids, and later envisioned plastids of the major groups of photosynthetic eucaryotes to have arisen from different groups of cyanobacteria (Mereschowsky, 1910). This idea was revitalized by Margulis (Sagan, 1967; Margulis, 1970), and Raven (1970) proposed that multiple endosymbiotic events involving procaryotes containing distinct pigment complements gave rise to extant plastids. Cyanobacteria possess the same pigment complement as rhodophytes and thus seemed likely progenitors of rhodophyte plastids (Table 1). However, until recently, procaryotes possessing pigment complements corresponding to chlorophyte or chromophyte plastids eluded detection. The discovery of the prochlorophyte Prochloron didemni,

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a procaryote containing both chlorophylls a and b (Lewin and Withers, 1975), caused great excitement and led to the proposal that it could be related to the putative ancestor of the plastids of the Chl a/bcontaining plants. Similarly, the brownish photoheterotroph Heliobacterium chlorum was postulated to be a representative of the lineage that gave rise to the putative ancestor of the chlorophyll a/c-containing algal plastids (Margulis and Obar, 1985). However, molecular studies have not borne out these expectations. Although there is now general agreement that cyanobacteria gave rise to plastids, one of the major unresolved questions is whether the primary endosymbiotic event involved a single (monophyletic) or multiple (polyphyletic) cyanobacterial ancestors (see Gray, 1991). Molecular data bearing on this question include comparisons of the arrangement of gene clusters in cyanobacteria with those of plastid genomes from photosynthetic eucaryotes, and phylogenetic analyses of homologous plastid and cyanobacterial genes. Most of the sequence data shows the emergence of plastids from a single point within the cyanobacterial assemblage indicating a monophyletic origin (see Morden et al., 1992). Comparisons of gene arrangement in cyanobacteria and plastids also support monophyly. However, vehement advocates of a polyphyletic origin persist and the controversy remains unresolved. The origin of plastids is further clouded by the possibilities of lateral gene transfers and endosymbionts containing chimaeric genomes (Martin et al., 1992).

Secondary endosymbiotic events (Fig. 1) between photosynthetic eucaryote(s) and phagotrophichost(s) have been postulated, based largely on ultrastructural data, to contribute to the formation of ‘complex’ plastids containing more than two membranes (Tomas and Cox 1973; Gibbs, 1978, 1981a; Whatley et al., 1979). However, whether this occurred only once as suggested by Cavalier-Smith (1982, 1986) or several times (Whatley et al., 1979) remains unclear. Electron microscopic in situ hybridizations (McFadden, 1990a, b), as well as comparisons of SSU rRNA sequences (Douglas et al., 1991; Eschbach et al., 1991a; Maier et al., 1991) of the nucleus and nucleomorph of cryptomonad algae, have confirmed that this type of alga has been formed by secondary endosymbiosis. Furthermore, phylogenetic analyses of these sequences suggested that multiple secondary events have contributed to the formation of those algae bearing complex plastids. II. The Procaryotic Ancestry of Plastids and Their Subsequent Evolution A. Plastid Gene Content The complete sequences of the plastid genomes from liverwort, tobacco and rice (Ohyama et al., 1986; Shinozaki et al., 1986; Hiratsuka et al., 1989) have shown that these plastids are arranged very conservatively and contain approximately the same

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gene complement (see Sugiura, 1992). Common features include a 25–30 kb rRNA-encoding inverted repeat, genes for many plastid tRNAs, most RNA polymerase subunits, many ribosomal proteins and several translational factors. In addition, various subunits of the photosynthetic apparatus components including Rubisco, Photosystems I and II, the

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cytochrome b/f complex and ATP synthase are plastidencoded. A surprising feature is the presence of eleven genes (ndh genes) encoding subunits of a putative respiratory chain NADH dehydrogenase complex. The genomes of the non-photosynthetic plastids of the land plant, Epifagus virginiana, and the euglenoid

Chapter 5 Plastid Evolution alga, Astasia longa, have lost most of the genes involved in photosynthesis and chlororespiration and encode a reduced complement of products involved predominantly in translation such as ribosomal proteins, rRNAs and tRNAs (see Wolfe et al., 1991). Surprisingly, the plastid genome of Astasia longa encodes a transcriptionally active rbcL gene (Siemeister and Hachtel, 1990), suggesting that this gene product may perform a function other than in photosynthesis in this organism. Although the plastid genomes of these non-photosynthetic organisms are transcriptionally active, the genes encoding subunits of RNA polymerase are lacking from the plastid genome of Epifagus virginiana (Wolfe et al., 1992), indicating the probable presence of a second nuclearencoded RNA polymerase that must be imported into the plastid. Recent studies of transcription in ribosome-deficient plastids of barley support this hypothesis (Hess et al., 1993). Extensive mapping and sequencing of plastid genomes from the non-green lineages (for reviews, see Bohnert and Löffelhardt, 1992; Douglas, 1992b; Reith and Munholland, 1993; Valentin et al., 1993) have revealed many novel features. These genomes encode many additional genes that are not found on land plant genomes, but the ndh genes have not yet been detected. These additional genes encode extra tRNAs as well as proteins involved in such functions as electron transport, photosynthesis, DNA binding, protein secretion, transcriptional regulation, protection from heat shock effects, and biosynthesis of amino acids, chlorophyll, carotenoids, phycobilins and fatty acids. One interpretation of this phenomenon is that more of the genes originally present on the genome of the endosymbiont have been lost or transferred to the nucleus in the lineage leading to green algae and land plants than in those leading to rhodophytes, cryptophytes and chromophytes. Many genes encoding components of the cyanobacterial phycobilisome (for reviews, see Bryant, 1992; Tandeau de Marsac, 1991) and photosystems (for reviews, see Vermaas and Ikeuchi, 1991; Chitnis and Nelson, 1991; Bryant, 1992) have been isolated, sequenced and used to infer phylogenetic relationships among plastids. In recent years genes involved in other cyanobacterial processes have been identified which, if homologs exist on plastid genomes, might well give clues about possible plastid ancestors. Genes involved in sulfate uptake have been identified from Synechococcus sp. strain PCC 7942 (Laudenbach and Grossman, 1991) and Synechocystis sp.

95 strain PCC 6803 (Kohn and Schumann, 1993), a gene involved in protochlorophyllide reduction has been identified from Plectonema boryanum (Fujita et al., 1991) and a gene involved in control glutamine biosynthesis has been identified from Synechococcus sp. strain PCC 7942 (Tsinoremas et al., 1991). Homologs (mbpX, mbpY, frxC (chlL) and glnB) of some of these genes are present on the plastid genome of the liverwort Marchantia polymorpha (Ohyama et al., 1986), Chlamydomonas reinhardtii (Huang and Liu, 1992), gymnosperms and pteridophytes (Yamada and Yamamoto, 1992) and rhodophytes (Reith and Munholland, 1993). Although most of the genes identified on plastid genomes have counterparts in cyanobacteria, in some cases cyanobacterial homologs have not been identified. It is now possible to use plastid probes to detect cyanobacterial genes, rather than the reverse strategy which was used in the past. PCR-based approaches are also very powerful tools. In addition, cyanobacterial genes that have gone undetected are being uncovered by amino-terminal amino acid sequencing and comparison to translated plastid sequences. An example of this approach is the detection in the cyanobacterium Anabaena variabilis strain ATCC 29413 of the gene product of psaI (Ikeuchi et al., 1991), a gene that had previously been detected only in land plant plastid genomes. This gene has recently been detected on the plastid genome of the cryptomonad alga Cryptomonas sp. (Douglas, 1992b) and on the cyanelle genome of Cyanophora paradoxa (D. A. Bryant and V. L. Stirewalt, personal communication) as well.

B. Gene Clusters A number of gene clusters are conserved between cyanobacteria and the plastids of land plants and algae. These include the tRNA-containing rRNA operons, ribosomal protein operons, the ATPase operons, and operons containing genes for photosynthesis. Of special significance are clusters that appear to have evolved subsequent to endosymbiosis and are present in plastid genomes from several lineages but absent in cyanobacteria. Such arrangements provide very strong evidence for the monophyletic origin of plastids, since such similar organization occurring in separate lineages could otherwise be explained only by an extraordinary degree of convergent evolution. As will be noted in the following discussion, a common theme is the loss

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of genes from the larger procaryote-like operons in non-green plastids to the nucleus in the more advanced plants, the scrambling and scattering of the remnant operons and the introduction of introns into some of the remaining genes (Figs. 2–5). As more data become available, a pattern of gene content and organization from the eubacterial situation, to these morphologically simple algae and more advanced land plants begins to emerge.

1. Ribosomal RNA Operons One of the most notable characteristics of land plant and green algal plastid genomes is an inverted repeat of approximately 20–30 kb that encodes the rRNA and tRNA genes and often some protein-encoding genes (Palmer, 1991). In Euglena gracilis the rRNA genes (1 –5 copies) are tandemly repeated (see Hallick and Buetow, 1989). Most chromophyte plastids contain rRNA-encoding repeats, although they are inverted in orientation, are frequently much smaller (approximately 5 kb) and usually do not contain protein-encoding genes. Among the rhodophytes, the rRNA genes are reported to be present either as one copy, as an inverted repeat, or as two small, directly repeated copies of non-identical sequence (see Reith and Munholland, 1994). This latter organization can be interpreted to represent the ancestral plastid state from which identical inverted repeats arose by inversion and whose identities were then maintained by a copy-correction mechanism. Subsequent expansion to include protein-encoding genes occurred in some green algae and land plants (especially the Geraniaceae), and several examples have now been documented in chromophytes and chlorophytes (Shivji et al., 1992; see Kowallik, 1993). In a few cases (such as the legumes), one repeat has presumably been lost and plastid instability resulting in genomic rearrangements has occurred (Palmer et al., 1987). Information about the orientation of cyanobacterial rRNA operons is limited (Bancroft et al., 1989), but multiple copies do exist in some taxa and at least those of Synechococcus sp. strain PCC 6301 are non-identical (Douglas and Doolittle, 1984; Kumano et al., 1983). A scheme showing the distribution of rRNA operons in different plastid types, and suggesting possible mechanisms for their formation, is shown in Fig. 2. If the assumption that non-identical repeats are ancestral is incorrect, schemes invoking other losses and gains of repeat structure among the rhodophytes are also plausible.

Susan E. Douglas The transition from chlorophyte algae to land plants may have involved numerous rearrangements as well as several independent losses and gains of repeat structure. In addition to information about the number and orientation of rRN A operons, the size of the intergenic spacers between the 16S rRNA and 23S rRNA genes and the presence or absence of introns in the tRNA genes can be used to infer phylogenetic relationships (see Douglas, 1993). Cyanobacteria, cyanelles and the plastids of non-green algae and E. gracilis all contain short intergenic spacers within which reside the genes for uninterrupted and On the other hand, the spacers of some green algae and land plants are much larger and contain the same tRNA genes but possessing group II introns, a marker that has been used to trace the ancestry of land plants from within the Charophyceae (sensu van den Hoek et al., 1988) group of green algae (Manhart and Palmer, 1991).

2. Ribosomal Protein Operons In eubacteria, many of the ribosomal protein genes are organized into operons: e.g. str, S10, spc, alpha, rplKAJL, each encoding from 2 to 11 proteins (see Nomura et al., 1984). These ribosomal protein operons have been studied in a number of land plants and algae. However, with the exception of the str (Buttarelli et al., 1989) and the rplKAJL operons (Schmidt and Subramanian, 1992), very few cyanobacterial ribosomal protein operons have been studied. A recent report of the sequence of the secY gene from Synechococcus sp. strain PCC 7942 (Nakai et al., 1992) indicates that the S10/spc gene cluster exists in this organism and that the secY gene is found at the 3' end as in Escherichia coli. Rhodophyte (see Reith and Munholland, 1993) and cryptophyte (see Douglas et al., 1992) plastids appear to have retained the most genes from the ancestral eubacterial str/S10/spc/alpha arrangement except that the str operon is now located downstream of the S10/spc/alpha operons (Fig. 3). Cyanelles of C. paradoxa contain almost as many genes and in the same arrangement as eubacteria except that the alpha operon is no longer downstream of the str/S10/spc operons (see Bohnert and Löffelhardt, 1992; also see Chapter 4) whereas land plant and green algal plastids contain substantially reduced ribosomal protein operons (see Subramanian et al., 1990, 1991). This can be interpreted as a differential loss or transfer of

Chapter 5 Plastid Evolution

ribosomal protein genes to the nucleus in the different plant lineages (Douglas, 1991). Indeed, the absence of the rpl22 gene from the plastid genome of legumes but its presence in most other land plant plastid genomes, suggests that some such transfers have taken place relatively recently (Gantt et al., 1991). In addition to containing fewer ribosomal protein genes, the remnant operons of land plant plastids have often been extensively scrambled and in some cases, trans-splicing between exons encoded on opposite strands of the plastid DNA must occur in order to obtain a functional polypeptide (Zaita et al., 1987). In other cases, the operons have been so disrupted that ribosomal protein genes are found next to genes involved in different functions, and introgression between two or more gene clusters has occurred (see Subramanian et al., 1991). In cyanobacteria, rps2 is found at a separate locus from the rpoBC1C2 and atpA gene clusters, but in many

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plastids, including the cyanelle of C. paradoxa (see chapter 4) it is now found between these two clusters (Fig. 4). The assembly of such a gene cluster after endosymbiosis has occurred, and its retention in a recognizable form in all of the major plant lineages gives strong support for the monophyletic origin of plastids (see Reith and Munholland, 1993). Additional information on ribosomal protein gene clusters from cyanobacteria would greatly enhance our understanding of the origins of plastids.

3. ATPase Operons The genes for the subunits of the are organized into two clusters in cyanobacteria, the atpB cluster containing atpB and atpE, and the atpA cluster containing atpI, atpH, atpG, atpF, atpD, atpA and sometimes atpC (Cozens and Walker, 1987). The structure of the atpB operon has been largely

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unaltered in all plastids except for the overlapping of the atpB and atpE genes by 4 bp in the land plant lineage. The atpA operon has remained largely intact in the plastid genomes of the diatom Odontella sinensis (Pancic et al., 1992) and the rhodophytes Antithamnion sp. (Kostrzewa and Zetsch, 1992) and Porphyra purpurea (Reith and Munholland, 1993). Preliminary sequence data indicate that a similar structure is also present on the plastid genome of Cryptomonas sp. (Douglas, 1992b) and the cyanelle genome of C. paradoxa (D. A. Bryant and V. L. Stirewalt, personal communication; also see chapter 4) although atpI is no longer present in the latter. In land plant plastid genomes, atpG and atpD are no longer present on the plastid genome, and an intron interrupts atpF (Fig. 4). In the green algae C. reinhardtii and C. moewusii, genes for the ATPase subunits (atpB, atpE, atpA, atpH, atpI and atpF) (Woessner et al., 1987, Turmel et al., 1988) and RNA polymerase (Fong and Surzycki, 1992) are scattered over the genome, presumably because of numerous rearrangement events. Cyanelles, chromophytic and rhodophytic plastids and the cyanobacterium Anabaena sp. strain PCC 7120 share a 4 bp atpF/atpD overlap (see Pancic et al., 1992). Since the atpD gene is lacking from land plant plastids, it is not possible to determine whether an overlap also existed in this lineage. Conversely, cyanobacteria, cyanelles (D. A. Bryant and V. L. Stirewalt, personal communication), cryptophytes (Douglas, unpubl.) and chromophytes (Jouannic et al., 1992; Leitsch and Kowallik, 1992) do not contain overlapping atpB/atpE genes whereas some, but not all, land plants do. It thus seems that the atpB/atpE overlap is a derived character whereas the atpF/atpD overlap is an ancestral one (Pancic et al., 1992).

4. Photosystem Operons The genes for Photosystems I and II have been extensively characterized in cyanobacteria (see Sherman et al., 1987; Bryant et al., 1990; Bryant, 1992), land plant plastids (see Sugiura, 1992), cyanelles (Bohnert and Löffelhardt 1992; also see chapter 4), and the plastids of rhodophytes (see Reith and Munholland, 1993) and chromophytes (Shivji et al., 1992; Kowallik, 1993). Many cyanobacterial gene clusters such as psaAB, psbCD and psbEFLJ have been retained in plastid genomes, whereas others such as those for phycobiliproteins and Rubisco have been lost or reduced.

Once again, one sees the post-endosymbiotic assembly of certain plastid photosynthetic gene clusters from disparate cyanobacterial operons. In land plants, cyanelles and the rhodophyte P. purpurea, the genes psbB, ORF31 , psb N and psbH are clustered whereas neither psbB, psbH nor psbN are linked in most cyanobacteria or Prochlorothrix hollandica (Golden et al., 1993). Linkage of psbN and psbH has been shown in Synechocystis sp. strain PCC 6803, however, indicating that this may be the ancestral condition (Mayes and Barber, 1991). Thus, genes from at least three operons have come together in plastids. Such a complex event is very unlikely to have been duplicated and supports a single origin of all plastids. A large ORF (283–290 amino acids) found upstream of psbB in both the cyanelle and P. purpurea (Fig. 5) may have been present in the ancestral plastid but lost in the green plastid lineage. A rearrangement has presumably resulted in the movement of psbH to a position upstream in the cluster from the cyanelle (D. A. Bryant and V. L. Stirewalt, personal communication). Once again, a pattern in the organization of gene clusters from different lineages is evident. The petBD genes of cyanobacteria and P. hollandica are distant from the psbB gene, as in cyanelles (D. A. Bryant and H. J. Bohnert, personal communication) and chromophyte plastids (Kowallik, 1993). In land plant plastids, petB and petD are located downstream of the psbBNH cluster and posttranscriptional processing of the primary transcript results in the differential accumulation of the psbB and the psbH and petBD gene products (Barkan, 1988). Interestingly, conserved 93-bp perfect repeat sequences are found upstream of the psbH and pet BD genes of P. hollandica that may result in a similar co-regulation of these genes as is found in land plants (Greer and Golden, 1992). The structure of the psbDC operons in cyanobacteria and plastids also supports a single origin of plastids from cyanobacteria. Recent studies have shown that a 17-bp overlap between the two genes is present in cyanobacteria and the plastids of rhodophytes (Maid and Zetsche, 1992), chromophytes (see Kowallik, 1993), and land plant plastids.

5. Other The ndh genes of land plant plastids are found in two main clusters: 5' ndhH-ndhA-ndhI-ndhG-ndhE-psaCndhD 3' and 5' ndhC-ndhK-ndhJ 3' (terminology of

Chapter 5 Plastid Evolution

Sugiura, 1992). Recentstudies have shownthatsome of these genes are found in the same relative order on cyanobacterial genomes. For example, the arrangements 5' ndhC-ndhK-ndhJ3' and 5' ndhA-ndhI-ndhGndhE 3' are found in Synechocystis sp. strain PCC 6803, and the psaC and ndhD genes are clustered but transcribed from opposite strands (Steinmüller et al., 1989; Ellersiek and Steinmüller, 1992). The arrangement 5' ndhA-ndhI-ndhG-ndhE 3' is also found in the filamentous cyanobacterium P. boryanum (Takahashi et al., 1991). The absence of ndh genes from the genomes of all of the investigated non-green plastids is very puzzling, and must represent a common loss very soon after their divergence from the green plastid lineage.

C. Plastid Transcription Although the eubacteria-like RNA polymerase subunits of plastids are expressed and transcription is

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from eubacteria-like promoters, information is too sparse to link plastids to any particular group of eubacteria on the basis of polymerase structure and promoter sequence similarity alone. Consensus promoter motifs have been identified for many land plant plastid genes, but very few promoters have been identified from other plastids. In both cyanobacteria and rhodophytes, common promoter motifs (TGTTA at–35; TCTTTTA at–10) have been found upstream of the transcription initiation sites of phycoerythrin genes (Roell and Morse, 1993), and reinvestigation of this region in Cryptomonas sp. (Reith and Douglas, 1990) revealed a similar motif (TCTTA at–35; GCTTTA at–10). The determination of the transcription initiation sites and promoters from more plastid genes may show further similarities between cyanobacteria and plastids. Transcription maps of chloroplast genomes (Woodbury et al., 1988; Kanno and Hirai, 1993) have shown that most of the plastid genome is transcribed and that many genes

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102 are transcribed polycistronically (as in eubacteria). Gene expression in land plantplastids is thought to beregulatedmainlypost-transcriptionally. However, the recent detection of genes encoding transcriptional regulatory proteins in cyanelle genomes (D. A. Bryant, personal communication) and the plastid genomes of non-green plastids (Kessler et al., 1992; Douglas 1992b; Reith and Munholland, 1993) suggests that some gene expression may be controlled at the transcriptional level in these plastids, as in eubacteria. Translational control may be a phenomenon that is mainly seen in green plastids.

D. Plastid Introns Introns are so rare among eubacteria that it is difficult to find a link between plastids and any particular group of eubacteria using this character. The only introns identified in eubacteria are the group I introns gene (Kuhsel et al., 1990) and found in the the and genes (Reinhold-Hurek and Snub, 1992). The intron in is very ancient and was presumably present in the ancestor of chloroplasts since it has been identified in some, but not all, algal plastid genomes. No group I introns have yet been demonstrated in any other genes from the cyanelle genome (D. A. Bryant, personal communication) or the plastid genomes from nongreen algae (see Douglas, 1993; Reith and Munholland, 1993) and, with the exception of a single rhodophyte gene, no group II introns have been detected in plastid genomes of non-green algae. The intron in the rhodophyte Rhodella violacea (Bernard et al., 1992) is found near the 5' end of the rpeB gene and although it possesses some of the features ofgroup II introns, it is considerably shorter and lacks several of the secondary structural features characteristic of this class of intron. Since this intron has not been detected in homologous genes from cyanobacteria (see Bryant, 1992), acryptophyte (Reith and Douglas, 1990) or two other rhodophytes (Apt and Grossman, 1993; Roell and Morse, 1993) it may represent a case of intron gain. Indeed, with the exception of the group I intron found in the gene (which was obviously present at an early stage of evolution), careful analysis of existing data indicates that introns in other plastid genes probably arose late in evolution (see Palmer and Logsdon, 1991). As mentioned previously (Section II B 1), the group II introns in the rRNA spacer-located genes for

and of land plants and certain green algae are homologous and have been used as a phylogenetic indicator of the ancestor of land plants (Manhart and Palmer, 1990). However, proteinencoding genes from the plastids ofgreen algae, land plants and euglenoids contain both group I and group II introns that are not evolutionarily related since, in a given gene, they may occur in different locations or even be of a different type. These introns probably arose fairly late in chloroplast evolution in the green algal progenitors of land plants. The presence of a group II ‘twintron’ (a group II intron inserted within another group II intron) in the plastid psb F gene from E. gracilis (Copertino and Halick, 1991) suggests that group II introns can spread to new locations. Protein-encoding genes from the plastid of E. gracilis are also characterized by multiple introns of a novel category (Christopher and Hallick, 1989) that resemble group II introns but are smaller and have a very high A + T content. The distribution of introns in plastid genes may give clues about the evolution of the plastid genome. A progressive increase in intron possession is seen in the genes of the str operon (see Douglas, 1991), that contributes to our understanding of plastid relationships. In the ancestral eubacterial state, there were probably no introns in any of the genes of the str operon (Fig. 3). There are no introns in the cyanelle (Kraus et al., 1990), rhodophyte (Reith and Munholland, 1993) or cryptomonad (Douglas, 1991) operons but the tufA gene of E. gracilis contains two introns. In C. reinhardtii the operon structure has been disrupted, and in land plants not only are there introns in the rps12 gene, but the exons are on opposite strands of the genome and require transsplicing for correct expression (Zaita et al., 1987).

E. Plastid Gene Sequences Comparison of gene sequences is a potentially useful method forestimating evolutionaryrelatedness. While it is not within the scope of this chapter to discuss the various parameters affecting phylogenetic inference (see Felsenstein, 1988), several factors are of vital significance. It is important that the genes being compared: 1) are true homologs, i. e., they have shared a common ancestor; 2) can be aligned unambiguously; 3) have a large number of positions for comparison; 4) are part of a reasonably broad database; 5) are subject to similar rates of nucleotide substitution; and 6) have not been laterally transferred.

Chapter 5 Plastid Evolution The following discussion will concentrate on some molecules for which the preceding criteria hold. However, severalpointsmustbeconsideredinrelation to these criteria. Firstly, chloroplast genes have an approximately five-fold lower substitution rate than nuclear genes (Wolfe et al., 1987) and substitution rates also differ between taxonomic groups (Britten, 1986). Secondly, substitutional bias may distort phylogenetic analyses when genes with different A + T contents are being compared (Lockhart et al., 1992; Howe et al., 1993). Since cyanobacteria display a broad range of A + T compositions, and plastid and cyanelle genes generally display a high A + T composition, this bias may obscure true relationships, depending on the taxa included in the study and the gene being analyzed. Thirdly, well-documented insertions have been implicated as more reliable indicators of evolutionary relatedness than similarities in nucleotide or amino acid sequences (Meyer et al., 1986). However, the rarity of such insertions limits their usefulness and the significance attributed to them can sometimes cause misleading conclusions, as in the case of the insertion in the carboxy terminus of the psbA gene product (D1 protein) of Photosystem II (see Gray, 1989). Thephylogeniesbased on Rubisco sequences will be discussed albeit they may be misleading in part due to the likelihood of lateral transfer. For a recent review and presentation ofgene trees based on plastid SSU rRNA, atpB, psbA, rbcL, rbcS, and tufA genes, the reader is referred to Morden et al. (1992). Only representative trees inferred from SSU rRNA and rbcL gene sequences will be presented in this discussion.

1. Ribosomal RNA a. 5S rRNA Conflicting results arise from phylogenetic analyses of plastid 5S rRNA sequences, undoubtedly because the molecules are so short and highly constrained (Halanych, 1991; Steele et al., 1991). Plastids from rhodophytes have been shown alternatively both to cluster with green plastids (van den Eynde et al., 1988) and to form a stable group separate from green plastids (Sommerville et al., 1992). In the latter study, the position of the cyanelle could not be reliably inferred and the topology of the tree was dependent on the sequence alignment and the method of tree-building. An earlier study (Maxwell et al., 1986) showed cyanelle 5S rRNA to have the highest

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percent similarity (78%) to the cyanobacterium Synechococcus lividans II and a slightly lower percent similarity (77%) to the cyanobacterium S. lividans III and the liverwort Marchantia polymorpha.

b. SSU rRNA Small subunit ribosomal RNA (SSU rRNA) sequences appear to be reliable molecules for the investigation of plastid origins since they are present in all organisms; sequence alignment is relatively straightforward (Gray et al., 1984); they contain a large number ofnucleotide positions for comparison (approximately 1500); a large database exists (over 600 sequences); they appear to exhibit a constant rate of nucleotide substitutionamongthetaxainvestigated (excluding Euglena and Chlamydomonas species); and there is no evidence of lateral transfer. SSU rRNA sequence comparisons showed Heliobacterium chlorum to be a member of the Gram-positive bacteria (sensu Woese, 1987) and to bear no close relationship with plastids from the chrysophyte Ochromonas danica (Witt and Stackebrandt, 1988), contrary to predictions based on chlorophyll content (Table 1). Similar analyses indicated that the prochlorophyte Prochlorothrix hollandica bore no specific relationship to landplant, chlorophyte or euglenoid plastids, or to the cyanelle of Cyanophora paradoxa, but rather it fell within the cyanobacterial line of descent (Giovannoni et al., 1988; Turner et al., 1989). This is further supported by phylogenetic analyses of SSU rRNA (Urbach et al., 1992) and partial rpoC1 (Palenik and Haselkorn 1992) gene sequences from the three known prochlorophyte genera Prochloron didemni and the more recently isolated P. hollandica (BurgerWiersma et al., 1986) and Prochlorococcus marinus (Chisholm et al., 1988). These analyses indicate that the prochlorophytes are spread throughout the cyanobacterial assemblage and show no specific affiliations to the chlorophyte lineage (for further discussion of the prochlorophytes and their properties, see chapter 3). These studies raise the possibility that either Chl b has arisen multiple times, as has been suggested by Ragan and Chapman (1978) based on its distribution in eucaryotes, or that the ability to synthesize Chl b has been laterally transferred. An alternative explanation put forward by Bryant (1992) is that the common ancestor of cyanobacteria and prochlorophytes had the ability to synthesize both Chl b and phycobilins, possibly under different

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environmental conditions. If such a eubacterium existed, it is possible that it was the ancestor of plastids, thus obviating the need for the independent evolution of Chl b in several different plant lineages (Bryant, 1992). With the recent indication of a Chl clike pigment in the prochlorophyte P. marinus (Chisholm et al., 1992) and in the plastid of the Chl b-containing alga Mantoniella squamata (Wilhelm, 1987), it is even conceivable that Chl c was also present in the common ancestor of plastids and was then lost from cyanelles and the plastids of rhodophytes and most chlorophytes (Douglas, 1992b; Kowallik, 1993). Relationships among plastids have also been investigated using SSU rRNA sequences (Douglas

Susan E. Douglas and Turner, 1991; Giovannoni et al., 1993). There is a general agreement that, although all plastids seem to have arisen from a single source in the cyanobacterial assemblage, there was an early split between the green and the non-green lineages and that the SSU rRNA of E. gracilis is most closely related to that of chromophytes. The study of Douglas and Turner (1991) also shows the cyanelle to be the earliest branch of the non-green lineage (Fig. 6), whereas the study of Giovannoni et al. (1993) shows the cyanelle to form the earliest branch of all the plastids. Phylogenetic analysis of SSU rRNA sequences excluding a cyanellar representative (Markowicz and Loiseaux-DeGöer, 1991) still supported all plastids as arising from within the

Chapter 5 Plastid Evolution cyanobacteria, but found the putative common ancestor of rhodophytes and chromophytes differed from that of green algae and land plants. However, these two putative ancestors originated in such close proximity that their branching order could not be determined. Unfortunately, all three ofthese studies used different methods of phylogenetic reconstruction, and it is difficult to resolve which (if any) are correct. However, the inclusion of a cyanelle sequence does seem to be important for recovering the correct tree topology by overcoming artifacts induced by sequences that have different rates of nucleotide substitution (see Giovannoni et al., 1993).

c. LSU rRNA Due to their larger size, LSU rRNA sequences may allow better resolution of the relationships between and among eubacteria and plastids. However, with the exception of one sequence from a brown algal plastid (Loiseaux-DeGöer, 1992), all other representatives are from green plastids, and phylogenetic analyses based on this molecule must await the acquisition of further sequence information.

2. ATP Synthase Subunit Beta (atpB) As previously mentioned, the genes encoding the subunits of the ATP synthase are found in two distinct clusters, the atpA cluster and the atpB cluster, in photoautotrophic eubacteria and plastids. Substantial sequence data exist for the atpB gene from which phylogenetic inferences may be made. The conclusions mirror those derived from SSU rRNA sequences in that all plastids arise monophyletically from the cyanobacteria and there is an early separation of green plastids, non-green plastids andcyanelles (Douglas, 1992a; Leitsch andKowallik, 1992; Morden et al., 1992).

3. Photosystem II Protein D1 (psbA) The psbA gene, encoding the D1 thylakoid protein from the Photosystem II reaction center, has been sequenced from several cyanobacteria (including Prochlorothrix hollandica) and plastids. The deduced amino acid sequences show a 7-amino acid deletion at the carboxy terminus of the protein that is shared by land plants, green algae and prochlorophytes but that is absent in other cyanobacteria, the cyanelles of C. paradoxa and the plastids of rhodophytes and

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chromophytes (Winhauer et al., 1991). This feature has been cited as evidence in support of the polyphyletic origin of plastids (Janssen et al., 1989; Morden and Golden, 1989; Maid et al., 1990; Scherer et al., 1991; Winhauer et al., 1991) although an alternative explanation that embraces a monophyletic origin was also offered by Scherer et al. (1991) and a recent phylogenetic analysis of this gene is also consistent with a monophyletic origin (Morden et al., 1992). It is interesting that the psbA gene of E. gracilis is truncated at a position very near to the gap (see Scherer et al., 1991). Thus this portion of the gene, to which so much significance has been attributed, may be highly variable and of dubious value as a phylogenetic marker. The D1 protein appears to restrict amino acid exchanges to a maximum of 19% (Winhauer et al., 1991) probably because of functional constraints on this structural component of the Photosystem II reaction center complex. Due to this high conservation, it is doubtful whether enough phylogenetically informative sites are present to be useful. In fact, in the case ofthe C. paradoxa sequence, the nucleotide similarity places it closest to green plastids, whereas the seven amino acid deletion places it with cyanobacteria, rhodophytes and chromophytes (Janssen et al., 1989). Another factortobe considered is the occurrence of multiple copies of this gene in various taxa. Several non-identical copies have been found in cyanobacteria and two non-identical copies have also been found in P. hollandica (see Golden et al., 1993). Landplantplastid genomes usually contain a single psbA gene although tandemly duplicated psbA genes have been found in two species of pine (Lidholm et al., 1991). In the plastids ofcertain algae such as Olisthodiscus luteus (Reith and Cattolico, 1986), Ochromonas danica (Shivji et al., 1992), Chlamydomonas eugametos and C. moewusii (Lemieux et al., 1985), the psbA gene is contained on the inverted repeat and is thus likely to be subject to copy-correction and perhaps a lower rate of nucleotide substitution than those genes found in the singlecopy regions (Wolfe et al., 1987).

4. Ribulose-1,5-Bisphosphate Carboxylase (Rubisco) Ribulose-1,5-bisphosphate carboxylase is the enzyme responsible for the fixation of carbon dioxide in plastids and in photosynthetic eubacteria (see Chapters 14 and 15). The holoenzyme is made up of large and

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small subunits and the genes encoding these subunits, rbcL and rbcS respectively, have been cloned and sequenced from a variety of plants, algae and photosynthetic eubacteria. The gene for the large subunit is plastid-encoded in all plants investigated and is highly conserved. As with psbA, rbcL is found on the inverted repeat on Olisthodiscus luteus, Ochromonas danica, Chlamydomonas reinhardtii and C. moewusii (Reith and Cattolico, 1986; Shivji et al., 1992; Lemieux et al., 1985). The gene for the small subunit is less well-conserved and is found on the plastid genome in non-green plants but in the nuclear genome in land plants, green algae and E. gracilis. The utility of Rubisco sequences as a

Susan E. Douglas phylogenetic indicator in land plants hasbeenrecently reviewed (Clegg, 1993). However, its use in discriminating deeper branchpoints may be obscured by the possibility that there was lateral transfer of the gene early in the evolution of plastids. Phylogenetic analyses of either rbcL (Assali et al., 1990; Douglas et al., 1990; Valentin and Zetsche, 1990a, b, c; Morden and Golden, 1991; Singh, 1991) or rbcS (Boczar et al., 1989; Assali et al., 1991; Morden and Golden, 1991) sequences showed the plastids of non-green plants to be most closely related to the proteobacteria, whereas the plastids of green plants and algae were most closely related to cyanobacteria and the proteobacteria (Fig. 7). While

Chapter 5 Plastid Evolution such evidence may be interpreted to mean plastids arose polyphyletically from two separate groups of bacteria, the possibility exists that the Rubisco genes were laterally transferred at some point prior to endosymbiosis or shortly afterward, in the lineage leading to rhodophytes (see Douglas, 1992b; Morden et al., 1992). The possibility of lateral transfer in purple bacteria (Dickerson, 1980), the existence of a reverse transcriptase-like sequence in the plastid genome of a green alga (Kück, 1989), of Rubisco genes on a transmissible plasmid in the proteobacterium Alcaligenes eutrophus (Andersen and Caton, 1987) and of plasmids in rhodophytes (Goff and Coleman, 1988; Villemur, 1990) lend credence to such a suggestion. An alternative explanation is that the endosymbiont(s) participating in the primary endosymbiotic event(s) possessed chimaeric genome(s) with genes from a number of different eubacterial sources (Assali et al., 1990; Markowicz and Loiseaux-DeGöer, 1991; Loiseaux-DeGöer, 1992; Martin et al., 1992). Investigation of Rubisco genes from a wider variety of cyanobacteria may determine whether extant species contain genes from other eubacteria and perhaps resolve the question of when lateral transfer might have occurred.

5. Elongation Factor Tu (tufA) The tufA gene, encoding elongation factor Tu, is plastid-encoded in algae but has undergone transfer to the nucleus within the green algal lineage leading to land plants (Baldauf and Palmer, 1990). Phylogenetic analysis of the derived amino acid sequences showed plastids and the cyanelle of C. paradoxa arising from cyanobacteria (Baldauf et al., 1990, see Morden et al., 1992), in agreement with trees based on SSU rRNA sequences. A major difference,however,istheclustering oftheeuglenoids Astasia longa and E. gracilis with green algae, a feature also characteristic of Rubisco-based trees.

6. Others In addition to the four completely sequenced land plant plastid genomes, the plastid genomes from P. purpurea, Cryptomonas sp. and E. gracilis and the cyanelle genome from C. paradoxa have been extensively sequenced. Phylogenetic trees have been constructed using the derived amino acid sequences of ribosomal protein genes (Evrard et al., 1990). However, such analyses suffer from the small size of

107 both the genes and the database. Some ribosomal protein genes have been sequenced from the cyanelle of C. paradoxa (see Löffelhardt et al., 1989; Bohnert and Loffelhardt, 1992; for more information on the cyanelles of C. paradoxa, see Chapter 4) and the plastids of E. gracilis (Christopher and Hallick, 1988; Christopher et al., 1988), Cryptomonas sp. (Douglas and Durnford, 1990; Douglas, 1991), Gracilaria tenuistipata (Kao and Wu, 1990; Kao et al., 1990), Cyanidium caldarium (Maid and Zetsche, 1992) and several land plants (see Subramanian et al., 1991) but the number of eubacterial, particularly cyanobacterial, representatives is small. Since the rate of evolution of different ribosomal proteins varies (Wittman-Liebold et al., 1990), specific proteins can be either plastid- or nuclear-encoded depending on the organism, and nuclear-encoded proteins may be of mitochondrial (Martin et al., 1990) rather than plastid (Smooker et al., 1990) origin, caution should be exercised when inferring phylogenetic relationships based on these proteins. Purple bacteria and cyanobacteria may have shared a common ancestor that possessed two types of reaction centers from which Photosystems I and II evolved (see Olson and Pierson, 1987). Studies ofthe genes involved in these photosystems may therefore help to identify the nature of the photosynthetic eubacterium involved in the formation of plastids. The Photosystem II psbA and psbD gene products are related to the L and M reaction center proteins of purple bacteria whereas the Photosystem I reaction centers are similar to those of green sulfur bacteria. Thus far, the sequence of psaB has been determined for the brown alga Pylaiella littoralis (Assali and Loiseaux-DeGöer, 1992) and the sequence of psaE has been determined for the red alga Porphyra purpurea (Reith, 1992). An unrooted phylogenetic tree based on five psaB sequences (Assali and Loiseaux-DeGöer, 1992) shows P. littoralis to be most closely related to the cyanobacterium Synechococcus sp. strain PCC 7002, and on a separate branch from E. gracilis and land plant plastids. Recently, the psaA and psaB sequences have been determined from Synechocystis sp. strain PCC 6803 (Smart and Mclntosh 1991), Anabaena variabilis strain ATCC 29413 (Toelge et al., 1991) and Synechococcus vulcanus (Shimizu et al., 1992). Additional sequence data from reaction center genes from photosynthetic bacteria should be useful in assessing the evolutionary origins of plastid genomes.

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F. Nuclear Gene Sequences Plant nuclear-encoded genes that may have been transferred from the endosymbiont during the establishment of the plastid as an organelle may also give clues about plastid origins. Plastids and cyanobacteria share a number of basic biochemical pathways and several plant enzymes have chloroplastic and cytosolic isozyme pairs (for reviews, see Weeden, 1981; Fothergill-Gilmore and Michels, 1993). In most cases, the chloroplast-specific isozyme exhibits a higher degree of similarity to the cyanobacterial enzyme than to its cytosolic counterpart, indicating that it originated from the endosymbiont genome. However, the isozymes of triosephosphate isomerase and perhaps other enzymes appear to have originated by duplication of an ancestral nuclear gene (Feierabend et al., 1990). A number of cyanobacterial genes that have nuclear-encoded plastid counterparts have recently been sequenced. These include genes involved in pigment biosynthesis, such as glutamate 1-semialdehyde aminotransferase (Grimm et al., 1991), phytoene synthase (Chamovitz et al., 1992) and phytoene desaturase (Chamovitz et al., 1991); fatty acidbiosynthesis(seeWadaetal, 1990); carbohydrate metabolism such as glyceraldehyde 3-phosphate dehydrogenase (GAPDH; Cerff et al., 1992), glucose6-phosphate dehydrogenase (Scanlan et al., 1992) and ADP-glucose pyrophosphorylase (Charng et al., 1992); electron transport such as ferredoxin-NADP oxidoreductase (Schluchter and Bryant, 1992); translation such as ribosomal proteins L10 and L12 (Sibold and Subramanian, 1990); nutrient uptake (Laudenbach and Grossman, 1991; Omata, 1991; Luque et al., 1992); and metal sequestration (Briggs et al., 1990; Shi et al., 1992). With the exception of GAPDH (see below), the databases for many ofthese genes are very small. However, future sequence determinations may allow more of these molecules to be used for phylogenetic analysis. Plants contain two forms of GAPDH, one active in glycolysis and located in the cytosol (encoded by gapC) and one active in the Calvin cycle that is located in the chloroplast (encoded by gapA and gapB). Both chloroplastic and cytosolic enzymes are nuclear-encoded but sequence comparisons show the chloroplastic GAPDHs to be more similar to GAPDHs from thermophilic bacteria than to their cytosolic counterparts (Shih et al., 1986; Brinkmann

Susan E. Douglas et al., 1987), providing evidence of gene transfer from the genome of the endosymbiont to the genome of the host. The GAPDH genes of land plant chloroplasts, gapA and gapB, are approximately 80% similar to one another, and are thought to have arisen from a duplication event prior to the emergence of angiosperms ( Brinkmann et al., 1989). Until recently, no GAPDH sequences have been available from cyanobacteria or non-green plastids. Unexpectedly, the cyanobacterium Anabaena variabilis has been shown to contain three genes encoding GAPDH (Cerff et al., 1992). Phylogenetic analysis shows one of the cyanobacterial GAPDH genes (gapA) to be more closely related to plastid GAPDHs than those from other eubacteria, but a second gene (gapA') is most closely related to those from other eubacteria. A third gene (gapC) is most closely related to the glycolytic, i. e. cytosolic, GAPDH (see Section III C 2). The chloroplastic GAPDH of the rhodophyte Gracilaria verrucosa is encoded by a single nuclear gene (gapA) and phylogenetic analysis of these sequences shows rhodophyte plastids as a sister group to land plant plastids (Zhou and Ragan, 1993). This relationship is confirmed by studies of the gapA and gapC genes from another rhodophyte, Chondrus crispus (Liaud et al., 1992). Together with the fact that the G. verrucosa GAPDH transit peptide has the Block II consensus motif of land plant GAPDH transitpeptides (Brinkmann et al., 1989) suggesting common ancestry of the genes, these data indicate that the two plastid groups arose from a single endosymbiotic event (Zhou and Ragan, 1993). III. Secondary Endosymbiosis in Plastid Evolution

A. Electron Microscopic Studies The three membranes surrounding the plastids of Euglena sp. have been variously interpreted to represent the remains of a phagocytosed green alga (Gibbs, 1978) or green algal plastid (Whatley et al., 1979). Similarly, the four membranes surrounding chromophyte and cryptophyte plastids were thought to represent the vestiges ofa phagocytosed eucaryotic endosymbiont (Gibbs, 1981a; Whatley et al., 1979). Electron microscopic examination of nuclear envelopes in several of these algae (Gibbs, 1962) showed continuity between the outer envelope ofthe

Chapter 5 Plastid Evolution nucleus and the outermost membrane of the chloroplast endoplasmic reticulum (CER). Subsequent analyses showed ribosomes on this membrane and vesicles between the CER and theplastid envelope proper (Gibbs, 1979, 1981b). It was suggested that nuclear-encoded proteins destined for the plastid were synthesized on these membrane-bound ribosomes andtransported intotheplastidviavesicles. Using a variety of ultrastructural methods, characteristics indicativeof secondaryendosymbiotic events have been investigated in cryptomonads and Chlorarachnion (for review, see Sitte et al., 1992). These included the demonstration of DNA in the nucleomorph by DAPI-staining and anti-DNA antibodies, and the presence of RNA in the ‘fibrillogranular body’ of the nucleomorph by the RNase-gold method. All of this information suggests that the nucleomorph is a remnant ofan endocytosed eukaryotic nucleus.

B. Hybridization Studies The possibility of secondary endosymbiosis as the evolutionaryprocess responsible forthe formation of cryptomonad plastids has been further investigated by in situ hybridization using probes specific for eubacteria-like or eucaryote-like rRNA on sections prepared for electron microscopy (McFadden, 1990a, b). The data showed eucaryote-like rRNA in the nucleus, nucleomorph and periplastidal spacebetween the CER and plastid envelope, but eubacteria-like rRNA within the plastid proper. Similarly, chromosomes isolatedfromthenucleomorphandresolved by pulsed field gel electrophoresis were shown to encode eucaryote-like rRNA by Southern hybridization (Eschbach et al., 1991b).

C. Gene Sequences Whereas comparisons of plastid and eubacterial gene sequences can give clues regarding the eubacterial ancestor(s) thatgave rise tophotosynthetic eucaryotes in the primary endosymbiotic event(s), comparisons of nuclear-cytoplasmic genes can give an indication of the variety of host cells that may have been involved in both the primary and secondary endosymbiotic events. Two types of nuclear genes that have been extensively studied, those encoding ribosomal RNA and those encoding glyceraldehye3-phosphatedehydrogenase (GAPDH), arediscussed below.

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1. Ribosomal RNA

a. 5S rRNA Phylogenetic analyses of cytoplasmic 5S rRNA showed that algae didnot form amonophyletic group (Hori and Osawa, 1986;Hori et al., 1989; van de Peer et al., 1990). The rhodophytes, chromophytes, cryptophytes, dinoflagellates and green plants were all found on separate branches. Interestingly, C. paradoxa and E. gracilis shared a branch in these studies and also in earlier analyses by Maxwell et al. (1986).

b. SSU rRNA Similarly, phylogeneticanalysesofcytoplasmic SSU rRNA (containing more than ten times as many nucleotide positions as 5S rRNA) showed that algae were found on several different branches of the evolutionary tree. Chromophytes occupied aseparate branch from green algae, dinoflagellates and euglenoids, and were most closely related to oomycetes (Gunderson et al., 1987). Rhodophytes were found to occupy a separate branch from dinoflagellates, chromophytes, euglenoids and chlorophytes (Bhattacharya et al., 1990; Bird et al., 1991). A recent analysis of dinoflagellate sequences has shown them to be most closely related to the Apicomplexa (Sadler et al., 1992). Definitive proof of secondary endosymbiosis has come from the phylogenetic analysis of nuclear and nucleomorph SSU rRNA sequences from cryptomonads (Douglas et al., 1991; Eschbach et al., 1991a; Maier et al., 1991). In these studies, the nucleomorph sequence occupied abranchseparatefrom thenuclearsequence, indicating that two separate eucaryotes contributed to the formation ofthe cryptomonad cell (Fig. 8). The nucleomorphsequence wasfound onthesamebranch as rhodophyte sequences, confirming the identity of the eukaryotic endosymbiont as red algal-like (Gibbs, 1981 a). In addition, the nuclear sequence occupied a branch separate from chromophyte sequences, indicating that chromophytes have not evolved directly from cryptomonads, contrary to ahypothesis advocated by Cavalier-Smith (1986).

c. LSU rRNA The LSU rRNA contains approximately twice the

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number of nucleotide positions as the SSU rRNA, but the database of complete LSU sequences is small and few complete sequences from photosynthetic eucaryotes have been determined. Phylogenetic trees have been based on partial sequences, determined for specific regions by reverse transcription of the RNA molecule (Baroin et al., 1988), and thus may not contain as much phylogenetic information as complete SSU rRNA sequences. However, as with evolutionary trees based on SSU rRNA sequences, there was a stable separation between chlorophytes, rhodophytes, chromophytes and dinoflagellates (Perasso et al., 1989). The grouping of cryptophytes and rhodophytes is not in agreement with trees based on S SU sequences (Douglas et al., 1991), unless the rRNA molecule sequenced was from the nucleomorph rather than the nucleus. This could occur if the primers used for reverse transcription preferentially selected the nucleomorph rRNA over the nuclear rRNA.

