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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in an oxygen-evolving Photosystem II preparation lacking the three extrinsic proteins in the oxygen-evolving system. Biochim Biophys Acta 890: 32–38 Svensson B, Vass I, Cedergren E and Styring S (1990) Structure of donor side components in Photosystem II predicted by computer modeling. EMBO 7: 2051–2059 Svensson B, Vass I and Styring S (1991) Sequence analysis of the D1 and D2 reaction center proteins of Photosystem II. Z Naturforsch 46c: 765–776 Tae G-S and Cramer WA (1989) Lumen-side topography of the subunit of the chloroplast cytochrome b-559. FEBS Lett 259: 161–164 Tae G-S and Cramer WA (1992) Truncation of the COOHterminal domain of the psbE gene product in Synechocystis sp. PCC 6803: requirements for Photosystem II assembly and function. Biochemistry 31:4066–4074 Tae G-S, Black MT, Cramer WA, Vallon O and Bogorad L (1988) Thylakoid membrane protein topography: Transmembrane orientation of the chloroplast cytochrome b-559 psbE gene product. Biochemistry 27: 9075–9080 Takahashi Y and Katoh S (1986) Numbers and functions of plastoquinone molecules associated with Photosystem II preparations from Synechoccus sp. Biochim Biophys Acta 848: 183–192 Tamura N and Cheniae G (1985) Effects of Photosystem II extrinsic proteins on microstructure of the oxygen-evolving complex and its reactivity to water analogs. Biochim Biophys Acta 809: 245–259 Tamura N, Radner R, Lantz S, Cammarata K and Cheniae G (1986) Depletion of Photosystem II-extrinsic proteins. II. Analysis of the Photosystem II/water-oxidizing complex by measurements of N,N,N’,N’,-tetramethyl-p-phenylenediamine oxidation following an actinic flash. Biochim Biophys Acta 850:369–379 Tanaka S and Wada K (1988) The status of the cysteine residues in the extrinsic 33-kDa protein of spinach Photosystem II complexes. Photosynth Res 17: 255–266 Tang XS, Fushimi K and Satoh K (1990) D1-D2 complex of the Photosystem II reaction center from spinach. Isolation and partial characterization. FEBS Lett 273: 257–260 Tang X-S, Chisholm DA, Dismukes GC, Brudvig GW and Diner B A (1993a) Spectroscopic evidence from site-directed mutants of Synechocystis PCC 6803 in favor of a close interaction between histidine 189 and redox-active tyrosine 160, both of the polypeptide D2 of the Photosystem II reaction center. Biochemistry 32: 13742–13748 Tang X-S, Sivaraja M and Dismukes GC (1993b) Protein and substrate coordination to the manganese cluster in the photosynthetic water oxidizing complex: and ENDOR spectroscopy of the state multiline signal in the thermophile cyanobacterium Synechococcus elongatus. J Am Chem Soc 115:2382–2389 Tang X-S, Diner BA, Larsen BS, Gilchrist ML, Lorigan GA and Britt RD (1994) Identification of histidine at the catalytic site of the photosynthetic oxygen evolving complex. Proc Natl Acad Sci USA 91: 704–708 Tavitian BA, Nabedryk E, Mantele W and Breton J (1986) Lightinduced Fourier transform infrared (FTIR) spectroscopic investigations of primary reactions in Photosystem I and Photosystem II. FEBS Lett 201: 151–157 Taylor MA, Nixon PJ, Todd CM, Barber J and Bowyer JR

(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.

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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 understanding of the structure‚ function‚ and genetics of the cytochrome complex. Special emphasis will be placed on the role of the Cyt complex in cyanobacteria‚ the possible molecular basis for functional differences between Cyt and complexes‚ and on advances gained from recent mutational studies not covered in earlier reviews.

II. Role of the Cytochrome Cyanobacteria

et al. (1988)‚ and Scherer (1990). The consequence of this for genetic analysis is that there appears to be no mechanism for propagation of cyanobacteria in the absence of a functional Cyt complex. This limits potential mutations to nonlethal ones and eliminates the approach‚ used successfully in cyanobacteria for analysis of PS II (Chapter 8) and PS I (Chapter 10)‚ in which null mutations are complemented with second mutations that can restore Photosynthetic growth. Yet to be characterized or engineered cyanobacterial strains may possess alternative oxidases‚ such as a quinol-oxidase (QOX; Fig. 1)‚ or other electron transfer mechanisms that may bypass the Cyt complex. This issue is discussed further in Section V D and in Chapter 13 in this volume.

Complex in

Figure 1 illustrates the electron transfer components required for photoautotrophic growth in cyanobacteria (shaded boxes) as well as some of the specialized‚ facultative‚ or hypothetical pathways unique to particular strains (light boxes). Return electron flow from PS I to the Cyt complex involves the PsaE protein of PS I (Yu et al.‚ 1992‚ 1993; Zhao et al.‚ 1993) and possibly the ferredoxinNADP+ oxidoreductase (FNR; Cramer et al.‚ 1987; Knaff and Hirasawa‚ 1991; Schluchter and Bryant‚ 1992). Many questions remain unresolved in regard to cyclic electron transport and cyanobacterial strain specific differences might exist (Section V C). NADH dehydrogenase (NDH; Alpes et al.‚ 1989; Ogawa et al.‚ 1991; Takahashi et al.‚ 1991; Ellersiek and Steinmüller‚ 1992; Schluchter et al.‚ 1993)‚ sulfide-quinone reductase (SQR; Arieli et al.‚ 1991; Shahak et al.‚ 1992)‚ and uptake hydrogenase (HUP; reviewed in Scherer et al.‚ 1988) are alternative mechanisms possessed by some cyanobacteria for reduction of the plastoquinone pool (Fig. 1). The known‚ respiratorypathway in cyanobacteriarequires oxidation of the quinone pool by the Cyt complex and ultimately by molecular oxygen at a terminal cytochrome oxidase (COX; Häfele et al.‚ 1988; Nicholls et al.‚ 1992) although the obligate role of the latter has recently come into question (see Chapter 13). The cytochrome complex appears to be part of a common set of elements essential for growth of cyanobacteria by photosynthesis or respiration as reviewed by Peschek (1987)‚ Scherer

III. Relation to Quinol-Cytochrome c Oxidases in Chloroplasts‚ Mitochondria‚ and Other Bacteria Cyt (ubiquinone or menaquinone-Cyt c oxidoreductase) or (plastoquinol-Cyt or plastocyanin oxidoreductase) complexes are nearly ubiquitous in energy transducing membranes. They function in all cases as quinol oxidases‚ Cyt c or plastocyanin reductases‚ and use the released redox potential energy to generate a transmembrane gradient ofprotons. Cyt complexes have been found in all mitochondria (Hauska et al.‚ 1983) and in diverse bacterial groups including Rhizobium sp. (ThönyMeyer et al.‚ 1991)‚ denitrifying bacteria (Trumpower‚ 1991)‚ Thermus sp. strains (Kuila and Fee‚ 1986)‚ and in Photosynthetic purple (Gabellini et al.‚ 1982)‚ green (Knaff and Malkin‚ 1976)‚ and heliobacteria (Liebl et al.‚ 1990). Cyt complexes were thought to reside exclusively in the Photosynthetic thylakoid membranes of chloroplasts (Hurt and Hauska‚ 1981)‚ algae (Wynn et al.‚ 1988)‚ and cyanobacteria (Krinner et al.‚ 1982)‚ but a Cyt complex more similar to the has been discovered in the nonphotosynthetic‚thermophilic Bacillus sp. strain PS3 (Kutoh and Sone‚ 1988). Cyt complexes have also been reported to occur in the cytoplasmic membranes of at least some cyanobacterial strains (Kraushaar et al.‚ 1990; Section VI B). Escherichia coli is among the seemingly exceptional bacteria that possess neither a Cyt nor a complex (Anraku and Gennis‚ 1987). The structural‚ functional‚ and phylogenetic