2. GAPDH As mentioned previously (see Section II F), the gapC gene from the cyanobacterium Anabaena variabilis shows the closest relationship to the genes for the glycolytic form of GAPDH (Cerff et al., 1992). This has important implications for the origins of plastids, since it is possible that higher eucaryotes possess a laterally transferred eubacterial gene for the glycolytic as well as the Calvin cycle GAPDHs. If this gene originated from a cyanobacterial endosymbiont destined to become a plastid, the possibility exists that higher eucaryotes became secondarily nonphotosynthetic through loss of their plastids. The demonstration of an extrachromosomal element of presumed plastid origin in Plasmodium sp., a member of the non-photosynthetic Apicomplexa (Howe, 1992; for review, see Palmer, 1992) lends support to this idea. Since the dinoflagellates appear to be most

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closely related to the Apicomplexa (Sadler et al., 1992), it is possible that this extrachromosomal element is the remnant of a plastid genome that was present in the common ancestor of dinoflagellates and the Apicomplexa. In addition, the sharing of an intron at identical positions in both the maize chloroplast GAPDH and a nematode cytosolic GAPDH genes (Quigley et al., 1988), suggesting that it was present in their common ancestor, is consistent with the view that this common ancestor was photosynthetic. Nuclear genomes contain DNA sequences of chloroplast and mitochondrial origin (Timmis and Scott, 1983), but the fact that they may be more highly chimaeric than previously supposed presents another important factor in the interpretation of gene trees. IV. Conclusions and Future Prospects Comparisons of gene arrangement, expression and sequences from photosynthetic eubacteria and plastids have consistently demonstrated the cyanobacterial ancestry of plastids. However, the eubacterial representatives have been limited to a few wellcharacterized species of cyanobacteria and have seldom included representatives of more divergent cyanobacterial genera or of other photosynthetic groups such as green and purple sulfur bacteria. Since phylogenetic analyses of different genes have indicated that plastid genomes may be chimaeric, having received genetic contributions from more than one eubacterial source, our knowledge of the evolution of plastids would be greatly enhanced by the study of genes from more divergent cyanobacterial taxa and other photosynthetic eubacteria. The preservation of gene clusters in widely separate plant lineages and the post-endosymbiotic assembly of certain operons point toward a single monophyletic origin of plastids (Fig. 9). A pattern can be seen wherein gene content is reduced in the green plastids compared to the non-green plastids. Similarly, the progressive gain of introns and scrambling of operons in green plastids relative to non-green plastids is evident. It appears that rhodophyte, cryptophyte and chromophyte plastids are more closely related to each other than to chlorophyte and land plant plastids whether SSU rRNA, Rubisco, atpB, tufA or psbA sequences are compared. However, it is uncertain whether the plastids of euglenophytes and the cyanelle of C. paradoxa are more closely related to the green

or the non-green lineage. Although gene sequences cannot yet unequivocally prove monophyletic vs. polyphyletic origins, improved statistical methodologies and ever-increasing databases may resolve this debate in the future. Phylogenetic studies using rRNA sequences have confirmed suggestions, based on ultrastructural information, that secondary endosymbiotic events were responsible for the formation of algae containing complex plastids and demonstrated that the secondary endosymbiont giving rise to cryptophytes and chromophytes was red alga-like (Fig. 9). Increased information on additional molecular characters are beginning to give clues about the relationships between photosynthetic and non-photosynthetic eucaryotes. Molecular markers constitute a complementary approach to the large array of morphological and biochemical markers and with the acquisition of more data, an evolutionary picture of the origin of plastids congruent with both approaches will emerge. Acknowledgments I express my gratitude to Ford Doolittle for introducing

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me to the cyanobacteria and their importance in evolutionary biology. I thank Drs Michael Reith, Don Bryant, and Veronica Stirewalt for sharing data prior to publication, and Mark Ragan, Sandie Baldauf and Michael Reith for helpful discussions and reviews of this manuscript. The support of fundamental research by the National Research Council in past years has been instrumental in providing an environment in which studies of the evolutionary origins of algae have flourished, and is gratefully acknowledged. This is NRCC publication number 34841. References Andersen K and Caton J (1987) Sequence analysis of the Alcaligenes eutrophus chromosomally encoded ribulose bisphosphate carboxylase large and small subunit genes and their gene products. J Bacteriol 169: 4547–4558 Anderson SL and McIntosh L (1991) Partial conservation of the 5' ndhE-psaC-ndhD 3' gene arrangement of chloroplasts in the cyanobacterium Synechocystis sp. PCC 6803: implications for NDH-D function in cyanobacteria and chloroplasts. Plant Mol Biol 16: 487–499 Apt KE and Grossman AR (1993) Characterization and transcript analysis of the major phycobiliprotein subunit genes from Aglaothamnion neglectum (Rhodophyta). Plant Mol Biol 21: 27–38 Assali NE and Loiseaux-DeGöer S (1992) Sequence and phy logeny of the psaB gene of Pylaiella littoralis (Phaeophyta). J Phycol 28: 209–213 Assali NE, Mache R and Loiseaux-DeGöer S (1990) Evidence for a composite phylogenetic origin of the plastid genome of the brown alga Pylaiella littoralis (L) Kjellm. Plant Mol Biol 15: 307–315 Assali NE, Martin WF, Sommerville CC and Loiseaux-DeGöer S (1991) Evolution of the Rubisco operon from prokaryotes to algae: Structure and analysis of the rbcS gene of the brown alga Pylaiella littoralis. Plant Mol Biol 17: 853–863 Baldauf SL and Palmer JD (1990) Evolutionary transfer of the chloroplast tufA gene to the nucleus. Nature 344: 262–265 Baldauf SL, Manhart JR and Palmer JD (1990) Different fates of the chloroplast tufA gene following its transfer to the nucleus in green algae. Proc Natl Acad Sci USA 87: 5317–5321 Bancroft I, Wolk CP and Oren EV (1989) Physical and genetic maps for the genome of the heterocyst forming cyanobacterium Anabaena sp strain PCC 7120. J Bacteriol 171: 5940–5948 Barkan A (1988) Proteins encoded by a complex chloroplast transcription unit are each translated from both monocistronic and polycistronic mRNAs. EMBO J 7: 2637–2644 Baroin A, Perasso R, Qu L-H, Brugerolle G, Bachellerie J-P and Adoutte A (1988) Partial phylogeny of the unicellular eukaryotes based on rapid sequencing of a portion of 28S ribosomal RNA. Proc Natl Acad Sci USA 85: 3474–3478 Bernard C, Thomas JC, Mazel D, Mousseau A, Castets AM, Tandeau de Marsac N and Dubacq JP (1992) Characterization of the genes encoding phycoerythrin in the red alga Rhodella

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118 of Plants. Vol. 7B, pp 26–111. Academic Press, New York. Villemur R (1990) Circular plasmid DNAs from the red alga Gracilaria chilensis. Curr Genet 18: 251–257 Wada H, Sakamoto T and Murata N (1990) Cyanobacterial genes for plant-type desaturases of fatty acids. In: Quinn PJ and Harwood JL (eds) Plant Lipid Biochemistry, Structure and Utilization. Ninth International Symposium on Plant Lipids, Kent, England, pp 453–455. Portland Press Ltd., London Wasmann CC, Löffelhardt W and Bohnert HJ (1987) Cyanelles: Organization and molecular biology. In: Fay P and Van Baalen C (eds) The Cyanobacteria, pp 303–324. Elsevier, Amsterdam Weeden NF (1981) Genetic and biochemical implications of the endosymbiotic origin of the chloroplast. J Mol Evol 17: 133– 139 Whatley JM, John P and FR Whatley (1979) From extracellular to intracellular: The establishment of mitochondria and chloroplasts. Proc Roy Soc Lond B204: 165–187 Wilhelm C (1987) The existence of chlorophyll c in the Chl b– containing light-harvesting complex of the green alga Mantoniella squamata (Prasinophyceae). Bot Acta. 101:7–10 Winhauer T, Jäger S, Valentin K and Zetsche K (1991) Structural similarities between psbA genes from red and brown algae. Curr Genet 20: 177–180 Witt D and Stackebrandt E (1988) Disproving the hypothesis of a common ancestry for the Ochromonas danica chrysoplast and Heliobacterium chlorum. Arch Microbiol 150: 244–248 Wittman-Liebold B, Köpke AKE, Arndt E, Krömer W, Hatakeyama T and Wittman H-G.(1990) Sequence comparison and evolution of ribosomal proteins and their genes. In: Hill WE, Dahlberg A, Garrett RA, Moore PB, Schlessinger D and Warner JE (eds) The Ribosome: Structure, Function and

Susan E. Douglas Evolution., pp 598–616. ASM Publications, Washington, DC Woese CR (1987) Bacterial evolution. Microbiol Rev 51: 221– 271 Woessner JP, Gillham NW and Boynton JE (1987) Chloroplast genes encoding subunits of the complex of Chlamydomonas reinhardtii are rearranged compared to higher plants: Sequence of the atpE gene and location of atpF and atpI genes. Plant Mol Biol 8: 151–158 Wolfe KH, Li W-H and Sharp PM (1987) Rates of nucleotide substitution vary greatly among plant mitochondrial, chloroplast and nuclear DNAs. Proc Natl Acad Sci USA 84: 9054–9058 Wolfe KH, Morden CW and Palmer JD (1991) Ins and outs of plastid genome evolution. Curr Opinions Genet Develop 1: 523–529 Wolfe K H , 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 Woodbury NW, Roberts LL, Palmer JD and Thompson WF (1988) A transcription map of the pea chloroplast genome. Curr Genet 14: 75–89 Yamada K and Yamamoto N (1992) Distribution of the liverwort chL (frxC) homologue among land plants. Plant Mol Evol Newsl 2: 38–40 Zaita N, Torazawa K., Shinozaki K and Sugiura M (1987) Trans splicing in vivo: Joining of transcripts from the ‘divided’ gene for ribosomal protein S12 in the chloroplasts of tobacco. FEBS Lett 210: 153–156 Zhou Y-H and Ragan MA (1993) cDNA cloning and characterization of the nuclear gene encoding chloroplast glyceraldehyde-3-phosphate dehydrogenase from the marine red alga Gracilaria verrucosa. Curr Genet 23: 483–489

Chapter 6 Supramolecular Membrane Organization Elisabeth Gantt Department of Botany, University of Maryland, College Park, MD 20742, USA Summary I. Introduction II. Localization of Intrinsic Membrane Proteins and Enzymes A. External Layers 1. Glycocalyx and Outer Membrane 2. Peptidoglycan Layer 3. Transporters 4. Carotenoid-Binding Proteins B. Plasma Membrane 1. Carbon Transporters 2. Respiratory Enzymes C. Plasma-Thylakoid Membrane Contacts D. Thylakoid Membranes 1. General Morphology 2. Phycobilisomes 3. Photosystem I 4. Photosystem II 5. Cytochrome Complex 6. Cytochrome 7. ATP Synthase 8. Ascorbate Peroxidase 9. Carotenoid Synthetic Enzymes 10. Other Complexes E. Photosynthetic Membrane Topography III. Future Focus Acknowledgments References

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Summary In cyanobacteria the outer membrane, plasma membrane and thylakoid membrane represent three structurally and functionally distinct membranes. Common themes are emerging from studies of thylakoid membranes which show that the major functional components, including Photosystem I, Photosystem II, cytochrome NADH dehydrogenase, cytochrome oxidase, and ATP synthase, exist as multisubunit complexes within the membrane. These integral membrane complexes traverse the 4–5 nm lipid bilayer and extend into the stromal and lumenal spaces–thus virtually doubling the membrane thickness. In situ it appears that cytochrome cytochrome oxidase, and Photosystem II preferentially occur as dimers and Photosystem I as trimers. Determination of the density of the supramolecular complexes per thylakoid area and the spatial relationships among them can provide useful insight into membrane functional. Calculations suggest that at least 50% of the intramembrane area is occupied by photosystems I and II, cytochrome and of ATP synthase alone. Topographic mapping of the membranes can be pursued with gold-antibody labeling with a resolution of approximately 15–20 nm. Studies on complexes of the outer membrane, the plasma membrane, and the D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 119–138. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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composition and influence of the peptidoglycan layer are still scarce and should be extended. The uniqueness of cyanobacterial membrane structure continues to emerge. Differences in the structure ofthe outer wall layers have been noted when compared to other eubacteria; however, essential thylakoid features are also found in prochlorophytes and eucaryotic algae oxygenic plants. These features emphasize that cyanobacteria and prochlorophytes should be considered a distinctive grouping of the eubacteria. I. Introduction Molecular biology techniques have provided extensive new information on the structure of cyanobacterial proteins in the last ten years, and this has resulted in considerable insight into the regulation and the interrelatedness of membrane components. This is particularly important because the cyanobacteria successfully compete with other organisms in many environments such as lithospheric conditions in the colonization of bare rock, subzero lakes in Antarctica, hotsprings, oceans, fresh-water lakes, and in nutrient-excessive sewage waters. As procaryotes they are viewed as simple organisms, yet when one considers the multiple metabolic and synthetic functions occurring in only one compartment, the cyanobacterial cell is far from simple. Although the general ultrastructural features have long been defined, the structural basis, i.e. the functional integration ofmulticomponentcomplexes within membranes, is still not well understood. Less than one dozen species, mostly moderate thermophiles (growing around 45 °C) or mesophiles (growing around 25 °C), have been studied in any detail. In these studies cyanobacteria are considered to be model systems for understanding green plants. The cyanobacteria are indeed a good model system for the basic structure of photosynthetic membranes, but they are appropriate for study in their own right because of the complexity of functions operating within a photosynthetic, oxygen-evolving cell. The two main goals of this chapter are to provide an overview of cyanobacterial membrane structure and to highlight the importance of topographic relationships of major protein complexes in photosynthetic membranes. Current models of membrane structures are generally presented as twodimensional representations with the size and shape of the proteins being derived from determinations Abbreviations: Chl – chlorophyll; Cyt – cytochrome; EM – electron microscopy; FAD – flavin adenine dinucleotide; FNR – erredoxin NADP+ oxidoreductase; PAGE – poly– acrylamide gel electrophoresis; PS–photosystem; SDS – sodium dodecylsulfate.

from SDS-PAGE and amino acid sequences (either deduced from gene sequences or determined directly). Yet, it is realized that components in biologically active membranes are involved in photon capture and transfer of excitation energy, transfer of electrons between donors and acceptors, and synthesis and functional insertion of apoproteins and cofactors. It is thus essential to view membranes in three dimensions, where distance between components, their density per area, and their directional orientation are essential parameters. This chapter is not intended to be a comprehensive review, but rather should be considered an update on the structural nature and the localization of supramolecular membrane complexes; it focuses on recent information, especially that having appeared since the publication of the last volume on cyanobacteria by Fay and VanBaalen (1987). Other chapters inthis volume will elaborate on the specifics of the structures of phycobilisomes (Chapter 7); photosystems II (Chapter 8); Photosystem I (Chapter 10); cytochrome (Chapter 9); ATP synthase (Chapter 11); and components of the respiratory apparatus (Chapter 13); other recent reviews include those on PS I by Golbeck and Bryant (1991), Almog et al. (1992), and Bryant (1992); on PS II by Pakrasi and Vermaas (1992) and Satoh (1992); and on phycobilisomes by Bryant (1991). In a recent review on the environmental effects on cyanobacterial structure Stevens and Nierzwicki-Bauer (1991) raised several important considerations on the interrelationship ofthe plasma membrane with the thylakoid membranes, and the points at which thylakoid membranes grow. These considerations are intimately related to the intended direction of the present chapter. The data on supramolecular membrane complexes selected for this chapter includes not only those of cyanobacteria but also those of membrane components which are expected to occur in cyanobacteria but have not yet been adequately described for this group. Rather than presenting an exhaustive review of available information on membrane complexes, the intention of the author was to highlight selected aspects of the latest research in order to allow other

Chapter 6 Membrane Organization researchers to consider membrane structures as part of an integrated system, i.e. membranes as functional entities. II. Localization of Intrinsic Membrane Proteins and Enzymes

A. External Layers 1. Glycocalyx and Outer Membrane Cyanobacterial cells are normally surrounded by an external, carbohydrate-enriched glycocalyx that

121 appears as a fibrous sheath. A glycocalyx is a simple means of protecting the cell against desiccation. In a liquid environment it probably also serves as a distinct boundary which retards solute loss and at the same time is effective in the sequestration of essential nutrients and solutes from the surrounding environment. The glycocalyx is closely associated with the outer membrane (see Figs. 1 and 2), and may, in fact, be involved in synthesis of the glycocalyx subunits. A periplasmic space between the outer membrane and the plasma membrane is divided by the peptidoglycan layer, thus creating an outer periplasmic space and an inner periplasmic space. It is very probable that these two spaces serve somewhat

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different functions. By EM the periplasmic space exhibits virtually no electron density (Fig. 2); however, the periplasmic space is known to contain proteins involved in transport activity (Nikaido and Saier, 1992). For example, a rhodanese-type of protein involved in sulfur metabolism has been identified in Synechococcus sp. strain PCC 7942 (Laudenbach et al., 1991). The periplasmic space also contains channels that facilitate transport between the outer membrane and the plasma membrane.

2. Peptidoglycan Layer In cyanobacteria, as in Gram-negative bacteria, the peptidoglycan layer provides mechanical protection and by its rigidity also determines the cell shape. In electron micrographs of thin sectioned cells, the peptidoglycan layer is a darkly staining band (10 nm, on average) between the outer and the plasma membrane (Fig. 2). Although its structural appearance is similar to that of Gram-negative bacteria, the cyanobacterial peptidoglycan layer is thicker (Golecki, 1988). Furthermore its chemical composition is closer to that of Gram-positive bacteria. In fact, compositional analyses of the outer cyanobacterial layers have lead Jürgens and Weckesser (1985) to conclude that cyanobacteria have a distinctive cell wall organization that is not typical of either the Gram-positive or the Gram-negative bacteria.

3. Transporters In Gram-negative bacteria, both the outer membrane and the plasma (cytoplasmic) membrane contain special protein complexes, that are also referred to as channels or transporter complexes, that facilitate the passage of solutes and small molecules across these membranes. Studies of bacterial transporters, recently reviewed by Nikaido and Saier (1992), have shown that amino acid sequences are strikingly similar to many transporters of animal cells, which suggests commonality in both structure and function. Although various terminologies may be used, three types of channels are recognized: (i) passive diffusionchannels with porin-proteins allowing passage of ions and small nutrient molecules; (ii) channels with specific binding sites allowing facilitated diffusion of slowerpenetrating compounds; and (iii) concentrating channels that function in sequestering vitamins and chelators and other small molecules, and that provide connections of external receptors with the plasma membrane through the rigid wall-layer (Nikaido and Saier, 1992). Channels in cyanobacteria are equally significant for cell function, but studies of such channels in cyanobacterial cells are only beginning. In cyanobacteria proteins and lipopolysaccharides are major constituents of the outer membrane, while carotenoids and lipids are minor components (Omata and Murata, 1984; Jürgens and Weckesser, 1985; Murata and Omata, 1988). However, it has often

Chapter 6 Membrane Organization been difficult to ascertain the exact composition of the individual external layers. As determined by SDS-PAGE two major proteins, and of ten one minor protein, are prominent but variable among species: 50 and 54 kDa in Fischerella sp. strain PCC 7414 (Pritzer et al., 1989); 53 and 62 kDa in Gloeobacter violaceus strain PCC 7421 (Schneider and Jürgens, 1991) and 61, 67, and 94 kDa in Synechocystis sp. strain PCC 6714 (Juergens and Benz, 1988). These polypeptides are associated with the peptidoglycan layer, and although they differ in molecular mass from those of enteric bacteria (36–38 kDa) they may have porin-like functions. Pores were obtained when outer membrane proteins (61 and 67 kDa) of Synechocystis sp. strainPCC 6714 were reconstituted into lipid bilayer membranes (Juergens and Benz, 1988) (Table 1). Whereas some activity was obtained in channel conductance experiments, the pores did not have the long lifetime found for bacterial pores. A comparison was made with bacterial-derived porins and those from Synechococcus sp. strain PCC 6301 by Zalman (1982) in Nikaido’s laboratory. They showed the 50 kDa cyanobacterial porin to be functionally competent by a liposome swelling assay and found that the channels formed were slightly

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larger than those of E. coli. However, the cyanobacterial porin had a lower efficiency, in that the observed diffusion rate per unit weight was lower than that for E. coli. The sulfate transporter in cyanobacteria is probably very similar to those previously described in other eubacteria. Insertional inactivation and sulfate uptake studies have provided evidence for a sulfate permease complex (Table 1) in Synechococcus sp. strain PCC 7942 (Laudenbach and Grossman, 1991). The products of three genes (sbpA, cysT, and cysW) seem to be involved in forming a sulfate-transport channel. A sulfate-binding protein (37.7 kDa gene product) is involved on the periplasmic side, while two proteins with masses of 30.4 and 30.7 kDa extend into the periplasmic region and are inserted in the plasma membrane (Fig. 1). Evidence for a nitrate transport system in cyanobacteria is described in Chapter 16.

4. Carotenoid-Binding Proteins Numerous laboratories have reported the occurrence of carotenoids in the outer membrane (Omata and Murata, 1984; Juergens and Weckesser, 1985; Jürgens and Benz, 1988). A zeaxanthin-rich protein that

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forms a dimer from two 23 kDa polypeptides (DiversePierluissi and Krogmann, 1988) differs from some of the outer-membrane-localized, carotenoid-binding proteins because it is soluble in the absence of detergents and thus is probably not integral to the membrane. Since some carotenoids co-isolate with some of the porin protein preparations, it is possible that some porin channel complexes may contain carotenoids. A 45 kDa carotenoid-binding protein, that is somewhat smaller than porin proteins (Table 1), has been localized in the outer membrane by immunolabeling (Fig. 3). In the cytoplasmic membrane a different carotenoid-binding protein with a mass of 42 kDa was also found (Fig. 4) by immunolabeling (Reddy et al., 1989; Engle et al., 1991). Dichroic measurements suggest that the carotenoids are perpendicularly oriented with respect to the plane of the membrane (Jürgens and Mäntele, 1991) and

Elisabeth Gantt possibly provide a protective function against high light exposure. However, direct evidence for a protective function is still lacking. Proteins of the outer membrane that have been sequenced in bacteria exhibit low overall hydrophobicity and thus may be readily exportable into the periplasm. In fact, the signal sequence of a thylakoidlumen protein (PsbO) of Anabaena sp. strain PCC 7120 can direct proteins fused with alkaline phosphatase to the periplasmic space (Borthakur and Haselkorn, 1989). This suggests that secretion of proteins into the periplasmic space and to the outer membrane occurs by a similar mechanism.

B. Plasma Membrane Direct connections between the plasma membrane and the outer membrane are predictable from a functional standpoint. Although they have been

Chapter 6 Membrane Organization included in the model of Fig. 1, actual physical connections have been difficult to demonstrate by EM. As in bacteria, oxidative phosphorylation, the transport of electrons, proton pumping and ATP generation are also interdependent in cyanobacteria and involve ATP synthase, respiratory enzymes, and various types of transporters.

1. Carbon Transporters Movement of carbon as and or into a cell is a principal requirement for photosynthetic autotrophs. A 42 kDa polypeptide has been suggested as being involved with the inorganic carbonconcentrating mechanism in cyanobacteria (reviewed in Kaplan et al., 1991; see Chapter 15). In the plasma membrane a 42 kDa polypeptide accumulates in cells grown under conditions. However, proof that the 42 kDa polypeptide is a transporter is not fully accepted because uptake of inorganic carbon still occurs in a mutant that does not show excessive accumulation of this polypeptide. A smaller protein with a mass of about 8 kDa may also play a role in carbon uptake as suggested from mutants constructed by Ogawa (1992). The hydropathy pattern suggest the occurrence of two hydrophobic regions in the 8 kDa polypeptide-suggesting that it could be a transmembrane protein. However, its location in the plasma membrane has yet to be directly demonstrated.

2. Respiratory Enzymes Enzyme complexes of the respiratory chain are present in both the plasma membranes and thylakoid membranes, even though these membranes are different in function and composition (Murata and Omata, 1988; Peschek et al., 1989b). A close interaction of respiratory and photosynthetic electron transport chains in thylakoid membranes suggests that functionally similar, if not identical, electron carriers may be utilized in both membranes (Scherer, 1990). However, the existence of a respiratory chain involving complexes of amitochondrial-type complex I (NADH:ubiquinone oxidoreductase), Cyt and cytochrome oxidase in the photosynthetic membrane as well as in cytoplasmic membranes is not universally accepted. Their occurrence is expected because cyanobacteria, most of which are obligate phototrophs, generate ATP from respiration in darkness. The respiratory functions in the plasma and photosynthetic membranes are being elucidated as

125 evidenced by Peschek’s (1987) summary in the last cyanobacterial volume and by subsequent studies in Peschek’s laboratory and in several other laboratories (reviewed in Chapter 13). Analyses of purified membranes from over 20 cyanobacterial species have shown that both cytoplasmic and photosynthetic membranes contain complete respiratory assemblies with NADPH dehydrogenases (Berger et al., 1991; Walker, 1992) plastoquinone, cytochrome complexes (Kraushaar et al., 1990) and aerobic type) cytochrome oxidases (Peschek et al., 1989b; Moser et al., 1991; Nicholls et al., 1992). As in mitochondria the cytochrome oxidase is a transmembrane complex (~ 100–120 kDa) that occurs as a dimer (Brunor and Wilson, 1982). The expected size and orientation of the complexes is shown in the plasma membrane (Fig. 1) and the thylakoid membrane (Fig. 8).

C. Plasma-Thylakoid Membrane Contacts The plasma membrane and thylakoid membrane system share many supramolecular complexes as indicated in Fig. 1, but the photosystems are restricted to the thylakoids. Only in rare exceptions does the plasma membrane also serve as a photosynthetic membrane. Such an exception is found in Gloeobacter violaceus strain PCC 7421, a simple photoautotroph in which thylakoids are absent and phycobilisomes are attached directly to the plasma-photosynthetic membrane (Guglielmi et al., 1981). It is interesting to note that this organism, when compared to most conventional cyanobacteria, has a much slower growth rate. Possible connections between plasma membranes and thylakoid membranes are indicated from EM analyses. Contact points between the plasma membrane and thylakoid membranes have been noted several times (Kunkel, 1982; Balkwill et al., 1984; Stevens and Nierzwicki-Bauer, 1991). [Editor's note: Although the continuity of the cytoplasmic and thylakoid membranes is still questioned by many workers, there are examples in the literature, e.g., Arthrospira jenneri (see fig. 4 of Wildman and Bowen, 1974) that suggest that thylakoids are infoldings of the cytoplasmic membrane.] In most species, the thylakoids typically end very close to the plasma membrane, as in Synechocystis sp. strain PCC 6714 (Fig. 5). These areas, although suggestive of direct continuity, do not exhibit clear continuities between the cytoplasmic and thylakoid membranes. Biochem-

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ical analyses by Peschek et al. (1989a) have demonstrated that chlorophyll precursors but not Chl a occur in the plasma membrane. It is possible that apoprotein formation of the Chl-binding complexes and Chl a insertion takes place at the contact sites. It is also interesting that a 33 kDa polypeptide which belongs to the oxygen-evolving complex, is present in both thylakoid and plasma membranes, although this complex is not functional in the plasma membrane (Smith et al., 1992).

D. Thylakoid Membranes 1. General Morphology Phycobilisomes generally cover the thylakoid membranes on the stromal (protoplasmic) side. Thylakoid membrane organization in cyanobacteria has been regarded as simple because grana-like regions do not occur, thus making it unlikely that PS

II is sequestered in special regions as in green plants. The thylakoid membranes traverse the cytoplasm (Figs. 3–6), but their exact arrangement depends on the species and sometimes on the physiological state (Stevens and Nierzwicki-Bauer, 1991). Concentric layering of thylakoids is commonly seen in thinsection views of small rod-shaped unicells like the Synechococcus species and also in filamentous species like Phormidium persicinum (Fig. 7). In other filamentous species, as in Calothrix and Nostoc species, the thylakoid arrangement is sometimes more dense and irregular and often exhibits anastomoses between neighboring thylakoids (Damerval et al., 1991; Stevens and NierzwickiBauer, 1991).

2. Phycobilisomes Phycobilisomes, the major light harvesting antennae of cyanobacteria, are attached to the stromal side of

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the thylakoids (Bryant, 1991) (see Chapter 7). It is well known that they primarily transfer excitation energy to PS II which has been directly verified in isolated phycobilisome-PS II complexes (Gantt, 1986). Direct energy transfer from phycobilisomes to PS I is possible, and in fact, PS I-enriched phycobilisome preparations of Synecochoccus sp. strain PCC 6301 indeed show such transfer (Mullineaux, 1992). Such transfer to PS I is predominantly associated with light-state 2 and is dependent upon the presence of allophycocyanin B in the phycobilisome (Zhao et al., 1992). Also, phycocyanin can transfer energy directly to Chl as recently shown in an allophycocyanin-deficient mutant (Su et al., 1993). It is significant to note, however, that in PS I-deficient mutants there is no decrease of the phycobiliprotein (and phycobilisome) content (Shen et al., 1993); furthermore, efficient energy coupling is high and without any loss through fluorescence. This would not be expected if there were unattached phycobilisomes in such mutants.

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Phycobilisome size and shape varies with species (Gantt, 1980; see Chapter 7), and is often dependent on the light conditions in which cells are grown. In many cyanobacterial species the phycobilisomes are hemidiscoidal (Figs. 6 and 8). In several filamentous species the phycobilisome morphology differs in that the core structure is more complex and the phycobilisomes are cylindrical in shape (Isono and Katoh, 1982; Glauser et al., 1992). A change in shape can be affected by changing the light-growth conditions—e.g., hemidiscoidal phycobilisomes might occur in cells grown in red light and cylindrical phycobilisomes might occur in cells grown in green light (Ohki and Fujita, 1992). Furthermore, the cylindrical phycobilisomes have a larger size. It may be inferred that the number of PS IIs increases as the phycobilisome size increases, because in the red alga Porphyridium cruentum the phycobilisome size is larger and so is the of PS II to phycobilisome ratio (Gantt, 1986). On the other hand, in the same species the PS II:phycobilisome ratio is greatly enchanced

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when cells were grown in red light, with the result that the phycobilisomes remained the same size but the PS II centers per phycobilisome increased (Cunningham et al., 1990). Often the phycobilisomes occur in closely spaced, parallel rows and thus, thylakoid-membrane models also show subtending rows of putative PS II particles (Giddings et al., 1983); these suggest that thylakoid formation occurred with a polar directionality. Phycobilisomes can actually be found in various orientations on the same membrane as seen in Fig. 6 in which small groups of phycobilisomes are oriented at various angles, suggesting that membrane formation has occurred in patches.

3. Photosystem I PS I is a large multiprotein complex (Table 1; see Chapter 10 for details). This reaction center is composed of a heterodimer of 82- and 83 kDa polypeptides that bind P700, two phylloquinones and the [4Fe-4S] cluster denoted These polypeptides are hydrophobic and traverse the

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thylakoid membrane about 8 to 9 times (Krauss et al., 1993). The genes (psaA and psaB) coding for these polypeptides predict polypeptides that are highly similar in cyanobacteria and green plants, except for a short extension of the predicted polypeptide at the ammo-terminus of PsaA (Golbeck and Bryant, 1991). In fact, the PS I reaction center complex of cyanobacteria is comparable to the core complex of green plants. Its molecular mass of about 250 kDa is accounted for by the eight intrinsic membrane polypeptides (PsaA, PsaB, PsaF, PsaI, PsaJ, PsaK, PsaL, and PsaM) and three water-soluble proteins (PsaC, PsaD, and PsaE). The entire complex has an actual molecular mass of about 340 kDa when all 11 polypeptides, 100 Chl molecules, and about 10–15 are included (Golbeck and Bryant, 1991; Bryant, 1992; Krauss et al., 1993). On SDS-PAGE the apparent molecular mass of this complex (~150 kDa) is smaller (Hefti et al., 1992) than when calculated from the protein:chlorophyll weight ratio or when deduced from gel exclusion chromatography (Rögner et al., 1990). Isolation of the PS I complex by detergent

Chapter 6 Membrane Organization

solubilization, usually with or dodecylmaltoside, has yielded stable samples that have proven suitable for structural examination for a number of thermophilic and mesophilic species including Mastigocladus laminosus, Phormidium laminosum, Synechococcus sp., Synechococcus sp. strain PCC 7002 and Synechocystis sp. strain PCC 6803. The description here will concentrate on Synechococcus sp., because it has been most extensively characterized and appears to possess the same essential features as those found in other species (Boekema et al., 1989; Rögner et al., 1990; Böttcher et al., 1992;Hefti et al., 1992). Analyses of negatively stained PS I complexes of Synechococcus sp. reveal that in vitro particles can be found as monomers or trimers, depending upon the salt concentration during isolation (Bald et al., 1992). The trimers, which after isolation have a disc-shaped appearance (Fig. 7), may aggregate along their broad faces into chains of two or more, and it is the trimeric aggregates that form crystals (Krauss et al., 1993). The initial size estimates of an isolated Synechococcus sp. monomer —i.e., 15 nm long, 6.4 nm high, and 10.6 nm wide (Rögner et al., 1990)—have been re-

129

evaluated because negative staining may subject proteins to drying and distortion. A recent structural model of the monomer was developed by examination of two-dimensional crystals (Böttcher et al., 1992). PS I appears as a highly asymmetric, conical complex with an overall width of approximately 10 nm. On one side it is flattened and wide but then tapers into a narrow, flat side. The larger side, designated as the one facing into lumen of the thylakoid, has a shallow cavity about 3 nm in diameter with a depth of 1 nm (Fig. 8). The general features of this model have been confirmed and extended by a high resolution model derived from X-ray diffraction studies recently published for the PS I trimers of the same organism (Witt et al., 1992; Krauss et al., 1993). Elucidation of the crystal structure at 6 Å resolution has allowed the precise placement of the Fe-S clusters and on the stromal side, and hence on the side opposite the large, flat side with the 3 nm depression. This is of special significance in that it determines with certainty the orientation of the PS I complex within the membrane. The longest axis (13 nm) of a PS I monomer is on the lumenal side. This is the side that contains the shallow cavity, the probable

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130 plastocyanin (or docking site (Fig. 8). On the protoplasmic side, where subunits PsaC, PsaD, and PsaE are attached, the complex projects about 4nm out of the lipid bilayer (~4–5 nm) and thus almost doubles the thylakoid thickness that had been assumed from electron micrographs of sectioned thylakoids. Antenna complexes, composed of polypeptides that bind chlorophylls and xanthophyll carotenoids and that are peripheral to the PS I reaction center core, are common in green plants and all groups of eucaryotic algae (Green et al., 1991; Wolfe et al., 1994). However, LHC I and LHC II complexes have not been found in cyanobacteria. An important question yet to be answered is: In what state, as a single unit or as an aggregate, does the PS I complex exist in the membrane? An answer to this question is difficult to obtain because the in vivo state is disturbed by any kind of probing. Freezefracture analysis, which is perhaps the least disruptive method, has not proven to be useful since PS I is virtually impossible to identify in a thylakoid membrane. The choice ofsalt and ionic strength used for isolation of complexes can influence the aggregation state of PS I complexes. Trimers are favored in low ionic strength (< 70 mM NaCl), while monomers are favored in high ionic strength (150 mM) (Bald et al., 1992). Detergent treatment is also thought to influence the aggregation state (Ford and Holzenburg, 1988). Evidence favoring the view that PS I exist only as monomers in vivo was provided from analysis of thylakoid membranes that had been enriched in PS I, by removal of PS II and cytochrome complexes, with mild detergent treatment (Hefti et al., 1992). However, the packing of the particles shown was very tight. The packing, in fact, was so tight that the question arises as to how other major membrane components (PS II, and cytochrome could be accommodated in the same membrane region. Persuasive arguments and evidence show that PS I exists as trimers in vivo in thylakoid membranes (Hladik and Sofrova, 1991). A trimeric spatial arrangement is common in crystals–even when crystals are prepared from monomers (Almog et al., 1991). Evidence for the preponderance of trimers in cyanobaeterial thylakoids also comes from experiments in which membranes were treated with hydrophobic, protein cross-linking agents (Pospisilova et al., 1990; Hladik and Sofrova, 1991). Trimers

were preserved if thylakoids were exposed to crosslinkers prior to solubilizing them under conditions that normally produced monomers. Examination of spatial distances of photosystems in thylakoids of a phycobilisome-containing red alga also provides evidence for clustering of PS I (Mustardy et al., 1992). In isolated membranes only 25% of the PS I present (determined by P700 per membrane area) were consistently labeled when probed with colloidal gold particles directly conjugated to anti-PS I antibodies. PS I clusters of three or four were considered likely because close-packing of the reaction centers would impose a steric hindrance that would impede attachment ofthe label that was about the same size (10 nm gold plus 5 nm antibody) as each PS I (13 nm widest diameter) monomer.

4. Photosystem II The PS II reaction center of cyanobacteria is a large complex composed of numerous proteins with a large number of membrane-spanning regions (Pakrasi and Vermaas, 1992; Satoh, 1992; see Chapter 8). It consists of the reaction center core and core antennae, but appears to lack any chlorophyll-xanthophyll bindingproteins that are normally found as peripheral antennae in green plants and algae. At least twelve of the gene products comprising the cyanobaeterial reaction center II are known to be highly conserved in cyanobacteria and green plants (Erickson and Rochaix, 1992; Pakrasi and Vermaas, 1992; Satoh, 1992). It is well known that subunits D1 and D2 comprise the central heterodimer to which are bound the essential cofactors: the P680 dimer, the intermediate electron acceptor pheophytin, the non-heme iron and the four manganese atoms of the water-oxidation complex. However, the placement of the remaining subunits in relationship to D1 and D2 and with one another has not been clarified. An interesting model for the topography of the PS II complex derives from studies of chemical crosslinking performed on thylakoids of Synechocystis sp. strain PCC 6803 mutants (Ikeuchi et al., 1992). In the provocative model proposed, ten of the subunits are spatially related to one another. Interestingly, D1 (PsbA) is only associated with D2 (PsbD). In a Ushaped configuration CP43 (PsbC) is bound to D2 in one location while PsbK is peripheral to it. Forming the other side ofthe U and directly attached to D2 are CP47 (PsbB) and PsbE-PsabF (cytochrome

Chapter 6 Membrane Organization Peripheral to CP47 is PsbH on one side and PsbL is peripheral to PsbF. These ten apoproteins account for a combined mass ofapproximately 190 kDa, which together with 45 chlorophylls and other cofactors yield a molecular mass (minus detergent) of about 250 kDa for a PS II monomer (Table 1) in agreement with previously determined values for Synechosystis sp. strain PCC 6803 and Synechococcus sp. (Rögner et al., 1990; Dekker et al., 1988). The shape of isolated PS II particles, visualized by negative staining, is elliptical with dimensions of 15.5 by 10.5 nm for monomers and 18.5 by 15 nm for dimers whether obtained from cyanobacteria or as reaction center II particles of spinach (Rögner et al., 1987; Irrgang et al., 1988). In isolated preparations the particles often aggregate into long chains, although this probably does not reflect the conditions within the membrane. A three-dimensional structure of the PS II complex of cyanobacteria has not been published, nor is the structure known for purified PS II reaction center of green plants. However, the first PS II structure, including the peripheral chlorophyll a/b-antenna complex, has just been proposed for higher plants (Holzenburg et al., 1993). The analysis was made from negatively stained thylakoid regions that exhibited two-dimensional crystalline arrays. It is suggested that the lumenal side is quite flat and has a central cavity of about 2–3 nm. Four domains surround the cavity, one of which is suggested as being involved with the water oxidation site. The other three domains seem to be connected within the membrane plane (4.5 nm width) and narrow as they extend into the protoplasmic space. Eight projections on the periphery, which are probably the subunits of the chlorophyll a/b-binding complex, appear to surround the central core. Assuming the absence of peripheral antennae in cyanobacterial PS II, the cyanobacterial monomer could be envisaged to have a lumenal, long axis of about 17 nm and a width of about 10 nm. A recent study on isolated PS II particles from a cyanobacterium (Synechococcus sp. strain PCC 6803) and from spinach by Boekema et al. (1993) suggests a very different configuration from that proposed by Holzenburg et al. (1993). They show a dimeric association of two asymmetric particles, each approximately 7.5 nm wide and 5.5 nm high, with a cavity between the adjacent particles. The stromal sides of the particles are relatively flat and almost even with the lipid bilayer, but have a 3nm projection extending into the lumenal space (Fig. 8).

131 As shown in Figure 8 the cyanobacterial PS II complexes in vivo are thought to exist in closely associated pairs, usually referred to as PS II dimers. Evidence for dimers primarily comes from freezefracture studies, in which 10 nm particles exposed in thylakoids are often seen to occur as pairs (Giddings et al., 1983;Mörschel and Schatz, 1987). The isolation of PS II particles and their visualization in reconstituted lipid vesicles (Mörschel and Schatz, 1987) provide further support for such a dimeric structure.

5. Cytochrome

Complex

The cytochrome is a membrane-intrinsic complex that is expected to be positioned between PS I and PS II because of its central function in electron transport between the photosystems in thylakoids (see Chapter 9). The existence of a comparable redox complex in the plasma membrane has been verified in purified plasma membranes from four cyanobacteria. Probing of immunoblots with antisera to four subunits of the complex from chloroplasts provided evidence of homologous complexes in both thylakoids and cytoplasmic membranes (Kraushaar et al., 1990). These complexes are not functionally identical, because the plasma-membrane complex exhibited sensitivity to the mitochondrial inhibitors antimycin A and rotenone while the thylakoid complexes were insensitive. However, it should be noted that in all cases examined only a single set of genes encoding the subunits of the complex have been identified (see Chapter 9). A monomeric Cyt complex isolated from thylakoids of Synechocystis sp. strain PCC 6803 has a molecular mass of 110 ± 20 kDa (Bald et al., 1992; Table 1). The complex includes subunits with the following apparent molecular masses: Cyt f, 35 kDa; Cyt 20 kDa and Subunit IV 13.5 kDa; the Reiske iron-sulfur-protein, 29 kDa; and three small polypeptides of 6.6,4 and 3.3 kDa of as yet unknown function. The EM images of the negatively stained complex (8.3 nm height, 4.4 nm width, 6 nm length) provide a calculated particle mass of 100 ± 30 kDa. Within the thylakoid membrane the cytochrome complex probably exists in the dimeric state (Fig. 8). In spinach the complex has been isolated as a dimer, with a mass of 200 kDa, and such preparations have approximately five-times greater activity than preparations of monomers (Cramer et al., 1992). A dimeric complex of 320 kDa isolated from

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132 Synechocystis sp. strain PCC 6714 (Tsiotis et al., 1992) appears larger, but it is very possible that the differences in apparent molecular mass are due to differing amounts of bound detergent in the various preparations.

6. Cytochrome Cytochrome is a membrane complex that occurs only in photosynthetic organisms that evolve oxygen (see Chapter 8). It exists as a heterodimer of two subunits of about 8 and 4 kDa, and although small, both appear to be a transmembrane proteins (Cramer et al., 1986; Cramer et al., 1992). This complex has been suggested to function in noncyclic electron transport between the reducing side of PS II and the oxidizing side of PS I (Ortega et al., 1989), or it could provide the electrons for the quenching of (Cramer et al., 1992) in the PS II reaction center. The complex has a close physical relationship with PS II and is present in isolated PS II reaction center core preparations (Nanba and Satoh, 1987). Furthermore, it is essential for the functional assembly of the PS II core (Pakrasi and Vermaas, 1992), but its exact function and whether it is exclusively restricted to PS II remain enigmatic.

7. ATP Synthase ATP synthase associated with thylakoid membranes is similar to that in plasma membranes and in bacterial and mitochondrial membranes in that it consists of two structurally distinct complexes, and (see Chapter 11). In cyanobacteria the thylakoid ATP synthase is generally regarded as essentially equivalent to the coupling factor of higher plants because of certain common features including the subunit composition of (Hicks, 1988; Feng and McCarty, 1990) and probably also of with a peripheral location on the membrane, is hydrophilic and contains the nucleotide binding sites. It has five different subunits with the following approximate molecular weights: and 15 kDa. By image analysis of electron micrographs it can be shown that three pairs of the and subunits form the club-shaped head of while at least one of each of the smaller subunits is clustered in the center near the base (Boekema et al., 1992). The mass of about 360 kDa, may vary slightly with the species but is expected to be nearly the same in cyanobacteria (see Chapter 11).

is hydrophobic and is integral to the membrane (Fig. 8) where it acts as a proton channel. When purified from spinach it has four subunits with masses of approximately 20, 18, 16, 8 kDa (Feng and McCarty, 1990). The mass of (Table 1) of~150 kDa is based on a 1:1:1:12 stoichiometry of the respective subunits above. EM images show that the base piece with a diameter of about 11 nm, is somewhat wider than the head piece (Mörschel and Staehelin, 1983; Pedersen and Amzel, 1993). The actual topography of the complex within the membrane, and its interaction through a connection stalk with remain to be elucidated (however see Chapter 11 for progress in these areas).

8. Ascorbate Peroxidase Superoxide radical production is an inevitable result of oxygenic photosynthesis–particularly due to the activities of PS I. Although the superoxide radicals are converted by superoxide dismutase to hydrogen peroxide and dioxygen, the hydrogen peroxide requires removal because of its inhibitory effect on An ascorbate peroxidase is effective in the scavenging ofhydrogen peroxide in cyanobacteria (Tel-Or et al., 1986) and in green plants (Miyake et al., 1991) by using a photoreductant produced in the thylakoids as the electron donor. Whereas enzymes involved in scavenging of superoxide radicals and hydrogen peroxide are not necessarily membranebound, a thylakoid-bound ascorbate peroxidase was recently found in spinach that may be the primary scavenging system in chloroplasts (Miyake and Asada, 1992). It is thought to be a 32 kDa integral membrane protein, because neither chelating nor chaotropic agents effected its removal and it could only be solubilized by detergents. A thylakoid-bound ascorbate peroxidase system is also expected in cyanobacteria, in which it may have first developed during evolution as cyanobacteria began to adapt to an increasingly oxidizing atmosphere.

9. Carotenoid Synthetic Enzymes Enzymes of the carotenoid biosynthetic pathway are membrane-bound beginning with phytoene desaturase (see Chapter 18 for details). The phytoene desaturase enzyme in the biosynthetic pathway of carotenoids has been localized in thylakoids by immunocytochemistry in Anabaena sp. and Synechocystis sp. (Serrano et al., 1990). For localization of

Chapter 6 Membrane Organization phytoene desaturase, an antibody was used that had been produced against a desaturase fusion protein produced from a Rhodobacter capsulatus gene. The antibody recognized a 65 kDa polypeptide when blots of SDS-PAGE gels of whole-cell extracts were immunodecorated. Confirmation of these results with antibodies produced to cyanobacterial fusion proteins will be of interest, because the sequence homology of phytoene desaturase of non-oxygenic and oxygenic plants is very low, whereas it is significantly higher among cyanobacteria and higher plants (Bartley et al., 1991). The cyanobacterial genes for phytoene synthase and lycopene cyclase have also been cloned but the localization of the respective proteins within the plasma membrane or the photosynthetic membrane remains to be ascertained (Cunningham et al., 1993).

10. Other Complexes Proteins involved with nitrogen metabolism, such as ferredoxin-nitrate reductase and ferredoxin-nitrite reductase, are known to be tightly attached to thylakoid membranes (Guerrero and Lara, 1987) even though they are not necessarily integral to the membrane. Assimilation of inorganic nitrogen compounds is dependent on reducing compounds from photosynthesis, but specific membrane complexes have yet to be characterized. A Type-1, NADH:ubiquinone oxidoreductase (complex I) in cyanobacteria has also been found (Berger et al., 1991). It is smaller than the bovine mitochondrial complex, that consists of 41 subunits with a total molecular mass of about 670–890 kDa (Walker, 1992). The intrinsic portion of the complex is L-shaped (Fig. 8) and is oriented parallel to the membrane plane and with the extrinsic portion projecting into the protoplasmic space. Its role in thylakoids remains to be clarified but a number of functions have been proposed (see Scherer, 1990, and Chapter 13). Mi et al. (1992a, b) suggested that the NADH dehydrogenase might be the re-entry point for electrons in cyclic electron transport. However, more recent results suggest that a cyclic pathway for electron transport exists even in mutants which do not contain a functional NADH dehydrogenase (Schluchter et al., 1993; Yu et al., 1993). This pathway is absolutely dependent upon the presence of the PsaE subunit of the PS I complex. Cross-linking studies indicate that the PsaE subunit makes contact in thylakoids with an integral

133 membrane protein of approximately 35–40 kDa which is not a part of the PS I complex; this component does not appear to be a subunit of the cytochrome complex (U. Mühlenhoff and D. A. Bryant, personal communication). Finally, it is interesting to note that ferredoxin oxidoreductase (FNR) has recently been shown to be a component of cyanobacterial phycobilisomes in Synechococcus sp. strains PCC 7002 and 7942 (Schluchter and Bryant, 1992). Approximately two molecules of FNR occur in each phycobilisome. Cyanobacterial FNR contains an additional domain, added at the amino terminal of the catalytic domains that bind FAD and (see Karplus et al., 1991), that is highly homologous to the CpcD protein of phycobilisomes. CpcD is a phycocyanin-associated linker polypeptide whose function is to limit peripheral rod-length heterogeneity and which is associated with the core-distal trimer of phycocyanin in the peripheral rods (see Chapter 7). In higher plants such as barley, FNR can be isolated as a component of the PS I complex under appropriate extraction conditions (Andersen et al., 1992). Hence, cyanobacteria and higher plants differ with respect to the localization of this important electron transport component, and it is interesting to speculate whether phycobilisomes and FNR form some sort of ‘supercomplex’ in thylakoids with PS I and PS II complexes.

E. Photosynthetic Membrane Topography The thylakoid membrane model shown in Fig. 8 is a synthesis of the current knowledge and presents a plausible topographic arrangement of the major supramolecular complexes. The sizes and shapes are based on protein structures derived from images of electron density maps or electron microscopy (ATP synthase, PS I, PS II) or derived from known molecular masses and best estimates (cytochrome transporters). Such a construction is useful for considering the size relationships of the complexes in the membrane. Further, it imposes certain constraints on where adjacent complexes might be placed, and what their density might be within the cyanobacterial membrane. Meaningful estimates for cyanobacteria are complicated by the difficulty of determining the number of phycobilisomes per unit thylakoid area and by the different values reported in the literature for the ratio of PS II centers per phycobilisome. Determinations of this ratio made

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134 whole cells indicate a PS II to phycobilisome ratio of 1:1 (Kawamura et al., 1979; Fujita and Murakami, 1987), but a ratio of 2:1 according to freeze-fracture data (Giddings et al., 1983; Mörschel and Schatz, 1987). The density of the complexes can be approximated if the phycobilisome number per unit thylakoid area is known along with the PS I and PS II stoichiometries. The bestexample exists forthe red algaPorphyridium cruentum in which the phycobilisome density has been directly observed and for which the PS I and PS II contents have been directly correlated (Mustardy et al., 1992). In this species conservative calculations show that at least 50% of the thylakoid area is occupied by PS I, PS II, and the cytochrome complex, if one assumes that (ATP synthase) and cytochrome occur with PS II in a 1:1:1 molar ratio. A ratio of one ATP synthase per cytochrome complex was obtained by Leong and Anderson (1986) for pea thylakoids, while Fujita and Murakami (1987) found a 1:1 ratio of cytochrome to PS II for the cyanobacterium Synechocystis sp. strain PCC 6714. In Synechococcus sp. strain PCC 6301 it can also be calculated that in cells grown in white light 50% or more ofthe thylakoid area is occupied by the four major photosynthetic complexes (PS I, PS II, and For this calculation, it was assumed that there is one PS II reaction center per phycobilisome (Kawamura et al., 1979; Fujita and Murakami, 1987) and that the complexes occur in the ratio PS I:PS with a phycobilisome density on the thylakoids of (Khanna et al., 1983). Figure 8 shows PS II predominantly as a dimer in accordance with available freeze-fracture data. However, since physiological measurements do not indicate a 1:1 molar ratio ofphycobilisomes to PS II centers, one must strongly entertain the possibility that approximately 50% of the phycobilisomes are ‘silent’ and perhaps not attached to PS II centers. Since there is no evidence that phycobilisomes are energetically uncoupled in cells under normal growth conditions, the idea that some phycobilisomes are attached to PS I may be considered, but there is no direct evidence for this in PS I-deficient mutnats (Shen et al., 1993). In Fig. 8 attachment between PS I and phycobilisomes is also suggested. Howerver, it should be pointed out that it is unlikely that the same phycobilisomes canbe freely shiftedspatially between PS I and PS II complexes or functionally coupled simultaneously to both PS I and PS II complexes.

III. Future Focus Are membranes composed of functional domains? What are the composition and sizes of such functional domains? Questions of this nature now are addressable at the level of individual complexes such as enzymes and photosynthetic reaction centers. However, at the level of membranes, the complexity greatly increases, although techniques are becoming available to study membrane components in situ. For example, a beginning has been made in the topographical mapping of red algal thylakoid membranes (Mustardy et al., 1992) by immunoelectron microscopy. By using this approach it will be feasible to determine the density of integral protein complexes with a resolution of 15–20 nm. Determination of the spatial organization of functionally interdependent complexes can also include the supramolecular intrinsic membrane complexes as well as those which are known to interact with such complexes, such as the superoxide dismutase complex and associated peroxidase around PS I. By using specific gold-particle sizes to tag specific antibodies, doubling labeling studies can establish the spatial and stoichiometric relationships ofPS I and PS II, ofPS I or PS II and Cyt and any other components for which antibodies are available. The nature of the sites assuring a functional connection between the phycobilisome and PS II remains to be elucidated. Also, if phycobilisomes are functionally connected to PS I, a separate connecting site would be expected. The stromal surface ofthe PS II complex is rather flat, while the stromal surface of a PS I monomer is dome-shaped and much higher. Studies on the structural and functional components of cyanobacterial membranes have been quite limited when compared to those on other eubacteria– especially enteric bacteria. Although the cyanobacteria are grouped with other eubacteria, it is important to recognize that the cyanobacteria, along with the prochlorophytes, form a distinctive grouping (see Chapters 1, 3 and 5). Although they are procaryotes, their photosynthetic complexes have far greater similarity to those of oxygen-evolving plants rather than to those of anoxygenic photosynthetic bacteria (purple bacteria, green sulfur bacteria, heliobacteria, etc.). Some features of the outer membrane layers are also emerging as unique to cyanobacteria.

Chapter 6 Membrane Organization Acknowledgments I am grateful to the many investigators who kindly made their reprints available, to Drs K. Ohki, Y. Fujita, L. Sherman, and E. J. Boekema for use of their electron micrographs. I am indebted to Drs M. Rögner and W. Cramer for helpful discussion on the orientation ofcomplexes presented in the thylakoid model, and to Beatrice Grabowski for her helpful comments on the manuscript. Dr H. Nikaido provided information on porins, and to him and his colleagues L. Zalman,and R.E.W.Hancock, Iextendmythanks for sharing unpublished results. References Almog O, Shoham G, Michaeli D and Nechushtai R (1991) Monomeric and trimeric forms of Photosystem I reaction center ofMastigocladus laminosus. Proc Natl Acad USA 88: 5312–5320 Almog O, Shoham G and Nechushtai R (1992) Photosystem I: Composition, organization and structure. In: Barber J (ed) The Photosystems: Structure, Function and Molecular Biology, pp 444–470. Elsevier, Amsterdam Andersen B, Scheller HV and Müller BL (1992) The PS I-E subunit of Photosystem I binds ferredoxin: oxidoreductase. FEBS Lett 311: 169–173 Bald D, Kruip J, Boekema EJ and Roegner M (1992) Structural investigations on cytochrome and PS I-complex from the cyanobacterium Synechocystis PCC6803. In: Murata N (ed) Research in Photosynthesis, Vol I, pp 629–632. Kluwer, Dordrecht Bartley GE, Viitanen PV, Pecker I, Chamovitz D, Hirschberg J and Scolnik PA (1991) Molecular cloning and expression in photosynthetic bacteria ofsoybean cDNA coding for phytoene desaturase,and enzyme of the carotenoid biosynthetic pathway. Proc Natl Acad USA 88: 6532–6536 Balkwill DL, Stevens SE and Nierzwicki-Bauer SA (1984) Use of computer-aided reconstructions and high-voltage electron microscopy to examine microbial three-dimensional architecture. Biotechniques 2: 242 Berger S, Ellersiek U and Ferguson SJ (1991) Cyanobacteria contain a mitochondrial complex I-homologous NADH dehydrogenase. FEBS Lett 286: 129–132 Boekema EJ, Boonstra AF, Dekker JP and Rögner M (1993) Electron microscopic structural analysis of Photosystem I, Photosystem II and the cytochrome complex from green plants and cyanobacteria. J Bioenerget Biomembr, in press Boekema EJ, DekkerJP, Rögner M, Witt I, Witt HT and van Heel (1989) Refined analysis of the isolated Photosystem I complex from the thermophilic cyanobacterium Synechococcus sp. Biochim Biophys Acta 974: 81–87 Boekema EJ, Harris D, Böttcher B and Gräber P (1992) The structure ofthe ATP-synthase from chloroplasts. In: Murata N (ed) Research in Photosynthesis, Vol II, pp 645–652. Kluwer, Dordrecht Borthakur D and Haselkorn R (1989) Nucleotide sequence of the

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Chapter 6 Membrane Organization cyanobacteria. Plant Cell Physiol 32: 33–43 Mörschel E and Schatz G (1987) Correlation of photosystem-II complexes with exoplasmatic freeze-fracture particles of thylakoids of the cyanobacterium Synechococcus sp. Planta 172: 145–154 Mörschel E and Staehelin LA (1983) Reconstitution of cytochrome and ATP synthase complexes into phospholipid and galactolipid liposomes. J Cell Biol 97: 301–310 Moser D, Nicholls P, Wastyn M and Peschek G (1991) Acidic cytochrome of unicellular cyanobacteria is an indispensable and kinetically competent electron donor to cytochrome oxidase in plasma and thylakoid membranes. Biochem Intern 24: 757– 768 Mullineaux CW (1992) Excitation energy transfer from phycobilisomes to Photosystem I in a cyanobacterium. Biochem Biophys Acta 1100: 285–292 Murata N and Omata T (1988) Isolation of cyanobacterial plasma membranes. Meth Enzymol 167: 245–251 Mustardy L, FX Cunningham, E Gantt (1992) Photosynthetic membrane topography: Quantitative in situ localization of photosystems I and II. Proc Natl Acad Sci USA 89: 10021– 10025 Nanba O and Satoh K (1987) Isolation of a Photosystem II reaction center consisting of D-l and D-2 polypeptides and cytochrome b-559. Proc Natl Acad Sci USA 84: 109–112 Nicholls P, Obinger C, Niederhauser H and Peschek GA (1992) Cytochrome oxidase in Anacystis nidulans: stoichiometries and possible functions in the cytoplasmic and thylakoid membranes. Biochim Biophys Acta 1098: 184–190 Nikaido H, and Saier MH (1992) Transport proteins in bacteria: common themes in their design. Science 258: 936–942 Ogawa T (1992) Identification and characterization of the Ict A/ Ndhl gene-product essential to inorganic carbon transport of Synechocystis PCC6803. Plant Physiol 99: 1604–1608 Ohki K and Fujita Y (1992) Photoregulation of phycobilisome structure during complementary chromatic adaptation in marine the cyanophyte Phormidium sp. C86. J Phycol 28: 803–808 Omata T and Murata N (1984) Isolation and characterization of three types of membranes from the cyanobacterium (bluegreen algae) Synechocystis PCC 6714, Arch Microbiol 139: 113–116 Ortega JM, Hervas M and Losada M (1989) Location of cytochrome b-559 between Photosystem II and Photosystem I in noncyclic electron transport. Biochim Biophys Acta 975: 244–251 Pakrasi HB and WFJ Vermaas (1992) Protein engineering of Photosystem II In: Barber J (ed) The Photosystems: Structure, Function and Molecular Biology, pp 231–257. Elsevier, Amsterdam Pedersen PL and Amzel LM (1993) ATP synthases: structure, reaction center mechanism, and regulation of one of nature’s most unique machines. J Biol Chem 268: 9937–9940 Peschek GA, (1987) Respiratory electron transport. In: Fay P and Van Baalen C (eds) The Cyanobacteria, pp 119–161. Elsevier, New York Peschek GA, Hinterstoisser B, Wastyn M, Kunter O, Pineau B, Missbichler A and Land J (1989a) Chlorophyll precursors in the plasma membrane of a cyanobacterium, Anacystis nidulans. J Biol Chem 264: 11827–11832 Peschek GA, Wastyn M, Trnka M, Molitor V, Fry IV and Packer L (1989b) Characterization of the cytochrome c oxidase in

137 isolated and purified plasma membranes from the cyanobacterium Anacystis nidulans. Biochemistry 28: 3057– 3063 Pospisilova L, Hladik J and Sofrova D (1990) Topographical study of the pigment-protein complexes of the cyanobacterial Photosystem 1. J Photochem Photobiol B: Biol 5: 401–412 Pritzer M, Weckesser J, and Jürgens UJ (1989) Sheath and outer membrane components from the cyanobacterium Fischerella sp. PCC 7414. Arch Microbiol 153: 7–11 Reddy KJ, Masamoto K, Sherman D and Sherman LA (1989) DNA sequence and regulation of the gene (cbpA) encoding the 42-kilodalton cytoplasmic membrane carotenoprotein of the cyanobacterium Synechococcus sp. strain PCC 7942. J Bacteriol 171: 3486–3493 Rögner M, Dekker JP, Boekema EJ and Witt HT (1987) Size, shape and mass of the oxygen-evolving Photosystem II complex from the thermophilic cyanobacterium Synechococcus sp.FEBS Lett 219: 207–211 Rögner M, Muehlenhoff, Boekema EJ and Witt HT (1990) Mono-, di- and trimeric PS I reaction center complexes isolated from the thermophilic cyanobacterium Synechococcus sp.. Size shape and activity. Biochim Biophys Acta 1015: 415–424 Satoh K (1992) Structure and function of Photosystem II reaction center In: Murata N (ed) Research in Photosynthesis, Vol II, pp 3–12. Kluwer, Dordrecht Scherer S (1990) Do photosynthetic and respiratory electron transport chains share redox proteins? Trends Biochem Sci 15: 458–462 Schluchter WM and Bryant DA(1992)Molecular characterization of oxidoreductase in cyanobacteria: cloning and sequence of the petH gene of Synechococcus sp. PCC 7002 and studies on the gene product. Biochemistry 31: 3092–3102. Schluchter WM, Zhao J, and Bryant DA (1993) Isolation and characterization of the ndhF gene of Synechococcus sp. strain PCC 7002 and initial characterization of an interposon mutant. J Bacteriol 175: 3343–3352 Schneider S and Jürgens UJ (1991) Cell wall and sheath constituents of the cyanobacterium Gloeobacter violaceus. Arch Microbiol 156: 312–318 Serrano A, Giminez P, Schmidt G and Sandmann G (1990) Immunocytochemical localization and functional determination of phytoene desaturase in photoautotrophic prokaryotes. J Gen Microbiol 136: 2465–2469 Shen G, Boussiba S and Vermaas WFJ (1993) Synechocystis sp. PCC 6803 strains lacking Photosystem I and phycobilisome function. Plant Cell 5: 1853–1863 Smith D, Bendall DS and Howe CJ (1992) Occurrence of a Photosystem II polypeptide in non-photosynthetic membranes of cyanobacteria. Mol Microbiol 6: 1821–1827 Stevens SE Jr and Nierzwicki-Bauer S (1991) The cyanobacteria. In: Stolz JF (ed) Structure of Phototrophic Prokaryotes, pp 15– 47. CRC Press, Inc, Boca Raton Su X, Fraenkel PG and Bogorad L (1992) Excitation energy transfer from phycocyanin to chlorophyll in an apcA -defective mutant of Synechocystis sp PCC 6803. J Biol Chem 267: 22944–22950 Tel-Or E, Huflejit M and Packer L (1986) Hydroperoxide metabolism in cyanobacteria. Arch Biochem Biophys 246: 396–402 Tsiotis G, Lottspeich F and Michel H (1992) Isolation and characterization of cytochrome from Synechocystis

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Chapter 7 Phycobilisome and Phycobiliprotein Structures Walter A. Sidler Institut für Molekularbiologie und Biophysik, Eidgenössische Technische Hochschule, CH-8093, Zürich, Switzerland Summary I. Introduction II. Phycobilisomes A. Electron Microscopy of Phycobilisomes B. Isolation of Phycobilisomes C. Phycobilisome Components 1. Variability of the Polypeptide Composition 2. Phycobiliproteins 3. Pigments 4. Linker Polypeptides and Phycobilisome Assembly Oxidoreductase(FNR) 5. Ferredoxin: D. Energy Transfer in Phycobilisomes E. Regulation of Phycobilisome Composition III. Phycobiliproteins Constituting the Phycobilisome Core A. The Allophycocyanin Family 1. Allophycocyanins (APCs, ApcA, ApcB) 2. Allophycocyanin-B (AP-B, ApcD) Subunit (ApcF) 3. B. the Core-Membrane Linker Phycobiliprotein (ApcE) C. Determination of the Core Size by the D. Allophycocyanin Complexes and Their Arrangement in the PBS Core E. Energy Transfer in Allophycocyanin and the PBS Core IV. Phycobiliproteins Constituting the Rod Elements of PBS A. The Phycocyanins 1. Constitutive Phycocyanins 2. Inducible Phycocyanins 3. Phycoerythrocyanin (PEC) 4. R-Phycocyanin (R-PC or R-PC-I) 5. R-Phycocyanin-ll (R-PC-II) 6. Synechococcus sp. strain WH 7805 Phycocyanin (R-PC-III) 7. Synechococcus sp.strainWH8501 Phycocyanin(R-PC-IV) 8. Amino Acid Sequences and Phylogenetic Relationships of the C-PC Family 9. The Crystal Structure of C-Phycocyanin a. The C-Phycocyanin Monomer and Its Subunits b. Trimeric C-Phycocyanin c. Hexameric C-Phycocyanin d. The Crystal Structure of Phycoerythrocyanin e. Chromophore Structure and Common Principles of Chromophore-Protein and Protein-Protein Interaction f. a Modified Amino Acid Residue in Phycobiliproteins 10. Energy Transfer in the PBS Rods

D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 139–216. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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B. Phycoerythrins 1. Phycoerythrins in Cyanobacteria and Red Algae a. C-Phycoerythrin-l b. C-Phycoerythrin-ll c. B-Phycoerythrin from Red Algae 2. Comparison of Phycoerythrins with Other Phycobiliproteins 3. The Phycoerythrin subunits 4. The Crystal Structure of B-Phycoerythrin from Red Algae 5. Phycoerythrin in the Light-Harvesting Antenna of Cryptomonads 6. Comparison of Cyanobacterial C-Phycoerythrin, Rhodophytan B-Phycoerythrin and Cryptophytan Phycocyanin-645 and Phycoerythrin-545 7. The Phylogenetic Relationship of Phycocyanin-645 Subunits 8. Specialization and Diversification of Phycoerythrins During Evolution V. Linker Polypeptides, the Skeleton of the PBS A. Interaction of Linker Polypeptides with Phycobiliproteins B. PBS-Core Linker Polypeptides the Small Core Linker Polypeptides 1. the Core-Membrane Linker Polypeptide 2. C. The Rod and the Rod-Core Linker Polypeptides the Small Rod Linker Polypeptide 1. the Rod Linker Polypeptides 2. Rod-Core Linker Polypeptides 3. D. Functional Domains of and and Binding Specificity of E. Rod-Linker Polypeptides for Phycoerythrin Complexes F. Phycobiliprotein-Linker Polypeptide Complexes from the Phycobilisome of Mastigocladus laminosus VI. Organization of the Genes Encoding the Phycobilisome Elements A. Genes Involved in Adaptation to Changes in Environmental Conditions B. The cpcE and cpcF Genes C. Genes Encoding Phycobilisome Components in the Cyanelles of Cyanophora paradoxa, Red Algae, and Cryptomonads D. Genetic Analysis of the Elements of the PBS for Mastigocladus laminosus E. The pec and cpc Operons of Mastigocladus laminosus F. The apc Operons in the Genome of Mastigocladus laminosus Acknowledgments References

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Summary Phycobilisomes serve as the primary light-harvesting antennae for Photosystem II in cyanobacteria and red algae. These supramolecular complexes are primarily composed of phycobiliproteins, a brilliantly colored family of water-soluble proteins bearing covalently attached, open-chain tetrapyrroles known as phycobilins. In addition, phycobilisomes also contain smaller amounts ‘linker polypeptides,’ most of which do not bear chromophores. These components are absolutely required for proper assembly and functional organization of the structure. Phycobilisomes are constructed from two main structural elements: a core substructure and peripheral rods that are arranged in a hemidiscoidal fashion around that core. The core of most hemidiscoidal phycobilisomes is composed of three cylindrical subassemblies. The peripheral rods radiate from the lateral surfaces of the core substructure which are not in contact with the thylakoid membrane. Absorbed light energy is transferred by very rapid, radiation-less downhill energy transfer from phycoerythrin or phycoerythrocyanin (if present) to C-phycocyanin and then to allophycocyanin species that act as the final energy transmitters from the phycobilisome to the Photosystem II or (partially) Photosystem I reaction centers. This chapter focuses on important recent developments concerning the structure and function of phycobilisome architecture and their constituent phycobiliproteins. Studies with the phycobilisomes and phycobiliproteins of the cyanobacterium Mastigocladus laminosus as will be emphasized. During the last decade tremendous progress has been made in the molecular biology of cyanobacteria—through gene

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sequencing, genetic analyses, and studies on gene regulation during adaptation. Many specialized functions of phycobilisome components have been revealed through the construction and characterization of deletion or insertional mutants and mutants harboring site-specific changes in phycobiliproteins. New phycobiliproteins, that play an important role in open-sea photosynthesis, have been discovered from marine cyanobacteria. Previously known to occur in red algae only, have been discovered in these marine cyanobacteria. The complete amino acid sequences for this last class of phycobiliproteins have now been determined. The structures of new chromophores from cryptomonad phycobiliproteins have been determined. Enzymes catalyzing site-specific bilin-attachment to apo-biliproteins have been described and their genes sequenced. The amino acid sequences of all components constituting the phycobilisomes of some cyanobacterial strains have been determined, and analyses of these data have revealed phylogenetic relationships. Structural and functional domains of the linker polypeptides have been recognized, and the special roles of the multifunctional, large core-membrane linker phycobiliprotein in assembling the phycobilisome core and in energy transfer were discovered. The crystal structures for several phycobiliproteins have now been determined at near-atomic resolution. These studies provide not only the protein structures, but additionally provide the details of chromophore-protein interactions and the basis for understanding energy-transfer mechanisms and kinetics.

I. Introduction Cyanobacteria, for which the fossil record dates back at least three billionyears, are the oldest oxygenevolving organisms (for reviews see Brock, 1973; Stanier, 1977; Stanier and Cohen-Bazire, 1977; Rippka and Herdman, 1985; Fay and Baalen, 1987). Cyanobacterial oxygenic photosynthesis made a basic contribution to the development of the present oxygen-enriched atmosphere, and with this oxygen

they practically ‘poisoned’ the world of anoxygenic photosynthetic bacteria, some of which nonetheless survived in anoxygenic ecological niches such as the bottom of ponds. The cyanobacterial photosynthetic apparatus houses photosystems I and II as well as the cytochrome complex and the ATP synthetase (see Chapters 8–11 for details). By means of endosymbiosis, cyanobacteria are considered to be the precursors) of the chloroplasts of all oxygen-evolving eucaryotic organisms, including the red and green

Abbreviations: AA – amino acids; or of cryptomonad phycocyanin-645; subunit of allophycocyanin; of allophycocyanin B; apcX– gene encoding APC-related protein or core component of the phycobilisome; APC I – first allophycocyanin-enriched peak eluted on ion-exchange chromatography; APC II – second allophycocyanin-enriched peak eluted on ion-exchange chromatography; APC – allophycocyanin; AP-B – allophycocyanin B; subunit of phycocyanin; subunit of phycoerythrocyanin; subunit of R-phycocyanin (red algae); B-PE – B-phycoerythrin from red algae (hexameric aggregate with a b-PE – b-phycoerythrin from red algae (trimeric aggregate without allophycocyanin subunit with an estimated molecular mass of 16.5 kDa; of allophycocyanin; BChl – bacteriochlorophyll; of phycocyanin; of cryptomonad phycocyanin-645; of phycoerythrocyanin; of Rphycocyanin (red algae); CpeC, CpeD – gene product encoded by cpeC, cpeD, etc.; cpcX– gene encoding phycocyanin and related proteins; C-PC – C-phycocyanin; cpeX – gene encoding phycoerythrin subunit or related protein; C-PE – C-phycoerythrin; Chl – chlorophyll; DEAE – diethylaminoethyl, ion-exchange group; oxidoreductase; fs – femtosecond; gamma subunit ofphycoerythrin in red algae and PE-II of cyanobacteria; HPLC – high performance liquid chromatography; linker polypeptide; linker phycobiliprotein; LHCII – peripheral light-harvesting antenna complex of Photosystem II, a chlorophyll a/b complex; LHP – light-harvesting polypeptide; polypeptide; linker polypeptide; – refers to a linker polypeptide (L) having a mass of Y, located at a position X in the phycobilisome, where X can be R (rod), RC (rodcore junction), C (core) or CM (core-membrane junction) and n is the number ofthe linker polypeptide when more than one linker have the same mass. When necessary the abbreviation for a linker is appended to that of its associated phycobiliprotein (nomenclature according to Glazer, 1985). mpeX – gene encoding marine phycoerythrin II subunit or related protein; ORF – open reading frame; PBS – phycobilisome(s); PC – phycocyanin; PC-645 – phycocyanin-645 from the cryptomonad Chroomonas sp.; PCB – blue-colored phycocyanobilin chromophore of phycobiliproteins; PCC – Pasteur Collection of Cyanobacteria, Institut Pasteur, Paris; PCR – polymerase chain reaction; PE – phycoerythrin; PE-545 – phycoerythrin-545 from the cryptomonad Cryptomonas maculata; PEB – red-colored phycoerythrobilin chromophore of phycobiliproteins; pecX – gene encoding phycoerythrocyanin subunit or related protein; PEC – phycoerythrocyanin; pI – isoelectric point of a protein (net charge = 0 at the pH-value of the solvent); PTH – phenylthiohydantoin (derivative of an amino acid); PUB – phycourobilin chromophore of phycoerythrins; PXB – cryptoviolin chromophore of the subunits of phycoerythrocyanin (PEC); RC – reaction center; REP – repetitive domains on the core-membrane linker polypeptide with amino acid sequences similar to each other domain and to linker polypeptides ofthe phycobilisome; Rubisco – ribulose-l,5-bisphosphate carboxylase/oxygenase; SDS-PAGE – polyacrylamide gel electrophoresis in the presence of sodium dodecylsulfate; SEC – size-exclusion chromatography.

142 algae, cryptophytan algae and plants; all have similar photosynthetic apparatuses (Bryant 1987, 1992; Stevens and Bryant, 1988; see Chapter 5). In the light reactions of oxygenic photosynthesis in cyanobacteria, the red and blue wavelengths of the visible light are mainly absorbed by cyclic tetrapyrroles, the chlorophylls, while the green, yellow, and orange wavelengths are mostly absorbed by open-chained tetrapyrrole pigments, the phycobilins. Pigments collect or harvest the light energy, trap the excitation energy at the ‘special pair,’ and finally transduce the light energy into stably separated charges. Pigments alone, however, are not able to perform the primary steps in the present form of photosynthesis. Proteins are essential elements that orient the pigments, give them the appropriate conformation and physical separation, and modulate the absorption properties needed for the special steps in the light reactions. Thus, analyses of protein structure are essential for an understanding of photosynthesis. The following presentation will mainly focus on research in the analysis of protein composition and structure of antenna pigmentpolypeptide complexes involved in cyanobacterial photosynthesis. Great structural variability is found among the supramolecular light-harvesting complexes (also referred to as light-harvesting organelles or antennae) in photosynthetic purple and green bacteria, cyanobacteria, algae and plants (Zuber and Brunisholz, 1991). This structural variability presumably reflects the different solutions to the same problem: how to collect most efficiently light of differing wavelength compositions (qualities) and intensities. The light-harvesting antennae and reaction centers (RC) of purple bacteria are located within the chromatophores, intracytoplasmic membranes differentiated from but connected to, the cytoplasmic membrane of the cell. Light-harvesting polypeptides surround the reaction centers forming the core antenna (LHC I) present in all purple bacteria photosynthetic membranes and absorbing at 870/890 nm (Rhodospirillum rubrum) or 1015 nm (Rhodopseudomonas viridis). In many species of purple bacteria additional and variable antennae (LHC II) surround the LHC Ireaction center complexes (Zuber and Brunisholz, 1991). The antenna complexes of green photosynthetic bacteria are located in unique, extramembranous, cigar-shaped bodies known as chlorosomes. These structures, which are approximately 100 ×30 ×12

Walter A. Sidler nm, are attached to the inner surface of the cytoplasmic membrane and may best be described as ‘sacks of bacteriochlorophyll’ (Staehelin et al., 1980; Staehelin, 1986). Chlorosomes are enclosed by a 3.5 nm-thick galactolipid-protein envelope. The chlorosomes of the green gliding bacterium Chloroflexus aurantiacus contain three proteins with apparent masses of 18 kDa, 11 kDa and 5.6 kDa; BChl c is the major antenna pigment of chlorosomes, although smaller amounts of BChl a also occur in these antennae. Although similar in function, the chlorosomes of green sulfur bacteria contain nine polypeptides and are structurally distinct from those of Chloroflexus aurantiacus (Chung et al., 1994). Excitation energy is transferred from the BChl c (BChl d or BChl e in some species) in the extramembrane chlorosome, through a BChl acontaining baseplate structure to a membrane-bound antenna complex and finally to the reaction centers (Gerola et al., 1988; Zuber and Brunisholz, 1991). The light-harvesting antenna system in green plants is composed of two types of Chl-protein complexes. The first type are the core-antenna complexes, that are tightly associated with the PS II or PS I reaction center complexes. These include the CP43 and CP47 Chl a-protein complexes (see Chapter 8); in PS I the antenna Chls are an integral part of the reaction complex itself (see Chapter 10). The second type of antenna system is the light-harvesting complex of PS II (LHC II), a Chl a/b -xanthophyll complex that is not essential for the structure and function of the PS II core complex (Thornber, 1986; Thornber et al. 1988), but which has essential regulatory functions (Anderson and Andersson, 1988). LHC II is the most abundant of the pigmented complexes in plants and contains about 50% ofthe total chlorophyll on Earth and one-third of the protein of plant thylakoids. The three-dimensional organization of the LHC IIb complex has been analyzed by electron microscopy and electron diffraction studies of two-dimensional crystals (Kühlbrandt and Wang, 1991). Cyanobacteria, cyanelles of some biflagellated protozoans such as Cyanophora paradoxa, and the chloroplasts of red algae do not contain LHC II (although the latter containrelatedproteins associated with PS I; Wolfe et al., 1992). Their light-harvesting antenna complexes for PS II, and to some extent for PS I, are large multiprotein organelles that have a molecular mass of about Da (depending on the organism and growth conditions) that are located on the stromal side of the thylakoid

Chapter 7 Phycobilisome and Phycobiliprotein Structures membranes. These so-called phycobilisomes (PBS), first described by Gantt and Conti (1966a,b; 1969), are the main light-harvesting antennae in these organisms. Phycobilisomes, like chlorosomes, are extramembranous antenna structures. Phycobiliproteins, a brilliantly colored family of watersoluble proteins bearing covalently attached openchain tetrapyrroles known as phycobilins, are directly involved in light absorption and energy transfer to the PS II reaction centers in the thylakoid membrane. PBS absorb visible light in the wavelength range 450–665 nm and extend the spectral range for photosynthetic light harvesting to the region between the red and blue absorption bands of Chls a and b. The PBS structure and composition are variable in the course of adaptation processes to varying conditions of light intensity, light quality and nutrient availability (see Chapter 21). Cryptomonads, a group of biflagellated eucaryotic algae with features characteristic of protozoa, contain phycobiliproteins in their intrathylakoidal lumen that is formed by a splitting of the thylakoid membrane (Gantt, 1979, 1980a; Wehrmeyer, 1970; MacColl and Guard-Friar, 1987a; also see Chapter 5). Electron micrographs of cryptomonads do not show structures similar to the PBS observed in cyanobacteria and red algae. Only one phycobiliprotein type, that may be either red-, purple- or blue-colored, is present in each cryptomonad. Interestingly, aChla/c light-harvesting complex, related to LHC II of higher plants, has been found in addition to the phycobiliproteins in the chloroplasts of cryptomonads (Ingram and Hiller, 1983; Rhiel et al., 1987; Sidler et al., 1988; Rhiel et al., 1989). Phycobilisomes and phycobiliproteins have been extensively reviewed. Additional specialized information on various aspects of these proteins and the structures which they form can be found in the following: Bogorad, 1975; Bryant, 1987, 1988, 1991; Cohen-Bazire and Bryant, 1982; Gantt, 1975, 1980b, 1986, 1988; Glazer, 1976, 1981, 1982, 1983; 1984, 1985, 1987, 1988, 1989; Holzwarth, 1991; MacColl, 1982; MacColl and Guard-Friar, 1987b; Mörschel and Rhiel, 1987; Rosinski et al., 1981; Rüdiger, 1975, 1980, 1994; Scheer, 1981, 1982, 1986;Tandeau de Marsac, 1977, 1983, 1991; Tandeau de Marsac and Houmard, 1993; Tandeau de Marsac et al., 1988, 1990; Wehrmeyer, 1983a, 1983b, 1990; Zilinskas and Greenwald, 1986; Zuber, 1978, 1983, 1985, 1986, 1987,1988; Zuber et al., 1985, 1987).

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II. Phycobilisomes

A. Electron Microscopy of Phycobilisomes In electron microscopic studies, PBS of red algae and cyanobacteria have been found to be regularly arranged inparallel rows onthe thylakoid, and freezefracturing showed them to be associated with membrane particles, possibly PS II (Bryant et al., 1979; Mörschel and Mühlethaler, 1983; Mörschel and Rhiel, 1987). The regularity of the PBS arrays suggests that the Chl-protein complexes of PS II, to which most of the PBS are presumably attached, must likewise be regularly displaced in the plane of the thylakoid membrane. Electron microscopic analyses of freeze-fractured cells from PBScontaining organisms provided strong evidence that each PBS is associated with a pair of membrane complexes approximately 10 nm in diameter (Staehelin et al., 1978; Giddings et al., 1983). These data are strongly supported by biophysical measurements indicating that two PS II reaction centers compete for the excitation energy of one PBS in Synechococcus sp. strain PCC 6301 (Manodori et al., 1985). Different morphological types of PBS have been described in cyanobacteria and red alga (Wehrmeyer 1983a): (1) hemidiscoidal; (2) hemiellipsoidal; (3) bundle-shaped; and (4) block-shaped. Block-shaped structures have only been reported thus far for the red alga Griffithsia pacifica (Gantt and Lipschultz, 1980). The only organism known to have bundleshaped PBS is the cyanobacterium Gloeobacter violaceus, which has no thylakoid membrane (Guglielmi et al., 1981). The PBS of Gloeobacter violaceus, which form a cortical layer on the inner surface of the cytoplasmic membrane, consist of a bundle of six rods. Each rod is 50–70 nm in length, 10–12 nm in diameter and is composed of eight to twelve disc-shaped subunits about 6 nm in thickness. Hemiellipsoidal PBS were the first type to be isolated and examined (Gantt and Lipschultz, 1972). Originally this type of PBS had been reported to occur only in red algae such as Porphyridium cruentum (Gantt and Lipschultz, 1972) and Gastroclonium coulteri (Glazer et al., 1983). However Guglielmi and Cohen-Bazire (1984) were also able to identify hemiellipsoidal PBS in cyanobacteria. Electron microscopic studies on this type of PBS with its large size (about Da) produces a superposition of stain layers in the electron

144 micrographs which are difficult to interpret. Face views from such isolated PBS were studied by Gantt et al. (1976), while top-view electron micrographs of Phorphyridium cruentum PBS on thylakoid membranes were presented by Staehelin (1986). Lange et al. (1990) proposed that the hemiellipsoidal PBS structure consisted of two rows of six rods; they proposed that these PBS were composed of halves resembling two adjacent hemidiscoidal PBS with width, length and height of 55–65 nm, 18–22 nm, and 35–40 nm, respectively. A new morphological type of PBS was recently discovered by Wehrmeyer et al. (1988) in the cyanobacterium Phormidium persicinum. From its dimensions (width, 65–80 nm; height, 35–40 nm; thickness, 20 nm) and mass (about this PBS represents an intermediate class between hemidiscoidal and hemiellipsoidal PBS. Hemidiscoidal PBS are the most common and best described PBS structures from various cyanobacteria (Bryant et al., 1979; Nies and Wehrmeyer, 1980, 1981; Rosinski et al., 1981; Mörschel and Rhiel, 1987), in the red algae Rhodella violacea (Mörschel et al., 1977) and Porphyridium aerugineum (Gantt et al., 1968) and in the cyanelle of the dinoflagellate Cyanaphora paradoxa (Giddings et al., 1983). The hemidiscoidal PBS (e.g., those of Calothrix sp. strain PCC 7601) exhibits a welldefined structure in electron micrographs (see Fig. 1 A, B). Hemidiscoidal PBS can be described as organelles, about 70 nm along the base, 30–50 nm in height and 14–17 nm in width, attached to the stromal side of the thylakoid membrane (Mörschel et al., 1977; Bryant et al., 1979; Glazer, 1984). These PBS have a mass of 4.5 to Da and contain 300– 800 covalently bound phycobilin chromophores (Zuber, 1987). Their bases are physically and energetically coupled predominantly to the PS II complexes that are embedded in the thylakoid membrane. In electron micrographs (Fig. 1 A, B), two discreet PBS subdomains are visible: the ‘core’ and the ‘peripheral rods’. The first subdomain, the PBS core, is seen in front view as either two or three circular objects arranged side-by-side or stacked to form a triangle. This subdomain is formed of either two (e.g., Synechococcus sp. strain PCC 6301; Glazer, 1982) or more commonly three cylindrical subassemblies (e.g., Calothrix sp. PCC 7601; see Fig. 1 A, B). Each of these core cylinders is composed of four stacked discs of about 3.5 nm in thickness

Walter A. Sidler (Bryant et al., 1979). In PBS with bicylindrical cores, these cylinders lie side-by-side on the surface of the thylakoid membrane, with each cylinder presumably making a close contact to one of two PS II particles embedded in the thylakoid membrane. The nature of the additional physical contacts to PS I are not yet known. In PBS containing tricylindrical cores the third cylinder is stacked onto the basal two, producing a pyramidal structure. Recent studies indicate that some cyanobacteria produce PBS with additional core elements attached to the three-cylinder core, thus effectively forming ‘four-cylinder’ cores (see Section III B and C). From the core of typical hemisicoidal PBS, six cylindrical rods, forming the second and more peripheral subdomain, radiate outwards (Fig. 1 A, B). Stacks of disks, each 6 nm in thickness and 11 nm in diameter, make up the rods. The length ofthe rod cylinders depends upon the source organism and the cell growth conditions (light quantity and quality, nutrient availability) and can vary between 12–36 nm (Bryant et al., 1979; Glazer et al., 1979). Electron microscopic analyses show a subdivision of the 6 nm-thick disks into two discs with a thickness of ~3 nm thickness. Electron microscopy and X-ray crystallography have shown that the 3 nm-thick disks represent trimers, the fundamental building blocks of the peripheral rods (Bryant et al., 1976; Schirmer et al., 1985). Pairs of these trimers are stacked together face-to-face to produce two to six 11 ×6 nm hexameric subassemblies per rod (Schirmer et al., 1986; Ficner et al., 1992). The number and the size of peripheral rods are not correlated with the core size. In the PBS of the cyanobacterium Phormidium persicinum ten peripheral rods (two bundles of five laterally associated rods) radiate from a three-cylinder core (Wehrmeyer et al., 1988).

B. Isolation of Phycobilisomes Methods for isolating intact PBS are a prerequisite for electron microscopy of PBS and for studies of their subcomplexes. The original method, yielding very pure PBS, was developed by Gantt and Lipschultz (1972). PBS were stabilized in highmolarity potassium phosphate buffer (0.75 M Kphosphate, pH 6.8 at room temperature), released from the thylakoids by treatment with Triton-X-100, and isolated by ultracentrifugation on sucrose step gradients prepared with 0.75 M K-phosphate (for a

Chapter 7 Phycobilisome and Phycobiliprotein Structures

review see Gantt, 1980b, 1988).Nies and Wehrmeyer (1980), however, required K-phosphate concentrations as high as 0.9 M for stabilization of PBS from Mastigocladus laminosus and for their isolation by ultracentrifugation. Füglistaller et al. (1984) developed a method for the isolation of large quantities of PBS from Mastigocladus laminosus in 0.9 M potassium phosphate buffer. PBS were released from the membranes by solubilization in 2% (v/v) Triton-X-100 and were precipitated with 15% (w/v) polyethylene-glycol6000.After centrifugationPBS could been recovered as a pellet from the resulting interphase. In general, proteolytic degradation of polypeptides in intact PBS in high-molarity phosphate buffer does not occur. However, at lower ionic strength PBS dissociate into their subcomplexes and without the addition of proteinase inhibitors, linker polypeptides are degraded rapidly by proteolysis. A concise review of available methods has been published (Glazer, 1988). Despite the highly polar and hydrophilic nature of PBS polypeptides, revealed by amino acid and gene sequence analysis (see below), a hydrophobic interaction mechanism for the formation and stabilization of the PBS structure is indicated by the fact that the structures are more stable at room temperature than at 4 °C and that high salt concentrations are required for their stabilization.

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C. Phycobilisome Components 1. Variability of the Polypeptide Composition The polypeptides making up the PBS may be grouped into three classes: (1) phycobiliproteins; (2) linker polypeptides; and (3) PBS-associated proteins (e.g., oxidoreductase = FNR). The polypeptide composition of PBS varies widely among strains of cyanobacteria. Moreover, for a single strain it also depends upon the environmental conditions such as nutrient availability, temperature, light quality and light intensity as shown in Table 1 for four different cyanobacteria. The hemidiscoidal PBS of Synechococcus sp. strain PCC 7002 are made up of 12 different polypeptides; the PBS of the thermophilic cyanobacterium Mastigocladus laminosus and the closely related mesophilic Anabaena sp. strain PCC 7120 are assembled from 16–18 different polypeptide types; and the PBS of Calothrix sp. strain PCC 7601, performing complementary chromatic adaptation (see Chapter 21) are assembled of 16 polypeptides when grown in green and white light conditions, of 15 polypeptides when grown under red light conditions, and of 13 polypeptides when grown under sulfurlimited conditions. (It should be noted that it has not been established whether

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oxidoreductase, a component of the PBS of Synechococcus sp. strains PCC 7002, 6301, and 7942 (Schluchter and Bryant, 1992) is a structural component of the PBS of Mastigocladus laminosus, Anabaena sp. PCC strain 7120, or Calothrix sp. strain PCC 7601 (see Section II C, 5). It should be noted that the degree of PBS compositional variability, which reflects the ability of an organism to adapt to environmental changes, varies from strain to strain. Adaptation mechanisms occurring within the peripheral rods of PBS are presently better understood than those that occur in the core.

2. Phycobiliproteins Cyanobacteria, red algae and cryptomonads frequently appear in masses known as blooms. Upon cell lysis, a brilliantly colored cell liquor is released. One hundred years ago, Molisch identified the redcolored, water soluble ‘Rhodophyceen Rot’ as a

Walter A. Sidler

chromophore-carrying protein (Molisch, 1894). The colors of the phycobiliproteins originate mainly from covalently bound, open-chain tetrapyrrole chromophores known as phycobilins. As noted above, PBS consist of both pigmented phycobiliproteins (~80% of the PBS by mass) and non-pigmented (~20% of the PBS by mass) linker polypeptides (Tandeau de Marsac and Cohen-Bazire, 1977; Glazer, 1984). On the basis of their visible absorption properties, the phycobiliproteins have been assigned to four spectroscopic classes: (1) Phycoerythrocyanin (PEC, see Fig. 2) which is primarily found in certain filamentous, heterocyst-forming cyanobacteria such as Fischerella sp. (also known as Mastigocladus laminosus), Anabaena sp. and Nostoc sp. (Bryant, 1982); (2) Phycoerythrins (PE, 565–575 nm; see Fig. 18 below). PEC and PE are found at the core-distal ends of the peripheral rods. (3) Phycocyanins (PC, see Fig. 2). PC constitutes the portion of the peripheral rods

Chapter 7 Phycobilisome and Phycobiliprotein Structures

adjacent to the core. (4) Allophycocyanin (APC, see Fig. 2) which forms the major component of the PBS core substructure. The allophycocyanin family includes minor phycobiliproteins such as an polypeptide (denoted and a polypeptide (denoted which form complexes with APC (Lundell and Glazer, 1981; Suter et al., 1987; Rümbeli et al., 1987a; Bryant et al., 1990). Although the prefixes to biliprotein classes originally indicated the type of source organism: C-, cyanobacterial; B-, Bangiophycean; and R-, Rhodophytan; these designations are now used to denote spectral properties of phycobiliproteins.

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Phycobiliproteins from cyanobacteria and red algae are hetero-monomers consisting of two different subunits, and which are present in equimolar stoichiometry and which differ in molecular mass (160–184 ammo acid residues each), amino acid sequence, and chromophore content. The fundamental assembly unit for all PBS is a stable phycobiliprotein trimer forming a toroidalshaped aggregate with a diameter of 11 nm and a thickness of 3–3.5 nm with a central hole 3 nm in diameter as described in Section IV A, 9a (see Figs. 13,14, and21 below). Hexamers are formedby faceto-face aggregation of the trimeric disks with or without the inclusion of a linker-polypeptide in the

148 central cavity (Schirmer et al., 1986, 1987). Instead of a linker polypeptide, PE hexamers of some marine cyanobacteria (Wilbanks and Glazer, 1993b) and red algae (Koller and Wehrmeyer, 1977; Glazer and Hixson, 1977) contain a third type of phycobiliprotein, the subunit (about 240–300 amino acid residues) in the central cavity of the hexamer (Ficner and Huber, 1993). subunits are bifunctional phycobiliproteins that act as light-harvesting phycobiliproteins and as linker-polypeptides. A fourth unique type of phycobiliprotein is the phycocyanobilin (PCB)-carrying core-membrane linker phycobiliprotein, formerly known as the ‘anchor protein.’ Two copies of this multifunctional protein (mass 70–128 kDa) are present per PBS core. Cryptomonads contain phycobiliproteins, related to that are associated with a fifth type of phycobiliprotein, the cryptomonad subunit family. A sixth phycobiliprotein type is phytochrome, a photoregulatory receptor protein found in low amounts in plants (Rüdiger, 1980).

3. Pigments The brilliant colors of the phycobiliproteins originate from covalently attached, linear tetrapyrrole prosthetic groups, known as phycobilins (Rüdiger, 1975,1980, 1994; Scheer, 1981).As many as three chromophores may be bound to a single or polypeptide (Glazer, 1985, 1989). Phycobilin chromophores are generally bound to the polypeptide chain at conserved positions either by one cysteinyl thioether linkage through the vinyl substituent on the pyrrole ring A of the tetrapyrrole (Fig. 3 A, B, C, E) or occasionally by two cysteinyl thioether linkages through the vinyl substitutes on both the A and the D pyrrole rings (Fig. 3 D, F; Glazer, 1985; 1989). In the phycobiliprotein structure they are maintained in an extended conformation through interactions with the protein (Scheer, 1981; Duerring et al., 1990). Nine different bile pigments with different numbers and arrangements of conjugated double bonds are known spectroscopically and structurally. Four main types are present in cyanobacteria and red algae: the blue-colored phycocyanobilin (PCB); the red-colored phycoerythrobilin (PEB); the yellow-colored phycourobilin (PUB); and the purple-colored phycobiliviolin (PXB; also named cryptoviolin). In addition to these four main types, five additional chromophores have recently been demonstrated to occur in the cryptomonad phycobiliproteins. Three

Walter A. Sidler chromophores have been found in the cryptomonad strain CBD phycoerythrin-566: Cys-bilin 618, DiCys-bilin 584, and Cys-bilin 584. A common and novel element in these new structures is an acryloyl substituent at C-12 of ring C in the open-chained tetrapyrroles. (Wedemayer et al., 1991). The green phycobilin chromophore of the cryptomonad phycocyanin has been determined by magnetic resonance spectroscopy to be mesobiliverdin and the purple, doubly linked 50/61 chromophore to be 15,16-dihydrobiliverdin (Wedemayer et al., 1992). All phycobilins are related to biliverdin, which is their biosynthetic precursor. Phycobilin structure, biosynthesis and regulation by light was recently reviewed by Rüdiger (1994; also see Chapter 17). Biliverdin is an oxidative degradation product of heme, theprostheticgroup in hemoglobin,myoglobin and cytochromes. Not only are the structures of phycobilins related to heme, but a surprising homology was also discovered between the protein structure of myoglobin and phycobiliprotein subunits (Schirmer et al., 1985). The different absorption properties of the phycobilins are predominantly caused by differences in the number of conjugated double bonds and the side chains in the tetrapyrrole prosthetic groups. NMR studies on bilin peptides (Glazer, 1985) revealed that the singly-linked forms of these four bilins are all isomers containing two keto groups, seven carbon-carbon double bonds and one carbon-nitrogen double bond. Five (phycourobilin) to nine (phycocyanobilin) of these double bonds may occur in conjugation (see Fig. 3), reflecting the long wavelength absorption maxima of these bilins. At least three elements are combined to generate the remarkable spectroscopic diversity exhibited by the phycobiliprotein family: (1) chemically distinct chromophores with varying numbers of double bonds but at conserved sites of attachment within the primary structure of the proteins; (2) chemically distinct chromophore-protein linkages (bilins may be singly or doubly linked to the polypeptide chain); (3) distinctive chromophore environments contributed by the different polypeptide chains. Other elements that may contribute to spectroscopic diversity include chromophore-chromophore (exciton) interactions and spectroscopic changes brought about by environmental perturbation by specific linker polypeptides.

Chapter 7 Phycobilisome and Phycobiliprotein Structures

4. Linker Polypeptides and Phycobilisome Assembly PBS are not only composed of phycobiliproteins but contain also a small quantity (5–10) of non-pigmented proteins, the linker polypeptides, that amount to about 10–20% of the total protein content of PBS (Tandeau de Marsac and Cohen-Bazire, 1977). These proteins are described in detail in Section V below. Linker polypeptides induce a face-to-face aggregation of PE, PEC and PC trimers, and additionally cause the tail-to-tail joining of hexameric assemblies to form larger aggregates such as peripheral rods and core-cylinders. These proteins also serve to connect the rods to the core, and last but not least direct the assembly of the PBS core and its attachment to the thylakoid surface. Small linker polypeptides (CpcD and ApcC) terminate the rod- and cylinder-stacking

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reactions. The linker polypeptides are probably located mainly in the central cavity of the hexamers. Two to four of these stacked hexamers form the PBS rods. APC trimers, held together by the 70–128 kDa core-membrane linker phycobiliprotein, a PCB carrying polypeptide, form the core cylinders (see Section III C and Fig. 8 below). Six to eight rods are coupled to the core cylinders by rod-core linkers with masses of 29–34 kDa. By interacting directly with the chromophores or by causing changes in chromophore environments indirectly, linker polypeptides can modulate the spectral properties of the different PBS subassemblies.

5.

Oxidoreductase (FNR)

Polypeptide analyses of PBS by SDS-PAGE has shown that many cyanobacterial phycobilisomes

150 contain substoichiometric amounts of polypeptides with masses of approximately 45–50 kDa. In Synechococcus sp. strains PCC 7002 and PCC 7942, such proteins have been shown to cross-react strongly with antibodies to FNR and/or were directly shown by amino-terminal sequence analysis to be FNR (Schluchter and Bryant, 1992). The amino-terminal domain of FNR was shown to be 78% similar to the (CpcD). FNR might be attached to peripheral rods of PBS by this N-terminal domain at positions topologically equivalent to the binding sites (Schluchter and Bryant, 1992; Zhao et al., 1992). Approximately two copies of FNR occur per phycobilisome in Synechococcus sp. strain PCC 7002 (J. Zhao and D. A. Bryant, personal communication). Recent studies have shown that the amino-terminal domain of the FNR of Anabaena sp. strain PCC 7119 also resembles CpcD, although the flexible hinge region connecting this domain to the catalytic portion of FNR was somewhat longer than for the FNR of Synechococcus sp. PCC 7002 (Fillat et al., 1993; W. M. Schluchter and D. A. Bryant, personal communication). The significance of this localization of FNR is not presently understood, but one function could be to tether FNR near the reducing side of the PS I reaction center. This could effectively reduce the diffusion distance for ferredoxin to reach the enzyme and thereby reduce the production of toxic superoxide radicals formed by oxidation of reduced ferredoxin by oxygen. Such a localization of FNR has not yet been shown to occur in Mastigocladus laminosus PBS, although a somewhat larger polypeptide observed in PBS preparations was sometimes caused by contaminating Rubisco large subunits (Glauser, 1991).

D. Energy Transfer in Phycobilisomes Isolated intact PBS exhibit a fluorescence emission maximum of ~680 nm (Gantt and Lipschultz, 1973). The absorbed light energy harvested at the periphery of the PBS is transferred to the PS II RC-complex by radiationless excitation energy transfer with an efficiency of > 95% (Manodori et al., 1984, 1985; Glazer, 1989). This implies that the energy transfer mechanism must proceed rapidly in order to avoid energy losses by competing radiative or non-radiative decay processes. Light energy is absorbed mainly by the peripheral rods, where the shortest wavelength absorbing phycobiliproteins (PE or PEC) are located. The light energy absorbed by PE or PEC is transferred

Walter A. Sidler by radiation-less, dipole-induced dipole resonance energy transfer (‘Förster energy-transfer mechanism’; Förster, 1965, 1967) to C-phycocyanin (C-PC) and then to allophycocyanin (APC) as energy is transmitted to PS II (and partially) to PS I reaction centers through the terminal emitters of the PBS (Suter and Holzwarth, 1987; Glazer, 1989; see also Sections IV A, 14c). Since a typical hemidiscoidal PBS carries 300–800 phycobilin chromophores, the PBS greatly augments the limited absorption crosssection of the approximately 50 Chl a molecules that are associated with each PS II reaction center. Energy transfer from phycobilisomes to Chl a associated with PS II in the thylakoid membrane is very fast. The fluorescence rise-time from Chl a occurs 150 picoseconds (ps) after the excitation of PE in cells of the red alga Porphyridium cruentum , and 120 ps after the excitation of PC in the cyanobacterium Synechococcus sp. strain PCC 6301 (Yamazaki et al., 1984). Energy transfer from the periphery of the PBS rods to the terminal emitters or/and in the core occurs in 56±8 ps based upon picosecond fluorescence techniques applied to PBS of Synechocystis sp. strain PCC 6701 (Glazer et al., 1985a, b). Based upon the results described above and as shown diagramatically in Fig. 4, it is generally believed that a directional and highly efficient energy transfer occurs, from the rods to the core of the PBS and finally to the chlorophyll proteins of the thylakoids. This unidirectional energy transfer between heterogeneous components of the PBS is a consequence of the energy difference of absorption between PE (PEC), PC, APC and the terminal emitters in the core and the modulation of the spectroscopic properties of these phycobiliproteins by the different linker polypeptides as described in Section V (see Glazer, 1989 for additional details).

E. Regulation of Phycobilisome Composition The composition of PBS in cyanobacteria and red algae change in response to environmental changes in light intensity, light quality (only cyanobacteria) and nutrient availability (Table 1; see also Chapter 21). Cyanobacteria, like virtually all other photosynthetic organisms, generally increase their cellular contents of antenna proteins and pigments in response to low light intensity (for additional information and reviews, see Bryant, 1982, 1987, 1988; Tandeau de Marsac and Houmard, 1993; Grossman, 1990; Grossman et al., 1993). Cyanobacteria and red algae

Chapter 7 Phycobilisome and Phycobiliprotein Structures

grown under high light have the lowest, and cells grown under low light the greatest, amounts of phycobilins per cell and the highest number of PBS per of thylakoids (Wehrmeyer, 1990). The PBS composition of some cyanobacteria (e.g., Calothrix sp. strain PCC 7601) is regulated by light quality. This phenomenon of structural response to different spectral light conditions is referred to as chromatic adaptation (Bogorad 1975; Tandeau de Marsac, 1977, 1983, 1991; Tandeau de Marsac and Houmard, 1993; Grossman, 1990; Grossman et al., 1993; see Chapter 21). Only cyanobacteria with PE perform complementary chromatic adaptation, and strains containing PEC do not exhibit this phenomenon (Bryant, 1982). The molecular mechanism of regulation of PBS phycobiliprotein composition by light quality has been subject to intensive studies at the genetic level for several years (for reviews see: Conley et al., 1988; Grossman et al., 1986, 1988, 1993; Grossman, 1990; Tandeau de Marsac, 1983, 1991; Tandeau de Marsac et al., 1988; Tandeau de Marsac and Houmard, 1993) and is summarized in Section VI (also see Chapter 21). The typical hemidiscoidal PBS of Calothrix sp. strain PCC 7601 cells grown in green-enriched light (e.g., cool-white fluorescent light) contain APC, CPC and C-PE I. An additional cpc gene set, encoding the ‘red-light inducible,’ blue-colored, and redabsorbing PC, is present which is expressed instead of the red-colored, green-absorbing C-PE when cells are grown in red light. Specific linker polypeptides are expressed for each C-PE hexamer or each C-PC 2 hexamer. A third C-PC gene set and corresponding

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linker polypeptides is exclusively expressed upon sulfur-starvation (Mazel and Marlière, 1989). These results indicate that the natural environmental conditions, to which these organisms are exquisitely able to adapt, can be directly imprinted in the amino acid sequences of their most abundant proteins. The PBS of Calothrix sp. strain PCC 7601 have a molecular mass of Da (from cells grown in white or green light). They consist of 88% pigmented and 12% non-pigmented proteins. The Calothrix sp. PBS can be described as hemidiscoidal organelles (71 nm along the base, about 50 nm in height and 1214 nm in thickness) that are attached to the stromal side of the thylakoid membrane (Rosinski et al., 1981). In electron micrographs of Calothrix sp. PBS, the two PBS-subdomains are clearly visible (Fig. 1 A, B). Under all light conditions a constitutive C-PC hexamer is attached to the core-proximal end of the rods. During red light growth, two additional bluepigmented C-PC hexamers are added (Fig. 1 B), whereas during growth under green or white fluorescent light three additional red-pigmented CPE hexamers are added to the peripheral rods (Fig. 1 A). A second type of chromatic adaptation has been described for Phormidium sp. strain C86 (Wehrmeyer, 1990; Westermann et al., 1993). Different PBS structure types were found in cells grown under green or red light. The PBS of cells grown in green light were nearly hemiellipsoidal with 10 rods, had a molecular mass of had a molar ratio of PE:PC:APC of 7.3:1:1, and were similar to PBS described for Phormidium persicinum. Under red-

152 light growth conditions, cells contained hemidiscoidal PBS that had six peripheral rods, a molecular mass of and a molar ratio of PE:PC:APC of 1.2:3.3:1. Different linker polypeptides compositions were found in the two PBS types. The third type of adaptation was observed upon alteration of light intensity combined with temperature changes in the thermophilic cyanobacterium Mastigocladus laminosus (58–60 °C optimal growth temperature). A ‘minimal phycobilisome’ type with an apparent molecular mass of Da was obtained at 37 °C under high-light illumination (50 and a ‘maximal phycobilisome’ type with an apparent molecular mass of Da with PEC was obtained at 48 °C under low-light conditions (10 Reuter and Nickel-Reuter, 1993). In its natural environment in Iceland, Phormidium laminosus completely out-competes Mastigocladus laminosus below 50 °C (W. Sidler, unpublished results). Cyanobacterial PBS are not only antenna complexes for light harvesting, but they can also be used as storage materials for reduced carbon and nitrogen (Bryant, 1987; see Chapter 21). Furthermore, adaptation of cyanobacteria to the environmental factors described above is sometimes accompanied by the induction of cell differentiation into heterocysts or hormogonia as observed with Calothrix sp. PCC 7601 (Rippka and Herdman, 1985; Tandeau de Marsac et al., 1988, 1990). It is suggested that a very complicated network of antagonistic and/or cooperative effectors are involved in the photoregulation of both complementary chromatic adaptation and cell differentiation processes (Tandeau de Marsac et al., 1988, 1990; Grossman, 1990; Tandeau de Marsac, 1991; Tandeau de Marsac and Houmard, 1993; Grossman et al., 1993).

III. Phycobiliproteins Constituting the Phycobilisome Core

A. The Allophycocyanin Family Allophycocyanins (APCs) assemble the PBS core with the assistance of three types of linker polypeptides: (1) the large core-membrane linker phycobiliprotein two copies per PBS); (2) rodcore linker polypeptides probably one polypeptide per peripheral rod); and (3) with small linker polypeptides (see Section V). Complexes

Walter A. Sidler containing APC are the longest wavelength absorbing and fluorescing in the PBS. The PBS core receives the excitation energy from the peripheral rods and transfers it to the Chl-proteins of the thylakoid membrane (Table 2). APC occurs mainly in the trimeric form A portion of the APC-containing complexes containing different APC subunits, either the subunit, the subunit or the phycobiliprotein-domain of the protein as described below. One part (about one half) of the APC trimers are believed to interact with the repetitive linker domains (REP domains) of the polypeptide while the other part of the APC trimers may interact with and the carboxyl-terminal domains of the and are involved in rod-core linkages. APC subunits with different migration properties on SDSPAGE have been reported by Reuter and Wehrmeyer (1988, 1990) from PBS of Mastigocladus laminosus and the red alga Rhodella violacea (Reuter et al., 1990) and denoted as and Interestingly, no heterogeneity in the amino-terminal amino acid sequences of and from Mastigocladus laminosus could be detected (Esteban et al., 1990), and until now only a single set of genes encoding the and subunits were found in this cyanobacterium (Esteban, 1993; Esteban et al., 1994). Thus, post-translational modifications (e.g., oxidation of Met (Sidler et al., 198la) may cause these heterogeneities and the possible role of these and subunits still must be determined. A second gene (apcA2), encoding an subunit (denoted with 59% sequence identity to was found in Calothrix sp. strain PCC 7601 (Houmard et al., 1988), but the function and location of this subunit is not known. The different APC subunits from Mastigocladus laminosus are uniformly 160 amino acid residues in length, but the subunits vary considerably (158 to 167 residues; Fig. 5). When conserved residues in all four APC subunits are considered, only 19% of the positions are invariant. The subunit shows a high degree of sequence identity (52%) with and is 36% identical to the phycobiliprotein-domain of the highest identity value for this domain to other phycobiliproteins. The subunit of Mastigocladus laminosus also exhibits a high degree of sequence similarity to the subunit (48% identity in the amino-terminal 150 residues). The homology within the differentAPC or subunits is smaller than in

Chapter 7 Phycobilisome and Phycobiliprotein Structures

153

154 the PC family (60–67%; Section IV A, 8 and Fig. 12). Sequence identities between the and subunits of the same phycobiliprotein decrease from APC (38%) to C-PC (26%) and PEC (21%). and are 32% identical, and 63% identical, and are 40% identical, and and 67% identical. These comparisons illustrate the divergent phylogenetic development of phycobiliprotein and subunits (also see Section IV A, 8 and Fig. 12).

Walter A. Sidler with The absorption maximum ofthis complex red-shifts to 653 nm, but the fluorescence emission maximum remains at 658– 662 nm (Füglistaller et al., 1987; Reuter and Wehrmeyer 1988). Identical data were obtained for the complex reconstituted with the protein isolated from Mastigocladus laminosus (Betz et al., 1993; Gottschalk et al., 1993) and the overexpressed in E. coli (Betz et al., 1993).

2. Allophycocyanin-B (AP-B, ApcD)

1. Allophycocyanins (APCs, ApcA, ApcB) Upon DEAE-ion exchange chromatography of phycobiliproteins from dissociated PBS (Sidler et al., 1981a; Füglistaller et al., 1984) the early eluting APC I is a fraction enriched with different APC complexes containing and and the late eluting APC-II is a fraction enriched in trimericAPC without The absorption maximum of the trimeric APC-II from Mastigocladus laminosus is at 651 nm (see Fig. 2), and the fluorescence emission maximum occurs at 658–662 nm at 22°C (Füglistaller et al., 1987; Rümbeli and Zuber, 1988; Reuter and Wehrmeyer, 1988). Allophycocyanins contain exclusively one PCB chromophore per subunit (see Figs. 5 and 6). From amino acid sequences the PCBs are shown to be singly bound to and 82 (Fig. 5; position 84 in the amino acid sequence, when aligned with C-PC). Assuming that the threedimensional structure oftrimeric allophycocyanin is similar to that of C-PC, the PCB chromophore at position is presumed to be located at the periphery of the trimer while the PCB chromophore at position would lie near the inner cavity of the trimer. However, no crystal structure for APC is available yet, although crystals of allophycocyanin have been reported (Bryant et al., 1976; Bryant et al., 1981). The trimeric forms a complex

Allophycocyanin-B (AP-B) was first purified from the unicellular cyanobacterium Synechococcus sp. strain PCC 6301 as a trimeric complex with the composition (Glazer and Bryant, 1975). AnAP-B containing complex, was subsequently described in which an subunit is substituted by a structurally similar but distinctive subunit (Ley et al., 1977; Lundell and Glazer, 1981, 1983b). The complex has a long-wavelength absorbance maximum at 654 nm and a fluorescence emission maximum at 679 nm at 25 °C. In Mastigocladus laminosus this complex was described as by Füglistaller et al. (1987) and Suter et al. (1987) but as by Reuter and Wehrmeyer (1988; also see Wehrmeyer, 1990). The subunit is encoded by the apcD gene and the nucleotide sequence for the gene from Mastigocladus laminosus shows 160 amino acid residues when the N-terminally processed Met is excluded (Fig. 5, Esteban 1993; Esteban et al., 1994). AP-B was originally proposed to function as a terminal energy emitter from phycobilisomes and to play a role in energy transfer from the PBS to the Chl-proteins of the thylakoids (Glazer and Bryant, 1975; Ley et al., 1977; Lundell and Glazer, 1981). Molecular biology has allowed this proposal to be

Chapter 7 Phycobilisome and Phycobiliprotein Structures tested. Synechococcus sp. PCC 7002 mutants, in which the apcD gene coding for was insertionally inactivated, have been constructed and characterized (Maxson et al., 1989; Bryant, 1991; Zhao et al., 1992). When grown in white light, ApcD-less mutants, in which was probably replaced by in PBS, showed a similar growth rate (~2% slower) as well as a correct and stable PBS assembly. However, the apcD mutant cells had a 35% greater doubling time than wild-type cells when grown ingreenlight. Room-temperaturefluorescence induction measurements with whole-cells and fluorescence inductionmeasurements in thepresence of DCMU showed that the mutant is unable to perform state transitions and is impaired in energy transfer from PBS to PS I that should normally occur in cells in state 2 (Zhao et al., 1992). The conclusion from these studies is that AP-B plays a critical role in energy transfer from PBS to PS I and in the partitioning of light energy between the PS I and PS II reaction centers (also see Section III E below).

3.

Subunit (ApcF)

Glazer and coworkers isolated a complex with the composition from the cores of PBS of Synechococcus sp. strain PCC 6301 (Yamanaka et al., 1982; Lundell and Glazer, 1983a; 1983b). It was part of a ‘half-core’-complex of PBS of this cyanobacterium and contained 50% of the total APC and all of the polypeptide. It was proposed that the phycobiliprotein domain of the replaces an subunit in the complex As found in Synechococcus sp. strain PCC 6301 (Yamanaka et al., 1982; Lundell and Glazer, 1983a; 1983b), this complex in Mastigocladus laminosus has a long wavelength absorption maximum and fluorescence emission maximum at 25 °C; Reuter and Wehrmeyer, 1990). The complete amino acid sequence of subunit of Mastigocladus laminosus was found to have 169 amino acid residues (Rümbeli et al., 1987a; Esteban, 1993). A 1:1 molar complex of the and subunits was isolated from a Synechococcus sp. strain PCC 7002 in which the cpcBA genes had been deleted (Bryant et al., 1990). This complex had an absorption maximum at about 616 nm, but the aggregation state of the complex was not determined; a reconstituted preparation of the purified subunit had an absorbance maximum at 618 nm.

155

Mutants, in which the apcF gene was insertionally inactivated, have been constructed in Synechococcus sp. strain PCC 7002 (Zhao et al., 1992). As found for the subunit described above, the highly conserved polypeptide was not obligately required for PBS assembly. The fluorescence emission maximum of PBS from the apcF mutant was shifted to 667 nm instead of678 nm. The apcF mutant grew more slowly than the wild-type strain in white or green light and at different light intensities (Zhao et al., 1992). Room temperature fluorescence induction experiments in the presence or absence of DCMU, and low temperature fluorescence emission spectra collected from cells in either State I or State II, suggest that the apcF mutant is impaired in energy transfer to PS II and that this impairment is much greater in cells in State II than in State I (Zhao et al., 1992; see Section III E below).

B. the Core-Membrane Linker Phycobiliprotein (ApcE) The core-membrane linker phycobiliprotein, denoted orApcE, is the largest chromoprotein in the PBS and has a molecular mass that varies from 70–128 kDa depending on the organism (see Section V B, 2). It is present in two copies per PBS. Lundell et al., (1981b) suggested the to be a new type of biliprotein and to be one of the terminal energy emitters of the PBS. Redlinger and Gantt (1981a, 1982) made similar suggestions based upon studies of the polypeptide of the PBS of the red alga Porphyridium cruentum. Low-temperature fluorescence studies of Mimuro et al. (1986a; see also Mimuro and Gantt, 1986) indicated the presence of two independent terminal emitters in the PBS of the cyanobacterium Nostoc sp. strain MAC. Moreover, these workers suggested that the phycobiliprotein provided the pathway forenergy to PS II. Basedupon the immunological properties of the (Zilinskas and Howell, 1987), as well as the amino acid sequence of its phycobiliprotein domain (see below), the protein represents a differentphycobiliprotein family. Some properties of the isolated APC-complex involving the and subunits were described in Section II A, 3 above. Genes encoding for the in cyanobacteria and red algae have been cloned and sequenced from Cyanophora paradoxa (Bryant, 1988), Calothrix sp. strain PCC 7601 (Houmard et al., 1990), Synechococcus sp. strain PCC 6301 (Capuano et al., 1991),

156 Synechococcus sp. strain PCC 7002 (Bryant, 1991), and Aglaothamnion neglectum (Apt and Grossman, 1993b) and Mastigocladus laminosus (Esteban, 1993; Esteban et al., 1994). Analyses of the deduced amino acid sequences revealed a number of remarkable structural and functional features of these polypeptides. The proteins were shown to be multifunctional, hybrid polypeptides, that can be divided into 3 to 5 domains, each consisting of approximately 220 amino acids. The amino-terminal portions of these proteins contain phycobiliprotein domains with a cysteine binding site for a single PCB chromophore. Site-directed mutagenesis experiments have shown that Cysl86 of the Synechococcus sp. strain PCC 7002 ApcE protein binds the chromophore (Bryant, 1991; Gindt et al., 1992; J. Zhou and D. A. Bryant, personal communication). Although the chromophore-binding cysteine within this domain is shifted towards the carboxyl terminus by about 38 residues when compared to the relative placement of Cys84 in other phycobiliproteins, the overall PCB binding pocket for the chromophore is highly homologous in amino acid sequence to that found in otherphycobiliproteins. In structural terms, this displacement causes the actual binding site for the chromophore to be shifted to the opposite surface of the binding pocket. The significance of this difference is not known, although only minor adjustments of the chromophore-protein interactions might be required to accommodate this change. The amino-terminal ‘phycobiliprotein’ domains of ApcE proteins are homologous to the amino acid sequences of the and subunit families of phycobiliproteins (about 35% identity; see Fig. 12 below). This domain of the polypeptide is about 65 amino acid residues longer than typical phycobiliproteins and contains a small insertion of approximately 50 to 72 residues forming a specific domain (LOOP). This ‘loop’ domain has been suggested to be involved in the attachment of the PBS to PS II and in the attachment ofthe PBS to the thylakoid membrane surface (Bryant, 1988, 1991). The carboxy-terminal portion of the contains two to four (depending on the size) ‘repeat’ or REP domains. These domains are similar in sequence to one another and are likewise similar to conserved domains ofthe rod and rod-core linker polypeptides. The REP domain structures are likely to be responsible for interactions with APC trimers that are required for the assembly the PBS core as

Walter A. Sidler discussed below (see Fig. 7 A). Finally, the polypeptide includes 2 to 5 sequence segments that form the connections between the phycobiliprotein and REP domains. These domains, denoted as ‘ARMs,’ might also form the connections between the different PBS core cylinders and provide for interactions in order to connect the two halves of the PBS (Bryant, 1988, 1991; Houmard et al., 1990; Capuano et al., 1991, 1993; see Fig. 7 A). Redlinger and Gantt (1981a, 1982) suggested that the protein anchors the PBS to the thylakoid membrane (hence the name ‘anchor polypeptide,’ sometimes used to describe this protein). Protein chemical analysis by partial proteolysis with trypsin of reconstituted PBS core complex and of native PBS from Anabaena variabilis strain M3 demonstrated that the is mainly embedded in the APC core and also demonstrated the partitioning of the polypeptide into four APC-binding domains (Isono and Katoh, 1987). Isono and Katoh (1987) also suggested that the is mainly involved in the assembly ofAPC into the PBS core structure. In the deduced amino acid sequences of the apcE genes (Bryant, 1988, 1991; Houmard et al., 1990; Capuano et al., 1991), no hydrophobic domain, which was expected to protrude into the thylakoid membrane, was present. Nonetheless, Triton X-114 phase partitioning experiments suggest that the highly polar and basic polypeptide nonetheless behaves as a hydrophobic protein and partitions into the detergent micelles (D. A. Bryant, personal communication). This result raises the possibility that the polypeptide is posttranslationally modified causing the protein to become more hydrophobic (e.g., it could be acylated). Further studies on this point are required. In the unicellular cyanobacterium Synechococcus sp. strain PCC 7002 the apcE gene encoding the polypeptide has been both insertionally inactivated or completely deleted (Bryant, 1991). No intact PBS could be isolated from either mutant, although both APC and PC were synthesized in essentially normal amounts. These and other studies show that the polypeptide plays a central role in the PBS assembly and architecture (Isono and Katoh, 1987). It is probably responsible for the attachment ofthe PBS to the thylakoid membrane (Gantt, 1988), and it transfers excitation energy from the PBS to the Chl a associated with the PS II core antenna (Redlinger and Gantt, 1982; Mimuro et al., 1986a; Glazer, 1989; Gindt et al., 1992; Zhao et al., 1992).

Chapter 7 Phycobilisome and Phycobiliprotein Structures

The apcE gene, encoding the of Mastigocladus laminosus is 3627 bp long and the deduced amino acid sequence predicts a polypeptide of 1134 amino acids with a molecular mass 127.6 kDa (Esteban, 1993; Esteban et al., 1994b). These results confirm the estimated molecular mass for this

157

polypeptide of 120 kDa by SDS-PAGE (Reuter and Wehrmeyer, 1990; Glauser et al., 1992a). It is presently the largest to have been sequenced. The amino acid sequence identity to the other known polypeptides is as high as 79%–94%. Fragments of the of Mastigocladus laminosus have been

158 determined by protein sequence analysis (Rümbeli et al., 1988). The calculated pIs for the REP domains are 5.9–9 and are typically basic as are other linker polypeptides. The calculated pIs for ARMs were calculated to be 10.4–11.4, while the pIs of the phycobiliprotein domains (pI= 4.6) differ significantly with (Esteban, 1993). The phycobiliprotein domain of this starts at residue 18 and ends at approximately residue 236, and this domain exhibits 21–35% identity to all other and phycobiliprotein subunits; the putative cysteine residue to bind the PCB chromophore occurs at residue 196. This phycobiliprotein domain is divided into two parts by a 72-amino acid residue long insertion (74–145). As noted above, it has been suggested that this loop may form a structure that serves to anchor the PBS to the thylakoid (Bryant, 1988; Capuano et al., 1991). The spacing ARM1 (residues 237–286) is adjacent to the phycobiliprotein domain and is followed by the first of four repeat domains (REP 1, residues 287–104; REP2, residues 548–663; REP3, residues 746–864 and REP4, residues 977–1094). Each REP domain is about 120 residues long and are suggested to provide the binding domains that interact with APC trimers. The REP domains show about 24–34% sequence identity to similar domains within the and linker polypeptides of Mastigocladus laminosus (Glauser et al., 1992b). The sequence identity among the REPs themselves is higher (42–53%). Higher identity is found between REP1 and REP 3 and between REP2 and REP4, possibly indicating similar functions for these pairs of domains. The high similarity of the REPs to each other suggests they may form a special linker polypeptide family enclosed within the The carboxyl-terminal portions of REP3 and REP4 are 39% and 37% identical to the small core linker polypeptide This indicates that these domains may possibly be involved in a cylinder-terminating function in the PBS. No relevant identities exist between ARMs 1–5, the domains between the REPs (ARM1, 237–290; ARM2, 405– 547; ARM3, 664–745; ARM4, 865–976; ARMS, 1095–1134). In Calothrix sp. strain PCC 7601, the apcE gene predicts a protein with a mass of 120 kDa, but SDSPAGE analyses of PBS from this species indicates that the polypeptide actually found in the PBS only has a mass of about 94 kDa (see Section III C and Fig. 24 below). The reason for this discrepancy is not clear at present, but it appears that the

Walter A. Sidler polypeptide is posttranslationally processed during or after PBS assembly in this cyanobacterium. It seems most likely that the fourth REP domain of this protein is missing in the assembled PBS. The fourth REP is expressed in Mastigocladus laminosus and also in Anabaena sp. strain PCC 7120, as indicated by the molecular weight determination for the polypeptides of these species by SDS-PAGE (see Fig. 24 below). This REP is suggested to bind additional APC, as already demonstrated by proteinbiochemical experiments with Anabaena variabilis (Isono and Katoh, 1987). If each fourth REP-domain of the two per PBS binds an additional APChexameric subassembly (i.e., the equivalent of one half of a core cylinder), this would theoretically add the equivalent of a fourth cylinder of APC per PBS. Characterization of the of Mastigocladus laminosus and other cyanobacteria by SDS-PAGE showsmainly and a minor amount The occurrence of the two forms is not regulated by light intensity (Esteban, 1993). The peptide making up the difference between these two polypeptides, probably released due to proteolysis of the subunit, was isolated from the PBS and identified by sequence analysis (Esteban, 1993; Gottschalk et al., 1994). The 23 kDa fragment contained REP4 and formed a reconstitution product with yielding a protein with an absorbance maximum at 651.5 nm and a fluorescence emission maximum at 663 nm. (Esteban, 1993;Gottschalk et al., 1994). An identical reconstitution product was obtained with the ARM4-REP4 and REP4 only from Mastigocladus laminosus (K. Locher, Diplomarbeit, ETH Zürich, 1993). A novel aspect of the architecture of the PBS from Mastigocladus laminosus and Anabaena sp. can be visualized in Fig. 7. The fourth REP-domain of the two polypeptides that occur per PBS bind an additional APC hexameric subassembly. Glauser et al. (1992a) proposed a model in which these additional APC hexamers constitute the bases of two peripheral rods. In this model the fourth repeat of binds an APC ‘hexamer’ as the first unit of an additional rod. In an alternative and perhaps more probable interpretation, the additional APC (two trimers) bound to each fourth repeat domain may be defined as core elements and not as peripheral rod elements. Two such APC portions (four trimers) would amount to the equivalent of a complete fourth core cylinder, but due to the symmetry of the two per PBS, these two APC portions (half cores) cannot be assembled

Chapter 7 Phycobilisome and Phycobiliprotein Structures

on the same side of the PBS into one typical core cylinder, but must be located as half-cylinders on both sides of the third (top) cylinder (Figs. 7 and 8). Thus, a novel PBS-type was found through comparative structural studies of Mastigocladus laminosus and Anabaena sp. PBS. The detailed molecular architecture of these PBS, however, is still not yet completely understood. Electron micrographs showing two peripheral APC substructures in addition to the tricylindrical core have been obtained from reconstituted PBS cores from Anabaena variabilis strain M3 (Isono and Katoh, 1987) andAnabaena sp. strain PCC 7120 (Fig. 7 B).

C. Determination of the Core Size by the The number of REP domains of the (which can indirectly be deduced from the molecular mass)

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determines the PBS core structure (i.e., the number of core cylinders formed by linking ofAPC trimers; Fig. 8). Cores comprised of two cylinders (e.g., those of Synechococcus sp. strain PCC 6301) contain the equivalent of eight APC trimers, and hence the polypeptide is correspondingly composed of two linker repeat domains (Capuano et al., 1991). Similarly, the polypeptide in tricylindrical cores, that contain twelveAPC trimer equivalents, has three linker-repeat domains (Bryant, 1988). The PBS cores of Anabaena variabilis strain M3, Anabaena sp. strain PCC 7120 and Mastigocladus laminosus have APC contents that are equivalent to that expected for a ‘four-cylinder’ core, and this APC is assembled by an with four REP domains (Figs. 7 and 8). In Calothrix sp. strain PCC 7601 the deduced amino acid sequence of the apcE gene product predicts a 120 kDa with four REP domains (Houmard et al., 1990). However, the carboxyl-

160 terminal part of this protein, containing the domain with similarity to is not present in the as it is in Mastigocladus laminosus (Glauser et al., 1992c). SDS-PAGE of PBS from Calothrix sp. strain PCC 7601 grown under green or red light contain polypeptides with an apparent mass of 94 kDa (see Fig. 24 below). This polypeptide must be processed at the carboxyl-terminus, since its amino-terminal amino acid sequence is identical to the deduced amino-terminus from the sequence of the apcE gene, except for the removal of the initiator methionine residue (Glauser et al., 1992c). Thus, only three REP domains can be present in the PBS, and electron micrographs of these PBS show only typical threecylinder cores (see Fig. 1 A, B). Therefore, the carboxyl-terminal fourth REP domain must be posttranslationally cleaved from a larger precursor, and no additional APC (forming a fourth-cylinder equivalent) is bound by the The function of the fourth REP encoded in the Calothrix sp. strain PCC 7601 apcE gene is not clear yet. Capuano et al. (1991) have recently suggested that both fourth linkerrepeat domains of the two ApcE proteins of Calothrix sp. strain PCC 7601 may act as a second type of ‘rodcore linker’ for the attachment of peripheral rods. These domains are proposed to bind two (of the six total peripheral rods) basal phycocyanin hexamers (not APC!) to each side of the three-cylinder core. However, a PBS containing the might be expected to bind eight peripheral rods. Conditions under which the fourth REP is not cleaved off and performs the proposed function are not known. Capuano et al. (1993) integrated the Calothrix sp. strain PCC 7601 apcE gene into the chromosome of Symchococcus sp. strain PCC 7942, which contains a and which normally assembles PBS with cores made up of two cylindrical substructures. About 5% of the PBS of this Synechococcus sp. (PIM9S1E) strain contained two heterologous Calothrix sp. strain PCC 7601 polypeptides and formed tricylindrical cores. Although both proteins and were able to exist in one PBS, heterogeneous and unstable PBS were made since each promoted the assembly of a different core substructure. Interestingly, the polypeptide was not proteolytically processed to the 94 kDa species when expressed in Synechococcus sp. strain PCC 7942.

Walter A. Sidler

D. Allophycocyanin Complexes and Their Arrangement in the PBS Core The molecular organization and function of the core in hemidiscoidal PBS is complex and is not as easy to understand as the rod structures (see Sections IV and V below). Glazer and coworkers dissociated the PBS cores of Synechococcus sp. strain PCC 6301 (and the AN112 mutant of this strain) into several well-defined multiprotein subcomplexes (Yamanaka et al., 1982; Lundell and Glazer, 1983a, b, c). These authors suggested that each ofthe two core cylinders in the PBS of Synechococcus sp. strain PCC 6301 is composed of four different subcomplexes:

Gingrich et al. (1983) succeeded in isolating a series of subcomplexes from the tricylindrical core of Synechocystis sp. strain PCC 6701, which resembled those isolated from the bicylindrical core of Synechococcus sp. strain PCC 6301. These authors suggested that the upper third core cylinder was composed of two and two subcomplexes. Anderson and Eiserling (1986) partially dissociated cores of Synechocystis sp. strain PCC 6701. They were able to isolate a ‘half core’ subcomplex containing 50% of the total APC and the total amount of the and polypeptides but no and Their results indicated that core subcomplexes 1 and 4 described above must be located at the periphery of the basal two core cylinders. Similar complexes have been isolated from the PBS from Mastigocladus laminosus (Füglistaller et al., 1987; Rümbeli and Zuber, 1988; Reuter and Wehrmeyer, 1988, 1990, Wehrmeyer, 1990; see Table 2). It was proposed that subcomplexes 2 and 3 are arranged adjacent to one another in each core cylinder. However, the unambiguous positioning of these four subcomplexes relative to one another has not been established to date, since it has not been possible to determine whether the order of the subcomplexes in the cylinder is 1, 2, 3, 4 or 1, 3, 2, 4 (Bryant, 1988). Based upon the observations described above, in

Chapter 7 Phycobilisome and Phycobiliprotein Structures

combination with results from analyses of mutants lacking core polypeptide components in Synechococcus sp. PCC 7002 performed by Bryant and coworkers (Bryant, 1988, 1991; Gindt et al., 1992), the basic model ofa hemidiscoidal PBS containing a tricylindrical core surrounded by six rods is shown in Fig. 9 A. The organization of the core subcomplexes is one of two discussed by Bryant (1988) and assumes an antiparallel orientation of the two lower core cylinders. A similar model was also adapted for the hemidiscoidal PBS of Mastigocladus laminosus (Zuber et al., 1987). However the hemidiscoidal PBS of Mastigocladus laminosus and Anabaena sp. strain PCC 7120 exhibit some significant differences from this basic structure as discussed above (see Fig. 9 B; Bryant, 1991; Bryant et al., 1991; Glauser et al.,

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1992a,b; Esteban, 1993; Esteban et al., 1994). Reuter and Wehrmeyer (1988, 1990; Wehrmeyer, 1990) isolated additional complexes containing and subunits, and their core analyses lead to a different model for the core structure, in which complex is located at the end of a cylinder. In theirmodel occupies a peripheral position with the small located between the peripheral and the second APC trimer of the cylinders (Reuter and Nickel-Reuter, 1993).

E. Energy Transfer in Allophycocyanin and the PBS Core The complex organization ofthe PBS core caused by the heterogeneity of the various complexes of APC

Walter A. Sidler

162 species with linker polypeptides and the phycobiliprotein renders energy transfer difficult to investigate and to understand. However, the two PCB chromophores in the APC trimer have different absorption spectra with maxima at about 600 nm and 650 nm, and energy transfer from the 600 nmabsorbing PCB species to the 650 nm-absorbing PCB species occurs mainly by a Förster dipoleinduced dipole energy transfer mechanism in about 440 femtoseconds (fs) (Sharkov et al., 1992). Bryant and coworkers have used molecular genetics to determine the energy transfer pathway in the tricylindrical PBS core of Synechococcus sp. PCC 7002 and especially examined the roles of the and polypeptides as terminal emitters (Maxson et al., 1989; Gindt et al., 1992; Zhao et al., 1992). These authors have constructed a strain harboring a site-directed mutation in ApcE polypeptide) in which the chromophore-binding cysteine residue 186 has been replaced by a serine. In another mutant the apcD gene coding for was insertionally inactivated (Maxson et al., 1989; Bryant, 1991), and double mutants combining both mutations have also been generated (Bryant, 1991; Zhao et al., 1992). It seems that the PCB chromophore is non-covalently associated with the polypeptide when Cys 186 is changed to Ser; the fluorescence lifetime of this species is much shorter than for the wild-type, and the emission maximum is red-shifted to about 715 nm (Gindt et al., 1992). The results ofthese experiments and others suggest that the core-membrane linker phycobiliprotein is the main terminal emitter in the PBS core; this polypeptide probably transfers about 75% of the excitation energy harvested by the PBS rods to the PS II reaction center in the thylakoid membranes. The second terminal emitter in the PBS core, AP-B, distributes the remaining excitation energy between PS I and PS II. When cells are in State 2, AP-B directs the energy to PS I, and when cells are in State I, the energy is directed towards PS II (probably by transfer first to the protein (Zhao et al., 1992). An important aspect of the proposal of Zhao et al. (1992)—the nature of the physical contacts between PBS and PS I and PS II that are required for the above-described energy transfer model—remains unsolved. The diagram that follows summarizes the proposed energy transfer paths for phycobilisomes of Synechococcus sp. PCC 7002 (see Zhao et al., 1992):

Considering the recent results concerning the structure and the results obtained from reconstitution experiments with the fourth and APC (Gottschalk et al., 1994), it mustfinally be emphasized that the molecular and functional organization ofthe core must be reinvestigated with respect to the role and influence of each REP of the onthe different APC complexes. A complete and reasonable model of the PBS core will only come through knowledge of the structural and spectral properties of the different APC complexes associated with the corresponding and ARMs. Such knowledge might be achieved by further reconstitution experiments with isolated or overproduced REP and ARM domains of the phycobiliprotein.

IV. Phycobiliproteins Constituting the Rod Elements of PBS The peripheral rods of PBS are composed of type hexameric subassemblies and contain PC, PE or PEC in association with the appropriate rod-linker polypeptides and an hexameric subassembly, composed of an essential PC type (see below) and its associated rod-core linker polypeptide at the core-proximal end of the rod. The position of these phycobiliprotein subassemblies within the rod is determined by their linker polypeptides and correlates with their absorption properties such that complexes absorbing higher energy wavelengths are more distal from the core. This organization facilitates unidirectional energy transfer through therods (Zuber, 1987; Glazer, 1989, see Section IV A, 10).

A. The Phycocyanins Although many cyanobacteria do not synthesize either PEC or C-PE, all naturally occurring cyanobacteria have been found to produce PC (Bryant, 1982).

Chapter 7 Phycobilisome and Phycobiliprotein Structures Hexameric complexes ofPC and linkerpolypeptides, of the types or are required to assemble the peripheral rods; typically, one complex of the latter type and two to four of the former type are found in a rod. Phycocyanobilin (PCB) is the chromophore typically found associated with the subunits of PC. The blue-colored, deeply red-fluorescent C-phycocyanin (C-PC) is the predominant PC form and contains three PCB chromophores per monomer. Amino acid sequence analyses of C-PC showed these to be located at Cys and (Frank et a1., 1978; Williams and Glazer, 1978; Freidenreich et al., 1978; Rumbeli et al., 1987b; Ducret et al., 1993). The typical absorption and fluorescence emission maxima of C-PC complexes are summarized in Table 2 (also see Figs. 2 and 26, and Wehrmeyer, 1990). Chromophore analysis in the C-PC trimer showed that PCB and PCB represent the two sensitizing chromophores located at the periphery of the light-harvesting complex, whereas PCB represents the fluorescing chromophore located in the central cavity of the trimer and hexamer (see Section IV A, 9). Adaptation processes to different light and nutrient conditions during evolution led to the development of a family of PCs differing in the amino acid sequences of the apoproteins and/or chromophore composition. Amino acid sequences as well as the crystal structures of C-PC and PEC have been determined. One, or in some cases two, of the peripheral, sensitizing chromophores PCB and/or PCB) have been replaced by PXB, PEB or PUB chromophores in each of the variants in order to adapt to conditions more enriched in blue and green wavelengths of light. The fluorescing PCB was conserved in all variants, however. The presence of PC at the rod-core linkage position is apparently essential for excitation energy transfer from the rods to the core. At the rod-core linkage positions in the PBS, PC and special rod-core linker polypeptides form stable rod-core complexes together with APC complexes:

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periphery of the PBS, but occur at the rod-core linkage position, and the rods of such organisms typically contain substantial amounts of PE. Thus, the replacement of PCB by PEB in R-PCs does not extend the absorption range of the PBS, but increases the absorption capacity for green light and might play a role in more efficient transfer of light energy to the APC-containing core. As a consequence of their now-recognized variation in chromophore composition, PCs can no longer be characterized by a ‘typical’ absorption spectrum. The common features ofPCs are an of 162 amino acid residues with one bilin binding site at and a of 172 amino acid residues and two phycobilins binding sites at positions (residue 82) and (residue 152; a PCB or PEB chromophore together with 10 residues inserted at position 151, when compared to APC subunits). The bilin apparently must be a PCB for reasons of efficient energy transfer to APC. As another common feature, PCs always interact with an to form the first hexamer at the base of a peripheral rod as a part of the so called rod-core complex (see Fig. 10 and Table 2). The interaction of PC with the polypeptide in this complex typically causes a large redshift of~17 run in absorption and fluorescence emission maxima of PC (Glazer, 1982, 1984, 1989; de Lorimier et al., 1990b; Gottschalk et al., 1991; Glauser et al., 1993). Phycobiliprotein families were first defined by the spectral and immunological properties of the proteins (immunological cross-reactivity of biliproteins within the same family, as reviewedby MacColl and GuardFriar, 1987a). Amino acid sequence comparisons now provide a more reliable method for classifying biliproteins. The amino acid sequence identity between the corresponding subunits of the various PCs ranges from 60 to 90%. PC-645 of the cryptophyte Chroomonas sp., however, does not crossreact with antisera against the members of the PC family (Berns, 1967; Guard-Friar et al., 1986) and belongs to the PE family rather than to the PC family, as demonstrated by its amino acid sequence (Sidler and Zuber, 1988; Sidler et al., 1990a).

1. Constitutive Phycocyanins The replacement of a blue PCB on the subunit by apurple PXB in PEC extends the absorption range of PBS containing this pigment considerably into the green portion of the spectrum. The different R-PCs (I–IV) are not phycobiliproteins located at the

Phycocyanins involved in rod-core subcomplexes (C-PC, R-PC-I, R-PC-II or R-PC-III) are constitutive PCs and are expressed under all growth conditions under which PBS are formed. Some cyanobacteria,

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such as Synechococcus sp. strain PCC 7002, Mastigocladus laminosus or Anabaena sp. strain PCC 7120 contain only a single set of PC genes (cpcB and cpcA, encoding the and subunits, respectively). In such organisms, additional PC hexamers in the peripheral rods are also formed by this same set of constitutively expressed cpcBA gene products interacting withdifferent proteins.

2. Inducible Phycocyanins In some cyanobacteria that are able to perform complementary chromatic adaptation (so-called Type III adapters), such as Calothrix sp. strain PCC 7601 and Pseudanabaena sp. strain PCC 7409, additional genes encoding C-PC have been identified. For example, when grown in red light Pseudanabaena

Chapter 7 Phycobilisome and Phycobiliprotein Structures sp. strain PCC 7409 synthesizes two distinct PCs, PCI and PC2 (Bryant, 1981; Bryant and CohenBazire, 1981); these PCs are the products of distinct genes (cpcB1A1 encoding PC1 = constitutive PC; cpcB2A2, encoding PC2 = red-light inducible PC; Dubbs and Bryant, 1993; see Chapter 21). For Calothrix sp. strain PCC 7601 three copies of the genes encoding the subunits (cpcA1,cpcA2, cpcA3) and the subunits (cpcB1, cpcB2, cpcB3) were found and sequenced (Conley et al., 1986; Lomax et al., 1987; Mazel et al., 1988; Capuano et. al., 1988; Conley et al., 1988; Mazel and Marliere, 1989). Only two copies are necessary for the complementary chromatic adaptation. Under green light conditions only the cpc1 operon encoding the constitutive PC 1 is transcribed, while under red-light growth conditions the cpc2 operon encoding the inducible PC2 is also transcribed. It is not known if separate or mixed PC trimers are formed by the PC 1 andPC2 geneproductswhenboth arebeing produced. PC3, encoded by the cpc3-operon, is only expressed under sulfur-limited growth conditions (Mazel and Marlière, 1989). Different PC-associated linker polypeptides are also expressed with each cpc-operon (Mazel et al., 1988); interestingly, the proteins encoded by the cpc3 operon are devoid of sulfur containing amino acids except for the initiator methionine residues (which are posttranslationally removed) and for the chromophore-binding cysteine residues in the and subunits (Mazel and Marlière, 1989; see Chapter 21 for additional details).

3. Phycoerythrocyanin (PEC) Phycoerythrocyanin (PEC) is the shortest wavelengthabsorbing rod element 634 nm) of those cyanobacteria which neither contain PE nor perform complementary chromatic adaptation (Bryant, 1982). The majority of such organisms also have the ability to form heterocysts. The function of PEC is the extension of the light-harvesting capacity of PBS into the green portion of the spectrum under medium or low-light conditions. The PEC content of PBS is subject to regulation by light intensity: under low light PEC-expression is strongly induced (Bryant, 1982; Swanson et al., 1992a; Reuter and NickelReuter, 1993; Esteban, 1993). PEC forms the peripheral hexamers of PBS rods in those species synthesizing this biliprotein. PEC was first described by Bryant et al. (1976) and was later isolated from Mostigocladus laminosus

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by Füglistaller et al. (1981), MacColl et al. (1981) and Nies and Wehrmeyer (1981) and from Anabaena sp. strain PCC 7120 (Swanson et al., 1992a). The native hexameric complex with the small 8.9 kDa and the 35 kDa rod-linker polypeptides, (PEC I) was obtained in the breakthrough from DEAE-cellulose ion exchange chromatography (Füglistaller et al., 1981). PEC appears in solution as a fuchsia- or purple-colored phycobiliprotein, and in lyophilized form it is blue. PEC and subunits from Mastigocladus laminosus can be separated by size exclusion chromatography in 63 mM formic acid (Füglistaller et al., 1981). From its amino acid sequence PEC is very similar to C-PC (Bryant et al., 1978; Füglistaller et al., 1983). It forms crystals isomorphous to those of C-PC (Bryant et al., 1976; Rümbeli et al., 1985); its threedimensional structure is practically identical to that of C-PC (Duerring et al., 1990; see below). The subunit was found to contain a purple chromophore (phycobiliviolin, PXB; see Fig. 3 B) whereas the two PCB chromophores of the are identical to those found in PC (Bryant et al., 1976). The structure of the PXB chromophore was determined by correlative 500 MHz1H NMR analysis of the chromopeptide Cys(PXB)-Val-Arg (Bishop et al., 1987). The PXB chromophore has a special behavior among the chromophores of biliproteins because it exhibits a reversible photochemistry reminiscent of that of the photoreceptor, phytochrome, of higher plants (Siebzehnrübel et al., 1989; Scharnagel and Fischer, 1993; Maruthi Sai et al., 1992, 1993).

4. R-Phycocyanin (R-PC or R-PC-I) R-Phycocyanin (R-PC or R-PC-I) was isolated by Glazer and Hixson (1975) from the red alga Porphyridium cruentum as an trimer (The determined molecular mass of the complex was 103 kDa.). The absorption spectrum of R-PC shows two maxima: the lesser with a wavelength maximum of 555 nm and the greater with a wavelength maximum of 619 nm; the fluorescence emission maximum occurs at 640 nm and indicates efficient energy transfer from the shorter wavelength absorbing chromophore to a PCB chromophore. The subunit contains a single PCB while one of the chromophores, that at position Cysl55 normally occupied by a PCB in C-PC, is replaced by a PEB (Glazer and Hixson, 1975; Bryant et al, 1978; Ong

166 and Glazer, 1987). The and were separated by HPLC and the complete amino acid sequences of both subunits were determined by protein sequencing (Ducret et al., 1994). The highest sequence identity was found to C-PC from the red alga Cyanidium caldarium (Troxler et al., 1981). Crystals of R-PC diffracting to 3 nm have been obtained, but they exhibit a high mosaicity hampering structure analysis (W. Sidler, unpublished results). R-PC-I has only been characterized from red algae up to now; however, a cyanobacterial protein with similar chromophore content (2 PCBs and 1 PEB) has been detected in the marine Synechococcus sp. strain WH 7805 (see Section IV A, 6, below). R-PCI with and without a linker polypeptide was isolated from the red alga Anotrichium tenue (Watson et al., 1986). The linker polypeptide, possibly L RC, had an estimated molecular mass of 30 kDa. The complex, possibly an or an complex shows a redshift of 7–8 nm in absorption maximum (624– 625 nm) and of 10 nm in fluorescence emission maximum (643 nm).

5. R-Phycocyanin-ll (R-PC-II) R-Phycocyanin-II (R-PC-II) was isolated by Ong and Glazer (1987, 1988) from the marine Synechococcus sp. strains WH 8103, WH 8020 and WH 7803, as the first PEB-containing PCs of cyanobacterial origin (absorption and fluorescence spectra are shown in Fig. 18 C below). These marine Synechococcus sp. are strongly adapted to green light conditions and contain mainly PEs I and II as light-harvesting rod elements (see PE family, Section IV B). R-PC-II from Synechococcus sp. strain WH 8103 was isolated as a complex and as an without any linker polypeptide. The large redshift of 17 nm, typically found for PC complexes located at the rod-core linkage position within the PBS and originating from the PCB chromophore (Gottschalk et al., 1991; Glauser et al., 1993), was observed. The absorption spectrum of R-PC-II exhibits maxima at 533, 554 and 615 nm and that of the complex exhibits absorbance maxima at 533, 554 and 632 nm. The redshift in the fluorescence emission maximum was from 646 nm for the complex to 652 nm for the complex (Ong and Glazer, 1988). By analysis of the chromopeptides the position of the single PCB was assigned to Cys position and the two PEBs to and (Ong et al., 1984). The

Walter A. Sidler complete amino acid sequence has been deduced from nucleotide sequence analysis of the corresponding genes (de Lorimier et al., 1993). Thus, RPC-II is a PC which is adapted to green-light conditions by the replacement of both sensitizing PCB chromophores by PEBs. The third chromophore, PCB apparently is essential for energy transfer to APC and cannot be replaced by a PEB without generating an unfavorably small overlap of the fluorescence spectrum of PEB with the absorption spectrum of APC.

6. Synechococcus sp. strain WH 7805 Phycocyanin (R-PC-III) A different PC, denoted here as R-PC-III and containing PEB chromophores, was isolated from Synechococcus strain WH7805. This PC had an absorption maximum at 555 nm and a shoulder at 590 nm, and the PCB:PEB ratio measured in 8 M acidified urea solution was 2:1 (Ong and Glazer, 1988). Although the chromophore content is identical to R-PC described above, it is not clear that this protein has the same positioning of the PEB chromophore since the native absorption spectrum differs significantly from that of R-PC. Other details concerning this PC must await furthercharacterization of the protein.

7. Synechococcus sp. strain WH 8501 Phycocyanin (R-PC-IV). The PC isolated from marine Synechococcus sp. strain WH 8501 was the first PUB-containing PC to be characterized (Ong and Glazer, 1988). A PUB occurs on the subunit and probably occupies the bilin-binding site at position Cys84 since the aminoterminal sequence of the subunit resembles that of other PCs. The subunit of this protein carries two PCB chromophores that are probably bound in the usual positions. The absorption spectrum of this PC (Ong and Glazer, 1987, 1988) exhibits maxima at 490 and 592 nm, and the fluorescence emission maximum was at 644 nm.

8. Amino Acid Sequences and Phylogenetic Relationships of the C-PC Family Complete amino acid sequences of phycocyanins from all spectroscopic classes and from phylogenetically diverse organisms have been determined

Chapter 7 Phycobilisome and Phycobiliprotein Structures byprotein sequencing, andnumerous othersequences have been deduced from nucleotide sequences ofthe corresponding genes (references are listed in the legendto Fig. 11). The sequence identity between the and subunits of the same phycobiliprotein decreases in the order APC (38%), C-PC (26%), PEC (21%), and C-PE (23%). Based on these relationships, a divergent phylogenetic development of the and families from a singlesubunit ancestor molecule, that gave rise to these subunit families by a gene duplication event, is assumed. The sequence identity between R-PC-I and subunits from Porphyridium cruentum is 28% and is similar to the values for C-PC from Mastigocladus laminosus (28%), for C-PC from the red alga Cyanidium caldarium (29%), and for PEC from Mastigocladus laminosus (27%). C-PE and B-PE show slightly lower identities between the and subunits, 26% and 24% respectively (Sidler et al., 1989; Sidler et al., 1990b). The highest identity to RPC-I (83%) within the PC-family is found for the and of the thermophilic red alga Cyanidium caldarium and the lowest (61%) to PEC. The identity values forR-PC-I when compared to the phycoerythrin family range from 44% to 50%. In terms of evolutionary development, the homologies of R-PC-I and II to the PEs show that the PEs did not evolve from R-PC or PEC (Figs. 11 and 12). Thus, PEC, R-PC-I and R-PC-II are notintermediate forms between PC and PE, as might be imagined based upon the content of red and purple bilins in these proteins.

9. The Crystal Structure of C-Phycocyanin The first crystals of phycobiliproteins were described more than one hundred years ago (Cramer, 1862; Molisch, 1894, 1895). The diffraction properties of Mastigocladus laminosus C-PC crystals (Dobler et al., 1972) allowed the determination of the crystal structure first to 3 Å resolution (Schirmer et al., 1985) and later to 2.1 Å (Schirmer et al., 1987). Subsequently, the crystal structures of the hexameric C-PC from Synechococcus sp. strain PCC 7002 (formerly Agmenellum quadruplicatum strain PR-6; Schirmer et al., 1986) and from PC1 from Calothrix sp. PCC 7601 (Duerring et al, 1991) have been determined by Patterson search techniques using the Mastigocladus laminosus C-PC molecular model. Fundamental structural principles of phycobili-

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proteins as deducedfrom the crystallographic studies are illustrated in Figs. 13 and 14.

a. The C-Phycocyanin Monomer and Its Subunits The phycobiliprotein monomer looks like a boomerang that is approximately 80 Å from tip to tip. The and subunits have very similar tertiary structures and are related by a local twofold rotational axis; strong subunit binding interactions take place around this two-fold axis, as may be discerned from the side view of the monomer (see Fig. 13 C). The secondary structure elements of the C-PC subunits include and structures. No elements are present, although such structures were suggested by the Chou-Fasman (1978) secondary structure prediction method from calculations performed before any crystal structure data were available (Sidler et al., 1981b). The three PCB chromophores in a monomer are located at the periphery of the molecule (see Fig. 16 B). The center-to-center distances forthe three chromophores in a monomer are approximately 50 Å between the chromophores attached at and 48 Å between the chromophores attached at and and 34.6 Å between the chromophores attached at and The C-PC subunits have globin-like folding and possess closely related counterparts to the myoglobin A, B, E, F, G and H. According to the myoglobin nomenclature of helices, the eight helices ofa C-PC subunit were denoted as X,Y,A, B, E, F, G and H (Schirmer et al., 1985). The additional X and Y stick out from the globular part of the structure and mediate strong subunit-subunit interactions with the A and E helices of the associated subunit in the region of the twofold symmetry axis. The binding sites of the PCB chromophores Cys84 and in C-PC are at a position topologically equivalent to His E7, the heme-iron ligand of myoglobin. A very detailed comparison of the structures of the globin and PC protein families support the idea that the oxygen-binding proteins of higher organisms (oxygen consumption in respiration) and PCs ofoxygen-producing procaryotes are distantly related members of a common protein family, and that the major differences (the large differences resulting fromverydifferentfunctions of the proteins) between them are the result ofdivergent evolution from a common ancestor (Pastore and Lesk, 1990).

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b. Trimeric C-Phycocyanin The building blocks of the Mastigocladus laminosus C-PC crystals are trimeric discs about 110 Å in diameter with a thickness of 30 Å , that have a central hole approximately 30 Å in diameter. The high degree of amino acid sequence similarity among all phycobiliproteins, and the fact that these proteins are isolated mostly as trimeric or hexameric aggregates, suggests that the trimers and hexamers are the fundamental assembly units ofthe peripheralrod and core substructures of PBS. These trimers resemble a water wheel with the paddles formed by the monomers (see Fig. 13 C, D and Fig. 14; Schirmer et al., 1985). Two trimers form a heterohexamer approximately 60 Å in thickness by head-to-head (or face-to-face) aggregation, as found in C-PC from Synechococcus sp. strain PCC 7002 (Fig. 13 E, F; Schirmer et al., 1986). These data correspond well with measurements from electron microscopic studies. The aggregation of three monomers to form a trimer brings the PCB chromophore attached to the position of one monomer into close proximity (20.6 Å) to the PCB chromophore attached to the position of a second monomer

(Schirmer et al., 1987; Duerring et al., 1991). Moreover, the three PCBs attached to the positions project into the central cavity ofthe trimer and are separated by only 35 Å (Fig. 13 D and Fig. 14; also see Fig. 16 B). The chromophores attached to the positions remain at the periphery of the molecule; their nearest neighbors for energy transfer are the PCB attached to the same subunit at position (34.6 Å) and the chromophore attached to the position ofa second monomer (39.1 Å; Duerring et al., 1991). The distances between the various PCB chromophores (22Å to >50 Å) may be too large for exciton coupling and rather suggest that an inductive, Förster resonance-energy-transfer mechanism occurs (Förster, 1965, 1967).

c. Hexameric C-Phycocyanin Hexamers ofboth PC and PE are formed from the face-to-face (head-to-head) assembly of two discshaped trimers (Schirmer et al., 1986, 1987; Duerring et al., 1991; Ficner et al., 1992; Ficner and Huber, 1993; see Fig. 13 E, F). The central cavity of the CPC hexamer (Fig. 13 E)probablyprovides the binding site for a large portion of the rod-linker or rod-core linker polypeptides (Lundell et al., 1981a; Yu and

Chapter 7 Phycobilisome and Phycobiliprotein Structures Glazer, 1982; see Ficner and Huber, 1993). Possible interaction principles of the phycobiliproteins with the rod linker polypeptides have been postulated by Glauser et al. (1992b, 1993; see Section V). In the hexamer many chromophores are separated by less than 37 Å and are favorably oriented for energy transfer. The peripherally located PCBs attached to the positions become coupled to chromophores in the second trimer or are coupled to PCBs attached to the positions. Interhexamer energy transfer probably occurs via the chromophores attached to the positions; if the stacking of hexamers in rods ofphycobilisomes andthePC crystals are similar,thenthePCBsattached at the positions of two different hexamers would be essentially parallel to one another and separated by about 26 Å (Schirmer et al., 1987; Duerring et al., 1991).

d. The Crystal Structure of Phycoerythrocyanin The crystal structure of trimeric of Mastigocladus laminosus was determined from hexagonal prisms on the basis of molecular modeling with the C-PC structure (Duerring et al., 1990). Both the protein and chromophore structures of PEC and C-PC are very similar, as expected fromtheir-highly similar amino acid sequences (63% identity in the subunits and 67% in the subunits). On the subunit, however, thatbears thephycobiliviolin (PXB) chromophore instead ofa PCB chromophore as in CPC, a rather different structure was found for residues 116 to 123, the protein motif interacting with the 84 chromophore (Duerring et al., 1990). However, the major cause for the absorption of the PXB tetrapyrrole at shorter wavelength is the conjugated which is shorter than in PCB (see Fig. 3).

e. Chromophore Structure and Common Principles of Chromophore-Protein and Protein-Protein Interaction From the crystal structure of Mastigocladus laminosus PEC at 2.7 Å, the refined structure of Mastigocladus laminosus C-PC at 2.1 Å, and the Synechococcus sp. strain PCC 7002 C-PC crystal structure at 2.5 Å resolution, the exact orientation of all PCB and PXB chromophores was determined and a common principle of chromophore-protein interaction was postulated (Schirmer et al., 1987; Duerring et al., 1991). The geometries of the PXB

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and PCB chromophores are very similar (Fig. 15 A). They resemble a cleaved porphyrin ring, twisted by about 180° around the C-5 - C-6 and C-14 - C-15 bonds with a conformation of C-5-anti, C-9-syn and C-14-anti and a configuration C-4-Z, C-10-Z and C15-Z. The chromophores also interact in a similar way with the protein: they each arch aroundaspartate residues. The nitrogens ofpyrroles B and C are most probablyprotonatedandarewithinhydrogen-bonding distance of one of the carboxylate oxygens. This carboxylate oxygen has a position equivalent to the iron in heme. The propionate side chains of the chromophores form salt bridges with Arg and Lys residues. Thus, charge-charge interactions seem to play a central role in the spectral properties of the bilins. The known structures of the phycobiliproteins, including C-PC from Calothrix sp. strain PCC 7601 (Duerring et al. 1991), show common structural features at all levels: the number and folding of helices and turns similar to the globin fold; the symmetric association of and subunits around a non-crystallographic internal two-fold symmetry axis; the formation of the basic, stable trimeric aggregate; and the formation of hexamers from two trimers (in the crystals without linker polypeptides). Hexamers and rods are formed in vivo, however, by the inclusion ofa linker polypeptide. C-PC crystal structures have not yet proved the in vivo existence of a hexamer from the head-to-head aggregation of two trimers plus an The crystal structure of hexameric B-PE fromPorphyridium sordidum (Ficner and Huber, 1992; Ficner et al., 1992), however, that includes a 30 kDa subunit, also shows a head-tohead (face-to-face) aggregation. This is alsoprobably true for C-PC and PEC hexamers.

f. a Modified Amino Acid Residue in Phycobiliproteins The presence of a posttranslationally modified amino acid residue inthe subunitwasoriginallyreported by Minami et al., (1985). Klotz et al. (1986) soon after identified (NMA) at position 71 in the amino acid sequence of from Anabaen a variabilis as well as in the subunits of BPE and R-PC from red algae. A subsequent reinvestigation of the amino acid sequences of PEC, C-PC and APC from Mastigocladus laminosus and of C-PE of Calothrix sp. strain PCC 7601 confirmed the presence of NMA at position 72 of the subunits

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of each protein (Rümbeli et al., 1987a, b, c). During amino acid sequence analysis the PTHderivative of NMA coelutes with PTH-serine on HPLC and gives rise to methylamine in ninhydrin post-column amino acid analysis ofthe hydrolysates. Preliminary inspection of the X-ray structure of CPC indicated, that residue 72 points towards the chromophore within hydrogenbonding distance (Rümbeli et al., 1987b; see Fig. 15 B) and reinvestigation of the crystal structures of C-PC from Mastigocladus laminosus and Synechococcus sp. strain PCC 7002 confirmed the presence of NMA at the position (Duerring et al., 1988). Although NMA is absent in phycobiliproteins of certain organisms and may be dispensable in some circumstances (Klotz and Glazer, 1987), the function of this modification remains to be determined. Swanson and Glazer (1990a) showed that PBS of two mutants free of methylase activity, and thus with unmethylated Asn exhibit defects in energy transfer. Unmethylated PBS showed greater emission from PC and APC and lower fluorescence emission quantum yields. NMA at thus contributes significantly to the efficiency of directional energy transfer in PBS. Site-directed mutations in which the Asn residue was converted to either Asp or Gln have been created similarly which exhibited lower

Walter A. Sidler

fluorescence emission quantum yields (J. Zhou and D. A. Bryant, personal communication).

10. Energy Transfer in the PBS Rods Numerous excellent energy-transfer kinetic studies performed with phycobiliprotein are not referred to in this chapter due to space limitation, but these have been reviewed by Scheer (1982, 1986), Glazer (1989), Wehrmeyer (1990), Holzwarth (1991) and Rüdiger (1994). Mimuro et al. (1986a) proposed a model for the optical characteristics and the pathway of energy transfer in C-PC from Mastigocladus laminosus (see Fig. 16B). The model is based on the threedimensional structure of C-PC and the data from steady-state absorption, circular dichroism, fluorescence and fluorescence-polarization spectroscopy. In the C-PC trimer, the chromophore was denoted as a sensitizing (s) chromophore, transferring excitation energy to the fluorescing (f) chromophore, located in an adjacent monomer. Energy transfer from the PCB, a peripheral (s) chromophore to the 84 f-Chromophore ofthe same monomer was also postulated (Fig. 16 B). The chromophores are suggested to transfer the excitation energy towards the APC core. Mimuro et al., (1986b)

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176 also proposed that aromatic amino acid residues such as Tyr, Tyr and Phe and Tyr, might affect the electronic structure of the chromophores. Deconvoluted, single-chromophore spectra of the C-PC chromophores fromMastigocladus laminosus (Siebzehnrübel et al., 1987; Fischer and Scheer, 1988; Fischeret al., 1988; see Rüdiger, 1993) confirm PCB to be the chromophore absorbing the longest wavelength (622–624 nm; Fig. 16 A) whereas the PCB is the shortest wavelength absorbing chromophore (598–699 nm). The absorption maximumofthe PCB lies in between (616–618 nm). Similar results for the spectroscopic properties of the individual chromophores in C-PC have been obtained from detailed characterization of a sitespecific mutant in which the Cys residue was changed to Ser (Debreczeny et al., 1993). This mutation causes the loss of the PCB that would normally be attached at this position. It was concluded from those studies that the PCB chromophore absorbed maximally at 600–602 nm, the PCB chromophore absorbed maximally at 626–628 nm, and the PCB chromophore absorbed maximally at 624 nm. In contrast to the function of this chromophore in PC, Hucke et al. (1993) demonstrated by two-color femtosecond transient absorption spectroscopy (with time resolution better than 200 fs) that the chromophore is the longest wavelength absorbing chromophore in PEC. Sauer and Scheer (1988) calculated the individual energy transfer rates within PC monomers, trimers and hexamers and showed that energy transfer is a combination of exciton interaction between and chromophores of adjacent subunits and of excitation energy transfer by the Forster induction mechanism. Gillbro et al. (1993) observed rapid Förster energytransfer (500 fs lifetime) in PC trimers, that did not occur in PC monomers; they assigned this to energy transfer between the nearest neighboring and chromophores. In stacked hexamers of crystals, energy transfer should take place mainly between the strongly coupled chromophores (r = 21–26 Å). Energy transfer along the rods can be regarded as a random walk (trap- or diffusion-limited) along a one-dimensional array of f-chromophores, as suggested by Schirmer and Vincent (1987). The crystal structure of the PC hexamer from Synechococcus sp. strain PCC 7002 shows that the central

84 chromophores of two trimers lie exactly above one another and are separated by 34 Å and with their transition dipole moments nearly parallel to each other (Schirmer et al., 1986). If hexamers stack into peripheral rods in PBS in a manner similar to that which occurs in the PC crystals, then the chromophores of two hexamers would be separated by 21–26 Å (Schirmer et al., 1987; Duerring et al., 1991). Nonetheless, the distances between the peripheral or and the central chromophores are too long to allow strong excitonic coupling. The crystal structure of the PC hexamer suggests that light energy absorbed by the s-chromophores and is transferred first to the 84 f-chromophores within one hexamer and then transduced via these central chromophores along the phycobilisome rod to the PBS core by a Förster-rype, dipole-induced dipole transfer mechanism (Förster, 1965, 1967; see also Glazer, 1989).

B. Phycoerythrins The phycobiliproteins of the red-colored phycoerythrin (PE) family exhibit a great diversity in spectral properties as well as in their chromophore and subunit compositions. PEs carry only one or two chromophore types, PEB and/or PUB, instead of the four chromophore types found among members of the PC family (PCB, PXB, PEB, and PUB). Also in contrast to the situation found for PCs, the subunit chain length varies considerably for PEs. While PCs contain only non-pigmented linker polypeptides, PE hexamer complexes contain in their central cavity either non-chromophorylated proteins or chromophore-bearing As found for PCs, there exist inducible (in complementary chromatic adaptation) and constitutively expressed PEs. An evolutionary tendency to attach as many bilins as possible to PE complexes, a process that involves not only the PE subunits themselves but also the linker polypeptides (converting them into subunits), may be recognized. Cyanobacteria contain either PEC or PE in the PBS but never both, and many cyanobacteria such as in Synechococcus sp. strain PCC 7002 possess no red-colored phycobiliproteins (i.e., neither PE nor PEC) but only the blue-colored APC and C-PC (Bryant, 1982, 1988). PEC does not occur in red algae. Cryptomonads contain several spectral types ofphycobiliproteins, and they all derived from B-PE

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which has been modified with different red, purple, blue and green chromophores (Sidler and Zuber, 1988; Sidler et al., 1987; Wedemayer et al., 1991, 1992). PE is the only phycobiliprotein present in cyanobacteria, red algae as well as in cryptomonads, and is thus most suitable for comparative studies to reveal structural, functional and phylogenetic relationships among these organisms. During evolution PE evolved earlyfrom acommon ancestor ofthe present phycocyanins in cyanobacteria by gene duplication (Figs. 11 and 12; Glazer, 1983). PEC and R-PC, both ofwhich share some functional similarities to phycoerythrin and which share the trait of carrying chromophores extending the absorption envelope ofthe phycobilisome to shorter wavelengths, arenotevolutionaryintermediateforms between PC and PE. These proteins developed independently from within the phycocyanin family and probably also arose later in evolution (Figs. 11 and 12). PE finally dominated in the PBS ofred algal chloroplasts. In cryptomonads only the subunit is conserved from all phycobiliprotein types present in cyanobacteria and red algae; the cryptomonad biliproteins are assembledwith anunrelated class of chromoproteins (Glazer and Apell, 1977; Sidler et al., 1985, 1990a). The brilliant red colors of the phycoerythrins originate from red phycoerythrobilinpigments (PEB, Figs. 3 C, D) and from phycourobilin chromophores (PUB, Figs. 3 E, F) found in the PEs of Gloeobacter violaceus (Bryant et al., 1981), of the marine Synechococcus sp. (Ong et al., 1984), and in B-PE (Glazer and Hixson (1977) and R-PE (Klotz and Glazer, 1985) ofred algae. The conjugated system is shorter for PEB and PUB than for PCB, causing the major absorption bands for these chromophores in the visible region to be shifted to the shorter wavelengths of the bluegreen region of the spectrum. In the peripheral rods ofthe PBS, PE is located at the core-distal periphery of the structure.

near 565 nm (PE I; see Table 3). An exception to this generalization is the PE of Gloeobacter violaceus. Although originally reported to carry both PEB and PUB chromophores in the ratio 6:1 (Bryant et al., 1981), revisions ofthe extinction coefficients for the two chromophores are more consistent with a ratio of 5:1. The PUB chromophore has been shown to reside on the (Bryant et al., 1981). Recently, PEs from marine unicellular Synechococcus sp. and Synechocystis sp. strains containing the yellow phycourobilin (PUB) chromophores in addition to PEB have been characterized in detail (Alberte et al., 1984; Ong et al., 1984; Ong and Glazer, 1988, 1991; Swanson et al., 1991). associated with PE hexamers, which were previously believed to occur only in red algae, have also been found in these marine cyanobacterial PE-II types (Swanson et al., 1991; Wilbanks and Glazer, 1993a, b). Additionally, a sixth chromophore binding site occupied by PUB was found in a recently discovered PE (PE II) of two Synechococcus sp. strains (WH 8020 and WH 8103) at the position (seeTable 3), These PEs show spectra with the strongest short wavelength absorption properties among the known cyanobacteria and red algae (see Fig. 18). The more usual PE type (PE I) with five total binding sites for PEB and PUB is also present in the same PBS of these Synechococcus sp. strains (Ong and Glazer, 1988, 1991). The most abundant light-harvesting component of almost all red algal chloroplasts is B-PE. The spectroscopicproperties (absorptionand fluorescence emission spectra) of more than one hundred phycoerythrins from representatives of all the orders of the Bangiophyceae and Florideophyceae have been determined in a survey by Glazer et al. (1982; see also Yu et al., 1981b). On the basis of these spectroscopic analyses, PEs were subdivided into the five groups that are listed in part B ofTable 3.

1. Phycoerythrins in Cyanobacteria and Red Algae

The first PEs described were PE-I types with five PEBs. PE-I proteins with PUBs have been reported recently from marine cyanobacteria of the genera Synechococcus and Synechocystis (Ong and Glazer, 1988, 1991; Swanson et al., 1991); these proteins have a first absorbance maximum near 491 nm originating from the PUB chromophores (Fig. 18 B). From the amino acid sequence of C-PE-I from

PE chromophore contents differ for cyanobacteria living in either freshwater and soil and or marine environments. PEs from freshwater and soil cyanobacteria typically contain only PEB chromophores and exhibit absorbance spectra with maxima

a. C-Phycoerythrin-l

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Calothrix sp. strain PCC 7601 (Sidler et al., 1986; Mazel et al., 1986), the subunit is seen to contain 164 amino acid residues and to bind two PEBchromophores, accounting for a total molecular mass of 18,368 Da. The subunit contains 184 amino acid residues and three chromophores (molecular mass: 20,931 Da; Figs. 17 and 19A). The amino acid sequence identity between the and subunits is only 23%. As already determined spectroscopically (Glazer and Hixson, 1975; Muckle and Rudiger, 1977), the sequence analysis revealed that C-PE contains five red PEB chromophores. The PEB binding sites at positions 84 in and correspond to those found in PC, PEC and APC subunits. The third PEB binding site at position 155 in is identical to that for subunits of the PC family. In the subunit, one of the additional chromophores is inserted together with a pentapeptide *CAPCRD at position 143a (Figs. 17 and 19A). In the subunit a third chromophore is doubly bound to and (Lundell et al., 1984; Figs. 17 and 19A). Doubly bound PEB chromophores have been found in the R-PE subunit at identical Cys positions in a tryptic peptide with similar sequences (Nagy et al., 1984; Schoenleber et al., 1984; Klotz and Glazer, 1985). A unique insertion of 10 amino acid residues (in comparison with the C-PC ) is found at positions (Figs. 17 and 19A, B). A methylated Asn ispresent at position (see Section IV A, 9 f; Rümbeli et al., 1987b,c) as well as in other

PE-I types (Klotz and Glazer, 1987).

b. C-Phycoerythrin-ll Marine Synechococcus sp. strains contain two different phycoerythrins: a PE-II type in addition to the more typical PE-I (Alberte et al., 1984; Ong et al., 1984; Ong and Glazer, 1988, 1991; Swanson et al., 1991; Wilbanks and Glazer, 1993a,b). The absorption spectrum of PE-II (Fig. 18A) shows a majorabsorbance maximum at 492 nm that originates from PUBs and a less prominent maximum at 543 nm due to PEBs. PE-II carries six bilin chromophores per PE-I and II are strongly adapted for the absorption of blue-green light near 500 nm by the replacement of PEBs by PUBs When compared with cyanobacterial C-PE (class I) and B-PE from of the red alga Porphyridium cruentum, the amino acid sequence of PE-II from Synechococcus sp. WH 8020, translated from the DNA sequence of the corresponding genes (Wilbanks et al., 1991; Wilbanks and Glazer, 1993a,b), shows that four PEB and one PUB occupy homologous chromophore binding sites. The sixth bilin (PUB), however, is attached at a new bilin binding site created by an inserted Cys at position Adapted to the crystal structure of phycobiliproteins (Schirmer et al., 1985; Ficner and Huber, 1993) the new PUB attached to position is located in the loop of helices B and E (see Fig. 23) and corresponds to a

Chapter 7 Phycobilisome and Phycobiliprotein Structures

sensitizing, peripheral chromophore. The ammo-acid sequence identity of PE-II to C-PE from Calothrix sp. strain PCC 7601 (PE-I) amounts to 63% subunit), 61% and to B-PE of P.cruentum 54% 68% The sequence of the domain is most different from PE-I, probably as a consequence of the new and added function as a chromophore-protein-interaction domain. Instead of an Asn, a Gly residue was found at position thus, the presence of a methylasparagine at is not possible. Its absence however has notbeen confirmedbyprotein-chemical analyses. A special feature of C-PE-II from marine

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cyanobacteria is the occurrence of a subunit in the hexameric complex (Wilbanks and Glazer, 1993a, b; seebelow).APE-II carries40bilins— the highest known number of bilins per hexameric complex! Until recently subunits had exclusively been found in red algal PE complexes.

c. B-Phycoerythrin from Red Algae B-PE was first investigated in detail 100 years ago by Hans Molisch (1894) in an outstanding scientific investigation for that time. He determined B-PE to be a protein, described an isolation procedure by ammonium sulfate fractionation, crystallized B-PE

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reproducibly and characterized the crystals as hexagonal prisms in collaboration with the crystallographer Dr. F. Becke in Prague. B-PE is a multisubunit complex with the polypeptide composition The subunit structure and chromophore composition of B-PE and b-PE was determined by Glazer and Hixson (1977). However, it has not yet been determined if the trimeric PE, denoted b-PE, exists in this form in the PBS, or

Walter A. Sidler

if all b-PE is a product of dissociation of hexameric B-PE caused by proteolysis of the subunits of BPE. The apparent molecular masses of the and subunits, estimated by SDS-PAGE, are about 18 and 20 kDa, (18,991 Da and 20,315 Da, respectively, calculated from the amino acid sequence and including the PEB chromophores) and those of the subunits are about 30 kDa. From electron microscopy (Mörschel et al., 1980),

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preliminary X-ray diffraction analyses (AbadZapatero et al., 1977; Fisher et al., 1980), and from the known crystal structures of other phycobiliproteins (Schirmer et al., 1985, 1986; Duerring et al., 1990), it had already been suggested the B-PE structure is a disc formed from two trimers assembled face-to-face withthe subunit located in a central cavity. This structural was recently confirmed by X-ray diffraction analysis (Ficner and Huber, 1992, 1993; Ficner et al., 1992; see Figs. 14 and 21 and Section IV B, 4). The subunit of B-PE from the red algae Porphyridium cruentum is composed of 164 amino acid residues, just as the subunit of C-PE, and the subunit of 177 amino acid residues—7 fewer than the subunit of C-PE (see Fig. 19A). The sequence identity between and subunits of B-PE is only 24% (compared to 26% for C-PE). In fact, despite its occurrence in a different structural type of PBS and the presence of additional PEB and PUB chromophores on the subunits ofB-PE from Porphyridium cruentum, the homology between the eucaryotic BPE and procaryotic C-PE is very high: the sequence identity is 69% for the subunits and 65% for the subunits. The PEB binding sites in B-PE (positions see Fig. 19 A) and the at position are the same as in C-PE (Klotz and Glazer, 1987). The only significant difference in these two proteins is in their sizes; the subunit of B-PE is 7 residues shorter than the C-PE subunit due to the deletion of residues in a loop region near the carboxyl terminus.

PEssixchromophoresper heteromonomer. The additional chromophore binding site in PE II is due to the insertion of a single amino acid residue at position 75 inthe subunit (Wilbanks et al., 1991). The amino acid sequence identity of PE II is higher to PE I (63–66%) than to B-PE (54%; Wilbanks et al., 1991). Additional PE sequences havebeen determinedfromC-PE-I ofCalothrix sp. strain PCC 7601 (Sidler et al., 1986; Mazel et al., 1986), Synechocystis sp. strain PCC 6701 (Anderson and Grossman, 1990b), and Pseudanabaena sp. strain PCC 7409 (Dubbs and Bryant, 1987, 1991). Interestingly, B-PE and PE-II which include a subunit in their rod-complex (e.g., B-PE of Porphyridium cruentum and PE-II of Synechococcus sp. strain WH 8020) show a common deletion of 8– 9 residues inthe subunit Fig. 19 A). This is probably a steric adjustment to the presence ofthe subunits, which may be reclaiming space for one of their four chromophores. Aspartate residues are found at positions and in C-PE as they are in C-PC; these highly conserved residues were presumed to form importantcontacts withthethree correspondingPEB chromophores. This conclusion has been confirmed by the determination ofthe crystal structures for the PEB chromophores of B-PE from Porphyridium sordidum and Porphyridium cruentum (Ficner and Huber, 1992, 1993; Ficner et al., 1992). The same principles ofbilin-protein interaction found for PCB and C-PC are also observed for PEB-protein interactions (see Section IV A, 9e).

2. Comparison of Phycoerythrins with Other Phycobiliproteins

3. The Phycoerythrin subunits

C-PE-I is the largest molecule among the APC, PC and PE phycobiliprotein families, its subunits being 4 to 17 residues larger than related subunits. The introduction of additional bilin chromophores has obviously been accompanied by insertions of additional amino acid residues inthe and subunits of C-PE-I, or by changes in the amino acid sequence of a specific domain in the case of the subunit ofCPE-II. This has occurred because the bilins always require interactions with special protein structures to obtain the required structural conformations for functionality, as was first noticed from the threedimensional structures of C-PC and PEC from Mastigocladus laminosus (Schirmer et al., 1987). Class I PEs carry five chromophores and class II

In addition to the the subunits found in B-PE complexes of red algae and in PE-II of cyanobacteria are the second type of bifunctional polypeptide— combined phycobiliprotein and linker polypeptide. As for the linker polypeptides, subunits are presumed to occupy the inner space of the PE hexamercomplex andadditionally carryPEB and/or PUB chromophores. The subunits of PE-II of cyanobacteria were found to carry a single bilin (PUB) whereas subunits of red algae typically carry four bilins with PEB and PUB occurring in a variable ratio (Fig. 17; Klotz and Glazer, 1985; Wilbanks and Glazer, 1993b; Apt et al., 1993). Thus, the subunits function as shortest wavelength-absorbing biliproteins as well as linker polypeptides, in which role they contribute strongly to the stabilization of the

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Chapter 7 Phycobilisome and Phycobiliprotein Structures hexameric B-PE complex. Although crystallized with B-PE (Ficner et al., 1992) the three dimensional structure of the subunit, as well as of all other linker polypeptides, still remains to be determined. In X-ray diffraction patterns of phycobiliproteinhexamers with linker polypeptides or subunits, the contribution of the latter to the electron density map is averaged out due to rotational symmetry allowed by the surrounding biliprotein hexamer. The amino termini of the subunits isolated from the hemiellisoidal PBS of the red alga Porphyridium cruentum are naturally blocked (Glazer and Hixson, 1977). By ion-exchange chromatography at least three different B-PE complexes have been separated

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from the PBS of P. cruentum; three distinct subunits, each containing PUB and PEB chromophores and with different molecular masses (30–33 kDa) were found in these complexes (Sidler et al., 1990b). These subunits can be purified by HPLC (Swanson and Glazer, 1990b; Swanson et al., 1991) or by gel permeation chromatography in 50% formic acid (W. Sidler, unpublished results), subunits have also been reported from the B-PE of the hemisicoidal PBS from the red alga Rhodella violacea (Koller and Wehrmeyer, 1977; Koller et al., 1977, 1978; Mörschel et al., 1980). Klotz and Glazer (1985) purified four bilin-binding chromopeptides from the subunit of R-PE from the red alga Gastroclonium coulteri

186 (1 PEB and 3 PUB) and determined the sequences of these tryptic bilin-carrying peptides. Two subunits with masses of 31 and 33 kDa are present in the PBS of the red alga Aglaothamnion neglectum (Apt et al., 1993). The subunit carries 3 PUB and 1 PEB as found for the subunit of Gastroclonium coulteri (Klotz and Glazer, 1985), while the subunit carries 2 PUB and 2 PEB as found for the subunit of Porphyridium cruentum (Glazer and Hixson, 1977). The primary structures of subunits of both origins, from PE-II of cyanobacteria and B-PE from red algae, have been completely determined by gene sequencing (see Fig. 20). The mpeC gene from Synechococcus sp. strain WH 8020 encodes a polypeptide of 293 residues with a predicted molecular mass of 32.1 kDa (Wilbanks and Glazer, 1993b). One PUB is singly bound to The calculated pI of 8.9 is typically high as found for other linker polypeptides. Red algae are eucaryotic algae containing a chloroplast. The subunits of these organisms are encoded on the nuclear DNA, not on the plastid DNA as the and phycobiliproteins, (Egelhoff and Grossman, 1983) and thus have to be imported into the chloroplast. The gene encoding the subunits of the red alga Aglaothamnion neglectum includes an amino-terminal transit peptide in the 36 kDa subunit. The subunit is suggested to be transported into the plastid by a mechanism similar to that of higher plants (Apt et al., 1993). The mature subunit is a 33 kDa polypeptide with four bilin-binding sites at residues 94 (PUB), 210 (PUB), 247 (PUB) and 297 (PUB). The sequence homologies between linker polypeptides are clearly lower than between phycobiliproteins. Amino acid sequence alignments (Fig. 20) of C-PEassociated linker polypeptides (CpeC, CpeD, CpeE) with subunits from the cyanobacterium Synechococcus sp. strain WH 8020 and the red algae Aglaothamnion neglectum and Porphyridium cruentum and, and the subunits of cryptophytan phycobiliproteins show that the identity between CPE-associated linker polypeptides and cyanobacterial subunits is still significant (28%; Wilbanks et al., 1993b; 36%, as calculated by the GCG-software program, Devereux et al., 1984, 1987). However, sequence identities reach the limit of significance when cyanobacterial and eucaryotic subunits are compared (18%, calculated by the GCG-software program). Nevertheless, it can be concluded that the

Walter A. Sidler subunits derive from cyanobacterial C-PEassociated linker polypeptides, and it is possible that cryptomonad subunits are derivedfrom subunits. It is interesting to note that during evolution within the same phycobiliprotein rod complexes, the phycobiliprotein moiety remained structurally conserved (~70% identity between cyanobacterial and red algal PE subunits), whereas the linker polypeptides and moieties changed drastically (low identity in the range of 17%–35%).

4. The Crystal Structure of B-Phycoerythrin from Red Algae One hundred years after the first detailed description of PE crystals by Molisch (1894), the crystal structures of B-PE from the red alga Porphyridium sordidum and b-PE from Porphyridium cruentum were solved by Ficner and Huber (1992, 1993) and Ficner et al. (1992), based upon the amino acid sequence of B-PE from Porphyridium cruentum (Sidler et al., 1989). Initial attempts with B-PE crystals from Porphyridium cruentum (Sweet et al., 1977; Fisher et al., 1980) had been hampered by twinning problems (Fischer and Sweet, 1980). The three-dimensional structure of B-PE from Porphyridium sordidum (Figs. 14,21 and 22) is quite similar to that of C-PC and is built up from nine helical segments in the globin fold (see Section IV A, 9a and Fig. 13). The doubly bound PEB in the subunit is linked by ring A to 50 and by ring D to 61, i.e., in the inverted orientation than originally proposed by Schoenleber et al., (1984; see Fig. 3D). Following the common principle of phycobilin-protein interaction (Schirmer et al., 1987), the nitrogen atoms N22 and N23 of rings B and C of all five PEBs interact with carboxyl oxygens of aspartates (in some cases with structurally bound water molecules in between), bringing the chromophores into the typical extended-arch conformation (Fig. 15). In addition, the ionic and polar interactions between the subunits are also conserved and show an additional Ser and Asp interaction. The trimer-trimer specific interactions are conserved as well. The two additional chromophores of the B-PE monomer are located at the periphery of the trimer (and hexamer) and therefore function as sensitizing chromophores. This means there are 24

Chapter 7 Phycobilisome and Phycobiliprotein Structures

sensitizing chromophores in a hexamer—twice the number found in a PC hexamer. A similar interaction of which comes into hydrogen

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bonding distance to PEB is also expected (see Section IV A, 9f). The deletions and insertions found in the amino acid sequences of the B-PE subunits, as

188 compared to the sequence of C-PC, cause specific but local changes in the structure in correlation with the two additional PEB chromophores at positions and (see Fig. 20 C, D). In the subunit, the deletion at makes an additional protein-PEB interaction with possible. The inserted chromopeptide forms a binding pocket for this additional chromophore. In the subunit the insertion enlarges the loop between helix and and is directed towards the additional, doubly bound PEB. The Thr residue at 146c ofthe insertion forms a hydrogen bond with the propionate carboxylate ofthe PEB (Ficner and Huber, 1992, 1993; Ficner et al., 1992). Thus, all the structural changes found in the amino acid sequences of the BPE subunits, as compared to those of C-PC, can be explained by changes in the structure-function relationships of PEs compared with PCs (also see Fig. 22). A similar tertiary structure is assumed for C-PE from Calothrix sp. strain PCC 7601 (Sidler et al., 1986). The 10-amino acid residue insertion in the subunit of this C-PE is also located near the 161 PEB and probably interacts structurally and functionally with this chromophore. By analogy to the C-PC and B-PE structures, the sixth chromophore of class II PEs at is also suggested to be located at the periphery of the trimers and hexamers (Wilbanks et al., 1991), yielding 30 sensitizing chromophores in a hexamer of this class II PE (see Fig. 23).

5. Phycoerythrin in the Light-Harvesting Antenna of Cryptomonads Primary structure analyses of cryptomonad phycobiliproteins have been performed by protein sequence analyses with ‘phycocyanin’-645 (PC-645) from Chroomonas sp. and with phycoerythrin-545 (PE545) from Cryptomonas maculata (Glazer and Apell, 1977; Sidler et al., 1985, 1987, 1988, 1990a). Additionally, the sequence of the subunit of Cryptomonas sp. strain has been deduced from the nucleotide sequence of the plastid-encoded gene of this organism (Reith and Douglas, 1990). MacColl and coworkers (MacColl et al., 1973; MacColl and Guard-Friar, 1983; MacColl et al., 1983) and Jung et al. (1980) characterized PC-645 and determined the subunit distribution, number, and types of tetrapyrrole chromophores in PC-645

Walter A. Sidler from Chroomonas sp. On each subunit they found two phycocyanobilins PCB and one bilin that was recently shown to be a doubly linked 15,16-dihydrobiliverdin chromophore nm) at positions (Wedemayer et al., 1992). This chromophore had previously been assumed to be a cryptoviolin PXB chromophore. One greencolored, far-red-absorbing rnesobiliverdin 697 nm) chromophore was found on each subunit in the protein of PC-645 (Wedemayer et al., 1992). Thus, there are eight chromophores per dimer in PC-645 from Chroomonas sp.— the same number as in the protomers of APC, PC and PEC combined. Although PC-645 from Chroomonas sp. is much smaller than a PBS, its absorption spectrum shows that this complex is able to absorb light energy in a spectral range similar to that of the large PBS from cyanobacteria and red algae. This is due to the combination of one dihydrobiliverdin and two PCBs on each subunit and one green-colored, far-red-absorbing (697 nm) mesobiliverdin on each subunit within the same heterotetrameric PC-645 molecule. Thus, a new principle of light-harvesting by phycobiliproteins developed in cryptomonads. The large PBS was reduced to a B-PE-type subunit assembled with two ormore different 10 kDa subunits and modified with various chromophores resulting in a structure that is functionally similar to the PBS (Sidler et al., 1985, 1987, 1988, 1990a; Wedemayer et al., 1991; 1992). In cyanobacteria and red algae, as many as three different phycobiliprotein types (PEC or PE, PC and APC) are necessary for the same function. SDS-PAGE revealed that PC-645 contains three different subunits (Mörschel and Wehrmeyer, 1975; 1977): two subunits with masses of 9 kDa and 10 kDa respectively, and a subunit with a mass of 15 kDa produce an overall stoichiometry of As already mentioned above, PC-645 has a unique absorption spectrum that results from three different chromophore types and that is similar to that of a PBS from Mastigocladus laminosus. The existence of two structurally different subunits was confirmed by amino-terminal amino acid sequence analysis (Sidler et al., 1985). The complete amino acid sequence of the subunits of PC-645 from Chroomonas sp. have been determined; the subunit has 70 residues and the subunit has 80 amino acid residues (Sidler et al., 1990a). The green-colored biliverdin-like chromophores were found to be

Chapter 7 Phycobilisome and Phycobiliprotein Structures

attached to Cys-18 in both subunits (Fig. 20). Surprisingly, no significant similarity was recognizable between the and subunits ofthe PC-645, as between otherphycobiliprotein and subunits. This result will be discussed furtherbelow. In contrast to absence of homology between the and subunits, the sequence identity between the two subunits is about 50%. At position 4 of the sequence a hydroxylysine was identified (Sidler et al., 1985). Although PC-645 crystals that diffract to beyond 3.3 Å have been reported (Morisset et al., 1984), no structure for this protein has yet been determined.

6. Comparison of Cyanobacterial CPhycoerythrin, Rhodophytan B-Phycoerythrin and Cryptophytan Phycocyanin-645 and Phycoerythrin-545 Figure 22 B shows the comparison ofthe amino acid sequences for with and II from cyanobacteria, from a rhodophytan alga, PE-566 from cryptomonad strain CBD and the aminoterminal sequence of from Cryptomonas maculata. This enables a unique comparison of phycobiliproteins from phylogenetically very different organisms. The subunit contains 177 amino acid residues and thus is the same size as the red algal BPE subunit. The two blue-colored PCB chromophores are attached at positions 84 and 155, as found

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in other phycobiliproteins (Sidler et al., 1988). The purple-colored 15,16-dihydrobiliverdin chromophore is doubly bound at Cys-50 and Cys-61, similar to PEs of cyanobacteria and red algae which bind a PEB or PUB at these positions. In a Glu was identified instead of the Asn found at residues in biliprotein from cyanobacteria and red algae. In from the cryptomonad strain CBD however, the Asn is present (Wilbanks et al., 1989). The chromophore-protein interactions in the PC-645 subunit may be similar to those of cyanobacterial and red algal phycobiliprotein subunits. Asp 85, required forinteractionwiththe tetrapyrrolenitrogens of phycobilin rings B and D, is conserved in all of these subunits. No deletions or insertions are seen whenthe 645 and subunits are compared. The subunit of the blue-colored PC-645 from Chroomonas sp. shows the highest sequence identity (86%!) to the subunit of the red algal B-PE and the lowest identity (50%) to The identity to cyanobacterial C-PE (65%) is intermediate between these extremes. These results supports theories about phylogenetic development of the cryptomonad chloroplast from cyanobacteria via red algal chloroplasts (Tomas and Cox, 1973; Wilcox and Wedemayer 1984; Staehelin, 1986; Ludwig and Gibbs 1985; see Chapter 5). Cryptomonad and also show a high sequence identity (86%) to one another. The

190 high structural similarity ofthe subunits of C-PE, B-PE, PC-645 and PE-545 leads to the assumption that the cryptomonad PC-645 subunit is phylogenetically derived from a red algal PE rather than from a PC, as are probably all cryptomonad phycobiliproteins. A later modification by the substitution of different types of chromophores on the B-PE subunit yielded the different types of cryptomonad phycobiliproteins seen today. Thus, PC-645 originates from red algal PE that was modified by the addition of PCB and 15, 16-dihydrobiliverdin chromophores rather than from a PC that was modified by the addition of a 15, 16-dihydrobiliverdin chromophore. Puzzling results from immunological studies now have an obvious explanation. Whereas crossreactivity was found only within the different spectral biliprotein types APC, PC and PE in cyanobacteria and red algae, PC-645 strongly crossreacted with PEs from red algae and cyanobacteria (Berns, 1967; GuardFriar et al., 1986; MacColl and Guard-Friar, 1987a). The high structural identity of the and BPE subunits found by amino acid sequence analyses explains this result. The phylogenetic origins of rhodophytan and cryptomonad phycobiliprotein subunits may be described as follows:

Walter A. Sidler 1990). At present, the mechanism by which the subunit is translocated into the thylakoid lumen is unknown.

7. The Phylogenetic Relationship of Phycocyanin-645 Subunits The amino acid sequences of the PC-645 subunits and other cryptomonad biliprotein subunits pose the most intriguing mystery concerning the phylogenetic origin(s) of these unique chromopeptides. Extended analyses of PC-645 subunits using the UWGCG software package (Devereux et al., 1984, 1987) with all groups of known biliproteins were performed. The results show that cryptomonad PC-645 subunits are not closely related to the known phycobiliproteins, linker polypeptides, B-PE or any other light-harvesting Chl polypeptides from the thylakoid. For most comparisons the amino acid sequence identity values were only 15%–20%. As discussed above, recent results indicate that the subunits also have sequences unrelated to other phycobiliproteins.Moreover, has the same number ofamino acid residues as the B-PE subunit and shares the common deletion of 8–9 residues with the subunits of B-PE and PE-II; this deletion is typical of PEs that contain subunits. These observations suggest that the cryptomonad may be derived from the subunits.

8. Specialization and Diversification of Phycoerythrins During Evolution Phycobiliproteins from cryptomonad organisms do not form trimeric or hexameric aggregates but tetrameric complexes, and therefore it is not possible to build up PBS. Phycobiliprotein trimers are formed by an interaction of the amino-terminal parts from and subunits of neighbouring monomers, as was shown by Schirmer et al., (1985). In cryptomonad phycobiliprotein subunits, a corresponding aminoterminal domain was either deleted or was never present (assuming an exchange of the original subunit by another unrelated polypeptide as discussed below). The phycobiliproteins in cryptomonads are found in the lumenal space between the thylakoids (Gantt, 1979, 1980a; Rhiel et al., 1989), but surprisingly no amino-or carboxyl-terminal extension was found in the sequence deduced from the cpeB gene of Cryptomonas sp. strain (Reith and Douglas,

Sequence comparisons of PEs from cyanobacteria, red alga and cryptomonads have revealed an interesting evolutionary development of the PE family. During evolution the PEs in cyanobacteria have been optimized and specialized for green light conditions by forming a special light-qualitydependent adaptation mechanism, chromatic adaptation (see Chapter 21). Specialization for green growth light conditions was maximal in red algae which contain abundant amounts of PE. In cryptomonads, however, only the subunit (and perhaps a portion of the subunit) of PE survived during evolution. In contrast to the previous specialization of this chromoprotein to green light conditions, an inverse trend, in which a diversification of the phycobiliproteins to various light qualities, occurred again in cryptomonads during further evolution. The subunits and additional subunit proteins were

Chapter 7 Phycobilisome and Phycobiliprotein Structures modifiedbycombinationwithvarioustypes ofgreen-, blue-, purple-, and red-colored chromophores toyield biliprotein types no longer specifically adapted for green-light conditions but adapted for a broad spectral range as are the entire biliprotein complement of cyanobacteria.

V. Linker Polypeptides, the Skeleton of the PBS

A. Interaction of Linker Polypeptides with Phycobiliproteins The ability to isolate intact PBS from cyanobacteria and red algae provided the basis for the detection of unpigmented polypeptides belonging unambiguously to the PBS (Tandeau de Marsac, 1977; Mörschel, 1982; Redlinger and Gantt 1981b). On a weight basis, these polypeptides amount to 10%–20% ofthe total PBS protein. These colorless polypeptides have been denoted as ‘linker polypeptides’ (Lundell et al., 1981a,b; Lundell and Glazer, 1983a,b,c; Zilinskas and Howell, 1983). Glazer (1985) has proposed a systematic nomenclature and abbreviated symbols to represent the various types of linker polypeptides. Linker polypeptides are believed to be located mainly in the central cavity of the torus-shaped phycobiliprotein hexamers or trimers. Polypeptide analyses of PBS from cyanobacteria and red algae by SDS-PAGE show linkerpolypeptides of three molecular-mass categories (Fig. 24): one to two polypeptides in the 75–127 kDa range; a 29–35 kDa linker polypeptide family; and one or two small linker polypeptides at about 9–12 kDa. Linker polypeptides can be divided into four groups according to their function in the PBS: (1) linkers which are involved in the assembly of the peripheral rod substructure; (2) so-called rod-core linker polypeptides thatmediate the attachment of the peripheral rods to the PBS core; (3) linkers which participate in the assembly of the core substructure; and (4) a linker involved in the interaction of the PBS core with the thylakoid membrane. Small linker polypeptides (9–12 kDa), belonging to group 1, probably also terminate the stacking of hexamers by binding to the ends of the rods (Füglistaller et al., 1984; de Lorimier et al., 1990a). It may be that all linker polypeptides from the PBS are interconnected within the PBS to form a ‘skeleton’ within the PBS.

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Linker polypeptides modulate the spectral properties of thedifferentphycobiliprotein hexamers and trimers to various extents mainly by red-shifting the absorption and fluorescence maximum of the phycobiliprotein-linker polypeptides complexes (Glazer, 1984; 1985; 1987,1988; 1989; Rümbeli and Zuber, 1988; Glauser et al., 1993). These minor spectroscopic changes, which correlate with the location of the disc-shaped phycobiliprotein units within the rod and core substructures ofthe PBS, are believed to support unidirectional transfer of excitation energy from the periphery of the PBS to the core. The protein surface of linker polypeptides do not seem to exhibit a hydration envelope as is typical for globular proteins and thus they behave as if they were somewhat hydrophobic, e.g. they show a high tendency to aggregate. They are expected to be positively charged at physiological pH, since their calculated isoelectric points are typically greater than pH 9. In contrast, the phycobiliproteins are extremelywater-soluble andcarry significantnegative charge at physiological pH. These observations suggest that linker polypeptides and phycobiliproteins interact by a combination ofhydrophobic and chargecharge interactions. Riethman et al. (1987) reported that several of the linker polypeptides were glycoproteins based on their reactivity on blots with concanavalin A (the Clegg procedure). Additional studies indicated the presence of glucose and Nacetylgalactosamine on two of four linkers isolated from Synechococcus sp. strain PCC 7942 by electroelution from polyacrylamide gel slices (Riethman et al., 1988). However, in more carefully controlled studies and using much more sensitive detection methods, Fairchild et al. (1991) found no evidence for glycosylation of either linker polypeptides or phycobiliproteins from Synechococcus sp. strain PCC 7942 or the PEs of the red algae Gastroclonium coulteri and Porphyridium cruentum.

B. PBS-Core Linker Polypeptides

1.

the Small Core Linker Polypeptides

denotes the smallest core linker polypeptide in Mastigocladus laminosus (Füglistaller et al., 1984). Molecular masses of 8–13 kDa have been indicated for other organisms. Based upon detailed biochemical analysis of subcomplexes isolated from the PBS cores ofvarious cyanobacteria (Lundell et al., 1981 b;

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Walter A. Sidler

Chapter 7 Phycobilisome and Phycobiliprotein Structures Lundell and Glazer, 1983a, b, c; Gingrich et al., 1983; Anderson and Eiserling, 1986; Füglistaller et al., 1984, 1987) and genetic analyses of the core components from Synechococcus sp. PCC 7002 (Bryant, 1988; 1991; Maxson et al., 1989), the linker polypeptide is suggested to be associated with trimeric APC at the peripheries ofthe core cylinders. This polypeptide seems not to be absolutely required for PBS assembly, but its presence considerably improves both the stability and energy transfer properties of PBS (Füglistaller et al., 1987; Maxson et al., 1989; Bryant, 1991). Reuter and Wehrmeyer (1988,1990) locate of Mastigocladus laminosus between the first and second APC trimer of the PBS core cylinders. The carboxyl-terminal regions of REP4 and ARM5 of the from Mastigocladus laminosus are 39% and 37%, respectively, identical to the (Esteban, 1993, Esteban et al., 1994). This suggests that the linker protein probably has a similar function to these domains in the APC complexes of the fourth core cylinder. From spectroscopic analyses of APC core complexes from Mastigocladus laminosus it was deduced that the does not influence the orthe chromophores,butit shifts theabsorption maximum of the chromophore to a longer wavelength than the absorption maximum of the chromophore in trimeric complexes. increases the oscillator strength of the chromophores and turns the chromophores from sensitizing into weakly fluorescing chromophores (Füglistaller et al., 1987). Hydropathy plot analysis of the suggested a three-fold symmetry in the cyclically arranged amino acid sequence of indicating a similar interaction of each of these segments with the APC (Cys84-PCB chromophore domain). A hydrophobic carboxylterminal ‘tail’-segment might be located in the very center of the complex (Füglistaller et al., 1987). Maxson et al. (1989) produced a apcC mutant of Synechococcus sp. strain PCC 7002 lacking Themutant,which grewsomewhat moreslowlythan the wild type, formed PBS indistinguishable from wild-type PBS, except for a small decrease fluorescence emission at 680 nm. The PBS were less stable than those of the wild-type and were more easily dissociated at lowered ionic strength or at increased temperature. Nonetheless, the function of in the PBS seems less essential than that of or linkers proteins.

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in red algae behaves differently: a small corelinker polypeptide, denoted as and associated with an AP-B-enriched fraction, was found in the PBS of the red alga Porphyridium cruentum (Graham et al., 1994). The AP complex containing this linker polypeptide was isolated and analyzed by SDS-PAGE and directprotein-microsequencing ofthe gelbands. An amino acid sequence was found that is not related to that of any linker polypeptide sequence known from the functionally homologous linker polypeptides of cyanobacteria.

2.

the Core-Membrane Linker Polypeptide

denotes the largest, multifunctional linker phycobiliprotein in the PBS that also serves as the terminal emitter ofenergy in isolated PBS and as the transmitter of light energy to PS II in intact cells. Its structure, function and genetic analysis is discussed in Sections III B and C.

C. The Rod and the Rod-Core Linker Polypeptides linkerpolypeptides canbe divided into two groups differing in their molecular masses: group I consists of an 8 to 10 kDa polypeptide denoted in Mastigocladus laminosus; and group II, consists of polypeptides with masses of about 30 kDa.

1.

the Small Rod Linker Polypeptide

denotes the small linker polypeptide associated with PEC or PC complexes in the PBS rods of Mastigocladus laminosus and is believed to be bound at the core-distal end of the rods, thus minimizing the heterogeneity of the rod lengths (de Lorimier et al., 1990a). The complete amino acid sequences of both small linker polypeptides Füglistaller et al., 1984) and and partial amino acid sequences of the rod linker proteins and (Füglistaller et al., 1985, 1986a,b), revealed that the linker polypeptides were quite basic (e.g., has a net charge of +6 at pH 7). Only one gene coding an was found in the pec and cpc operons (Eberlein and Kufer, 1990; Kufer et al., 1992). It has to be assumed, that the same terminates the rods ending with PEC in PBS from cells grown in low light intensity and also ending with C-PC complexes in PBS from cells grown at high light intensity.

Walter A. Sidler

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2.

the Rod Linker Polypeptides

denote the linker polypeptides associated withphycobiliproteins at least one hexamer or more distant from the core substructure in the peripheral rods. (The first hexamer ofthe rod element always contains an linker and is involved in the rod-core linkage with APC. See below.). Cyanobacteria contain, depending on the length of their PBS rods, one or more proteins. The different rod linkers ofthe 30 kDa family are associated either with PE, PEC or PC and cause a red-shift of the absorption and fluorescence emission maxima ofthe phycobiliproteins to which they are bound in vitro (Lundell et al., 1981a; Yu et al., 198la). Mutational studies in Synechococcus sp. strain PCC 7002 have established that in vivo these linker polypeptides mediate the stacking of the neighboring phycobiliprotein hexamer to that proximal to the core (de Lorimier et al., 1990b). The complete amino acid sequences of the and from Mastigocladus laminosus are presented in Fig, 25 together with the sequences of the rod-core linker polypeptides. The sequences of the 34.5 kDa rod linker polypeptides and were completed by DNAsequencing (Eberlein and Kufer, 1990; Kufer et al., 1992). The seems to have a more minor influence on the energy transfer within the rod than the Bhalerao et al. (1991) generated a mutant strain (IMcpcI) of Synechococcus sp. strain PCC 7942 in which the protein was missing. In this mutant the protein could occupy the position protein with no detectable change in the energy harvesting or energy transfer characteristics of the rods.

3.

Rod-Core Linker Polypeptides

A key position in the PBS structure and energy transfer pathway is the rod-core junction. A special class of linker polypeptide with a molecular mass of about 30 kDa, the C-PC-associated rod-core linker polypeptide attach the peripheral rods to the PBS core and form different types of C-PC-to-APC interactions (Lundell et al., 1981a; Yu et al., 1981a; Yu and Glazer, 1982). It was generally believed that only one type of rod-core linker polypeptide is involved in the attachment of all peripheral rods to the PBS core. This appears to be true for many cyanobacteria including Synechococcus sp. strains PCC 6301 and 7002. However, in the cyanobacteria Mastigocladus laminosus and Anabaena sp. strain

PCC 7120 more than one type of linker polypeptide was found to attach the rods to the PBScore (Glauser et al., 1990, 1992 a, b; Bryant et al, 1991). The cpcG gene of Synechococcus sp. strain PCC 7002 (Bryant, 1988,1991) and four cpcG genes in Anabaena sp. strain PCC 7120 (Bryant et al., 1991) have been cloned and sequenced. The proposed function of the polypeptides in Synechococcus sp. strain PCC 7002 has been confirmed in vivo by analysis of a mutant in which the unique cpcG gene, encoding the polypeptide, was insertionally inactivated (Bryant, 1991). This mutant fluoresces brightly at wavelengths characteristic for free PC, indicating that PC, which is produced at normal levels, is not effectively coupled to the Chl a antenna of PS II. Additional analyses demonstrated that this mutant does not assemble any PC onto PBS cores (Bryant, 1991). The rod-core linker polypeptides are seen to play two main roles: firstly, they associate the peripheral rods with the PBS core; secondly, they impart a strong red-shift in the wavelength of maximum absorbance and fluorescence emission of the central phycocyanobilin chromophores, to enable an optimal rod-to-core energy transfer. A 22 kDa proteolytic degradation product, originating from a polypeptide of Mastigocladus laminosus, caused a major redshift of about 15 nm when associated with PC trimers (Gottschalk et al., 1991), indicating an essential functional role in excitation energy transfer. By DNA sequence analysis, Bryant et al. (1991) found four similar cpcG-genes in the cpc-operon from Anabaena sp. PCC 7120, whereas Synechococcus sp. strain PCC 7002 contained only one cpcG gene. Microanalytical protein amino acid sequence analyses of the linker polypeptides from the PBS of Mastigocladus laminosus revealed four different sequences, typical for rod core linker polypeptides. However on SDS-PAGE only three bands containing have so far been separated and identified. The band showed a heterogeneous ammo-terminal amino acid sequence (N/K at position 9 and R/D at position 28) and the stoichiometric ratio of : was 1:2:1, providing evidence for the possible existence of a fourth linker in Mastigocladus laminosus (Glauser et al., 1990; Bryant et al., 1991, Glauser et al, 1992a, 1992b). From the PBS of Anabaena sp. strain PCC 7120, three different linkers were also identified by this method. No amino-terminal sequence corresponding

Chapter 7 Phycobilisome and Phycobiliprotein Structures

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Walter A. Sidler

196 to the product of the cpcG3 gene was detected, but it seems highly unlikely that the cpcG3 gene is not translated (Bryant et al., 1991). The fourth (CpcG3) was identified only by DNA sequence analysis. The amino-terminal sequence identity within the of the same PBS was high. Still higher identity values were found between equivalent pairs of polypeptides from Anabaena sp. strain PCC 7120 and from Mastigocladus laminosus. In contrast to this, only one type of was identified in the PBS of Calothrix sp. strain PCC 7601in green-lightgrown cells (containing C-PE) or in red-light-grown cells (containing only C-PC as rod elements) by protein microsequence analysis (Glauser et al., 1992c). Thus, two types of PBS can be distinguished on the basis of the number of which occur in them: those with a single type of such as the PBS of Calothrix sp. strain PCC 7601 and Synechococcus sp. strain PCC 7002; and those with up to four different such as the PBS of Anabaena sp. strain PCC 7120 and Mastigocladus laminosus. The similarity of the primary structures for all is quite high, but similarity values are significantly lower when and proteins are compared, linker polypeptides form a special linker polypeptide family, and their amino-terminal amino acid sequences differ sufficiently from those of the family that such sequences may be used as convenient ‘fingerprints’ for the and families (Glauseret al., 1990). As a result of these studies, the classical hemidiscoidal PBS model, which has been adapted for Mastigocladus laminosus and also for Anabaena sp. strain PCC 7120, had to be reconsidered. Fig. 9 B shows a working model for the architecture of a PBS for Mastigocladus laminosus and Anabaena sp. strain PCC 7120. The working model is based on the determined stoichiometries ofthe linker polypeptides and phycobiliproteins in the PBS (Glauser et al., 1992a) and the core architecture implied by the 4REP (Esteban, 1993). A PC-to-APC ratio of 2:1 was obtained by laser densitometry of polypeptides from the PBS from Mastigocladus laminosus that had been separated on SDS-PAGE gels. Similar ratios were also obtained by spectroscopic quantification of the PC and APC fractions isolated from different batches of PBS from Mastigocladus laminosus and Anabaena sp. strain PCC 7120. A ratio of 2 :

(2+2) : 2 : 8 was obtained by laser densitometry of the separated rod and rod-core linker polypeptides from Mastigocladus laminosus by SDS-PAGE. Thus, eight rods may theoretically be attached to the PBS core. This arrangement of eight rods allows two occurrences each offour different PC-to-APC (‘rodto-core’) interactions involving the core subcomplexes 1 + 2, 1 + 3, 3 + 4, and the peripheral-rod APC hexamer. Each one of these interactions would be specifically mediated by one of the four rod-core linker polypeptides. Electron micrographs of reconstituted PBS cores of Anabaena variabilis strain M3 (Isono and Katoh, 1987) support the idea of additional APC bound to the core and electron micrographs of whole PBS from Mastigocladus laminosus (Fig. 1C) and Anabaena sp. strain PCC 7120 (Fig. 1D) show that these PBS are definitely different from the Calothrix sp. strain PCC 7601 PBS with six rods (Fig. 1 A,B) and do not contradict this model (Bryant et al., 1991; Glauser et al., 1992a). In order to confirm or refine further the novel PBS architecture of Mastigocladus laminosus and Anabaena sp., several important structural details of the PBS core architecture still remain to be determined or confirmed by additional experiments. These include the specificity of the rod-binding to the core, the determination of the number of rods bound to each core-cylinder, the composition ofthe individual rods as well as the total number of rods per PBS. Models with two rods bound to a AP-“hexamer” have been described previously (e.g., the two-core-cylinder and six peripheral rods model of Lundell and Glazer, 1983a, b, c). Interpretation of electron micrographs from Mastigocladus laminosus is difficult and electron microscopy alone is unlikely to resolve questions concerning the number of core cylinders and the number and the size ofthe rods as is possible with Calothrix sp. strain PCC 7601 PBS.

D. Functional Domains of Binding Specificity of

and

and

Most linker polypeptides have calculated isoelectric points in the pH range 8–11, and it is probable that electrostatic interactions with the negatively charged phycobiliproteins and hydrophobic (possibly aromatic) interactions are especially important in the interaction of these two types of proteins. The complete amino acid sequences of the linker polypeptides have been determined by nucleotide

Chapter 7 Phycobilisome and Phycobiliprotein Structures sequencing of the cpcG genes of Anabaena sp. PCC 7120 phycobilisomes (Bryant et al., 1991) and Mastigocladus laminosus PBS (Glauseretal., 1992b). The sequences of three cpcG genes, encoding polypeptides with 279, 247 and 254 amino acid residues, have been determined from the latter cyanobacterium (Fig. 25). A high degree of homology in the amino-terminal domains (190 amino acids) and an overall identity of 47–53% between the three gene products was found. Each of the three CpcG polypeptides is highly related to one of the four polypeptides from Anabaena sp. strain PCC 7120 (66–81% identity). It is striking that these crossspecies pairs of CpcG proteins have nearly identical numbers of amino acid residues. Possibly these pairs of polypeptides mediate the same specific type of PC-to-AP interaction in the similar PBS of Mastigocladus laminosus and Anabaena sp. strain PCC 7120. The similarity values for the CpcG1, CpcG2 and CpcGS polypeptides to the single polypeptide of Synechococcus sp. PCC 7002 (36– 41% identity; Bryant, 1991) are much lower. In all analyzed cyanobacterial and polypeptides, the initiator methionines have been posttranslationally removed. Amino-terminal sequence analyses of a partially degraded isolated from an intact complex demonstrated that the amino-terminal portion of the still binds to the (Glauser et al., 1993). This was also demonstrated with a 22 kDa proteolytic fragment from the protein, isolated from a crude phycobiliprotein extract from Mastigocladus laminosus and reassociated with (Gottschalk et al., 1991). Amino-terminal sequence analysis showed that the amino-terminal part was unaffected by proteolysis (Glauser et al., 1992b). Thus the amino-terminal part of the binds to the and presumably the carboxylterminal parts will bind to theAPC. Interestingly, the amino-terminal part bound to exhibits high sequence homology within the whereas the carboxyl-terminal parts of the polypeptides have different sizes and show low similarities to one another within the same organism (Fig. 25). This finding suggests that the may bind specifically to the APC-complexes ofthe core. This hypothesis was partially tested by reconstitution experiments of different rod and core elements (Glauser et al., 1993) and with from Mastigocladus laminosus overproduced in E. coli (Ruegsegger et al. 1993; Sidler et al., 1993).

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A common feature of and is the binding to PC and PEC hexamers (or trimers) presumably in the inner space of the C-PC and PEC hexamers. Despite the low overall sequence similarity of and similarity comparisons of and sequences in their amino-terminal domains revealed the existence of six motifs with high similarity, formed by the amino acid sequence motifs 47–59, 60–77, 80–96, 108–120, 145–166, 174–186 and 174–186 of and (Fig. 25). This could indicate functional and structural interaction sites with a corresponding motif 106–124 ofthe six phycobiliprotein subunits in the inner space ofthe phycobiliprotein-hexamer (Glauser et al., 1992b). Interactions could involve salt bridges and hydrophobic interactions, and the aromatic residues might be responsible for the red-shift observed in these complexes. Only eight and seven charged residues ofthe and subunits, respectively, seem to form neither salt bridges nor hydrogen bonds within the phycobiliprotein structure, as was seen from the crystal structures. Three of these side chains Lys Glu andAsp point towards the central channel of the trimeric complex and may be important for CPC linker association. The sequences of linker polypeptides show high degrees of amino acid sequence identity within their own linker family (34.5 kDa linker polypeptide family: 41%; 8.9 kDa linker polypeptide family: 31%). Lower identity values were found between these two LPP families: 28%. The identity to phycobiliproteins was very low on average—between 4% and 21% in different fragments. A hypothetical evolution of the genes encoding the phycobilisome proteins from Mastigocladus laminosus was proposed on the basis of the known sequences (Füglistaller et al., 1985). This scheme suggested that all polypeptides of the PBS may be derived from an ancestral phycobiliprotein subunit related to myoglobin. The 34.5 kDa linker polypeptide family was proposed to originate from a fusion ofthe genes and the corresponding intercistronic DNA sequence. However, homologies of 21% and more have also be found between phycobiliproteins and other proteins from the photosynthetic apparatus (Sidler et al., 1990a), and it may be possible that the linker polypeptides developed from an earlier (possibly non-globin) ancestor of the phycobiliproteins. In this context, the phylogenetic development of the large core-membrane linker phycobiliprotein that contains a phycobiliprotein-domain and

Walter A. Sidler

198 linker domains (the ‘REP-domains’) within the same polypeptide chain, must also be considered (Section III B,C). In present-day PBS the is a prerequisite for proper functioning and assembly of the PBS. The arrangement of the pecA, pecB, cpcA , and cpcB genes suggests that PEC evolved later by duplication of the cpc operon (cpcBACD Eberlein and Kufer, 1990; Kufer et al., 1992; W. Kufer, personal communication) and not from a common ancestor of the subunits as proposed by Füglistaller et al., (1985).

E. Rod-Linker Polypeptides for Phycoerythrin Complexes Federspiel and Grossman (1990) reported the DNA sequences of two genes (cpeC and cpeD) encoding PE-associated linker polypeptides. A third C-PE associated linker polypeptide in the PBS of Calothrix sp. strain PCC 7601 was detected by amino acid sequence analysis of a 33 kDa linker polypeptide, isolated from the complex (Glauser et al., 1992c). PBS of Calothrix sp. strain PCC 7601 cells grown in green light have a PE:PC ratio of three, and electron microscopy of such PBS suggested that most peripheral rods contained four biliprotein hexamers (see Fig. 1 A and Glauser et al., 1992c). Federspiel and Scott (1992) separated the and polypeptides by varying the pH during SDSPAGE and completed the nucleotide sequence of cpeE gene. The complete deduced amino acid sequences for the three PE-associated linkers of Calothrix sp. strain PCC 7601 exhibit 31% identity within the common 259 residues and show the same six conserved domains which were proposed by Glauser et al. (1992b) to interact specifically with the phycobiliprotein in the inner space of the hexamer and to play a functionally and structurally important role (see Section V D). The predicted sizes and molecular masses are: amino acids 31.8 kDa amino acids 27.9 kDa amino acids 27.6 kDa

F. Phycobiliprotein-Linker Polypeptide Complexes from the Phycobilisome of Mastigocladus laminosus Six different phycobiliprotein-linker polypeptide complexes from the PBS of Mastigocladus laminosus

have been characterized (Table 2). These include the following: and (III; Füglistaller et al., 1986); and and Glauser et al., 1992a). Complexes IV, V and VI contain polypeptides and complex VI even contains a core subcomplex. No complex with the (cpcG1 gene product) as the only linker polypeptide component has yet been isolated from the PBS of Mastigocladus laminosus. Two different complexes (IV and V) containing the and linker polypeptides have been obtained (Table 2). Analogous trimeric complexes with similar spectroscopic properties had been isolated from Anabaena variabilis (Yu et al., 1981a) and Synechococcus sp. strain PCC 7002 (de Lorimier et al., 1990b). The trimeric state of aggregation of these complexes, however, does not seem to correspond to that suggested in native PBS. Electron microscopic analyses (e.g., see Figs. 1A-D), the propensity for disproportionation ofthese complexes in high phosphate concentrations (Yu et al., 1981; de Lorimier et al., 1990b) and results from picosecond energy transfer kinetics that indicate a more efficient energy transfer in hexameric phycobiliprotein discs than in trimeric (Holzwarth et al., 1987; Sandström et al., 1988a, b), all show that the PBS rods are built up of hexameric phycobiliprotein-linker polypeptide complexes (Fig. 9). In contrast to this generally accepted structure of the PBS rods, another model for the architecture of the PBS rods in Mastigocladus laminosus, encompassing peripheral and internal trimeric phycobiliproteinlinker complexes, has recently been postulated (Reuter, 1989, Reuter and Nickel-Reuter, 1993). As shown in Table 2 and Fig. 26, the absorption and fluorescence emission maxima of the and complexes shift to longer wavelengths. Such a gradient in light absorption and emission is consistent with the positioning of the disc-shaped phycobiliprotein-linker polypeptide aggregates within the PBS rods and facilitates unidirectional energy transfer towards the core (Zuber, 1987; Glazer, 1989). By reconstitution experiments Glauser et al. (1993) showed that the PC-to-APC interaction in complex VI, mediated by the polypeptide is specific and cannot be reconstituted by the peripheral-rod complex or even by a similar rodcore complex This is an indication

Chapter 7 Phycobilisome and Phycobiliprotein Structures

that each of the four polypeptides in the PBS of Mastigocladus laminosus and Anabaena sp. strain PCC 7120 may attach two peripheral rods specifically to oneofpresumablyfourdifferentcorebindingsites (assuming eight total binding sites). The PC-linker polypeptide complex was successfully reconstituted with isolated and overproduced in E. coli. (Rüegsegger et al., 1993; Sidler etal., 1993). Inaddition, therod-core complex was reconstituted with isolated and the reconstituted The complex showed an absorption maximum at 616 nm and an emission maximum at 642 nm. The absorption maximum of

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the reconstituted complex was at 632.4 nm and the emission maximum at 645 nm. Thus, a redshift of 16.4 nm in the absorption maximum, characteristic for a functionally correct reconstituted complex was observed. The reconstituted complex showed an emission maximum at 659 nm which is only 3 nm lower than the emission maximum of

VI. Organization of the Genes Encoding the Phycobilisome Elements Extensive structural and functional analyses of PBS

200 via molecular genetics have been performed for Synechococcus sp. strain PCC 7002 (for a review, see Bryant, 1991) and for other cyanobacteria (e.g., see Houmard et al., 1986; Tandeau de Marsac et al., 1990; Grossman, 1990; Tandeau de Marsac and Houmard, 1993; Grossman et al., 1993). The number of transcriptional units encoding PBS components varies in the different cyanobacteria (about 5–9 units), and the distribution of genes within these units is also variable. The organization and transcription patterns for genes encoding PBS components of several cyanobacteria was recently summarized by Bryant (1991). The organization of the complete set of genes encoding structural components of PBS for Mastigocladus laminosus is shown in Fig. 27. The cpc operons are composed of genes encoding PBS rod components and polypeptides that are involved in chromophore attachment to the subunit (cpcE and cpcF genes; Bryant, 1988, 1991; Zhou etal., 1992; Swanson et al., 1992b; Fairchild et al., 1992). The cpcG gene(s), encoding the are contained in the large pec-cpc superoperon in Anabaena sp. strain PCC 7120 and Mastigocladus laminosus, whereas in ynechococcus sp. strain PCC 7002 the cpcG gene forms a separate transcription unit. In Calothrix sp. strain PCC 7601 the cpcG gene has not yet been cloned but is not encoded by one of the three cpc operons. In all cyanobacteria studied to date, the cpcA gene, encoding the subunit, is located downstream from the cpcB gene encoding the subunit. The same order of genes has been determined for the cpeBA operon coding for the and subunits (Mazel et al., 1986; Dubbs and Bryant, 1987,1991; Anderson and Grossman, 1990b; Bernard et al., 1992) and for the pecBA operon (Eberlein and Kufer; 1990; Swanson et al., 1992a). The genes encoding the PC or PEC subunits are typically followed by genes encoding the linker polypeptides and/or the genes for chromophore attachment to the alpha subunit. The reverse order is found in all apc operons analyzed so far; the apcA gene, encoding the is found 5' to the apcB gene that encodes (Bryant, 1988,1991). The apc operons, that have been shown to have variable gene composition, have been characterized from several cyanobacteria including Synechococcus sp. PCC 7002 (Bryant, 1988, 1991), Calothrix sp. strain PCC 7601 (Houmard et al., 1988, 1990), Synechococcus sp. strain PCC 6301 (Hournard et al., 1986), Anabaena variabilis (Johnson

Walter A. Sidler et al., 1988), and Synechocystis sp. strain PCC 6803 (Su et al., 1992). In all cyanobacteria examined thus far, the apcC gene, encoding the small linker polypeptide lies downstream from the apcB gene. The apcD and apcF genes encode the minor APC-related phycobiliprotein subunits and respectively. The apcD gene occurs as a monocistronic unit in Synechococcus sp. strain PCC 7002 (Maxson et al., 1989), Calothrix sp. strain PCC 7601 (Houmard et al., 1988), and Mastigocladus laminosus (Esteban, 1993; see Fig. 27). The apcF gene likewise occurs as a monocistronic unit in Synechococcus sp. strain PCC 7002 (Bryant, 1988; 1991) and Mastigocladus laminosus (Esteban, 1993; see Fig. 27). The apcE gene encodes the large coremembrane linker phycobiliprotein In Synechococcus sp. strain PCC 7002 (Bryant, 1988, 1991), the apcE gene is transcribed as a separate unit, whereas in Synechococcus sp. strain PCC 6301 (Capuano et al., 1991), Calothrix sp. strain PCC 7601 (Houmard et al., 1990), and Mastigocladus laminosus it is located on a transcriptional unit together with apcA, apcB and apcC. This arrangement was first reported for Nostoc sp. strain MAC by Zilinskas and Howell (1987). In cyanelles of Cyanophora paradoxa, the apcE gene is also found upstream from apcAB, but no apcC gene has been found in the cyanelle genome (Bryant et al., 1985; Bryant, 1988; also see Chapter 4). A second apcA2 gene, encoding a second subunit and with 59% sequence identity to not included in the apcEA 1BC operon but forming a monocistronic unit was found in Calothrix sp. PCC 7601 (Houmard et al., 1988). Its function and location within the PBS is not yet clear.

A. Genes Involved in Adaptation to Changes in Environmental Conditions In some cyanobacteria multiple gene sets for some types of phycobiliproteins and linker polypeptides have been found. For example, two cpcBA gene sets occur in Synechococcus sp. strain PCC 6301 (cpcB1A1 and cpcB2A2; Lind et al., 1987; Lau et al., 1987a, b; Kalla et al., 1988) and Pseudanabaena sp. strain PCC 7409 (cpcB1A1 and cpcB2A2; Dubbs and Bryant, 1987, 1993) or even three different cpcBA gene sets in Calothrix sp. strain PCC 7601 (Mazel et al., 1988; Capuano et al., 1988; Mazel and Marlière,

Chapter 7 Phycobilisome and Phycobiliprotein Structures

1989). In Synechococcus sp. strains PCC 6301 and PCC 7942, the duplicated phycocyanin genes encode identical polypeptides and are arranged as a tandemrepeat unit with the genes for the three rod linkers between the cpcB1A1 gene set and the downstream cpcB2A2EF gene set (Kalla et al., 1988; Kalla et al., 1989; Bhalerao et al., 1993). The occurrence of several different gene sets for the same type of PBS component is apparently the result of adaptation of these organisms to environmental conditions such as light quality (complementary chromatic adaptation) and nutrient availability (in particular, sulfur availability; Mazel and Marlière, 1989). The molecular mechanism of this regulation by light quality has been the subject ofintensive studies at the genetic level for several years (Conley et al., 1988; Grossman et al., 1986, 1988; 1993; Grossman, 1990; Tandeau de Marsac et al., 1988, 1990; Tandeau de Marsac, 1991; Dubbs and Bryant, 1991, 1993; Tandeau de Marsac and Houmard, 1993). In Calothrix sp. strain PCC 7601 three copies of the genes encoding the subunits (cpcA1, 2, 3) and the subunits (cpcB1, 2, 3) have been found on three different operons and sequenced (Conley et al., 1986, 1988; Lomax et al., 1987; Mazel et al., 1988; Capuano et al., 1988; Mazel and Marlière,

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1989). Only two copies are necessary for the complementary chromatic adaptation: under greenlight growth conditions, only the cpc1 operon encoding the constitutive PC 1 is transcribed together with the cpe operons, while under red-light growth conditions, the cpc2 operon, encoding the inducible PC2 is transcribed together with the cpc1 operon. Whether separate or mixed C-PC trimers are formed in Calothrix sp. strain PCC 7601 from the two subunit classes is not known. However, in Nostoc sp. strain MAC, evidence for the formation of mixed trimers was found (G. Guglielmi and D. A. Bryant, unpublished results). The cpc3 operon, encoding PC3 in Calothrix sp. strain PCC 7601, is transcribed only under sulfur-limited growth conditions but the cpc1, cpc2 and cpe operons are completely switched off under these conditions (Mazel and Marlière, 1989; see Chapter 21). Different sets of PC-associated linker polypetides are synthesized with PC2, PC3, and PE (see Grossman et al., 1993; Tandeau de Marsac and Houmard, 1993; and Chapter 21). The cpeBA operons of Calothrix sp. strain PCC 7601 (Mazel et al., 1986), Pseudanabaena sp. strain PCC 7409 (Dubbs and Bryant, 1987, 1991), and Synechocystis sp. strain PCC 6701 (Anderson and Grossman, 1990b) were not found to contain genes

202 for linker polypeptides. In Calothrix sp. strain PCC 7601, the three PE-associated rod-linker polypeptides have been found to occur on a separate cpeCDE operon that is only transcribed when cells are grown in green light (Federspiel and Grossman, 1990; Federspiel and Scott, 1992). In contrast, the mpeC gene for the bifunctional linker polypeptidephycobiliprotein of PE-II was found on the same transcription unit in an organism that does not perform chromatic adaptation but that is strongly adapted for growth under green light conditions (Wilbanks and Glazer, 1993a, b). DNA sequence analysis of Synechococcus sp. strain WH 8020, a unicellular, open-ocean Synechococcus sp., showed the genes for the and subunits of PE-I and PE-II as well as for the subunit of PE-II to be clustered in a large 15-kb region of the genome. Twelve open reading frames, the largest number found in such an operon, are included in this region (Wilbanks and Glazer 1993a).

B. The cpcE and cpcF Genes The cpcE and cpcF genes, now identified in several cyanobacteria (see Bryant, 1988, 1991; Zhou et al., 1992), are located on the cpc operon containing the genes encoding the constitutive PC subunits. These genes have been identified and sequenced from Synechococcus sp. strain PCC 7002 (Zhou et al., 1992), Calothrix sp. strain PCC 7601 (Mazel et al., 1988), Anabaena sp. strain PCC 7120 (Belknap and Haselkorn, 1987; Bryant et al., 1991), Pseudanabaena sp. strain PCC 7409 (Dubbs and Bryant, 1993) and Mastigocladus laminosus (Kufer et al., 1992). Mutational analyses in Synechococcus sp. strain PCC 7002 indicated that strains harboring interposon mutations in cpcE, cpcF, or both genes were defective for PCB attachment to the subunit (Zhou et al., 1992; Swanson et al., 1992). Overproduction of the CpcE and CpcF proteins in E. coli was used to show that the two proteins together form an enzyme, phycocyanobilin lyase, that can attach the PCB chromophore to the apoprotein (Fairchild et al., 1992). This enzyme can also catalyze the removal of PCB from the subunit and can catalyze an exchange reaction in which the PCB chromophore is transferred from an holoprotein of one species to an apoprotein of another species. The CpcE and CpcF gene products form an enzymatically active heterodimer (CpcE: CpcF = 1:1), and the enzyme can also catalyze the addition of phycoerythrobilin to the

Walter A. Sidler subunit. However, the enzyme shows a preference for phycocyanobilin over phycoerythrobilin, both binding affinity and in rate of catalysis, that is sufficient to account for the selective attachment of phycocyanobilin to the subunit (Fairchild and Glazer, 1994). Genes homologous to the cpcE and cpcF genes, denoted pecE and pecF (the pecF gene was originally designated ORF173), have been located downstream from the pecBAC operon of Anabaena sp. strain PCC 7120 (Swanson et al., 1992b). The PecE protein is predicted to be 253 amino acids in length and has an average identity of 47% to several CpcE proteins. The PecF protein is predicted to be 173 amino acids in length and is approximately 27% identical to several CpcF proteins. It is presumed that the PecE and PecF act in a fashion analogous to the CpcE and CpcF proteins; thus, these proteins would catalyze the attachment of the PXB chromophore to the subunit. Genes homologous to the cpcE and cpcF genes, denoted rpcE and rpcF, have also been located downstream from the rpcB-rpcA operon of Synechococcus sp. strains WH 8020 and WH 8103 (de Lorimier et al., 1993). The rpcE gene encodes a polypeptide of 265 amino acids that shows an average of ~40% sequence identity to three CpcE polypeptides and 34% identity to the PecE protein of Anabaena sp. strain PCC 7120. The rpcF gene encodes a polypeptide of 210 amino acids that is ~37% identical to CpcF proteins and 28% identical to the PecF protein of Anabaena sp. strain PCC 7120. The subunit of R-PC-II carries a phycoerythrobilin chromophore (see Section IV A, 5). It is presumed that the specificity for attachment of a phycoerythrobilin chromophore rather than a phycocyanobilin chromophore resides in the RpcE and RpcF proteins. Finally, additional homologs of the cpcE and cpcF genes of cyanobacteria have been identified in the vicinity of genes encoding phycoerythrins in Calothrix sp. strain PCC 7601 (Tandeau de Marsac et al., 1988), Pseudanabaena sp. strain PCC 7409 (Dubbs and Bryant, 1991), and Synechococcus sp. strain WH 8020 (Wilbanks and Glazer, 1993a). In Calothrix sp. strain PCC 7601 and Pseudanabaena sp. strain PCC 7409, open reading frames denoted orfZ were detected downstream from the cpeBA operon. These potential genes, that encode proteins of 202 and 205 amino acids, respectively, show significant homology to members of the CpcE family

Chapter 7 Phycobilisome and Phycobiliprotein Structures (see Fig. 5 of Wilbanks and Glazer, 1993a) and a closely related gene, denoted cpeZ, was located between the mpeBAC and cpeBA operons of Synechococcus sp. strain WH 8020. Two additional genes, denoted mpeU and mpeV, encode proteins of 297 and 301 amino acids, respectively, that are also members of the CpcE protein family (Wilbanks and Glazer, 1993a). It seems quite likely that the proteins encoded by these genes and open reading frames are involved in chromophorylation of PE subunits.

C. Genes Encoding Phycobilisome Components in the Cyanelles of Cyanophora paradoxa, Red Algae, and Cryptomonads Studies with inhibitors of protein synthesis demonstrated that the major pigmented polypeptides and the linker protein of the phycobilisomes are translated on 70S plastid ribosomes while some the subunit of PE and some linker polypeptides are translated on cytoplasmic 80S ribosomes (Egelhoff and Grossman, 1983). Similar results were found for Porphyridium aerugineum, Porphyridium cruentum, Cyanidium caldarium, and Cyanophora paradoxa (Egelhoff and Grossman, 1983; Grossman et al., 1983). Molecular cloning and nucleotide sequence analyses soonconfirmedthese results for Cyanophora paradoxa. The cpcB gene encoding the subunit was identified first (Lemaux and Grossman, 1984), and shortly thereafter the genes encoding the subunit and the and subunits ofAP were isolated and sequenced (Bryant et al., 1985; Lemaux and Grossman, 1985; see Chapter 4). Further nucleotide sequence analysis upstream from the apcAB genes demonstrated the presence of the apcE gene in the cyanelle genome as anticipated from the inhibitor studies (Bryant, 1988, 1991). A sixth gene, the apcD gene that encodes the subunit of phycobilisomes, was subsequently identified (Michalowski et al., 1990). A homolog of the apcF gene has recently been identified upstream from the cpcBA operon of the cyanelle (see Chapter 4). Biliprotein genes have also been identified on the plastid genomes of several red algae and cryptomonads. Apt and Grossman (1993a,b,c) have extensively characterized genes encoding phycobilisome components from the red alga Aglaothamnion neglectum. To date, nine genes have been cloned and sequenced: apcAB, cpcBA, cpeBA, apcE, apcF, and cpcG. An open reading frame that could encode a small, AP-associated linker polypeptide was detected

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downstream from apcF, but confirmation that this gene actually encodes a phycobilisome component has not yet been obtained. All of these genes have also been identified on the plastid genome ofred alga Porphyra purpurea except for cpcG, which has not yet been detected (Reith and Munholland, 1993). The genes encoding the R-PE subunits of Polysiphonia boldii (Roell and Morse, 1993) and the BPE subunits of Rhodella violacea (Bernard et al., 1992) have also been isolated and sequenced. Interestingly, the rpeB gene of the latter organism is split by an intron with characteristics of group II. Finally, Valentin et al. (1992) characterized a phycobiliprotein gene cluster in the plastid genome of Cyanidium caldarium that includes the apcF, ‘apcC’ and cpcG genes. The apcF-’apcC’ genes form a dicistronic transcription unit; the cpcG gene was found adjacent to this operon and is convergently transcribed towards a common stem-loop structure. These three genes occur with the same organization found in Aglaothamnion neglectum (Apt and Grossman, 1993c). Apt et al. (1993) recently isolated and characterized a cDNA encoding a subunit of the R-PE of Aglaothamnion neglectum. This represents the first isolation of a cDNA for a nuclear-encoded phycobilisome component from a red alga. The cDNA predicts a protein of 317 amino acid residues and demonstrates the occurrence of a presequence of 40 amino acids for chloroplast targeting. The fulllength protein has a predicted mass of 34.6 kDa and a predicted pI of 9.6. The subunit is predicted to contain sequences surrounding four cysteine residues that are highly similar to those offour chromopeptides (1 phycoerythrobilin and3 phycourobilins arebound) isolated from the subunit of the R-PE of Gastroclonium coulteri. The 40-amino acid presequence of the R-PE subunit, when fused to the pea Rubisco small subunit polypeptide, could direct the uptake of this polypeptide by pea chloroplasts. Although this polypeptide was notproperlyprocessed during import, the Rubisco small subunit could nonetheless be assembled into Rubisco holoenzyme. These results suggest that the uptake of proteins translated in the cytoplasm of red algal cells may occur by a targeting process that is similar to the process that occurs in higher plants. Very little information is presently available for the genes encoding phycobiliproteins in cryptomonads. As noted above, the cpeB gene, encoding the subunit of Cryptomonas sp. strain is encoded

204 in the plastid genome of this organism (Reith and Douglas, 1990). The gene predicts a protein of 177 amino acids that exhibits a very high similarity to the subunits of red algal PEs as well as cryptomonad PCs. Surprisingly, no amino-terminal or carboxylterminal targeting sequence was observed that could direct the polypeptide to the thylakoid lumen. The absence of a targeting signal suggests that a novel mechanism could be involved in directing cryptomonad biliproteins from the plastid to the thylakoid lumen. Jenkins et al. (1990) found evidence for at least three subunits in Cryptomonas sp. strain CS-24, and a gene encoding one of these was cloned and sequenced. The mature protein is predicted to be 76 amino acids in length and the gene additionally predicts the presence of a presequence of 52 amino acids. The targeting sequence appears to be a composite sequence with both a chloroplast targeting sequence and a lumenal targeting sequence; the presequence differs from the targeting sequence found for the stromally located subunit of B-PE encoded in the nuclear genome of Aglaothamnion neglectum (Apt et al., 1993). When the presequence is compared to presequences of thylakoid-lumen targeting sequences encoded in cyanobacterial, chloroplast, or eucaryotic-nuclear genomes, it seems likely that the gene encoding the subunit occurs in the nucleomorph genome (the remnant of the red algal nucleus) of the organism.

D. Genetic Analysis of the Elements of the PBS for Mastigocladus laminosus Genetic analyses ofthe genes encoding the structural components of PBS for Mastigodadus laminosus (Fig. 27) were carried out in three stages. Firstly, the genes encoding PEC and PC were identified and cloned using a hybridization probe encoding part of the subunit from Synechococcus sp. strain PCC 7002. The 5' part of the pecBACEF operon and the cpcBACDEF genes were cloned, mapped, and sequenced (Eberlein and Kufer, 1990; Kufer et al., 1992). Secondly, a PstI DNA library Mastigocladus laminosus was probed with a DNA fragment encoding the cpcG1 and cpcG2 genes of Anabaena sp. PCC 7120 (Bryant et al., 1991). A 4.5-kbp Pst1 fragment containing the 3' portion of the cpc operon, including the genes 5' cpcF-cpcG1-cpcG2-cpcG was cloned, mapped and sequenced (Glauser et al., 1992b). The fourth cpcG gene was not present in this 4.5-kbp PstI

Walter A. Sidler fragment, as confirmed by sequencing of the complete fragment. Although the fourth cpcG4 gene has not yet been cloned, Southern-blot hybridizations suggest the existence of this gene that is split into two parts on adjacent Pst Ifragments 8–9-kbp in size. Thirdly, the genes encoding all six PBS core proteins from Mastigocladus laminosus were cloned and sequenced (Esteban, 1993; Esteban et al., 1994). Three apc loci (see Fig. 27) were identified by probing Mastigocladus laminosus chromosomal DNA with different probes specific for the apcAB, apcE, apcD, and apcF from Synechococcus sp. strain PCC 7002 (Bryant, 1988, 1991) or for the apcD gene of Synechococcus sp. strain PCC 6301. The derived amino acid sequences of and are in complete agreement with the sequences determined by Edman degradation by Sidler et al. (1981).

E. The pec and cpc Operons of Mastigocladus laminosus The pec and cpc loci of Mastigocladus laminosus together form a rather large transcription unit compared to corresponding cpc operons of many other organisms (Bryant 1991). The pec operon contains five genes, pecBACEF, and the cpc operon consists of nine genes: cpcBACDEFG1G2G3. Nearly the same organization was found in Anabaena sp. strain PCC 7120, in which a fourth cpcG gene (cpcG4) occurs downstream from the cpcG3 gene (Belknap and Haselkorn, 1987; Bryant etal., 1991). Moreover, the pecBACEF genes of this organism are also found immediately upstream from the cpc operon (Swanson et al., 1992a). Indeed the pec and cpc operons together may be denoted as the ‘rod’ operons, because they include all genes necessary to encode the structural elements for the PBS rods of Mastigocladus laminosus (Fig. 27). The 5' end begins with the pecBACEF gene cluster, encoding the and two proteins (PecE and PecF) which are not structural elements of the rods but are probably involved in chromophore attachment to the subunit. The proteins encoded by the pecE and pecF genes are homologs of the CpcE and CpcF proteins that together form the phycocyanobilin lyase, and hence it is probable that the PecE and PecF proteins form the enzyme that attaches PXB to the subunit. The cpc gene cluster is separated by a 622 bp region containing the promoter for the PC encoding genes (Eberlein and Kufer, 1990; Kufer et

Chapter 7 Phycobilisome and Phycobiliprotein Structures al., 1992; W. Kufer, personal communication). This pec gene cluster is proposed to be a gene duplication of the downstream and adjacent cpcBACDEF gene cluster. The cpcD gene product encodes the unique rod-terminating linker protein This gene apparently did not undergo the gene duplication, and thus it may be assumed that the functions as rod-terminating linker polypeptide for rods ending either with PEC in low-light-grown cells or with PC in high-light-grown cells. A multigene family of cpcG genes occurs downstream from the cpcF gene. This gene cluster includes three of the four proposed cpcG genes that encode different rodcore linker polypeptides of the PBS from Mastigocladus laminosus (Glauser et al., 1990, 1992b). The gene encoding the postulated fourth rod-core linker polypeptide in Mastigocladus laminosus has not yet been isolated.

F. The apc Operons in the Genome of Mastigocladus laminosus The organization of the apc transcription units is presented in Fig. 27. The six genes encoding the core elements and are distributed in the genome of Mastigocladus laminosus on three transcriptional units. The first locus, including the apcEABC genes, encodes the essential core elements. In this operon, apcE encodes the the large, multifunctional linker phycobiliprotein. The apcA, apcB, and apcC genes encode and respectively. The apcD and apcF genes, encoding the specialized, minor core subunits and phycobiliprotein, respectively lie adjacent to ORFs that encode unknown proteins. Cells require much more of the APC and subunits than to assemble PBS. The molecular mechanisms which regulate the complex transcription of the apcEABC loci in various cyanobacteria are not yet understood (Houmard et al., 1986, 1990; Capuano et al., 1991; Bryant, 1991).

Acknowledgments I am grateful to D. A. Bryant for improving this chapter with changes and informative additions. I also thank Axel Ducret, Andreas Engel, Shirley Müller, and Ralf Ficner for supplying figures. The Swiss National Foundation provided funds that made our research in this field possible. This work is part

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of the habilitation of W. S. at the Eidgenössischen Technischen Hochschule, Zürich.

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Walter A. Sidler laminosus. J Photochem Photobiol B: Biol 18: 51–66 Reuter W and Wehrmeyer W (1988) Core substructure in Mastigocladus laminosus phycobilisomes: I Microheterogeneity in two of three allophycocyanin core complexes. Arch Microbiol 150: 534–540 Reuter W and Wehrmeyer W (1990) Core substructure in Mastigocladus laminosus phycobilisomes: II The central part of the tricylindrical core - contains the anchor polypeptide and no allophycocyanin B. Arch Microbiol 153: 111–117 Reuter W, Nickel C and Wehrmeyer W (1990) Isolation of allophycocyanin B from Rhodella violacea results in a model of the core from hemidiscoidal phycobilisomes of Rhodophyceae. FEBS Lett 273: 155–158 Rhiel E, Mörschel E and Wehrmeyer W (1987) Characterization and structural analysis of a chlorophyll a/c light harvesting complex and of Photosystem I particles isolated from thylakoid membranes of Cryptomonas maculata (Cryptophyceae). Eur J Cell Biol 43: 82–92 Rhiel E, Kunz J and Wehrmeyer W (1989) Immunocytochemical localization of phycoerythrin-545 and of a chlorophyll a/c light harvesting complex in Cryptomonas maculata (Cryptophyceae. Botanica Acta 102: 46–53 Riethman HC, Mawhinney TP and Sherman LA (1987) Phycobilisome-associated glycoproteins in the cyanobacterium Anacystis nidulans R2. FEBS Lett 215: 209–214 Riethman HC, Mawhinney TP and Sherman LA (1988) Characterization of phycobilisome glycoproteins in the cyanobacterium Anacystis nidulans R2. J Bacteriol 170: 2433– 2440 Rippka R and Herdman M (1985) Division patterns and cellular differentiation in cyanobacteria. Ann Microbiol (Inst Pasteur) 136A 33–39 Roell MK and Morse DE (1993) Organization, expression and nucleotide sequence of the operon encoding R-phycoerythrin and subunits from the red alga Polysiphonia boldii. Plant Mol Biol 21: 47–58. Rosinski J, Hainfeld JF, Rigbi M, and Siegelman HW (1981) Phycobilisome ultrastructure and chromatic adaptation in Fremyella diplosiphon . Ann Bot 47: 1–12 Rüdiger W (1975) Phycobiliproteide. Ber Deutsch Bot Ges 88: 125–139 Rüdiger W (1980) Plant Biliproteins In: FC Czygan (ed) Pigments in Plants 2nd ed, pp 314–351. G Fischer, Stuttgart Rüdiger W (1994) Phycobiliproteins and phycobilins. Progress in Phycological Research, in press Rüegsegger U, Sidler WA, Esteban A, Betz M and Zuber H (1993) Reconstitution of the rod-core complex using the overexpressed rod-core linker polypeptide from the cyanobacterium Mastigocladus laminosus. Eur J Biochem, submitted Rümbeli R and H Zuber (1988) Isolation and characterization of the components of the phycobilisome from Mastigocladus laminosus and crosslinking experiments In: Scheer H and Schneider S (ed) Photosynthetic Light-Harvesting Systems, pp 61–70. Walter de Gruyter, Berlin Rümbeli R, Schirmer T, Bode W, Sidler W and Zuber H (1985) Crystallization of phycoerythrocyanin from the cyanobacterium Mastigocladus laminosus and preliminary characterization of two crystal forms. J Mol Biol 186: 197–200

Chapter 7 Phycobilisome and Phycobiliprotein Structures Rümbeli R, Wirth M, Suter F and Zuber H (1987a) The phycobiliprotein of the allophycocyanin core from the cyanobacterium Mastigocladus laminosus. Characterization and complete amino acid sequence. Biol Chem Hoppe-Seyler 368: 1–9 Rümbeli R, Suter F, Wirth M, Sidler W and Zuber H (1987b) Isolation and localization of in phycobiliproteins from the cyanobacterium Mastigocladus laminosus. Biol Chem Hoppe-Seyler 368: 1401–1406 Rümbeli R, Suter F, Wirth M, Sidler W and Zuber-H (1987c) N-Methylasparagine in phycobiliproteins from the cyanobacteria Mastigocladus laminosus and Calothrix. FEBS Lett 221: 1–2 Rümbeli R, Frank G, Wirth M and Zuber H (1988) Isolation and partial amino acid sequences of the 89 kDa-anchorpolypeptide of phycobil isomes from Mastigocladus laminosus. Experientia 44: A60 Sandström Å, GillbroT, Sundström V, Wendler J and Holzwarth AR (1988a) Picosecond study of energy transfer within 18-S particles of AN 112 (a mutant of Synechococcus 6301) phycobilisomes. Biochim Biophys Acta 933: 54–64 Sandström Å, Gillbro T, Sundström V, Fischer R and Holzwarth AR (1988b) Picosecond time-resolved energy transfer within C-phycocyanin aggregates of Mastigocladus laminosus. Biochim Biophys Acta 933: 42–53 Sauer K and Scheer H (1988) Excitation transfer in Cphycocyanin. Förster transfer rate and exciton calculations based on new crystal structure data for C-phycocyanins for Agmenellum quadruplicatum and Mastigocladus laminosus. Biochim Biophys Acta 936: 157–170 Scharnagel C and Fischer S (1993) Reversible photochemistry in the of phycoerythrocyanin: characterization of chromophore and protein by molecular dynamics and quantum chemical calculations. Photochem Photobiol 57: 63–70 Scheer H (1981) Biliproteine. Angew Chem. 93: 230–250 Scheer H (1982) Phycobiliproteins: molecular aspects of photosynthetic antenna system In: FK Fong (ed) Light Reaction Path of Photosynthesis. Mol Biol Biochem Biophys 35: 7–45 Scheer H (1986) Excitation transfer in phycobiliproteins In: Staehelin LA and Arntzen CJ (eds) Photosynthesis III. Encyclopedia of Plant Physiology New Series, Vol 19, pp 372–336. Springer-Verlag, Berlin Schirmer T and Vincent M (1987) Polarized absorption and fluorescence spectra of single crystals of C-phycocyanin. Biochim Biophys Acta 893: 379–385 Schirmer T, Bode W, Huber R, Sidler W and Zuber H(1985) Xray crystallographic structure of the light-harvesting biliprotein C-phycocyanin from the thermophilic cyanobacterium Mastigocladus laminosus and its resemblance to globin structures. J Mol Biol 184: 257–277 Schirmer T, Huber R, Schneider M, Bode W, Miller M and Hackert ML (1986) Crystal structure analysis and refinement at 2.5 Å of hexameric C-phycocyanin from the cyanobacterium Agmenellum quadruplicatum. The molecular model and its implications for light-harvesting. J Mol Biol 188: 651–676 Schirmer T, Bode W and Huber R (1987) Refined threedimensional structures of two cyanobacterial C-phycocyanins at 2.1 and 2.5 Å resolution. A common principle of phycobilinprotein interaction. J Mol Biol 196: 677–695 Schluchter WM and Bryant DA (1992) Molecular characterization

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214 Photosynthesis, Vol II, pp 497–500. Kluwer, Dordrecht Sidler W, Rüegsegger U and Zuber H (1993) Overexpression of linker polypeptide from M. laminosus and reconstitution of the rod-core complex Experientia 49: A33/189 Siebzehnrübel S, Fischer R and Scheer H (1987) Chromophore assignment in C-phycocyanin from Mastigocladus laminosus Z. Naturforsch. 42c: 258–262. Siebzehnrübel S, Fischer R, Kufer W and Scheer H (1989) Photochemistry of phycobiliproteins: reciprocity of reversible photochemistry and aggregation in phycoerythrocyanin from Mastigocladus laminosus. Photochem Photobiol 49: 753–761 Staehelin LA (1986) Chloroplast structure and supramolecular organization of photosynthetic membranes In: Staehelin LA and Arntzen CJ (eds) Photosynthesis III. Encyclopedia of Plant Physiology New Series, Vol 19, pp 1–84. SpringerVerlag, Berlin Staehelin LA, Giddings TH, Badami P and Krzymowski WW (1978) In: Deamer DW (ed) Light Transducing Membranes: Structure, Function and Evolution, pp 335–355. Academic Press, New York Staehelin LA, Golecki JR and Drews G (1980) Supramolecular organisation of chlorosomes (Chlorobium vesicles) and their membrane attachment sites in Chlorobium limicola. Biochim Biophys Acta 589: 30– 45 Stanier RY (1977) The position of cyanobacteria in the world of phototrophs. Carlsberg Res Commun 42: 77–98 Stanier RY and Cohen-Bazire G (1977) Phototrophic prokaryotes: The cyanobacteria. Annu Rev Microbiol 31: 225–274 Stevens SE Jr and Bryant DA (1988) Light-Energy Transduction in Photosynthesis: Higher Plant and Bacterial Models. American Society of Plant Physiologists, Rockville MD Su X, Fraenkel PG and Bogorad L (1992) Excitation energy transfer from phycocyanin to chlorophyll in an apcA deficient mutant of Synechocystis sp. PCC 6803. J Biol Chem 267: 22944–22950 Suter F, Füglistaller P, Lundell DJ, Glazer AN and Zuber H (1987) Amino acid sequences of alpha-allophycocyanin B from Synechococcus 6301 and Mastigocladus laminosus. FEBS Lett 217: 279–282 Suter GW and Holzwarth AR (1987) A kinetic model for the energy transfer in phycobilisomers. Biophys J 52: 673–683 Swanson RV and Glazer AN (1990a) Phycobiliprotein methylation Effect of residue on energy transfer in phycocyanin and the phycobilisome. J Mol Biol 214: 787–796 Swanson RV and Glazer AN (1990b) Separation of phycobiliprotein subunits by reverse-phase high pressure chromatography. Analyt Biochem 188: 295–299 Swanson RV, Ong LJ, Wilbanks SM and Glazer AN (1991) Phycoerythrins of marine unicellular cyanobacteria. II. Characterization of phycobiliproteins with unusually high phycourobilin content. J Biol Chem 266: 9528–9534 Swanson RV, de Lorimier R and Glazer AN (1992a) Genes encoding the phycobilisome rod substructures are clustered on the Anabaena chromosome. Characterization of the phycoerythrocyanin operon. J Bacteriol 174: 2640–2647 Swanson RV, Zhou J, Leary JA, Williams T, de Lorimier R, Bryant DA and Glazer AN (1992b) Characterization of phycocyanin produced by cpcE and cpcF mutants and

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216 bacteria, pp 53–55. Plenum Press, New York Zuber H and Brunisholz R A (1991) Structure and function of antenna polypeptides and chlorophyll-protein complexes: principles and variability In: Scheer H (ed) Chlorophylls, pp 627–703. CRC Press, Boca Raton Zuber H, Sidler W, Füglistaller P, Brunisholz R and Theiler R (1985) Structural studies on the light-harvesting polypeptides from cyanobacteria and bacteria. In: Steinback KE, Bonitz S,

Walter A. Sidler Arntzen CJ, and Bogorad L (eds) Molecular Biology of the Photosynthetic Apparatus, pp 183–195. Cold Spring Laboratory Publications, Cold Spring Harbor, NY Zuber H, Brunisholz R and Sidler W (1987) Structure and function of light-harvesting pigment-protein complexes. In: Amesz J (ed) Photosynthesis (New Comprehensive Biochemistry, Vol 15), pp 233–271. Elsevier Science Publishers, Amsterdam

Chapter 8 The Use of Cyanobacteria in the Study of the Structure and Function of Photosystem II Bridgette A. Barry and Renee J. Boerner Department of Biochemistry, University of Minnesota, St. Paul, MN 55108, USA

Julio C. de Paula Department of Chemistry, Haverford College, Haverford, PA 19041, USA

Summary I. Introduction II. A Comparison of the Biochemical Properties of Cyanobacterial and Higher Plant Photosystem II A. The 33-kDa (PsbO) Extrinsic Protein B. The 43 Da (PsbC) Protein C. The 24-kDa and 18-kDa Extrinsic Proteins are not Present in Cyanobacteria D. Small Polypeptides in Plant and Cyanobacterial Photosystem II 1. Photosystem II Preparations from Plants 2. Photosystem II Preparations from Cyanobacteria 3. Low-Molecular-Weight Polypeptides in Reaction Center Complexes 4. Low-Molecular-Weight Polypeptides in Core Photosystem II Particles 5. Low-Molecular-Weight Polypeptides in Plant Photosystem II Membranes and Cyanobacterial Photosystem II Particles E. Cytochrome III. Site-Directed Mutagenesis Studies of the Donor Side of Photosystem II A. A Search for the Ligands to the Manganese Cluster 1. Mutations in the D1 Polypeptide (psbA Gene Product) 2. Mutations in the D2 Polypeptide (psbD Gene Product) 3. Mutations in the 47-kDa Protein (psbB Gene Product) B. The Location of the Redox Active Tyrosines IV. Biophysical Studies of Cyanobacterial Photosystem II A. Tyrosine Radical, has a Slightly Different EPR Lineshape in Plants and Cyanobacteria B. Difference FT-IR Studies of the Redox Active Tyrosine Residues in Photosystem II C. The Structure of the Manganese Complex in Plants and Cyanobacteria 1. Magnetic Resonance Studies of the State a. Electron Paramagnetic Resonance b. Electron Spin-Echo Envelope Modulation c. Electron-Nuclear Double Resonance 2. X-Ray Absorption Studies of the S States V. Concluding Remarks References

D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 217–257. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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Summary Oxygenic photosynthesis occurs in plants, green algae, and procaryotic cyanobacteria. Two chlorophyllcontaining photosystems cooperate to transfer electrons from water to Photosystem II is the membrane protein complex that carries out the light-catalyzed oxidation of water and reduction of plastoquinone. The reaction center is composed ofboth intrinsic and extrinsic proteins; the prosthetic groups involved in electron transfer include chlorophyll, pheophytin, quinone, tyrosine residues, and a manganese cluster. Cyanobacteria have emerged as a convenient system with which to study the structure and function of Photosystem II for two reasons. Firstly, isotopic labeling experiments are possible in this organism, facilitating many types of biophysical experiments. Secondly, site-directed mutagenesis is easily performed. This chapter will review what is known about the structure and function of Photosystem II with particular emphasis on the use of cyanobacteria in such studies. Areas in which there are significant differences between plants and cyanobacteria will be highlighted. I. Introduction In plants, green algae, and cyanobacteria, two chlorophyll-containing membrane protein complexes cooperate to transfer electrons from water to Light energy is used to drive this process. Photosystem II (PS II) is the reaction center that carries out the light-catalyzed oxidation ofwater and reduction of bound quinone (Fig. 1). The primary chlorophyll donor of PS II is called After light absorption, an electron is transferred from an excited state of to a pheophytin, which in turn reduces a bound plastoquinone molecule, called reduces a second quinone, which, unlike canfunction as a two electron acceptor. On the donor side of PS II, the chlorophyll cation radical, oxidizes a tyrosine residue, Z, which in turn oxidizes a cluster of four manganese atoms. This metal center is the catalytic site of water oxidation, and the cluster is able to accumulate the four oxidizing equivalents that are necessary in order to release from two molecules of water. The five sequentially oxidized forms of the cluster are called S states. is stable upon long-term dark adaptation, and is unstable Abbreviations: Chl – chlorophyll; DCMU – 3-(3,4-dichlorophenyl)-1,1-dimethylurea; DCPIP – 2,6-dichlorophenolindophenol; DPC – 1,5-diphenylcarbazide; – concentration at which 50% of activity is inhibited, relative to a control untreated sample; EPR – electron paramagnetic resonance; ESEEM – electron spin-echo envelope modulation; ENDOR – electronnuclear double resonance; EXAFS – extended X-ray absorption fine structure; FT-IR – Fourier transform infrared; NMR – nuclear magnetic resonance; PS I – Photosystem 1; PS II – Photosystem II; PAGE – polyacrylamide gel electrophoresis; RDPR – ribonucleoside diphosphate reductase; SDS – sodium dodecyl sulfate; XANES – X-ray absorption near-edge structure.

and spontaneously converts to thereupon releasing molecular oxygen. The reaction center also contains a nonheme iron atom and a stable tyrosine radical, PS II is made up of both integral membrane proteins and extrinsic proteins. Polypeptides that are required for oxygen evolution in both plants and cyanobacteria under physiological conditions are: the 47-kDa (CP47), the 43-kDa (CP43), D1, D2, the 33-kDa extrinsic protein, and two subunits, and of a cytochrome (Table 1). Plants also contain two extrinsic proteins of 24 and 18 kDa. These polypeptides sequester the calcium and chloride ions that are required for catalysis. Cyanobacterial PS II does not contain these two subunits, but has two different extrinsic proteins: a 9-kDa (or 12-kDa) protein and a cytochrome Both plant and Cyanobacterial PS II also contain small polypeptides kDa); the role of these low molecular weight subunits in water oxidation is often not known. In both plants and cyanobacteria, the hydrophobic polypeptides, D1 and D2, bind most ofthe prosthetic groups that are involved in electron transfer (Nanba and Satoh, 1987; Gounaris et al., 1989). There are regions of sequence homology between D1 and D2 and the L and M subunits of the reaction center from purple, nonsulfur bacteria (Deisenhofer and Michel, 1989). There are also functional similarities between acceptor side electron transfer in the two systems. In analogy with the role of L and M in the bacterial reaction center, it is widely accepted that D1 and D2 form the heterodimer core of the PS II reaction center. However, there is very little structural information about the donor side of PS II, since the bacterial reaction center, which does not oxidize water, does not provide a direct structural model (but

Chapter 8 Photosystem II

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of the properties of plant and cyanobacterial PS II. Insights derived from site-directed mutagenesis studies of the donor side of PS II will also be considered. We apologize to all those whose work we do not have space to mention.

II. A Comparison of the Biochemical Properties of Cyanobacterial and Higher Plant Photosystem II

A. The 33-kDa (PsbO) Extrinsic Protein see Svensson et al., 1990,1991; Ruffle et al., 1992). Many excellent reviews of PS II and of water oxidation have appeared in the last few years (for example, see Diner et al., 1991; Yocum, 1991; Debus, 1992; Erickson and Rochaix, 1992; Pakrasi and Vermaas, 1992; Rutherford et al., 1992; Vermaas, 1993). This review will concentrate on a comparison

The 33-kDa extrinsic protein ormanganese stabilizing protein (MSP) plays an intriguing role in PS II. The 33-kDa protein influences the properties of the manganese catalytic site. However, the exact mechanism by which these effects are mediated is not known and has been the subject of some controversy. In addition, there is some indication that the 33-kDa protein may play different roles in

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plants and cyanobacteria. This is possible since there are substantial differences in sequence between the typical cyanobacterial protein and the typical plant 33-kDaprotein (Erickson and Rochaix, 1992). In the next section, we will discuss the properties and role of the 33-kDa protein in plants and cyanobacteria. The protein sequence of the spinach 33-kDa protein has been obtained (Oh-oka et al., 1986). The gene (psbO) sequence has been obtained from higher plants, green algae, and cyanobacteria (for review, see Erickson and Rochaix, 1992). In plants and green algae, the psbO gene is encoded in the nucleus. The PsbO protein is synthesized with a transit sequence (or a leader peptide in the case ofcyanobacteria) that targets the polypeptide to the thylakoid lumen (for review, see Erickson and Rochaix, 1992). The genes encoding the 33-kDa proteins from spinach (Seidler and Michel, 1990), Arabidopsis thaliana (S. Betts, E. Pichersky, C. F. Yocum, unpublished results), wheat (Meadows and Robinson, 1991), and the cyanobacterium, Anabaena sp. strain PCC 7120 (Borthakur and Haselkorn, 1989), have been expressed in E. coli. The recombinant proteins from spinach (Seidler and Michel, 1990) and A. thaliana (S. Betts, E. Pichersky, C. F. Yocum, unpublished results) have been shown to be effective in rebinding to the spinach PS II reaction center. The spinach 33-kDa protein can be removed from the reaction center in vitro by several different types of washing procedures, including washes with alkaline Tris (Åkerlund and Jansson, 1981; Yamomoto et al., 1981), (Ono and Inoue, 1983), and urea (Miyao and Murata, 1984b). Once removed, it behaves as a soluble protein and can be rebound to PS II membranes (Miyao and Murata, 1983b). The stoichiometry of the 33-kDa polypeptide per plant reaction center has been reported to be either one (Murata et al., 1984; Miyao and Murata, 1989; Enami et al., 1991) or two (Andersson et al., 1984; Millner et al., 1987) on the basis ofCoomassie staining and crosslinking studies. Recent measurements using immunological detection indicate that the stoichiometry is two per tetrameric manganese cluster (Xu and Bricker, 1993). The dissociation constant has been reported to be 12 nM in the presence of the manganese cluster. Measurement of the binding constants for cyanobacterial systems is also of interest, since, in a purified PS II preparation from the thermophile Phormidium laminosum it has been reported that the 33-kDa protein cannot be removed by alkaline Tris-

washing (Stewart et al., 1985a). On the other hand, it has been reported that the 33-kDa protein can be removed from PS II particles of the thermophilic cyanobacterium Synechococcus vulcanus Copeland by washing (Koike and Inoue, 1985). Also, it has been reported that Tris-washing does not remove the 33-kDa polypeptide from thylakoid membranes of Synechocystis sp. strain PCC 6803 (Nilsson et al., 1990), although Tris-washing can remove the 33kDa protein from purified Synechocystis PS II particles (Noren et al., 1991). Interestingly, recent studies on the chlorophyll a/b-containing prochlorophyte, Prochlorothrix hollandica, show that in this organism the ‘33-kDa’ protein has an apparent molecular mass of 37 kDa and is hydrophobic, as assessed by phase-partitioning experiments (Mor et al., 1993). In the absence of the manganese cluster, the affinity of the spinach reaction center for the spinach 33-kDa has been reported to be lower, with a dissociation constant of 88 nM (Miyao and Murata, 1989). However, these experiments also predict a stoichiometry ofone per reaction center. Other experiments also suggest that loss of manganese lowers the affinity of both the plant and cyanobacterial reaction center for the 33-kDa protein (Ghanotakis et al., 1984c; Kavelaki and Ghanotakis, 1991; Boerner et al., 1992; Noren and Barry, 1992). The spinach 33-kDa protein has two cysteines that form adisulfide linkage, andthis disulfide isessential in maintaining the folded form of the protein. When the linkage is reduced, the protein is unable to rebind to the reaction center (Tanaka and Wada, 1988). Since sequence analysis shows that these two cysteines are conserved, cyanobacterial 33-kDa proteins are also likely to have a structurally important disulfide linkage (Philbrick and Zilinskas, 1988). The amino-terminal end of the spinach 33-kDa polypeptide has been identified as essential for rebinding to the reaction center through experiments in which the amino terminus is removed by protease treatment (Eaton-Rye and Murata, 1989). The aminoterminal end of the spinach 33-kDa protein has also been implicated as essential in rebinding by crosslinking experiments (Odom and Bricker, 1992). The binding site for the 33-kDa protein on the reaction center has not been precisely defined. Crosslinking experiments have been performed to address this question. In plants it has been reported that the 33-kDa protein can be crosslinked to PS II polypeptides of 22, 24, 26, 28, 29 and 31 kDa

Chapter 8 Photosystem II (Bowlby and Frasch, 1986). This complex was also found to contain 3–4 mol of Mn/mol protein. The 33kDa polypeptide has been crosslinked to the 47-kDa protein (Enami et al, 1987; Bricker et al, 1988; Enami et al., 1989b; Odom and Bricker, 1992)and to D1 and D2 (Mei et al., 1989). The latter crosslinked complex has been reported to contain manganese and to be active in oxidation of (Mei et al., 1989). Rebinding of the 33-kDa protein to purified (reaction center) particles has also been described (Gounaris et al., 1988). Recently, the 33kDa protein has been crosslinked to a subunit of cytochrome and to the 4.8-kDa product of the psbI gene (Enami et al., 1992). Immunoprecipitation has been used to assess nearest neighbor interactions in plants. These data suggest an interaction between a 24-kDa protein and the 33-kDa protein (Ljungberg et al., 1984). It has also been shown that, when bound to the reaction center, the 33-kDa polypeptide protects both the 43kDa (Isogai et al., 1985) and the 47-kDa (Bricker and Frankel, 1987) proteins against proteolytic attack. The 33-kDa protein prevents binding of a monoclonal antibody that recognizes an epitope on the 47-kDa protein (Bricker and Frankel, 1987; Bricker et al., 1988; Frankel and Bricker, 1992). A site for biotin labeling on the 47-kDa is also shielded by the binding of the 33-kDa protein to the reaction center (Bricker et al., 1988; Frankel and Bricker, 1992). One conclusion from the work described above is that the plant 33-kDa protein associates closely with the 47-kDa protein. This interaction may be through salt-bridges and involves a large extrinsic loop of the 47-kDa polypeptide (Frankel and Bricker, 1992; Odom and Bricker, 1992). This loop is predicted to connect hydrophobic helix V and helix VI (Bricker, 1990) (Fig. 2). However, the experiments described above indicate that the 33-kDa protein may also interact with other hydrophobic PS II proteins, and the binding site may be quite complex. Little is known about the 33-kDa protein binding site in cyanobacteria. The 33-kDa protein has been reported to be essential for oxygen evolution in green algae (Mayfield et al., 1987). However, recent evidence has shown that cyanobacteria in which the psbO gene has been deleted are still active in oxygen evolution. These mutant strains have been found to have a lower steady-state rate of oxygen evolution than control cells (Burnap et al., 1989; Bockholt et al., 1991; Burnap and Sherman, 1991; Mayes et al.,

221 1991; Philbrick et al., 1991; Burnap et al., 1992; Vass et al., 1992). The psbO deletion strains show a greater susceptibility to photoinhibition and a requirement for calcium in the growth media (Mayes et al., 1991; Philbrick et al., 1991). These results may imply that the PsbO plays substantially different roles in higher plants and cyanobacteria. The role of the 33-kDa protein in vitro has also been controversial. Early biochemical experiments using plant PS II preparations showed that low levels ofoxygen evolution could be observed after removal of the PsbO protein, although the manganese cluster was destabilized (Miyao and Murata, 1984b; Ono and Inoue, 1984). In low-chloride media it was observed that half of the bound manganese was lost. In the presence of high concentrations of chloride, manganese wasretainedbythereactioncenter (Miyao and Murata, 1984b). These results are vulnerable to the criticism that the low rates of oxygen evolution might arise from a small number of centers that retain the 33-kDa protein. Recent experiments using immunological detection have shown that it is possible to remove ninety-nine percent of the 33kDa protein by or NaCl-urea washes (Bricker, 1992). In these samples, the oxygen evolution rate was approximately twenty percent of the control rate, but was dependent on the presence of high concentrations of calcium and chloride in the media. Reconstitution experiments have been performed to address the question of whether the cyanobacterial and plant 33-kDa extrinsic proteins are interchangeable (Koike and Inoue, 1985). PS II particles were isolated from the thermophilic cyanobacterium S. vulcanus Copeland; PS II membranes from spinach were employed (see Section II D, 1 and 2 for review of biochemical preparations from plants and cyanobacteria). The 33-kDa proteins were removed from the reaction centers by washing with and this crude supernatant was used forreconstitution experiments without furtherpurification. In this way, the plant 33-kDa protein was rebound to the cyanobacterial reaction center, and, in turn, the cyanobacterial 33-kDa was rebound to the spinach reaction center. In these experiments, very low rates of oxygen evolution were observed from either type of preparation in the absence of the 33-kDa protein. The cyanobacterial 33-kDa protein could substitute for the spinach 33-kDa protein and restore twentyeight percent of control oxygen evolution rates to spinach reaction centers at 23 °C. The spinacfh 33kDa protein restored sixty percent of control rates to

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cyanobacterial reaction centers at 23 °C. However, thermophilic preparations show maximal rates at elevated temperatures, and the preparation reconstituted with the spinach 33-kDa protein showed no activity under these conditions. Also, restoration of activity was not strictly proportional to the amount of spinach 33-kDa protein rebound, when assayed by SDS-denaturing gel electrophoresis. Taken at face value, these experiments suggest that the binding determinants in the plant and the cyanobacterial 33kDa protein are similar. Determination of a binding constant through the use of purified proteins would be of use in evaluating these experiments. The 33-kDa protein seems to affect the manganese cluster, since, when it is removed,the and states are more stable. This is observed both in a cyanobacterium in which the psbO gene has been deleted (Burnap et al., 1992; Vass et al., 1992) and after biochemical removal of the 33-kDa protein from spinach preparations (Miyao et al., 1987; Vass et al., 1987). In either system, loss of PsbO is also

associated with a retardation or inhibition of the transition (Ono and Inoue, 1985; Miyao et al., 1987; Burnap et al., 1992; Vass et al., 1992). These studies are evidence for a close association of the manganese cluster and the 33-kDa protein in both plants and cyanobacteria. It has been suggested that the 33-kDa protein may provide amino acid residues as ligands to the manganese. Under oxidizing conditions, the isolated spinach 33-kDa was found to contain bound manganese (Abramowicz and Dismukes, 1984; Yamomoto et al., 1984). A sequence similarity to a manganese-binding region of superoxide dismutase was also noted for the spinach protein (Oh-oka et al., 1986). Comparison of additional sequences has indicated, however, that this region is not strictly conserved in all species (Philbrick and Zilinskas, 1988). Moreover, it is now clear that low levels of oxygen evolution are possible in the absence of the 33-kDa protein (Bricker, 1992). Therefore, if the 33kDa protein provides ligands to the cluster, they

Chapter 8 Photosystem II must be replaceable or non-essential. It has been reported that an EPR signal from the state of the manganese cluster (‘multiline signal,’ see Section IV C, 1a) can be generated after removal of the 33kDa protein from plant preparations (Miller et al., 1987; Styring et al., 1987), although there has been controversy over this issue (for example, see Hunziker et al., 1987). There is also disagreement overwhether the multiline signal, when observed, is significantly altered by removal of the 33-kDa protein (Miller et al., 1987; Styring et al., 1987). An X-ray absorption study ofa PsbO-depleted, but manganese containing, PS II preparation showed a manganese K-edge spectrum from the dark-adapted state that was very similar to the spectrum of a control sample (Cole et al., 1987). These spectroscopic results have been used to argue against the idea that the PsbO protein provides ligands to the cluster (Cole et al., 1987; Miller et al., 1987). It has been suggested that PsbO may be a calciumbinding protein (Wales et al., 1989; Burnap et al., 1990). This suggestion comes from sequence similarities between the 33-kDa protein and an intestinal calcium-binding protein (for a review of similarities with other calcium-binding proteins, see Yocum, 1991). Thereislittlequantitativeinformation about the affinity (or lack thereof) ofthe isolated 33kDa for calcium (Yocum, 1991). A weak affinity of a 32-kDa protein for has been noticed after SDS polyacrylamide gel electrophoresis and transfer to nitrocellulose (Webber and Gray, 1989a). However, this polypeptide was never positively identified as the manganese stabilizing protein. The affinity and number ofcalcium-binding sites in PS II reaction centers is still controversial (Yocum, 1991; Debus, 1992).

B. The 43 Da (PsbC) Protein The binding site may be different in plants and cyanobacteria. The indirect evidence to support this idea comes from studies of the role of the 43-kDa chlorophyll-binding protein. The gene for the 43kDa or CP43 protein (psbC gene product) is chloroplast-encoded in plants. Plant and cyanobacterial proteins are predicted to be approximately 77% homologous (Erickson and Rochaix, 1992). To determine the role of this protein in PS II, the psbC gene has been deleted in Synechocystis sp. strain PCC6803, and the effect of this mutation has been

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assessed. This mutant does not evolve oxygen, and the content of PS II proteins was found to be reduced dramatically in intact cells of this deletion mutant. As expected, no 43-kDa protein was detectable by immunoblot analysis. Optical measurements of electron transfer were performed. These studies showed that assembled PS II reaction centers were active in reduction of and oxidation of Z; the chlorophyll antenna size was approximately 30 chlorophylls per In this work, there was no quantitation of the content of bound plastoquinone per reaction center, and EPR studies ofthe lineshape of and in a purified preparation were not performed (Rögner et al., 1991). Biochemical studies of the role of the 43-kDa protein sometimes give a different result. For example, spinach preparations can be depleted of the 43kDa protein through treatment with the nonionic detergent, dodecyl maltoside, and ion exchange chromatography(Petersen et al., 1990). These spinach PS II preparations contain the 47-kDa protein, D1, D2, and cytochrome and they contain approximately thirty chlorophylls per reaction center. Reaction center antenna size was quantitated on the assumption that there are two cytochrome per reaction center (see Section II E). In these 43-kDa depleted samples, the content of plastoquinone was found to be reduced to only 0.15–0.20 per two cytochrome EPR measurements on this preparation showed that a tyrosine radical with a normal lineshape (Section IV A) was generated in the light in the majority ofcenters, but only when an exogenous acceptor was present. In the absence of acceptors, a narrower, (10 G) structureless signal was observed in the light. This signal was ascribed to a reduced pheophytin molecule. On the other hand, a second type of spinach PS II preparation containing CP47, D1, and D2 was isolated through the use of digitonin polyacrylamide gel electrophoresis after treatment with the chaotropic agent potassium thiocyanate. The antenna size of this preparation was approximately thirty chlorophylls per reaction center as assessedby the amplitude of photoreducible Plastoquinone quantitation via extraction showed 1.8 quinones per reaction center, in contrast to results of Petersen et al. (1990). The optical difference spectrum of Z could be observed in this preparation. The EPR signal of was reported to be present in this preparation when DCPIP was used as an acceptor, but with a modified,

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narrow lineshape (Yamaguchi et al., 1988). There was no spin quantitation of the signal. It is difficult to assess the suggestion that the narrow signal is a modified form of without further experimentation, since there are other radicals that give rise to a narrow g = 2.0 signal. Quantitation of plastoquinone in a third type of spinach PS II preparation containing CP47, D1, and D2 has given 0.8 plastoquinones per 17 chlorophylls or per reaction center. This sample was isolated through the use of nonionic detergents and sucrose density-gradient centrifugation. Antenna size was quantitated by the assumption that there are two pheophytins per reaction center (Akabori et al., 1988). However, a reaction center size of 17 chlorophylls seems small in comparison to the other CP43-depleted preparations that were described above. In contrast, cyanobacterial PS II preparations that lack the 43-kDa protein seem to retain A preparation from a thermophilic cyanobacterium lacking the 43-kDa protein has been characterized previously. These samples are prepared by electrophoresis of PS II preparations in the presence of the denaturing detergent sodium dodecyl sulfate. Therefore, during purification these cyanobacterial samples are exposed to much harsher conditions than the plant preparations described above. In one characterization of cyanobacterial samples that lack the 43-kDa protein, one mole of extractable plastoquinone was found per 67 chlorophylls. The chlorophyll antenna size was approximately 60 chlorophylls per reaction center, as determined by photoreduction of (Yamagishi and Katoh, 1985; Takahashi and Katoh, 1986). Thus, this preparation contains approximately one quinone per reaction center. A narrow EPR signal was observed in the light in this preparation. In this case, this signal was ascribed to a chlorophyll radical, not a modified form of No spin quantitation ofthe narrow signal was reported. The optical absorption spectrum of was not observed (Takahashi and Katoh, 1986). On the other hand, EPR characterization of the same preparation by another group has shown oxidation of Z and reduction of (Boska et al., 1986). In this case, had a normal EPR lineshape. Spin quantitation showed that there was one spin per 50 chlorophylls. The authors suggested that the discrepancy between these EPR results and those of Takahashi and Katoh (1986) may be due to the fact that the studies of Boska et al. (1986) used lower

temperatures to record the EPR data. Loss of the 43-kDa protein seems to affect the EPR lineshape of the tyrosine radicals in some of thesepreparations. This is an interestingphenomenon that should be better characterized in a system where isotopic labeling is possible. This may allow the origin of the narrow signals to be determined. Removal ofthe 43-kDa protein also seems to lead to loss of at least in some biochemical preparations. In fact, as described above, reported quinone contents per reaction center varies widely. Some of this variation maybedue to differences inthemethods that were used to measure the chlorophyll antenna size. On the other hand, it has been suggested that the 43-kDa protein provides part of the binding site for (Petersen et al., 1990). This suggestion explains loss ofquinone when the 43-kDa is removed, Rögner et al. (1991) disagree with this conclusion and have proposed that loss of in some of these preparations is not caused by removal of the 43-kDa protein but instead by the biochemical conditions used to effect this removal. This could either be caused by conformational destabilization or by removal ofother essential polypeptides that may play a role in quinone binding or function (Nagatsuka et al., 1991). However, an unexplored factor behind this variation could be a difference in the binding site for quinone in cyanobacteria and plants. Notice that the cyanobacterial preparation described in (Takahashi and Katoh, 1986) and (Yamagishi and Katoh, 1985) retains in spite of removal of the 43-kDa under rather harsh conditions (SDS), while there is a great deal ofvariability in the amount ofquinone associated with the spinach preparations.

C. The 24-kDa and 18-kDa Extrinsic Proteins are not Present in Cyanobacteria The 24- and 18-kDa proteins are found in plants and green algae (reviewed in Erickson and Rochaix, 1992). Unlike the 33-kDa protein, the plant 24-kDa and 18-kDa extrinsic proteins can be removed by NaCl washing. Once removed, these extrinsic proteins behave as soluble proteins and can be rebound to the reaction center (Miyao and Murata, 1983a). Binding of the 24-kDa protein seems to require the 33-kDa protein (Miyao and Murata, 1989); binding of the 18-kDa protein requires the 24-kDa protein (Miyao and Murata, 1983a). Since the stoichiometric ratios of the 33-, 24-, and 18-kDa proteins are 1:1:1 (Murata et al., 1984), the result that there are two copies ofthe

Chapter 8 Photosystem II 33-kDa protein per reaction center suggests that two copies ofthe 24- and 18-kDa proteins are also present (Xu and Bricker, 1993). No evidence has been found for association of the three extrinsic proteins in solution via sedimentation experiments (Miyao and Murata, 1989). Therefore, it has been suggested that a conformational change may occur upon binding of the 33-kDa protein to the reaction center. This conformational change would then create the binding sites for the 24- and 18-kDaproteins either on the 33kDa protein or on the reaction center (Miyao and Murata, 1989). Removal of the 24- and 18-kDa proteins leads to low rates of oxygen evolution unless millimolar concentrations of and chloride are provided in the assay media (Ghanotakis et al., 1984a; Ghanotakis et al., 1984d; Miyao and Murata, 1984a). Reconstitution of the polypeptides does not restore the oxygen evolution rate in the absence of (Ghanotakis et al., 1984d). Preparations to which the 24-kDa protein has been rebound, but the 18-kDa protein has not, show an elevated requirement for chloride (Akabori et al., 1984; Miyao and Murata, 1985). No gene products that are homologous to the plant 24- and 18-kDa proteins have been identified in cyanobacteria (reviewed in Erickson and Rochaix, 1992). Further, cyanobacteria have no polypeptides that crossreact with antibodies against the spinach 24- and 18-kDa proteins (Stewart et al., 1985a). Thus, it would be expected that cyanobacterial PS II preparations should show elevated requirements for and in order to obtain maximal activity. However, the situation is more complex than this simple argument would predict, since other ions can substitute in some cyanobacteria (for reviews of the function of in plants and cyanobacteria, see Yocum, 1991; Debus, 1992). In addition to their role in sequestering and the 24- and 18-kDa proteins also protect the manganese cluster from reduction by hydroquinone and make the cluster less accessible to reduction by hydroxylamine (Ghanotakis et al., 1984c; Tamura and Cheniae, 1985; Tamura et al., 1986). This effect may be via steric hindrance. Protection could also occur through a conformational change in the reaction center that is caused by the binding of the 24- and 18kDa proteins. It is not known what protein component, if any, performs this function in cyanobacteria. In a comparative study of membranes, the manganese

225 cluster of untreated Chlamydomonas reinhardtii thylakoids was found to be more accessible to hydroxylamine reduction than in Synechocystis sp. strain PCC 6803 thylakoids (Mor et al., 1993). However, there was no verification that the C. reinhardtii membranes retained the 24- and 18kDa proteins. In intact cells of Synechocystis sp. strain PCC 6803, it has been shown that the for reduction is and that the psbO deletion strain is 3.5 fold more accessible to the reductant (Burnap and Sherman, 1991). It is difficult to compare these values to previous experiments on salt-washed plant preparations (for example, Ghanotakis et al., 1984c), since the experimental conditions (e.g., incubation times) are different. Thus, there has not as yet been a comparative study of reduction of the manganese cluster by hydroxylamine in plant preparations and in cyanobacterial PS II particles. Also, it has not been determined if manganese is protected from reduction by hydroquinone in cyanobacteria, as it is in intact plant preparations (Ghanotakis et al., 1984c). While no extrinsic proteins homologous to the 24kDa and 18-kDa proteins have been found in cyanobacteria, two other extrinsic proteins of unknown function have been detected (Shen and Inoue, 1993). One of these proteins has a molecular mass of either 9 or 12 kDa depending on the cyanobacterial source. For example, a 9-kDa extrinsic protein has been found in PS II particles from the thermophilic cyanobacteria, Phormidium laminosum (Stewart et al., 1985a;Rolfe and Bendall, 1989), and a 12-kDa protein has been identified in S. vulcanus (Shen et al., 1992; Shen and Inoue, 1993). The 12kDa protein has been found to be homologous to the 9-kDa polypeptide (Shen et al., 1992). The second unique extrinsic protein is cytochrome a low-potential cytochrome that has been detected in several cyanobacterial strains including S. vulcanus (Shen et al., 1992; Shen and Inoue, 1993), P. laminosum (Bowes et al., 1983), Microcystis aeruginosa (Cohn et al., 1989), Aphanizomenon flos-aquae (Cohn et al., 1989), and Synechocystis sp. strain PCC 6803 (MacDonald et al., 1994). Like other PS II extrinsic proteins, the 9-kDa, 12kDa, and the cytochrome proteins can be removed by 1M or Tris treatment (Stewart et al., 1985a; Shen et al., 1992). Differences in binding affinity seem to exist between the homologous 9-kDa and

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the 12-kDa proteins, since 1M NaCl, 1M and low glycerol treatments can extract the 9-kDa, but not the 12-kDa protein from the thylakoid membrane (Stewart et al., 1985a; Stewart et al, 1985b; Rolfe and Bendall, 1989; Shen et al., 1992). Removal of the 9-kDa protein in P. laminosum has been associated with loss of oxygen evolution, and reconstitution with the 9-kDa protein was shown to restore partial activity (Stewart et al., 1985b; Rolfe and Bendall, 1989). However, these studies have been criticized because it is not clear whether the loss of oxygen evolution is due to the absence of the 9-kDa protein or to a glycerol effect (Shen et al., 1992). A different result is seen in S. vulcanus, where even after almost complete removal of the 12-kDa protein, oxygen evolution rates from 1000–1400 were still observed (Shen et al., 1992). This last result shows that the 12-kDa protein is not essential for oxygen evolution in this species. Reconstitution studies were recently carried out in S. vulcanus to determine the binding properties and function of the 12-kDa and cytochrome polypeptides (Shen and Inoue, 1993). It was found that cytochrome can bind to the thylakoid membrane in the absence of both the 33-kDa and the 12-kDa proteins. Such binding does not enhance oxygen evolution activity. The 12-kDa protein does not bind to the thylakoid membrane in the absence of the 33-kDa and cytochrome proteins. The 12kDa and cytochrome proteins enhanced oxygen evolution activity in the presence of the 33-kDa protein, implying a close binding interaction and functional interdependence among these three extrinsic proteins. It was proposed that the 12-kDa (and probably the 9-kDa) and cytochrome proteins are important in regulation of the oxygen evolution activity (Shen and Inoue, 1993). Other proposed roles for cytochrome include involvement in controlling the degree of PSI cyclic electron transfer and in noncyclic electron transfer between PS I and PS II (Cohn et al., 1989; Shen et al., 1992). It is clear that more work needs to be done to elucidate the roles of these extrinsic proteins in cyanobacterial PS II.

D. Small Polypeptides in Plant and Cyanobacterial Photosystem II In this section present knowledge about the function of low-molecular-weight components of PS II preparations from plants and cyanobacteria will be presented.

1. Photosystem II Preparations from Plants Three types of plant PS II preparations will be considered here:

Plant PS II membranes are purified grana membranes that contain PS II and the light harvesting complex (reviewed in Dunahay et al., 1984). Triton X-100 or digitonin is used to solubilize the stroma membranes, leaving a membrane fraction that can be pelleted at low centrifugal forces. These membranes have low levels of contaminating PS I, cytochrome and ATP synthase; the amounts depend on the preparation (Dunahay et al., 1984). Oxygen-evolving PS II core complexes are monodisperse in detergent and have fewer polypeptides than PS II membranes. Core-complex purification procedures often start with PS II membranes, followed by subsequent removal of the light harvesting complex and other accessory proteins, which are not required for oxygen evolution (reviewed in Ghanotakis et al., 1987b). A typical spinach PS II core preparation uses the non-ionic detergent, octyl glucoside, to solubilize PS II membranes, after which column chromatography is performed in order to remove nonessential components (Ghanotakis et al., 1987a). Some of these purification procedures have the effect of changing the properties of the acceptor side of PS II (Ghanotakis et al., 1987b). A method to purify a core complex directly from the thylakoid membrane through the use ofone detergent has also been described (Fotinou and Ghanotakis, 1990). Although core complexes typically lose the 24- and 18-kDa proteins (Ghanotakis et al., 1987b), three preparations that reduce the chlorophyll antenna size and retain these extrinsic proteins have been developed (Enami et al., 1989a; MacDonald and Barry, 1992; Mishra and Ghanotakis, 1993). The smallest antenna size reported for an oxygen-evolving core preparation is thirty-five chlorophylls per reaction center (van Leeuwen et al., 1991). The most resolved plant PS II preparation is the ‘reaction center complex.’ PS II reaction center complexes are no longer able to evolve oxygen, but are still capable of primary charge separation between

Chapter 8 Photosystem II theprimary chlorophylldonorandpheophytin(Nanba and Satoh, 1987). A reaction center complex can be purified by Triton X-100 detergent extraction and anion exchange chromatography (Nanba and Satoh, 1987). This treatment removes all of the protein components, except for D1, D2, cytochrome and one low-molecular-weight protein that is discussed below. Reaction center preparations that employ dodecyl maltoside or a combination ofoctyl glucoside and octyl thioglucoside have been shown to be more stable than the Triton-purified material (Akabori et al., 1988; Seibert et al, 1988;Ghanotakis et al., 1989; Fotinou and Ghanotakis, 1990). Treatment of Triton-purified reaction center complexes with a second detergent, octyl glucoside, and gel permeation high-performance liquid chromatography, yields D1-D2 particles with no cytochrome subunits or other small polypeptides (Tang et al., 1990). This isolated complex is active in reduction of pheophytin under steady state illumination.

2. Photosystem II Preparations from Cyanobacteria Three types of PS II preparations can be described for cyanobacteria:

227 these latter three methods, the Noren et al. (1991) preparation is the only one that retains high rates of oxygen evolution. We will define a second type of oxygen-evolving preparation from cyanobacteria, ‘cyanobacterial core particles.’ Cyanobacterial core particles are depleted of accessory pigments and other contaminating proteins. They are prepared from PS II particles by treatment with a second detergent, followed by ion exchange chromatography or sucrose gradient centrifugation (Bowes et al., 1983; Dekker et al., 1988; Koike et al., 1989). Like plant reaction center complexes, cyanobacterial (Gounaris et al., 1989; Ikeuchi et al., 1989a) reaction center complexes can be purified using Triton X-100 or lauryl maltoside. This treatment removes all of the protein components except for D1, D2, cytochrome and two low-molecularweight polypeptides (discussed below). The cyanobacterial reaction center preparations were found to retain more chlorophyll than the comparable plant preparations (Gounaris et al., 1989; Ikeuchi et al., 1989a). Ikeuchi et al. (1989a) attributed the higher chlorophyll to reaction center ratio to contaminating 47-kDa protein. However, no contaminating 47-kDa protein was found in the preparation of Gounaris et al. (1989), when assayed with an antibody against the spinach CP47 protein.

3. Low-Molecular-Weight Polypeptides in Reaction Center Complexes (see Tables 2 and 3) The least resolved cyanobacterial PS II preparation is typically monodisperse in detergent and consists ofprotein-detergent micelles (but see, Nilsson et al. 1992). These micelles contain PS II and phycobiliproteins, with minor amounts of contaminating PS I and ATP synthase. In a typical preparation thylakoid membranes are isolated, and a detergent is used to solubilize the thylakoid membranes. Many preparations rely on differential solubilization of PS II from the membrane (for example, see Stewart and Bendall, 1979; England and Evans, 1981; Miyairi and Schatz, 1983; Pakrasi and Sherman, 1984; Schatz and Witt, 1984; Smutzer and Wang, 1984; Frei et al., 1988; McDermott et al., 1988; Burnap et al., 1989; Kirilovsky et al., 1992; Nilsson et al., 1992). Only a few methods do not use this technique (Satoh et al., 1985; Rögner et al., 1990; Noren et al., 1991). Of

The psbI gene product is found tightly associated with the PS II reaction center in both cyanobacteria (Ikeuchi et al., 1989a) and plants (Ikeuchi and Inoue, 1988; Ikeuchi et al., 1989c; Webber et al., 1989c). The psbI gene products from spinach and the cyanobacterium, S. vulcanus, have apparent molecular weights of 4.8-kDa and 5.0-kDa, respectively, as determined by SDS-PAGE. This protein is predicted to contain one membrane spanning region (Ikeuchi and Inoue, 1988; Ikeuchi et al., 1989a). The function of the psbI gene product is not known, although its tight association with both higher plant and cyanobacterial reaction centers imply that it is important for PS II function or structure. The hydrophobic psbL gene product is found in cyanobacterial PS II reaction center complexes from S. vulcanus. This protein has an apparent molecular weight of 5 kDa, as determined by SDS-PAGE

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(Ikeuchi et al., 1989a). The psbL gene product is not found in higher plant reaction center complexes, but is found in oxygen-evolving core complexes from plants (Ikeuchi et al., 1989c). ThepsbL gene product is necessary for assembly and/or function of PS II, since its deletion in Synechocystis sp. strain PCC 6803 results in loss of PS II activity (Pakrasi and Vermaas, 1992). Based on reconstitution of spinach PS II preparations with crude protein extracts, either the psbL gene product or the 4.1-kDa protein (see Section II D, 4) has been suggested to be important for photoreduction of (Nagatsuka et al., 1991).

4. Low-Molecular-Weight Polypeptides in Core Photosystem II Particles Low-molecular-weight components of the PS II reaction center (Section II D, 2) are also present in core preparations, since core preparations are less well resolved (see Tables 2 and 3). Several additional small polypeptides are found in oxygen-evolving core particles. Proteins common to both cyanobacterial and plant core complexes are a 4.1-kDa protein (Ikeuchi et al., 1989b; Ikeuchi et al., 1989c; Koike et al., 1989) and the psbH gene product (Michel and Bennet, 1987; Ikeuchi et al., 1989c; Koike et al., 1989). The gene origin of the 4.1-kDa protein is not known. It is presumed to be encoded by the nuclear genome, since a homologous sequence is not found in the chloroplast genome (Ikeuchi et al., 1989c). The 4.1-kDa protein shows low homology when plant and cyanobacterial sequences are compared, and it is predicted to contain one hydrophobic, transmembrane region (Ikeuchi et al., 1989b). The 4.1-kDa protein or the psbL gene product has been suggested to be important for photoreduction of (Nagatsuka et al., 1991). The psbH gene product contains one transmembrane region and undergoes light dependent phosphorylation (Bennett, 1979) of a threonine residue found at position 2 in the mature protein of plants (Michel and Bennet, 1987). These corresponding first twelve amino acids are absent in the PsbH from cyanobacteria, and the cyanobacterial protein is not phosphorylated (Koike et al., 1989; Abdel-Mawgood and Dilley, 1990; Mayes and Barber, 1990). Possible roles for phosphorylation in the regulation of photosynthesis have been discussed in (Allen, 1992). Gene inactivation studies in Synechocystis sp. strain PCC 6803 show that PsbH is

not absolutely essential for PS II assembly or activity, although these deletion mutants have slower photoautotrophic growth rates and decreased oxygenevolution activity (Mayes et al., 1993). However, under high light intensity, it has been reported that these deletion mutants are no longer able to grow photoautotrophically (Pakrasi and Vermaas, 1992). The deletion mutants may have impaired electron transfer between and This result suggests that PsbH optimizes electron transfer between the two quinones, possibly by directly interacting with the binding site for (Mayes et al., 1993). An interaction of PsbH with the two PS II plastoquinone molecules has also been proposed by Packham (1988). A 5-kDa hydrophilic protein is found in higher plant core complexes, but this protein has not been identified in cyanobacterial preparations (Ljungberg et al., 1986b; Murata et al., 1988; Ikeuchi et al., 1989c). It is not known whether this hydrophilic protein is located on the inside or outside of the thylakoid membrane, and its function is not known. The absence of the 5-kDa hydrophilic protein in cyanobacteria may be related to the absence of the 24- and 18-kDa extrinsic proteins. There are several low-molecular-weight components that have been found in cyanobacterial core PS II particles and that have not been identified in higher plant core complexes. An example is the 3.9kDa psbK gene product, which is not found in higher plant core complexes, but is found in less well resolved plant PS II membranes (Murata et al., 1988; Schroder et al., 1988; Koike et al., 1989). PsbK contains one putative transmembrane region, and is not essential for PS II function, since deletion mutants in Synechocystis sp. strain PCC 6803 are able to grow photoautotrophically (Ikeuchi et al., 1991). Moreover, these deletion mutants grow more slowly under both photoautotrophic and photoheterotrophic conditions, implying that PsbK may be involved in more than just optimizing PS II activity (Ikeuchi et al., 1991). The psbM and psbN gene products are both approximately 4.7 kDa in mass and contain one putative transmembrane region (Ikeuchi et al., 1989b). Both proteins are present in cyanobacterial core particles, but neither protein has been detected in any preparation from higher plants. However, the psbM and psbN genes are both present in the chloroplast genome, and active transcription of psbN in peas has been observed (Ikeuchi et al., 1989b). The functions of PsbM and PsbN are not known. Deletion of the

Chapter 8 Photosystem II

psbN and psbH genes in Synechocystis sp. strain PCC 6803 still permits photoautotrophic growth (Mayes et al., 1993). A 5-kDa protein of unknown function and gene origin has been detected in core PS II particles from the cyanobacterium, S. vulcanus (Ikeuchi et al., 1989b). The amino-terminal sequence of the 5-kDa protein has been determined. This work has shown that this polypeptide is not homologous to any other

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known protein (Ikeuchi et al., 1989b), including the 5-kDa hydrophilic protein found in higher plants (Ljungberg et al., 1986b; Ikeuchi et al., 1989c; see discussion above). However, the 5-kDa protein may not be an intrinsic component of cyanobacterial PS II, since it has also been detected in PS I core preparations from S. vulcanus (Ikeuchi et al., 1989b).

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5. Low-Molecular-Weight Polypeptides in Plant Photosystem II Membranes and Cyanobacterial Photosystem II Particles Higher plant PS II membranes contain proteins that have not been identified in cyanobacterial PS II particles, including a 6.1-kDa protein and the psbR gene product (Ikeuchi et al., 1989c; see Tables 2 and 3). Since these two proteins are not found in plant oxygen evolving core complexes, they must not be essential for PS II oxygen-evolution activity. The 6.1-kDa protein from (Ikeuchi et al., 1989c) is the same subunit identified as the ‘6.5-kDa’ and ‘7kDa’ protein by (Schroder et al., 1988), The amino terminal sequence of all three species is the same, indicating that this protein may be susceptible to proteolysis near the carboxyl terminus. No homologous sequence has been found in the chloroplast genome, suggesting that the 6.1-kDa protein is encoded by the nucleus (Schroder etal., 1988; Ikeuchi et al., 1989c). The psbR gene product has been proposed to be a peripheral membrane protein based on its extraction

from the lumenal side of the membrane by Tris or detergent treatment. This polypeptide also shows low solubility in aqueous solutions (Ljungberg et al., 1986a). It has also been suggested that PsbR is an integral membrane protein with a single transmembrane region (Lautner et al., 1988). Studies have been conducted in potatoes using anti-sense RNA to decrease the levels of the 10-kDa PsbR protein. Depletion of this protein had the effect of lowering oxygen evolution rates and damping the period four oscillation pattern for oxygen evolution, as compared to nondepleted samples (Stockhaus et al., 1990). The psbJ gene product has been identified in cyanobacterial thylakoid membranes, but has not been detected in purified cyanobacterial PS II preparations. The gene product has not been detected in higher plants (Lind et al., 1993). Early termination of the psbJ gene in Synechocystis sp. strain PCC 6803 results in an approximately fifty percent decrease in PS II content, suggesting that PsbJ is important for the assembly and/or stability of the PS II complex (Lind et al., 1993).

Chapter 8 Photosystem II

E. Cytochrome Cytochrome is present in both the plant and cyanobacterial PS II complex. Cytochrome has two subunits, and which are encoded by the psbE and psbF genes, respectively (reviewed in Erickson and Rochaix, 1992). In plants, cytochrome is chloroplast-encoded (Herrmann et al., 1984). The are present in a 1:1 stoichiometry (Widger et al., 1985), and heme ligation occurs through histidine residues (Babcock et al., 1985). Since each subunit has only one histidine (Herrmann et al., 1984; Widger et al., 1985; Pakrasi et al., 1988), the heme must cross-link the two subunits (Cramer et al., 1986). It has been suggested that cytochrome may be a heterodimer of and subunits (Widger et al., 1985; Cramer et al., 1986). Both polypeptides are predicted to cross the membrane with one segment (Herrmann et al., 1984). The amino terminus of the subunit is exposed on the stromal side, and the carboxyl terminus of the subunit is exposed on the lumenal side of the thylakoid membrane (Tae et al., 1988; Tae and Cramer, 1989; Vallon et al., 1989). In a preliminary report, the amino terminus of the subunit was found to be exposed on the stromal side of the membrane, suggesting that the and subunits have parallel orientation in the membrane (Tae and Cramer, 1989). However, the suggestion that the subunit is parallel in orientation to the subunit may be in contradiction with a preliminary report on mutants in which the and subunits have been linked using genetic techniques (Pakrasi and Vermaas, 1992). Cytochrome is retained in highly resolved PS II reaction center preparations from either plants or cyanobacteria (see Section II D, 3). Also, deletion of the psbE and psbF genes disrupts assembly of active PS II complexes (Pakrasi et al., 1988; Pakrasi et al., 1990). When the putative histidine ligands to the heme are mutagenized by site-directed techniques, there is no assembly of active PS II centers (Pakrasi et al., 1991). These observations provide evidence that cytochrome plays an essential structural role in PS II. The functional role ofcytochrome is less clear cut (for discussions, see Cramer and Whitmarsh, 1977; Buser et al., 1992a; Nedbal et al., 1992; and references therein). The midpoint potential of cytochrome is heterogeneous and depends on

231 the preparation (Cramer and Whitmarsh, 1977; Thompson et al., 1989). Deletion studies have been performed to address the role of cytochrome in PS II. Truncations of the carboxyl terminus of the PsbE decreased the number of assembled centers, but assembled centers were active in oxygen evolution (Tae and Cramer, 1992). This result has been cited as evidence that the subunit of cytochrome is not required for water-splitting and does not provide ligands to the manganese cluster (Tae and Cramer, 1992; however, see Shukla et al., 1992 for a contrary view). A controversy also exists concerning cytochrome stoichiometry in PS II. In thylakoid membranes, most workers have found two cytochrome per PS II reaction center (e.g., Whitmarsh and Ort, 1984, but see Buser et al., 1992b for a contrary report). In PS II preparations of various kinds, either one (Ford and Evans, 1983; Ghanotakis et al., 1984b; Yamagishi and Fork, 1987; Miyazaki et al., 1989; Gounaris et al., 1990; Buser et al., 1992b) or two (Lam et al., 1983; Murata et al., 1984; Briantais et al., 1985; de Paula et al., 1985; Haag et al., 1990; Noren and Barry, 1992) cytochromes per reaction center have been reported. Some studies have reported intermediate values (Nanba and Satoh, 1987; Rögner et al., 1990; van Leeuwen et al., 1991). Unfortunately, these results are difficult to compare; different preparations, chemical reductants, and extinction coefficients for cytochrome have been employed. Also, different methods have been used to quantitate reaction center size. Interestingly, the studies of van Leeuwen et al. (1991) have found evidence for the release of cytochrome from the spinach reaction center during purification of PS II core preparations. This result suggests that loss of cytochrome or heme may underlie the measured variability in cytochrome stoichiometry. This hypothesis is supported by a recent study in which a cyanobacterial and a plant PS II preparation were found to have different cytochrome stoichiometries (MacDonald et al., 1994).

III. Site-Directed Mutagenesis Studies of the Donor Side of Photosystem II The procaryotic cyanobacteria have emerged as a convenient system for studying structure/function relationships in PS II. In particular, the unicellular

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organism, Synechocystis sp. strain PCC 6803, has been used extensively for such studies. This organism has the advantage of being easily transformable and undergoing gene replacement through homologous recombination (Williams, 1988). Also, Synechocystis sp. strain PCC 6803 can be grown photoheterotrophically, so mutations that completely inactivate the water-oxidizing complex can be recovered for study (Williams, 1988). Recently, two strains that lack PS I have been developed (Smart et al., 1991; Shen et al., 1993). These strains should be of utility in future mutagenesis studies on PS II. The disadvantage of using Synechocystis sp. strain PCC 6803 as a system in which to study PS II was the initial difficulty in obtaining good biochemical preparations of PS II from this organism. However, a variety of oxygen-evolving preparations have now been described (Burnap et al., 1989; Noren et al., 1991; Kirilovsky et al., 1992; Nilsson et al., 1992). In addition to these oxygen-evolving preparations, a non-oxygen-evolving corepreparation fromSynechocystis sp. strain PCC 6803 has also been developed (Rögner, 1990; Rogner et al., 1990). In this section, we will discuss the use of sitedirected mutagenesis to understand the function and structure of the donor side of PS II. For recent reviews of acceptor side mutations in cyanobacteria, see Diner et al. (1991) and Pakrasi and Vermaas (1992). For a review of deletion mutagenesis studies of PS II polypeptides, see Pakrasi and Vermaas (1992).

A. A Search for the Ligands to the Manganese Cluster The manganese cluster must be coordinated by amino acid residues. The cluster is believed to be tetranuclear (see Yachandra et al., 1993). Since the manganese must be either five- or six-coordinate (Pecoraro, 1988), the maximum number of ligands required for binding of four manganese is twenty-four. However, this number does not account for or other bridging species, water ligation, or the fact that some ligands may be bidentate. As described above, oxygen-evolving PS II preparations are made up of multiple subunits. At this point, very few of these can be definitely eliminated as a source of ligands to the metal cluster. The best positive evidence for manganese ligation points to D2 and D1. Recent work shows that mutations in CP47 influence the properties of the

water-splitting complex. The strategy in using site-directed mutagenesis to identify manganese ligands is to target a particular subunit and then make multiple mutations at residues that are predicted to lie in or near the lumen. Often, multiple mutations are introduced at each site. In most cases, the residues chosen for mutagenesis are those with side chains that are nitrogen- or oxygenrich. Recently, spin-echo measurements have shown that at least one histidine is a ligand to the manganese cluster in the state (Tang et al., 1994). Comparison with manganese model compounds suggests that carboxylates are also likely to be ligands to the cluster, since carboxylate ligands would have the effect of destabilizing the high oxidation states of the manganese (Pecoraro, 1988). Analysis of the effect of these mutations is more problematic. Potentially important residues for PS II function or structure can easily be identified by their photoheterotrophic phenotype. However, such a phenotype can be induced by a variety of changes in the reaction center, and providing definitive evidence that a given mutation affects only the binding of the metal cluster is difficult in the absence of crystallographic data. Also, the metal cluster may contain and/or chloride as integral components of the cluster (for example, see the model in Yachandra et al., 1993). In such a case, it may be difficult to distinguish changes in manganese binding from changes in the binding affinity of other components of the metal cluster. In spite of these reservations, it is important to point out that an X-ray structure of the manganese containing PS II reaction center is still probably years away. Site-directed mutagenesis studies on another integral membrane protein, bacteriorhodopsin, have shown that the cumulative effect of many mutational studies can yield important structural and functional information (for review, see Henderson et al., 1990). This is the strategy that several groups have now adopted in their approach to the water-oxidizing complex. In the following section, we review work to date that is designed to locate residues affecting the stability or assembly of the manganese cluster.

1. Mutations in the D1 Polypeptide (psbA Gene Product) The D1 polypeptide has been the target of many mutagenesis studies. Indirect evidence of several

Chapter 8 Photosystem II kinds suggests that the D1 polypeptide may provide ligands to the metal cluster (reviewed in Debus, 1992 and Rutherford et al., 1992). One line of evidence is derived from studies ofthe LF-1 mutant of Scenedesmus obliquus. This mutant has been found to have no functional manganese cluster, although it does retain some manganese and has a partially active reaction center (Metz et al., 1980). It is now known that the defect in this mutant is a failure to process the carboxyl terminus of D1, which is located on the lumenal side of the thylakoid membrane (Marder et al., 1984; Diner et al., 1988; Taylor et al., 1988a). The defect was found to be in a gene encoding a specific D1 protease (Taylor et al., 1988b; Fujita et al., 1989; Inagaki et al., 1989; Bowyer et al., 1992). One interpretation of the phenotype of the LF-1 mutant is that residues on the lumenal side of D1 bind manganese (see Metz and Seibert, 1984; Seibert et al., 1989).

233 Site-directed mutations have been generated at many of the oxygen and nitrogen containing residues that are predicted to fall on the lumenal side of D1 (Fig. 3). Tables 4 and 5 summarize the sites where substitutions either eliminate or impair photoautotrophicgrowth. This isthe minimumrequirement for consideration ofa residue as a ligand to a catalytic site. Of the sites in Tables 4 and 5, further characterization has identified the following set as most likely to influence the stability, assembly, or catalytic efficiency of the metal cluster: aspartate 170, aspartate 342, histidine 332, histidine 337, glutamate 189, glutamate 333 (Nixon and Diner, 1991; Chu et al., 1993). Mutations at histidine 190 also inhibit photoautotrophic growth (Table 5); it has been suggested that this residue is a hydrogen bonding donor to tyrosine Z (Diner et al., 1991; Roffey et al., 1994), as also suggested on the basis of modeling studies of D1 and D2 (Svensson et al.,

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1990). (Alternatively, modeling studies have suggested that D is hydrogen bonded to glutamine 164 of D2 and that Z is not hydrogen bonded (Ruffle et al., 1992)). Aspartate 59 and aspartate 61 are believed to directly or indirectly influence the binding of ions (Chu et al., 1993). The free carboxylate

of alanine 344, which is the carboxy terminus of the processed form of D1, has also been suggested to be a manganese ligand (Nixon et al., 1992; see below). In this section, we will discuss some of the sitedirected mutagenesis evidence that has led to these assignments. Chemical modification studies also provide a route for identification of ligands to manganese (see Preston and Seibert, 1991). Characterization of the effect of multiple mutations at D170 has been the most extensive. In one study, eleven different substitutions were made at this site (Diner and Nixon, 1992; Nixon and Diner, 1992). Analysis of the properties of intact cells indicated that the ability of the substituted residue to sustain steady-state oxygen evolution was correlated with a lower for the residue. Using a core preparation that does not evolve oxygen, it was observed that preparations containing residues with a lower and a higher for oxygen evolution had a lower for oxidation ofexogenous manganese by tyrosine This work implies that aspartate 170 is necessary for the assembly of the manganese cluster. Further, Nixon and Diner (1992) have suggested that aspartate 170 may ligate directly the first manganese atom to bind to the reaction center. In another study, two substitutions at D170 were characterized through the use ofan oxygen-evolving PS II preparation (Boerner et al., 1992). This work showed a substantial reduction in the content of bound manganese in thylakoid membranes and in

Chapter 8 Photosystem II PS II particles from the D170N D1 mutant. A perturbation of the catalytic efficiency of the manganese cluster was also observed in the D170E D1 mutant (Boerner et al., 1992). These results are consistent with the idea that this residue influences the assembly or stability of the cluster. Previous mutagenesis studies of other metalloproteins have shown that substitutions at a metal ligand often lead to loss ofactivity and loss of metal. Thus, the results of Boerner et al. (1992) are consistent with aspartate 170 providing a ligand to a metal atom in the cluster. However, the possibility of conformational changes leading to this phenotype cannot be eliminated. To date, this is the only published characterization of a putative manganese mutant through the use of an oxygen-evolving preparation. The carboxy-terminal alanine itself has been shown to be essential for photoautotrophic growth (Nixon et al., 1992). This was observed by generating a series of mutants harboring premature termination codons in the D1 polypeptide. In the wild type protein alanine 344 is the carboxy-terminal amino acid residue after processing. If it is deleted, the organism is unable to assemble a functional manganese cluster. Mutants in which glycine, methionine, serine, and valine are substituted at the sitecangrowphotoautotrophically. Mutantsinwhich tyrosine or lysine are substituted at 344 cannot grow photoautotrophically, but can evolve oxygen at low steady state levels. These data were explained by proposing that the free carboxy-terminus of alanine 344 ligates manganese (All of these mutants at A344 are processed, since they were constructed in a strain containing a mutated psb A gene encoding a truncated D1 polypeptide (Nixon et al., 1992).). However, mutations at this site leave a high-affinity binding site for the oxidation of manganese essentially unperturbed, when assayed in vitro using a non-oxygenevolving preparation. The authors suggested that in this mutant the cluster is unable to assemble after the binding of the first manganese. Alternatively, the cluster may be assembled but unable to advance normallythroughthe S states. Characterizationusing a manganese-containing preparation is necessary in order to distinguish between these possibilities (Nixon et al., 1992).

2. Mutations in the D2 Polypeptide (psbD Gene Product) D2 also contains carboxylates that may also serve as

235 ligands to the manganese cluster (Fig. 4). Negatively charged residues, as well as histidine, glutamine, and asparagine, have been targeted for mutagenesis. Of approximately thirty residues, the only one at which mutations gave rise to a photoheterotrophic phenotype was glutamate 68 (E69 corresponds to E68 in the numbering scheme in Fig. 4 and in Trebst, 1986; Pakrasi and Vermaas, 1992). Two substitutions were introduced at glutamate 68: glutamine (E68Q) and valine (E68V; Vermaas et al., 1990; Yu and Vermaas, 1993). Both strains required glucose for growth. The valine-substituted strain was completely inactive in oxygen evolution and had no detectable content of assembled PS II centers. DCMU binding showed that the glutamine mutant, on the other hand, contained assembled PS II centers. When compared to wild type, this mutant had an approximately threefold lower number of centers on a chlorophyll basis. Although the glutamine mutant could not grow photoautotrophically, intact cells ofthe E68Q mutant evolved oxygen at an initial rate that was a factor of 3.5 lower than wild-type, in reasonable agreement with the lower PS II content. However, oxygen evolution was rapidly inhibited. This effect was ascribed to photoinhibition (Van der Bolt and Vermaas, 1992), and addition of 1 mM manganese chloride was reported to stabilize activity (Vermaas et al., 1990). However, there was no detailed characterization ofthe electron transfer properties of this mutant. On the basis ofthe data described above, it was suggested that glutamate 68 of the D2 polypeptide is a ligand to manganese. An increased tendency for photoinhibition has also been noted in a proline 161 D2 mutant (Vermaas et al., 1990; Van der Bolt and Vermaas, 1992). In contrast to the E68Q mutant, manganese did not protect against photoinhibition in this mutant (Vermaas et al., 1990). The etiology of this increased susceptibility to photoinhibition for the proline 161 D2 mutant is not known. There have been no other reports describing other D2 residues as manganese ligands.

3. Mutations in the 47-kDa Protein (psbB Gene Product) CP47 (or the 47-kD protein, or PsbB) is a tightly bound, chlorophyll-containing antenna protein in PS II. Hydropathy plots suggest that the protein crosses the membrane six times. A large hydrophilic, lumenal loop is predicted to connect helix V and VI (reviewed in Bricker, 1990; see Fig. 2). This subunit contains

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twelve conserved histidines in the predicted hydrophobic regions (Fig. 2); site-directed mutagenesis has identified a subset of these histidines residues as chlorophyll ligands (Shen et al., 1993). Mutants in which the psbB gene has been deleted or insertionally inactivated fail to accumulate D1 and D2 in the membrane, suggesting thatCP47 is required for assembly ofa functional PS II complex (Vermaas et al., 1988a; Eaton-Rye and Vermaas, 1991). Several lines of evidence suggest an association between the 47-kDa protein and the 33-kDa extrinsic protein (see Section II A above). Some biochemical evidence for an association between CP47 and the manganese cluster itself has also been obtained. This work was performed through the use ofa monoclonal antibody that recognizes its epitope on the 47-kDa protein only after all four manganese have been removed. However, since conformational changes may

accompany manganese removal, these results do not necessarily imply that CP47 directly ligates the metal cluster (Bricker and Frankel, 1987; Frankel and Bricker, 1989). The extrinsic loop between helix V and VI contains many charged residues. In particular, it contains several pairs of basic residues. Deletion mutants in this loop have been generated (Eaton-Rye and Vermaas, 1991). A deletion mutant, does not assemble functional centers and cannot grow photoautotrophically. This mutant does accumulate immunologically detectable amounts of the 47-kDa, 43-kDa, D1 and D2 PS II proteins. A second deletion mutant, assembles a smaller number of assembled PS II centers when compared to wild type (Eaton-Rye and Vermaas, 1991). This second deletion mutant can still grow photoautotrophically. Recently, however, use of

Chapter 8 Photosystem II smaller deletions has indicated that the photoheterotrophic phenotype seen in the first deletion mutant, may be the result ofprotein conformational changes. This could be caused by the large deletion of amino acid residues (Haag et al., 1993). This more recent deletion study indicates thatregions near helix V and VI are essential for assembly of an oxygen-evolving PS II complex. Site-directed mutagenesis has been used to change a pair of conserved arginines to glycines (PutnamEvans and Bricker, 1992). These arginines, R384 and R385, are located in the region (R384-V392) deleted by Eaton-Rye and Vermaas (Eaton-Rye and Vermaas, 1991). Like the deletion mutant, intact cells ofthe RR384,385GG mutant grow photoautotrophically, but exhibit a lower specific activity for oxygen evolution. Thylakoid membranes from the mutant had oxygen evolution rates to DCPIP) that were only 15% ofcontrol rates, while the rate of electron transfer from DPC to DCPIP was approximately the same as control rates. From this characterization, it was concluded that this substitution destabilizes the manganese cluster. Also, the mutant is more susceptible to photoinhibition (Putnam-Evans and Bricker, 1992). However, mutations at a catalytic site might be expected to have a more profound effect on activity, so these effects may be due to a conformational alteration on the donor side of PS II (Putnam-Evans and Bricker, 1992). Analysis of electron transfer rates and metal content using a purified PS II preparation would aid in assessment of the effects of this mutation.

B. The Location of the Redox Active Tyrosines PS II contains two redox-active tyrosine residues, D and Z (Barry and Babcock, 1987; Boerner and Barry, 1993). In the oxidized and paramagnetic form, characteristic electron paramagnetic resonance (EPR) signals from these residues can be observed. The radical is very long-lived (Babcock and Sauer, 1973b), and its function is unknown. D is oxidized either by the and states (Babcock and Sauer, 1973a) or, in the absence of a functional manganese cluster, by (Buser et al., 1990; Vass and Styring, 1991; Noren and Barry, 1992). In turn, is reduced duringthe to transition in the dark (Styring and Rutherford, 1987). was identified as a tyrosine radical by isotopic labeling of tyrosine in the procaryotic cyanobacterium, Synechocystis sp. strain PCC 6803, and EPR spectroscopy (Barry and

237 Babcock, 1987). In addition to the stable signal described above, PS II also contains another free radical, Z, with a similar EPR line shape. This radical decays on a much faster time scale. This signal was originally observed upon illumination in manganese-depleted preparations (Babcock and Sauer, 1975a). Timeresolved EPR spectroscopy was used to show that the rise-time of this radical correlates with the reduction of which occurs on the microsecond time scale in such manganese depleted preparations (Conjeaud and Mathis, 1980; Boska et al., 1983; Yerkes et al, 1983). Recently, isotopic labeling and EPR spectroscopy have been used to demonstrate that the radical in manganese depleted material arises from a tyrosine residue (Boerner and Barry, 1993). EPR studies suggestthat is also an intermediate between manganese and in oxygen evolving particles (Babcock and Sauer, 1975b; Blankenship et al., 1975; Babcock et al, 1976; Cole and Sauer, 1987; Hoganson and Babcock, 1988). In addition, optical experiments have shown that a tyrosine is oxidized by and reduced by the manganese cluster in these preparations (Dekker et al., 1984c; Renger and Weiss, 1986; Koike et al., 1987; Saygin and Witt, 1987; Gerken et al., 1988). The finding that both the and species arise fromredox-active tyrosines suggests that site-directed mutagenesis experiments can be used to locate them. It was proposed that D and Z are symmetrically located on the polypeptides, D1 and D2 (Debus et al., 1988b; Vermaas et al., 1988b). This model assumes that there is symmetry in the D1 /D2 core ofPS II; this idea is based on the symmetry of the L/M core of the purple bacterial reaction center (Debus et al., 1988b; Vermaas et al., 1988b). Reference to a folding pattern for D1 and D2 (Trebst, 1986) (Figs. 3 and 4) led to the suggestion that Z is tyrosine 161 of D1 and D is tyrosine 160 of D2 (Debus et al., 1988b; Vermaas et al., 1988b). Sitedirected mutagenesis was used to substitute a nonredox-active phenylalanine at the 160 position ofthe D2 subunit. The EPR signal was not observed in intact cells of this mutant; this result is consistent with this residue giving rise to stable radical, (Debus et al, 1988a; Vermaas et al., 1988b). Three studies have characterized the effect of substitution of a non-redox-active phenylalanine at the 161 D1 position (Y161F D1 mutant) (Debus et al., 1988b; Metz et al., 1989; Noren and Barry,

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1992). The model described in Debus et al., 1988a and Vermaas et al., 1988b predicts that this residue gives rise to Z. The Y161F D1 mutant has a nonoxygen evolving phenotype (Debus et al., 1988b; Metz et al., 1989). Characterization of intact cells showed that the mutant exhibited the EPR signal of the tyrosineradical; therefore, the mutant reaction centers were assembled and partially functional. Electron transfer was found to be inhibited on the donor side of PS II (Debus et al., 1988b; Metz et al., 1989). The mutant was also found to have a very low content of tightly bound manganese, suggesting that tyrosine 161 may play a role in assembly of the manganese cluster (Noren and Barry, 1992; Blubaugh and Cheniae, 1990). Analysis through the use of a non-oxygen-evolving core preparation showed that the EPR signal of was not observed in the Y161F D1 mutant samples. Further, an optical characterization of the mutant preparation showed none of the characteristic UV absorption features of – Z (Metz et al., 1989). A new redox-active PS II species has been observed in the Y161F D1 mutant through the use ofthe PS II preparation of Noren et al., which gives oxygenevolving, wild-type particles (Noren and Barry, 1992; Noren et al., 1991). Instead of the control spectrum, a new light-induced EPR signal is observed in the Y161F D1 preparations (Noren and Barry, 1992). Spin quantitation shows that this signal is obtained in 60% of the centers. This four-line signal has a different line shape from Z+, but a similar g-value (Fig. 5). This new signal is also observed if either a tryptophan or a phenylalanine are substituted at tyrosine 160 ofthe D2 polypeptide. Characterization of PS II preparations isolated from the position 160 D2 mutants has shown that these substitutions alter the properties of tyrosine Z (Boerner et al., 1993). In these mutants, a light-induced signal is observed that has a different lineshape from but a similar lineshape to that of the new radical observed in the Y161F D1 mutation (Noren and Barry, 1992; Fig. 5). However, charge recombination kinetics between and an oxidized donor are not significantly perturbed by the D2 mutations. Up to 100% of the centers could generate the four-line signal, and the amplitude of the new signal was the same when either red light or white light was used for illumination (Boerner et al., 1993). The line shape and g-value of the new signal observed in these three mutants are incompatible

with its origin being any known prosthetic group in the reaction center. Therefore, it was proposed that the signal originates from an oxidized amino acid residue (M). Recent work has shown that is a tyrosine radical with an unusual structure (Boerner and Barry, 1994). This work shows that the lineshape of the radical is altered when is incorporated into PS II particles. However, the 7 G singlet obtained when the and tyrosine radicals are deuterated is not observed. Instead, a narrow signal (11 G) is obtained that still exhibits small hyperfine splitting. One interpretation of these results is that PS II contains a covalently modified tyrosine that can become redox active. Covalently modified amino acid residues are involved in electron transfer in other proteins (Janes et al., 1990; Ito et al., 1991; McIntire et al., 1991). The role of the modified tyrosine residue in electron transfer reactions in PS II remains to be elucidated. Recently, it has been shown that the control EPR spectrum is not observed when tyrosine, leucine, or aspartate substitutions are made at histidine 190 in D2 (Tommos et al., 1993; Tang et al., 1993a). Instead, a narrow, structureless EPR signal is observed. This signal was assigned to an altered form of tyrosine radical, although isotopic labeling was not performed to confirm this assignment. Substitutions

Chapter 8 Photosystem II at glutamine 164 of D2 slightly altered the EPR signal. These results may imply that the side chains of histidine 190 and glutamine 164 are in the immediate vicinity of redox active tyrosine D (Tommos et al., 1993; Tang et al., 1993a). Models of the donor side of PS II suggest that either this histidine (Svensson et al., 1990) or this glutamine (Ruffle et al., 1992) are hydrogen bond acceptors for IV. Biophysical Studies of Cyanobacterial Photosystem II

A. Tyrosine Radical, has a Slightly Different EPR Lineshape in Plants and Cyanobacteria The EPR lineshape of a tyrosine radical is sensitive to the environment of the redox-active species. A comparison of the EPR spectrum of and the tyrosine radical in ribonucleotide reductase (RDPR) illustrates this. The EPR spectrum of RDPR is a doublet with hyperfine splittings on the order of 20 G. On the other hand, the spectrum is a singlet with a peak to trough splitting of approximately 20 G. Several studies have shown that changes in either the spin-density distribution or the dihedral angle at the bond can alterthe EPR lineshape (reviewed by Barry, 1993; in the numbering scheme used here, is the ring carbon bound to the phenol oxygen). An ENDOR study of the radical in RDPR has shown that the spin-density in this radical is mainly located at The spectrum is dominated by a single coupling to one methylene proton, which has a dihedral angle of 33° with respect to the orbital. This deduced geometry at was found to agree with the X-ray diffraction structure ofthe B2 subunit of RDPR (Bender et al., 1989; Nordlund et al., 1990). An EPR study of the tyrosine radical in PS II was performed through the use of specifically deuterated tyrosines (Barry et al., 1990). This study, conducted with intact cells of Synechocystis sp. strain PCC 6803, assumed that the spin-density distributions in the RDPR and the radicals are similar. With this assumption, the difference in lineshape between RDPR and was explained by proposing that there is a 20° change in the dihedral angle between a methylene proton and the orbital of when the two tyrosine radicals are compared (Barry et al., 1990). These conclusions are in agreement with an

239 ENDOR study of the radical in spinach PS II membranes, which gave a 10 G coupling to one of the methylene protons (Rodriguez et al., 1987). A different conclusion has been reached by (Hoganson and Babcock, 1992). ENDOR measurements were conducted on specifically deuterated in Synechocystis sp. strain PCC 6803 PS II particles. This work showed that the coupling to one strongly coupled methylene proton is 8 G, not 10 G, as previously deduced from ENDOR studies ofspinach PS II membranes (Rodriguez et al., 1987). The results of Hoganson and Babcock also predict that the 3, 5 coupling is slightly different in spinach and cyanobacteria. This interpretation has been questioned recently (Rigby et al., 1994). At minimum, these results predict that should have a slightly different EPR lineshape in spinach and cyanobacteria. This is in fact the case, as illustrated in Fig. 6, which shows the EPR spectrum of in spinach PS II complexes and PS II particles from Synechocystis sp. strain PCC 6803.

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B. Difference FT-IR Studies of the Redox Active Tyrosine Residues in Photosystem II Cyanobacteria can be used to perform isotopic labeling experiments in photosynthetic proteins. This capability has been exploited in magnetic resonance studies (see Barry and Babcock, 1987; DeRose et al., 1991; Tang et al., 1993b, 1994). Another important use of isotopic substitution is in the assignment of vibrational lines, since vibrational spectroscopy is of great utility in studying structural changes in proteins. Difference or ‘reaction-induced’ FT-IR spectroscopy can be used to observe structural changes in both protein and prosthetic groups. This technique has been used to study the mechanism of proton pumping in bacteriorhodopsin (Braiman and Rothschild, 1988). When the reactions ofinterest are initiated by visible light, light-minus-dark difference spectra reflect conformational changes that accompany the light-driven events. Some of these conformational changes are likely to be ofimportance in facilitation of the electron transfer reactions. Infrared data on PS II have been reported. However, there has been no study that has combined specific isotopic labeling ofamino acids or prosthetic groups with this technique, so that definitive assignments could be made (Tavitian et al., 1986; Berthomieu et al., 1990; Nabedryk et al., 1990; Berthomieu et al., 1992; MacDonald and Barry, 1992; Noguchi et al., 1992). Difference FT-IR has recently been used to obtain structural information about the D and Z tyrosine residues in PS II (MacDonald et al., 1993). In this study, and labeling of tyrosine were used to identify vibrational lines of the tyrosine residue and the neutral tyrosineradical. The vibrational difference spectra of D and Z were found to be different from each other (Fig. 7), in particular, the spectrum exhibited vibrational mode at (positive) that was not observed in the spectrum of A difference in the strength of a hydrogen bonding interaction might explain the alterations observed. Such a hypothesis is appealing, since it could also help to account for the lower midpoint potential of D, when compared to Z (Boussac and Etienne, 1984; Metz et al., 1989). These results from infrared spectroscopy are also consistent with a recent EPR study of and in specifically deuterated PS II particles (Boerner and Barry, 1993). This EPR study has obtained evidence for structural differences between the two tyrosine radicals.

C. The Structure of the Manganese Complex in Plants and Cyanobacteria It is now widely accepted that the catalytic site for the four-electron photooxidation ofwater consists of protein-bound manganese ions, with and chloride as additional, possible co-factors (reviewed by Debus, 1992; Yocum, 1992). The structure of the manganese cluster in plant and cyanobacterial PS II preparations has been studied through the use of various spectroscopic techniques. The rationale for studies comparing the properties of plants and Cyanobacteria is three-fold. One, since both systems evolve oxygen, a discussion of the similarities and differences in the structure of the manganese cluster will elucidate those structural features that are essential for water splitting. Two, as illustrated in Section IV A and B, Cyanobacteria are ideal for spectroscopic experiments that depend on isotopic substitution for assignments. Three, as described in Section III, Cyanobacteria are

Chapter 8 Photosystem II now commonly used for site-directed mutagenesis studies that are aimed at identification of ligands to the manganese cluster. We will review magnetic resonance and X-ray absorption studies of the manganese cluster with a particular emphasis on interesting differences between cyanobacteria and plants. More complete reviews of biophysical studies of the manganese cluster can be found in (Debus (1992); Dekker, (1992); Sauer et al., (1992); and Vänngård et al.,(1992).

1. Magnetic Resonance Studies of the State Several magnetic resonance techniques can be used to investigate the structure ofthe manganese cluster inthe state. Three methods that have been applied to both plant and cyanobacterial preparations will be discussed: electron paramagnetic resonance (EPR), electron-nuclear double resonance (ENDOR), and electron spin echo envelope modulation (ESEEM).

a. Electron Paramagnetic Resonance When the state is produced by flash excitation of spinach chloroplasts, a complex EPR signal is observed (Dismukes and Siderer, 1981). This so called ‘multiline signal’ is centered at g = 2 and is comprised of more than 16 hyperfine lines that are spaced by an average of 87.5 G. These spectral characteristics are consistent with a mixed-valence complex of at least two interacting manganese ions (Dismukes and Siderer, 1981; Dismukes et al., 1982). The state multiline signal can also be produced by continuous illumination, if either the acceptor side (Hansson and Andréasson, 1982; Brudvig et al., 1983) or the donor side (Styring and Rutherford, 1988a) is limited to one turnover. For example, continuous illumination at 200 K produces a multiline signal, since the transition cannot proceed at this temperature (Styring and Rutherford, 1988a). As originally suggested (Dismukes and Siderer, 1981; Dismukes et al., 1982), the multiline EPR signal is now thought to arise from a spin 1/2 ground state of a mixed-valence multinuclear Mn complex (see Aasa et al., 1987; Hansson et al., 1987; Britt et al., 1992). A multiline spectrum of plant PS II membranes has been obtained at S-band (3.9 GHz) (Haddy et al., 1989). The results of this study also suggest that more than two interacting Mn ions give

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rise to the multiline spectrum. Oxygen evolving PS II particles from Synechococcussp. strainPCC6301 exhibita state multiline EPR signal that is very similar to the signal observed inplantPS IImembranes (Aasaetal., 1987). Multiline signals have also been observed in other cyanobacterial preparations (see McDermott et al., 1988; Noren et al., 1991; Kirilovsky et al., 1992; Nilsson et al., 1992; Tang et al., 1994). Since this signal is expected to be sensitive to the structure of the manganese cluster, these data argue that, in this form of the state, the metal cluster is similar in cyanobacteria and plants. However, the temperature at which the maximal multiline signal is formed has been reported to be higher for S. elongatus PS II than for spinach PS II membranes (McDermott et al., 1988). It was not determined whether this is a donor or acceptor-side phenomenon. Also, in the same study, it was argued that a high concentration of glycerol is necessary in order to observe the cyanobacterial multiline signal (McDermott et al., 1988). On the other hand, in plant preparations, the multiline signal can be observed in a variety of cryoprotectants (Zimmermann and Rutherford, 1986). Another EPR signal can also be observed from the state (Casey and Sauer, 1984; Zimmermann and Rutherford, 1984). This signal, with a single turning point at g = 4.1, is produced by continuous illumination of spinach PS II membranes at 130 – 140 K or by flash excitation. If the 130 – 140 K illuminated sample is incubated at200 K, the multiline signal appears in accord with the disappearance of the g = 4.1 signal (Casey and Sauer, 1984; Zimmermann and Rutherford, 1984; de Paula et al., 1985). The species giving rise to the g = 4.1 EPR signal is not an intermediate electron carrier between the Mn site and P680, because it is not observed duringthe transition (de Paula et al., 1985; Zimmermann and Rutherford, 1986). Under certain solvent conditions, such as the presence of sucrose in the suspension buffer, both the multiline and g = 4.1 signals can be photogenerated at 200 K in plant preparations (Zimmermann and Rutherford, 1986). In the dark and in the presence of ethylene glycol, the g = 4.1 signal is unstable between 140 and 160 K and converts to the multiline signal. This process has a negative entropy of activation (de Paula et al., 1987). This suggests that conversion of the ‘g=4.1’ state to the ‘multiline’ state involves ordering of the Mn site.

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Certain treatments that affect evolution rates also affect the equilibrium between the multiline and g = 4.1 forms ofthe state in plants (reviewed in Rutherford et al., 1992). For example, ammonia can bind to two sites near or at the evolving complex (Sandusky and Yocum, 1983, 1984). Binding of ammonia to a site at which chloride competes yields an state that exhibits the g = 4.1 EPR signal only (Beck and Brudvig, 1986; Andréasson, et al., 1988; Beck and Brudvig, 1988). Binding to a site at which chloride does not compete yields an state that shows an alteredmultiline signal. This signal exhibits 18 lines with an average hyperfine spacing of 67.5 G (Beck et al., 1986; Andréasson, et al., 1988). This effect is consistent with binding of to the multinuclear manganese complex (Beck et al., 1986; Andréasson, et al., 1988; Beck and Brudvig, 1988). Spin echo measurements have shown definitively that binds to manganese (Britt et al., 1989; see Section IV C, 1b). This altered multiline EPR signal can also be observed in cyanobacterial PS II particles (Aasa et al., 1987; Kirilovsky, et al., 1992). The frequency dependence of the g = 4.1 EPR signal indicates that it arises from a spin 5/2 spin state (Haddy et al., 1992; Vänngård et al., 1992; but see Pace et al., 1991), which is consistent with either a Mn(II) monomer or a Mn multimer. Moreover, EPR spectroscopy of oriented spinach PS II membranes treated with has shown that the g = 4.1 signal has a complex nuclear hyperfine pattern (Kim et al., 1990; Kim et al., 1992). From the number of lines and the magnitude of the observed hyperfine splittings, Kim et al.( 1990) argued that the g = 4.1 signal originates from a tetranuclear Mn complex. Magnetic susceptibility measurements also provide evidence that a multinuelear Mn complex gives rise to the g = 4.1 signal (Baumgarten et al., 1990). Taken together, the recent EPR evidence on the multiline and g = 4.1 signals suggests that both signals originate from the same site, the tetranuclear Mn complex. The two signals do not arise from the same spin manifold. Instead, these two forms of the state arise from different coupling configurations of the site. These two configurations differ in the strengths of the magnetic interactions between the Mn ions, such that a spin 5/2 state gives rise to the g = 4.1 signal and a spin 1/2 state gives rise to the multiline signal (de Paula et al., 1986a; Zimmermann and Rutherford, 1986; Hansson et al., 1987; Kim et

al., 1990; Haddy et al., 1992; Kim et al., 1992). When the 18- and 24-kDa extrinsic proteins are removed from spinach PS II membranes by NaCl washes, an effect on low temperature donation from the manganese cluster is observed (de Paula et al., 1986b). In this study, DCMU was used to limit the reaction center to a single turnover. A normal multiline could be generated by illumination at 200 K. (The authors concluded that this signal was generated in centers that retained calcium. It should be noted that the effect of calcium depletion on the S state transitions is controversial. For discussion, see Debus, 1992 and Yocum, 1992.) In untreated spinach PS II membranes, high potential cytochrome is oxidized by illumination at temperatures from 77K to 130K. In 18- and 24-kDa depleted samples, a chlorophyll radical is generated by illumination at these temperatures, since cytochrome is low potential and thus is already oxidized (de Paula et al., 1986b). Interestingly, illumination at 130K did not generate the g = 4.1 EPR signal, but a chlorophyll radical instead. Also, warming of this 130 K illuminated sample to 200 K did not generate the multiline signal (de Paula et al., 1986b). To explain these results, it was proposed that removal ofthe 18and 24-kDa proteins has an effect on the relative rate of electron transfer from chlorophyll and to Experiments on NaCl washed and then 18- and 24kDa protein reconstituted preparations were not reported. These experiments would distinguish between the effects of extrinsic polypeptide removal and other irreversible changes caused by manipulation of the sample through NaCl washing. Cyanobacterial PS II preparations lack the 18- and 24-kDa extrinsic proteins, so it might be expected that similar behavior would be seen in these preparations. Indeed, 140 K illumination of PS II particles from S. elongatus did not produce a g = 4.1 EPR signal (McDermott et al., 1988). Also, warming of 140 K illuminated samples did not produce the multiline spectrum. However, unlike the situation in spinach PS II membranes, no other donor side EPR signals from either chlorophyll or cytochrome were observed upon 140 K illumination of the cyanobacterial PS II particles (A signal at g = 1.6 was observed that was assigned to a perturbed iron quinone signal from the acceptor side.). Significantly, XANES studies showed that oxidation of manganese does occur when illumination is performed at this temperature, although neither the multiline or the g = 4.1 EPR signal could be observed (McDermott et

Chapter 8 Photosystem II al., 1988; see Section IV C, 2). Since chlorophyll was oxidized in this temperature range in 18- and 24kDa-depleted spinach preparations, the results of McDermott et al. (1988) suggest a difference in the kinetics of low temperature donation to when plants and cyanobacteria are compared. Alternatively, the results of de Paula et al. (1986b) may have been caused by some other effect of NaCl washing, besides 18- and 24-kDa protein removal, as previously suggested (McDermott et al., 1988). The data of McDermott et al. (1988) also imply that an EPR silent form of the state can be generated in cyanobacteria under these conditions. Thus, the behavior of this cyanobacterial preparation is not directly comparable to spinach preparations from which the 18- and 24-kDa extrinsic proteins have been removed. To date, the g = 4.1 signal has not been observed in any cyanobacterial preparation (Aasa et al., 1987; Kirilovsky, et al., 1992). It should be noted that, in most attempts to generate the signal, glycerol was used as the cryoprotectant, and little is known about the effects of cryoprotectants on signals in cyanobacterial PS II preparations.

b. Electron Spin-Echo Envelope Modulation Conventional EPR fails to give unambiguous information about the nature ofthe protein ligands to Mn in the evolving complex. This is because the nuclear ‘superhyperfine’ interactions arising from coupling of the electron spin on Mn with the ligand nuclear spin are too small to be resolved in a conventional EPR spectrum (Sauer et al., 1992). However, pulsed EPR techniques, particularly electron spin-echo envelope modulation (ESEEM), are capable of measuring superhyperfine couplings in the MHz scale, as might be expected from Ncontaining ligands to transition metal complexes. A general description of the technique, with applications to the evolving complex is given in (Sauer et al., 1992). ESEEM is particularly effective in characterizing interactions between a paramagnet and a nucleus containing a quadrupole moment, such as In such experiments, the assignments tometal-nitrogen interactions are made by comparing results in a samplecontainingthe naturally abundant which has a nuclear spin I=1, with a sample enriched in The ESEEM peaks in the sample reveal the nuclear hyperfine coupling, A, to the paramagnet.

243 The first application ofESEEM to the study ofthe Mn site was described by Britt et al. (1989). These workers probed the state of PS II membranes treated with and The spectra showed the presence of modulations by with A = 2.29 MHzandby with A = 3.22 MHz. This experiment definitively demonstrates that binds directly to the Mn site in the state (Britt et al., 1989). The ESEEM spectrum of the state in PS II particles from S. elongatus shows a peak at 4.8 MHz (DeRose et al., 1991). Spinach PS II membranes show a similar feature in the ESEEM spectrum (Britt et al., 1989). The 4.8 MHz peak was not observed in PS II particles from a S. elongatus culture grown on a medium. This isotopic labeling experiment provides evidence that the 4.8 MHz peak arises from modulations by a species near the Mn site. From chemical arguments, the first-coordination sphere ofMn is expected to consist ofmostly oxygen containing ligands (Pecoraro, 1988; Larson and Pecoraro, 1992). However, the ESEEM results suggest that at least one histidine residue is a ligand to manganese in the state in both plants and cyanobacteria. Recent work has shown that a 5 MHz peak in the ESEEM spectrum can be assigned to histidine. The magnitude of the hyperfine coupling provides evidence that one or both imidazole nitrogens are ligands. The number of ligands could not be estimated (Tang et al., 1994).

c. Electron-Nuclear Double Resonance ESEEM cannot detect hyperfine couplings to a nucleus without a quadrupole moment. This limitation prevents the analysis of proton couplings to the Mn site of the evolving center by ESEEM. Another magnetic resonance technique, electron-nuclear double resonance (ENDOR), can circumvent this limitation. In an ENDOR experiment, the EPR transition is saturated with large amounts of microwave power and a scanning radio-frequency field probes the sample. One observes ‘NMR’ transitions of the nuclei coupled to the electrons associated exclusively with the saturated EPR transition. Therefore, like ESEEM, ENDOR can give information about ligands in paramagnetic transition metal complexes. The advantage of ENDOR lies in the ability to detect nuclear couplings from a host of nuclei, including and (for a

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general treatment, see Wertz and Bolton, 1986). Two groups have used ENDOR to probe the ligand environment ofthe Mn site in the state (Kawamori et al., 1989; Tang et al., 1993b). Kawamori et al. have reported the ENDOR of the state multiline EPR signal in spinach PS II membranes, while Tang et al. have reported both and ENDOR of PS II particles from S. elongatus. Kawamori et al. observed six pairs of resonances that they attributed to protons coupled to the Mn site in the state: 0.37,0.69, 1.07, 1.41, 2.01, and 4.02 MHz. Two of these protons (at 2.01 and 4.02 MHz) were exchangeable with the aqueous environment, as determined by studies performed in Frequency shifts were observed when samples were suspended in a glycerol/sucrose containing buffer, giving resonances at: 0.53, 0.76, 1.19, 1.44, 2.41, 4.02 MHz. In contrast, Tang et al. observed only four pairs of resonances, at 0.5, 1.0, 2.4, and 4.9 MHz when cyanobacterial samples were suspended in glycerol; all were exchangeable. Since Tang et al. observe resonances at 0.75, 1.38, and 4.11 MHz that they assign to cyanobacterial they propose that the additional resonances at 0.76,1.44, and 4.02 MHz in the study of Kawamori et al. are resonances of spinach In addition to the 0.76, 1.44, 4.02 MHz resonances that are unique to the Kawamori et al. study, there are other discrepancies between the two groups. For example, Tang et al observe an additional resonance at 4.9 MHz, and the two groups observe differences in the effects of exchange. Experimental differences may account for some of these discrepancies. Tang et al. carried out exchange under illumination, while Kawamori et al. did not. Also, the two groups recorded spectra at different temperatures. Another experimental difference between the two studies, is that Tang et al., but not Kawamori et al., employed ‘thermal cycling’ of the samples to decrease the contribution of to the spectrum. At this point, it is not clear ifexperimental differences account for the discrepancies or if a significant structural distinction exists between the Mn sites of plants and cyanobacteria (Tang et al., 1993). However, if there is a significant difference in structure, it must have no large effect on the EPR lineshape ofthe plant and cyanobacterial multiline signals, which are similar. Tang et al. conclude from their ENDOR and EPR investigations that there is no evidence for

strongly coupled protons in the first coordination shell of the manganese cluster. These conclusions are consistent with ligation to Mn by largely deprotonated O-containing ligands, such as bridges and the carboxylate groups of aspartate or glutamate (Tang et al., 1993b). This conclusion has important repercussions for the mechanism of water oxidation. As pointed out by (Pecoraro, 1992), the protonation states of oxo-bridges can strongly influence the coordination and redox properties of a multinuclear manganese complex. In this regard, the possibility that the ENDOR spectrum of the multiline is different in plants and cyanobacteria is an important one and should be investigated further. Further information about the first coordination sphere ofMn in the state comes from ENDOR (Tang et al., 1993b). As proposed previously on the basis of ESEEM, ENDOR detects nitrogen resonances that are coupled to the Mn site, at 0.7 and 3.7 MHz. Tang et al., (1993b) attribute these resonances arise from one (or at most two ligands), which may be histidine residues.

2. X-Ray Absorption Studies of the S States X-ray absorption spectroscopy has the potential to answer many structural questions about the manganese cluster. The general pattern of absorption of X-rays by transition metal complexes consists of two useful features. First, a sharp rise in the absorption at a well-defined X-ray energy corresponds to removal ofan inner-shell electron. This region ofthe spectrum, called X-ray absorption near-edge structure (XANES) is characteristic of a given transition metal. Changes in the onset ofthe absorption rise can be correlated to changes in the oxidation state of the metal, in the electron density ofthe metal, and in the symmetry of the metal’s first coordination sphere. Most studies are performed in the K-edge region, which corresponds to promotion of a 1s electron of the metal to a higher orbital. Second, there are periodic modulations in the spectrum that arise from backscattering of photoelectrons by the immediate molecular environment of the transition metal. Fourier analysis ofthis region results in an X-ray absorption extended fine structure (EXAFS) spectrum that can determine the chemical nature and numbers of neighboring atoms (Sauer et al., 1992). XANES has the potential of determining the oxidation states of Mn ions in the evolving complex. Such studies have been performed in

Chapter 8 Photosystem II spinach chloroplasts, spinach PS II membranes, and spinach PS II core particles (Kirby et al., 1981; Goodin et al., 1984; Yachandra et al., 1986; Cole et al., 1987; Yachandra et al., 1987; George et al., 1989; Kusunoki et al., 1990; Penner-Hahn et al., 1990; MacLachlan et al., 1992; Ono et al., 1992; Riggs et al., 1992; Yachandra et al., 1993). The position and the shape of the Mn K-edge of spinach PS II membranes in the state can be fit to two Mn (III) and two Mn(IV) ions (Riggs et al., 1992; Yachandra et al., 1993). The state has also been studied in PS II particles from cyanobacteria (McDermott et al., 1988; Yachandra et al., 1993). While no major differences were observed, a small shift in the K-edge inflection energy was found upon comparison ofthe spectra of plant PS II membranes with S. elongatus PS II particles (McDermott et al., 1988; Sauer et al., 1992). This result suggests that there are no large structural differenceswhenthetwometalclusters are compared; but there may be a small difference in the ligand environment ofthe metal cluster (McDermott et al., 1988; Sauer et al., 1992; Yachandra et al., 1993). The edge position shifts to higher energy upon formation of the state in both plants and cyanobacteria (Goodin et al., 1984; Yachandra et al., 1987; McDermott et al., 1988; Kusonoki et al., 1990; MacLachlan et al., 1992; Ono et al., 1992). The magnitude of the change is similar. This edge shift has been attributed to the oxidation of Mn (III) to Mn (IV) (reviewed in Sauer et al., 1992). Ultraviolet absorption studies and NMR studies agree with the conclusion that the transition involves manganese oxidation (reviewed in (Dekker, 1992; Sharp, 1992). Shifts to higher energy occur upon illumination ofthe state at either 140 K and at 190 K, suggesting that the Mn site is oxidized to the same extent in both in the ‘g = 4.1’ and ‘multiline’ forms of the state (Cole et al., 1987; Yachandra et al., 1987). Cyanobacterial PS II behaved similarly upon illumination at the two temperatures (McDermott et al., 1988). This is of significance since the g = 4.1 signal has not been observed in cyanobacteria, and no other EPR signal from oxidized donor side was observed instead of the g = 4.1. This result implies that some forms of the state are EPR silent in cyanobacteria (McDermott et al., 1988). The transition is more controversial. When the S-states are advanced by flash excitation at 10 °C and then rapidly frozen, the Mn K-edge of the state is observed to be at a higher energy than that

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ofthe state (Ono et al., 1992). Ono et al conclude that the simplest possible interpretation of their data is that Mn is oxidized during the transition. However, since other factors besides oxidation state affect the K-edge energy, the authors cannot absolutely eliminate the possibility that the observed edge shift arises from configurational rearrangements of the manganese (Ono et al., 1992). Interestingly, the result of Ono et al. (1992) is not in agreement with K edge studies in which the state is trapped under continuous illumination at lower temperatures (Guiles et al., 1990b). In this work, no change in the K-edge was observed upon the transition. The amount of was monitored by observation of the disappearance of the multiline signal. The result of Guiles et al. (1990b) suggests that Mn is not oxidized during the under these conditions, but that the oxidizing equivalent resides on a site close to Mn (Sauer et al., 1992). The reason for the discrepancy between the Ono et al. (1992) and the Guiles et al. (1990b) results is not clear at this time. The discrepancy could be due to the difference in the illumination temperature. Contamination ofthe state with other S states is possible in the continuous illumination experiment. Resolution of this discrepancy is of great importance, since other spectroscopic techniques have also argued against oxidation of manganese on this transition. For example, it is has been suggested that a histidine may be oxidized upon the transition, but an EPR signal attributable to this spin center is observed only after treatments that inhibit oxygen evolution (reviewed in (Rutherford et al., 1992). Also, although the optical studies described in (Dekker et al., 1984a; Dekker et al., 1984b; Dekker et al., 1984c; Saygin and Witt, 1987; Kretschmann et al., 1988) support oxidation ofmanganese on the transition, the optical studies of Lavergne have suggested that manganese may not be oxidized on this transition (Lavergne, 1991, and references therein). NMR proton spin relaxation experiments also suggest that there is no metal oxidation upon this transition (Srinivasan and Sharp, 1986a; Srinivasan and Sharp, 1986b), as do studies of the EPR relaxation properties of (Styring and Rutherford, 1988b; Evelo et al., 1989). The state has also been probed by X-ray absorption spectroscopy. When the S-states are advanced by flash excitation at 10 °C and then rapidly frozen, the Mn K-edge of the state is observed to be at a lower energy than that of the state (Ono

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et al., 1992). This change is consistent with a two or three electron reduction of the manganese cluster. The simulated data show that the K-edge energy shifts up by 1 eV on the transition. The magnitude of this edge shift is similar to the changes observed upon the and the transitions in the same study (Ono et al., 1992). It should be noted that the conclusion that manganese is oxidized on this transition is in agreement with NMR and EPR studies (Srinivasan and Sharp, 1986a; Srinivasan and Sharp, 1986b; Styring and Rutherford, 1988b; Evelo et al., 1989). Ultraviolet absorption studies disagree on the spectral characteristics associated with the transition (reviewed in Dekker, 1992). Ono et al. (1992) attribute the K-edge change on the transition to oxidation of Mn (III) to Mn (IV), An attempt has been made to trap the state for spectroscopic study (Guiles et al., 1990a). Because the state is oxidized in the dark to the state by (Styring and Rutherford, 1987), it is difficult to trap a sample that is 100% unless chemical treatments are used. In the Guiles et al. (1990a) study, spinach PS II membranes were treated with hydroxylamine at low concentrations. At low concentrations, hydroxylamine is known to reduce the manganese cluster by two electrons. At high concentrations, hydroxylamine leads to the irreversible loss of manganese from the reaction center (reviewed in Mei and Yocum, 1992). In the XANES study of Guiles et al.( 1990a), the position ofthe K-edge ofthe dark-adapted, hydroxylamine-treated sample was similar, but not identical, to that of an untreated sample poised in the state. The small difference was attributed to the irreversible release ofmanganese in a small number of centers (Guiles et al., 1990a). Upon illumination of the hydroxylamine-treated sample, the Mn K-edge shifted to lower energy. The interpretation of these results was that the state was formed during illumination and then reduced by hydroxylamine. The new state, is believed to be akin to the state. The change in K edge energy when the is compared to the state was comparable to the shift observed by the same group upon the transition (Guiles et al., 1990a). The change upon the has been attributed to the oxidation of Mn (II) (reviewed in Sauer et al., 1992). A study of hydroxylamine effects on spinach PS II core particles, which lack the 18- and 24-kDa extrinsic proteins, gives a different result (Riggs et

al., 1992). In this work, samples were treated with either the reductant, hydroquinone, or the reductant, hydroxylamine, in the presence ofcalcium. Previous work has shown that both hydroxylamine and hydroquinone reversibly reduce the manganese cluster under these conditions (Mei et al., 1989; Mei and Yocum, 1991). Also, hydroxylamine and hydroquinonewereshowntoreducedistinctsubpopulations of manganese. K-edge shifts to lower energy were observed in the dark states of both hydroxylamine-treated and hydroquinone-treated samples, relative to untreated controls. The shifts were reversed upon illumination and subsequent dark adaptation (Riggs et al., 1992). The conclusion of this study is that hydroxylamine and hydroquinone both reduce the Mn site in the dark. However, the hydroxylamine-reduced and hydroquinone-reduced states differ in average Mn oxidation (Riggs et al., 1992). These results are in agreement with the earlier conclusions of Mei and Yocum on the basis ofbiochemical characterization and EPR spectroscopy. However, the conclusion that hydroxylamine reduces the manganese cluster in the dark is at variance with the earlier work in which thehydroxylamine-reduced state was believed to be similar to the state (Guiles et al., 1990a). Riggs et al. (1992) give two possible reasons for the discrepancy between their result and that of the Guiles et al. study. Firstly, the difference may be due to the fact that the preparations of Riggs et al. lack the 18- and 24-kDa polypeptides, while the preparations of Guiles et al. do not. Secondly, Riggs et al. propose that part of the difference could be due to interpretation of the data, since Guiles et al. observed small edge-shifts in the dark. However, Guiles et al. attributed the change to irreversible loss of Mn (II). It would be interesting to pursue this question with plant core preparations that retain the 18- and 24-kDa (see Section II D, 1), since salt washed and untreated samples could then be directly compared. Measurements on cyanobacterial PS II preparations would also be of great interest. EXAFS analyses ofthe oxygen-evolving complex have been performed in spinach chloroplasts, spinach PS II membranes, and spinach PS II core particles (Kirby et al., 1981;Goodin et al., 1984; Yachandra et al., 1986; Cole et al., 1987; Yachandra et al., 1987; George et al., 1989; Corrie et al., 1990; Kusunoki et al., 1990; Penner-Hahn et al., 1990; MacLachlan et al., 1992; Yachandra et al., 1993). PS II particles from the cyanobacterium S. elongatus have also

Chapter 8 Photosystem II been studied, and no major differences between plants and cyanobacteria have been observed (McDermott et al., 1988; Yachandra et al., 1993). This area has been reviewed recently (Sauer et al., 1992; Debus, 1992).

V. Concluding Remarks The structure and function of PS II are rapidly being elucidated. The progress that has recently been made in this area is due to the interdisciplinary nature of the approaches in use. In particular, cyanobacteria have been of tremendous benefit, since isotopic labeling and site-directed mutagenesis can both be performed through the use of this organism. Although cyanobacteria are increasingly used as a model system for plant water oxidation, we have described some differences between the two systems. For example, the polypeptide components are not identical, and some subunits may play slightly different roles in plant and cyanobacterial PS II. There is some indication that the redox active tyrosine radicals are in different environments in plants and cyanobacteria. Also, a difference in low-temperature donation to has been described. A small difference in inflection energy was observed in the X-ray absorption K-edge spectrum when the state of plants and cyanobacteria were compared. This change was attributed to a small change in the ligand environment ofthe manganese cluster. Interestingly, the g = 4.1 signal from the state of manganese cluster has never been observed in cyanobacteria. There is also good evidence for an EPR silent form of the state in cyanobacteria, and recent ENDOR studies may be consistent with a structural difference between the states of plants and cyanobacteria. Since both complexes are able to efficiently carry out water oxidation, the fact that the plant and cyanobacterial systems are slightly different makes comparative studies ofgreat value. Such comparative studies will elucidate the minimum requirements for oxygen evolution.

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depletion and F- substitution: Oxidation of a protein residue. Biochemistry 29, 10814–10822 Beck WF and Brudvig GW (1986) Binding of amines to the evolving center of Photosystem II. Biochemistry 25: 6479– 6486 Beck WF and Brudvig GW (1988) Ligand substitution reaction of the center of Photosystem II. Chemica Script a 28A: 93–98 Beck WF, de Paula JC and Brudvig GW (1986) Ammonia binds to the manganese site of the complex of Photosystem II in the state. J Am Chem Soc 108: 4018– 4022 Bender CJ, Sahlin M, Babcock GT, Barry BA, Chandrashekar TK, Salowe SP, Stubbe JA, Lindstrom B, Petersson L, Ehrenberg A and Sjoberg B-M (1989) An ENDOR study of the tyrosyl free radical in ribonucleotide reductase from Escherichia coli. J Am Chem Soc 111: 8076–8083 Bennett J (1979) Chloroplast phosphoproteins, the protein kinase of the thylakoid membranes is light-dependent. FEBS Lett 103:342–344 Berthomieu C, Nabedryk E, Mantele W and Breton J (1990) Characterization by FTIR spectroscopy of the photoreduction of the primary quinone acceptor in Photosystem II. FEBS Lett 269: 363–367 Berthomieu C, Boussac A, Mantele W, Breton J and Nabedryk E (1992) Molecular changes following oxidoreduction of cytochrome characterized by Fourier transform infrared difference spectroscopy and electron paramagnetic resonance: Photooxidation in Photosystem II and electrochemistry of isolated cytochrome and iron protoporphyrin IXbisimidazole model compounds. Biochemistry 31: 11460– 11471 Blankenship RE, Babcock GT, Warden JT and Sauer K (1975) Observation of a new EPR transient in chloroplasts that may reflect the electron donor to Photosystem II at room temperature. FEBS Lett 51: 287–293 BlubaughDJandCheniaeGM(1990)Kineticsofphotoinhibition in hydroxylamine-extracted Photosystem II membranes: Relevance to photoactivation and sites of electron donation. Biochemistry 29: 5109–5118 Bockholt R, Masepohl B and Pistorius EK (1991) Insertional inactivation of the psbO gene encoding the manganese stabilizing protein of Photosystem II in the cyanobacterium Synechococcus PCC7942. FEBS Lett 294: 59–63 Boerner RJ and Barry BA (1993) Isotopic labeling and EPR spectroscopy show that a tyrosine residue is the terminal electron donor, Z, in manganese-depleted Photosystem II preparations. J Biol Chem 268: 17151–17154 Boerner RJ and Barry BA (1994) EPR evidence that the radical, which is observed in three site-directed mutants of Photosystem II, is a tyrosine radical. J Biol Chem 269: 134– 137 Boerner RJ, Nguyen AP, Barry BA and Debus RJ (1992) Evidence from directed mutagenesis that aspartate 170 of the D1 polypeptide influences the assembly and/or stability of the manganese cluster in the photosynthetic water-splitting complex. Biochemistry 31: 6660–6672 Boerner RJ, Bixby KA, Nguyen AP, Noren GH, Debus RJ and Barry BA (1993) Removal of stable tyrosine, affects the structure or redox properties of tyrosine Z in manganesedepleted Photosystem II particles from Synechocystis 6803. J

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Chapter 8 Photosystem II Biochemistry 30: 440–446 Burnap R, Koike H, Sotiropoulou G, Sherman LA and Inoue Y (1989) Oxygen evolving membranes and particles from the transformable cyanobacterium Synechocystis sp. PCC 6803. Photosynth Res 22: 123–130 Burnap R, Webb R and Sherman LA (1990) Progress towards the production and analysis of splitting mutants of the cyanobacterium Synechocystis sp. PCC 6803. In: Baltscheffsky M (ed) Current Research in Photosynthesis, Vol I, pp 255– 258. Kluwer Academic Publishers, Dordrecht Burnap RL, Shen J-R, Jursinic PA, Inoue Y and Sherman LA (1992) Oxygen yield and thermoluminescence characteristics of a cyanobacterium lacking the manganese-stabilizing protein of Photosystem II. Biochemistry 31: 7404–7410 Buser CA, Thompson LK, Diner BA and Brudvig GW (1990) Electron-transfer reactions in manganese-depleted Photosystem II. Biochemistry 29: 8977–8985 Buser CA, Diner BA and Brudvig GW (I992a) Photooxidation of cytochrome in oxygen-evolving Photosystem II. Biochemistry 31: 11449–11459 Buser CA, Diner BA and Brudvig GW (1992b) Reevaluation of the stoichiometry of cytochrome in Photosystem II and thylakoid membranes. Biochemistry 31: 11441–11448 Casey JL and Sauer K (1984) EPR detection of a cryogenically photogenerated intermediate in photosynthetic oxygen evolution. Biochim Biophys Acta 767: 21–28 Chen JC, Meng BY, Fukuta M and Sugiura M (1990) Nucleotide sequence of the psbI gene of the cyanobacterium, Anacystis nidulans 6301. Nucl Acids Res 18: 4017 Chu H-A, Nguyen AP and Debus RJ (1993) Residues of the D1 polypeptide that influence the assembly or stability of the manganese cluster or the binding ofcalcium in Photosystem II. Biophys J 64: A216 Cohn CL, Sprinkle JR, Alam J, Hermodson M, Meyer T and Krogmann DW (1989) The amino acid sequence of lowpotential cytochrome from the cyanobacterium Microcystis aeruginosa. Arch Biochem Biophys 270: 227–235 Cole J and Sauer K. (1987) The flash number dependence of EPR Signal II decay as a probe for charge accumulation in Photosystem II. Biochim Biophys Acta 891: 40–48 Cole, J, Yachandra, VK, Guiles, RD, McDermott, AE, Britt, RD, Dexheimer, SL, Sauer, K and Klein, MP (1987) Assignment of the g 4.1 EPR signal to manganese in the state of the photosynthetic oxygen-evolving comples: An X-ray absroption edge spectroscopy study. Biochim Biophys Acta 890: 395– 398 Cole JL, Yachandra VK, McDermott AE, Guiles RD, Britt RD, Dexheimer SL, Sauer K and Klein MP (1987) Structure of the manganese complex of Photosystem II upon removal of the 33-kilodalton extrinsic protein: An X-ray absorption spectroscopy study. Biochemistry 26: 5967–5973 Conjeaud H and Mathis P (1980) The effect of pH on the reduction kinetics of P-680 in Tris-treated chloroplasts. Biochim Biophys Acta 590: 353–359 Corrie AR, Evans MCW, Hubbard JAM and Hasnain SS (1990) An EXAFS study of the manganese oxygen-evolving complex of Photosystem 2. In: Baltscheffsky M (ed) Current Research in Photosynthesis, Vol 1, pp 793–796. Kluwer, Dordrecht Cramer WA and Whitmarsh J (1977) Photosynthetic cytochromes. Ann Rev Plant Physiol 28: 133–72 Cramer WA, Theg SM and Widger WR (1986) On the structure

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Nordlund P, Sjoberg B-M and Eklund H (1990) Three-dimensional structure of the free radical protein of ribonucleotide reductase. Nature 345: 593–598 Noren GH and Barry BA (1992) The YF161D1 mutant of Synechocystis 6803 exhibits an EPR signal from a lightinduced Photosystem II radical. Biochemistry 31: 3335–3342 Noren GH, Boerner RJ and Barry BA (1991) EPR characterization of an oxygen-evolving Photosystem II preparation from the cyanobacterium, Synechocystis 6803. Biochemistry 30:3943– 3950 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 Oh-oka H, Tanaka S, Wada K, Kuwabara T and Murata N (1986) Complete amino acid sequence of 33-kDa protein isolated from spinach Photosystem II particles. FEBS Lett 197: 63–66 Ono T and Inoue Y (1983) Mn-preserving extraction of 33, 24 and 16-kDa proteins from Photosystem II particles by divalent salt-washing. FEBS Lett 164: 252–260 Ono T and Inoue Y (1984) restoration of evolving activity in Photosystem II particles depleted of 33, 24, and 16-kDa proteins. FEBS Lett 168: 281–286 Ono T and Inoue Y (1985) S-state turnover in the system of Photosystem II particles depleted of three peripheral proteins as measured by thermoluminescence. Removal of 33-kDa protein inhibits to transition.Biochim Biophys Acta 806: 331–340 Ono T-A, Noguchi T, Inoue Y, Kusunoki M, Matsushita T and Oyanagi H (1992) X-ray detection of the period-four cycling of the manganese cluster in photosynthetic water oxidizing enzyme. Science 258: 1335–1337 Pace RJ, Smith P, Bramley R and Stehlik D (1991) EPR saturation and temperature dependence studies on signals from the oxygenevolving center of Photosystem II. Biochim Biophys Acta 1058: 161–170 Packham NK (1988) Is the 9-kDa thylakoid membrane phosphoprotein functionally and structurally analogous to the ‘H’ subunit of bacterial reaction centres? FEBS Lett 231: 284– 290 Pakrasi HB and Sherman LA (1984) A highly active oxygenevolving Photosystem II preparation from the cyanobacterium Anacystis nidulans. Plant Physiol 74: 742–745 Pakrasi HB and Vermaas WFJ (1992) Protein engineering of Photosystem I I . In: Barber J (ed)The Photosystems: Structure, Function and Molecular Biology, Vol 11,pp 231–258. Elsevier Science Publishers, Amsterdam 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 of cytochrome in Photosystem II. EMBO 7: 325–332 Pakrasi H, Nyhus K and Granok H (1990) Targeted deletion mutagenesis of the beta subunit of cytochrome protein destabilizes the reaction center of Photosystem II. Z Naturforsch 45c: 423–429 Pakrasi HB, Ciechi PD and Whitmarsh J (1991) Site directed mutagenesis of the heme axial ligands of cytochrome b559 affects the stability of the Photosystem II complex. EMBO 10: 1619–1627 Pecoraro VL (1988) Structural proposals for the manganese

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(1988a) Characterization of the D1 protein in a Photosystem II mutant (LF-1) of Scenedesmus obliquus blocked on the oxidizing side. FEBS Lett 235: 109–116 Taylor MA, Packer JCL and Bowyer JR (1988b) Processing of the Dl polypeptide of the Photosystem II reaction center and photoactivation of a low fluorescence mutant (LF-1) of Scenedesmus obliquus. FEBS Lett. 237: 229–233 Thompson LK, Miller A-F, Buser CA, de Paula JC and Brudvig GW (1989) Characterization of the multiple forms of cytochrome b559 in Photosystem II. Biochemistry 28: 8048–8056 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 the immediate vicinity of on the D2 protein. Biochemistry 32, 5436–5441 Trebst A (1986) The topology of the plastoquinone and herbicide binding peptides of Photosystem II in the thylakoid membrane. Z Naturforsch 41c: 240–245 Vallon O, Tae G-S, Cramer WA, Simpson D, Hoyer-Hansen G and Bogorad L (1989) Visualization of antibody binding to the photosynthetic membrane: The transmembrane orientation of cytochrome b-559. Biochim Biophys Acta 975: 132–141 Van der Bolt FV and Vermaas W (1992) Photoinactivation of Photosystem II as studied with site-directed D2 mutants of the cyanobacterium Synechocystis sp. PCC 6803. Biochim Biophys Acta 1098: 247–254 van Leeuwen 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 Vänngård T, Hansson Ö and Haddy A (1992) EPR studies of manganese in Photosystem II. In: Pecoraro VL (ed) Manganese Redox Enzymes., pp 105–118. VCH Publishers, New York 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 Vass I, Ono T and Inoue Y (1987) Stability and oscillation properties of thermoluminescent charge pairs in the system depleted of or the 33-kDa extrinsic protein. Biochim Biophys Acta 892: 224–235 Vass I, Cook KM, Deak Z, Mayes SR and Barber J (1992) Thermoluminescence and flash-oxygen characterization of the IC2 deletion mutant of Synechocystis sp. PCC 6803 lacking the Photosystem II 33-kDa protein. Biochim Biophys Acta 1102:195–201 Vermaas WFJ (1993) Molecular-biological approaches to analyze Photosystem II structure and function. Ann Rev Plant Physiol Plant Mol Biol, in press Vermaas WFJ, Ikeuchi M and Inoue Y (1988a) Protein composition of the Photosystem II core complex in genetically engineered mutants of the cyanobacterium Synechocystis sp. PCC 6803. Photosynth Res 17: 97–113 Vermaas WFJ, Rutherford AW and Hansson Ö (1988b) 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, Charité J and Shen G (1990) Glu-69 of the D2 protein in Photosystem II is a potential ligand to Mn involved

Chapter 8 Photosystem II in photosynthetic oxygen evolution. Biochemistry 29: 5325– 5332 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 by a multi-gene family in pea. Plant Mol Biol 12: 439–451 Wallace TP, Stewart AC, Pappin D and Howe CJ (1989) Gene sequence for the 9-kDa component of Photosystem II from the cyanobacterium Phormidium laminosum indicates similarities between cyanobacterial and other leader sequences. Mol Gen Genet 216: 334–339 Webber AN and Gray JC (1989a) Detection of calcium binding by Photosystem II polypeptides immobilised onto nitrocellulose membrane. FEES Lett 249: 79–82 Webber AN, Packman LC and Gray JC (1989b) A 10-kDa polypeptide associated with the oxygen-evolving complex of Photosystem II has a putative C-terminal non-cleavable thylakoid transfer domain. FEES Lett 242: 435–438 Webber AN, Packman L, Chapman DJ, Barber J and Gray JC (1989c) A fifth chloroplast-encoded polypeptide is present in the Photosystem II reaction centre complex. FEES Lett 242: 259–262 Wertz JE and Bolton JR (1986) Electron spin resonance. Chapman and Hall, New York Westhoff P, Farchaus JW and Herrmann RG (1986) The gene for the 10,000 phosphoprotein associated with Photosystem II is part of the psbB operon of the spinach plastid chromosome. Curr Genet 11: 165–169 Whitmarsh J and Ort DR (1984) Stoichiometries of electron transport complexes in spinach chloroplasts. Arch Biochem Biophys 231: 378–389 Widger WR, Cramer WA, Hermodson M and Hermann RG (1985) Evidence for a hetero-oligomeric structure of the chloroplast cytochrome b-559. FEBS Lett 191: 186–190 Williams JGK (1988) Construction of specific mutations in Photosystem II photosynthetic reaction center by genetic engineering methods in Synechocystis 6803. Meth Enyzmol 167:766–778 Xu Q and Bricker TM (1993) 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, Guiles RD, McDermott A, Britt RD, Dexheimer SL, Sauer K and Klein MP (1986) The state of manganese in the photosynthetic apparatus 4. Structure of the manganese complex in Photosystem II studied using EXAFS spectroscopy. The state of the Photosystem II complex from spinach. Biochim Biophys Acta 850: 324–332 Yachandra VK, Guiles RD, McDermott AE, Cole JL, Britt RD,

257 Dexheimer SL, Sauer K and Klein MP (1987) Comparison of the structure of the manganese complex in the and states of the photosynthetic complex: An X-ray absorption spectroscopy study. Biochemistry 26: 5974–5981 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 Yamagishi A and Katoh S (1985) Further characterization of the two Photosystem II reaction center complex preparations from the thermophilic cyanobacterium, Synechoccus sp. Biochim Biophys Acta 807: 74–80 Yamagishi A and Fork DC (1987) Photoreduction of and cytochrome b-559 in an oxygen-evolving Photosystem II preparation from the thermophilic cyanobacterium Synechococcus sp. Arch Biochem Biophys 259: 124–130 Yamaguchi N, Takahashi Y and Satoh K (1988) Isolation and characterization of a Photosystem II core complex depleted in the 43-kDa-chlorophyll binding subunit. Plant Cell Physiol 29: 123–129 Yamomoto Y, Doi M, Tamura N Nishimura N (1981) Release of polypeptides from highly active Photosystem-2 preparation by Tris treatment. FEBS Lett 133: 265–268 Yamomoto Y, Shinkai H, Isogai Y, Matsuura K and Nishimura M (1984) Isolation of an Mn-carrying 33-kDa protein from an oxygen-evolving Photosystem-II preparation by phase partitioning with butanol. FEBS Lett 175: 429–432 Yerkes CT, Babcock GT, and Crofts AR (1983) A Tris-induced change in the midpoint potential of Z, the donor to Photosystem I I , as determined by the kinetics of the back reaction. FEBS Lett 158: 359–363 Yocum CF (1991) Calcium activation of photosynthetic water oxidation. Biochim. Biophys. Acta 1059: 1–15 Yocum CF (1992) The calcium and chloride requirements for photosynthetic water oxidation. In: Pecoraro VL (ed) Manganese Redox Enzymes., pp 71–84. VCH Publishers, New York Yu J and Vermaas WFJ (1993) Synthesis and turnover of Photosystem II reaction center polypeptides in cyanobacterial D2 mutants. J Biol Chem 268: 7407–7413 Zhang ZH, Mayes SR and Barber J (1990) Nucleotide sequence of the psbK gene of the cyanobacterium Synechocystis 6803. Nucl Acids Res 18: 1284 Zimmermann J-L and Rutherford AW (1984) EPR studies of the oxygen-evolving enzyme of Photosystem II. Biochim Biophys Acta 767: 160–167 Zimmermann J-L and Rutherford AW (1986) Electron paramagnetic resonance properties of the state of the oxygenevolving complex of Photosystem II. Biochemistry 25: 4609– 4615

Chapter 9 The Cytochrome

Complex

Toivo Kallas Department of Biology and Microbiology‚ University of Wisconsin-Oshkosh‚ Oshkosh‚ Wl 54901‚ USA Summary I. Introduction II. Role of the Cytochrome Complex in Cyanobacteria III. Relation to Quinol-Cytochrome c Oxidases in Chloroplasts‚ Mitochondria‚ and Other Bacteria IV. Polypeptides‚ Redox Centers‚ Substrate Binding Sites‚ and Subunit topology Complex A. Isolation and Composition of the Cytochrome B. Monomer‚ Dimer‚ Supercomplex? and Subunit IV Proteins C. The Cytochrome D. The Rieske Iron-Sulfur Protein E. The Cytochrome f Protein F. Additional Low Molecular Mass Subunits? G. In Vitro Reconstitution V. Electron and Proton Transfer Pathways A. Inhibitor Specificities B. Is There a Q-Cycle in the Cytochrome Complex? C. Role in Cyclic Electron Transport D. Are There Alternative Electron Transport Pathways in Cyanobacteria? E. Role in Redox-Sensing and Mediation of State Transitions VI. Three-Dimensional Structure and Biogenesis A. Structure Determination and Overall Topography B. Localization in Cyanobacteria and Biogenesis VII. Genetics and Mutational Analysis A. The pet Genes for Photosynthetic Electron Transport B. The Quinol-Oxidation Site C. The Quinone-Reduction Site D. The Rieske Iron-Sulfur Protein Complex from Cyanobacteria E. Prospects for Genetic Analysis of the Cytochrome VIII. Unresolved Questions and Perspective Acknowledgments References

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Summary The plastoquinol-cytochrome oxidoreductase (Cyt complex) catalyzes the rate limiting‚ quinol-oxidation step in oxygenic photosynthesis. Overall‚ it transfers electrons between the two photochemical reaction centers (PS II and PS I)‚ is required for cyclic electron flow around PS I‚ and establishes a transmembrane gradient of protons for ATP synthesis. Four polypeptides (Cyt subunit IV‚ the Rieske Fe-S protein‚ and Cyt f) encoded by the petBD and petCA operons‚ respectively‚ and four prosthetic groups (two bhemes‚ one c-type heme‚ and a 2Fe-2S center) catalyze these activities in vitro. Additional low-molecular-mass subunits may have roles in vivo.The Cyt complex in cyanobacteria provides the only known pathway for plastoquinol oxidation and appears to be indispensable for both photosynthesis and heterotrophy. This has D. A. Bryant (ed): The Molecular Biology of Cyanobacteria‚ pp. 259–317. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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precluded the propagation of inactivating mutations and complicated molecular genetic analysis. The related Cyt complex is found in all mitochondria and many bacteria. The Rieske‚ Cyt b‚ and Cyt polypeptides of complexes correspond to the Rieske‚ Cyt IV‚ and Cyt f‚ respectively‚ of Cyt complexes. The Cyt b subunit in the former has been split into separate and subunit IV polypeptides in the latter. Both complexes have binding sites for quinol oxidation or and quinone reduction or — each associated with a b-heme on opposite sides of the membrane. Modified Q-cycle models explain most of the experimental data on electron and proton transfer reactions in the Cyt complex but are more controversial in the Cyt complex. Other differences include the biophysical characteristics of prosthetic groups‚ turnover rates‚ and inhibitor specificities. The molecular basis for these differences has not been elucidated‚ although features such as the split Cyt and subunit IV proteins may be involved. Additional questions pertain to the pathway for electrons during cyclic flow‚ possible alternative electron acceptors‚ the role(s) of monomer or dimer forms in vivo‚ the role of the complex as a sensor of redox potential and mediator of state transitions‚ and mechanisms for attachment of hemes and the Rieske Fe-S center and assembly of the complex. Knowledge of the Cyt complex has been advanced greatly in recent years through intensive mutational analyses summarized in this chapter. Comparable studies have not‚ until very recently‚ been possible in the Cyt complex but will clearly help to elucidate its unique features and define molecular differences relative to the Cyt complex. Progress has also been impeded by the absence of three-dimensional structural information for either the Cyt or complexes. Very recently the structure of the water soluble‚ heme-binding domain of turnip Cyt f has been solved by X-ray crystallography at 2.8 Å resolution; the structure reveals several novel features including the use of the amino-terminus as an axial ligand for the heme. Such structures will provide a rational framework for subsequent mutational and biochemical studies and we can expect these combined approaches to begin to unravel the mysteries of membrane Cyt complexes. I. Introduction

Oxygenic photosynthesis in plant and algal chloroplasts and cyanobacteria requires three thylakoid‚ membrane-spanning protein complexes. These are the two photochemical reaction centers (Photosystem II [PS II; see Chapter 8] and Photosystem I [PS I; see Chapter 10]) and the Cyt complex (Cyt ). The latter transfers electrons between the two photosystems‚ from plastoquinol (PQH2) in the membrane to a soluble electron carrier (plastocyanin [PC] or cytochrome c553; see Chapter 12) located in the aqueous intrathylakoidal space. The Cyt complex is also required for cyclic electron flow around PS I. In these reactions the Cyt complex converts the redox potential energy of plastoquinol into a transmembrane electrochemical charge gradient of protons used for ATP synthesis

(see Chapter 11) and other energy requiring processes. The Cyt complex contains at least four essential proteins (Cyt subunit IV‚Rieske‚ andCytf‚encoded by the Photosynthetic electron transport genes‚ petB‚ petD‚ petC‚ and petA‚ respectively)‚ two b-hemes‚ a c-heme (Cyt f)‚ and a characteristic‚ high-potential 2Fe-2S center (Malkin and Aparicio‚ 1975) of the type first observed by Rieske et al. (1964). The related cytochrome bc1 complex (complex III) occurs in all mitochondria and many procaryotes including purple Photosynthetic bacteria (Hauska et al.‚ 1983). The early literature on aspects of Cyt and complexes has been reviewed by Rieske (1976)‚ Malkin and Bearden (1978)‚ Trumpower (1981)‚ Bendall (1982)‚ and Hauska et al. (1983). More recent reviews include those by Cramer et al. (1987)‚ Gabellini (1988)‚ Hauska et al. (1988)‚ O’Keefe (1988)‚ Trumpower (1990)‚ Knaff(1990)‚ Cramer et

Abbreviations: b-heme; b-heme; CM – cytoplasmic membrane; Cyt – cytochrome; DBMIB – 2‚ 5-dibromo-3-methyl-6-isopropylbenzoquinone; DMSO – dimethylsulfoxide; DNP-INT – 2-iodo-6-isopropyl-3-methyl-2'‚ 4‚ 4'trinitrodiphenyl-ether; EPR – electron paramagnetic resonance; ENDOR – electron nuclear double resonance; ESEEM – electron spinecho modulation; fd – ferredoxin; FNR – ferredoxin: oxidoreductase; FPLC – fast protein liquid chromatography; FQR – ferredoxin-quinone reductase; HPLC – high performance liquid chromatography; HQNO – 2-n-heptyl-4-hydroxyquinoline-N-oxide; LHC II – light-harvesting chlorophyll a/b complex II; MOA – methoxyacrylate; Muc – mucidin; Myx – myxathiazol; NDH – NADH dehydrogenase; NMR – nuclear magnetic resonance; NQNO – 2-n-heptyl-4-hydroxyquinoline-N-oxide; OM – outer membrane; ORF – open reading frame; PC – plastocyanin; PCR – polymerase chain reaction; pI – isoelectric point; PQ – plastoquinone; plastoquinol; Stg–stigmatellin; SU IV–subunit IV; TMAO–trimethylamine-N-oxide; TMPD – N‚ N‚ N’ N’-tetramethyl-p-phenylenediamine; UHDBT – 5-n-undecyl-6-hydroxy-4‚7 dioxobenzothiazol; UV – ultraviolet.

Chapter 9 The

Complex

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al. (1991)‚ Widger and Cramer (1991)‚ Malkin (1992)‚ Anderson (1992)‚ Knaff (1993)‚ and Hope (1993). A considerable part of current understanding of the Cyt complex has been inferred from studies on cytochrome complexes. Thus an important‚ incompletely resolved question is to what extent are these Cyt complexes similar‚ and in what ways do they differ? This chapter summarizes current unders