262

similarities of the Cyt and complexes have emerged from characterizations of a set of redox centers and core proteins common to both complexes (Hauska et at.‚ 1983‚ Cramer et al.‚ 1987)‚ the overall similarity of electron and proton transfer reactions (Rich‚ 1988; Trumpower‚ 1990b)‚ and DNA and protein sequence comparisons (Gabellini‚ 1988; Hauska et al.‚ 1988; Widger and Cramer‚ 1991). Table 1 lists the polypeptides and associated redox centers (two b-hemes‚ one c-heme‚ and the highpotential‚ Rieske 2Fe-2S center (Rieske et al.‚ 1964)) common to both Cyt complexes and essential for catalytic activity. Four polypeptides (Cyt subunit IV‚ Rieske‚ and Cyt f) are required for Cyt activity

Toivo Kallas

in vitro (Black et al.‚ 1987). Fig. 2 shows the possible orientation of each of these proteins in relation to the thylakoid membrane. Evidence for this model is discussed in Section IV below. Additional lowmolecular-mass subunits such as the 4.0 kDa PetG protein (Haley and Bogorad‚ 1989) may have roles in vivo. Mitochondrial Cyt complexes contain four to seven additional subunits (González-Halphen‚ 1988; Trumpower‚ 1990a). The roles of these ‘extra’ subunits are not yet clear‚ although recent evidence suggests that at least one is required for electron transfer (Graham et al.‚ 1992). Bacterial Cyt complexes have many fewer subunits. Three proteins (Cyt b‚ Rieske‚ and Cyt suffice for catalytic

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264 activity in complexes isolated from Paracoccus denitrificans (Yang and Trumpower‚ 1986)‚ Rhodospirillum rubrum (Kriauciunas et al.‚ 1989)‚ and Rhodobacter capsulatus (Robertson et al.‚ 1993). The Cyt complex from Rhodobacter sphaeroides contains an additional‚ apparently essential subunit of ~14 kDa(Andrews et al 1990; Purvis et al.‚ 1990; Yu and Yu‚ 1991). Photoaffinity labeling suggests a quinone-binding role for this subunit (Usui and Yu‚ 1991)‚ that appears to be lacking from other purple bacteria (Knaff‚ 1991). The Cyt f‚ Rieske‚ and IV proteins from Cyt complexes correspond‚ respectively‚ to Cyt Rieske‚ and Cyt b proteins from Cyt complexes. Widger and Cramer (1984) first noted in sequence comparisons that the Cyt and SU IV polypeptides correspond to the amino- and carboxylterminal segments‚ respectively‚ of the longer Cyt b protein from the complex. This ‘split Cyt b’ protein is a distinguishing feature of the Cyt complex‚ including the one from Bacillus sp. strain PS3 (Kutoh and Sone‚ 1988)‚ and may be responsible in part for the different sensitivities to inhibitors exhibited by Cyt and complexes (Sections V A and VII C). The Cyt (or f) subunit binds the mobile‚ Cyt c or plastocyanin electron acceptor protein as indicated in Table 1 and discussed below (Section IV E).

IV. Polypeptides‚ Redox Centers‚ Substrate Binding Sites‚ and Subunit topology

A. Isolation and Composition of the Cytochrome Complex Nelson and Neumann (1972) first isolated from chloroplasts a complex that contains Cyt and Cyt f‚ in a 2:1 molar ratio‚ and non-heme iron. This preparation in its redox center composition strikingly resembled the mitochondrial Cyt complex III (Hatefi et al.‚ 1962). The unique‚ high-potential‚ Rieske 2Fe-2S center (Rieske et al.‚ 1964)‚ a distinguishing feature of the Cyt complex‚ was subsequently found also in plant chloroplasts (Malkin and Aparicio‚ 1975). Characterization of the Cyt complex was advanced considerably by the discovery of plastoquinol-1-plastocyanin (Wood and Bendall‚ 1976) and duroquinol-plastocyanin (White et al.‚ 1978) oxidoreductase activities for specific measurements of electron transfer activity and by procedures

Toivo Kallas for the isolation of defined‚ catalytically active complexes from Photosynthetic membranes. The first preparation of this type from spinach chloroplasts (Hurt and Hauska‚ 1981)‚ contained five polypeptides (with apparent molecular masses of 34‚ 33‚ 23.5‚ 20‚ and 17.5 kDa)‚ one Cyt f‚ two Cyt one Rieske FeS center‚ and showed a turnover rate of (mol Cyt reduced per mol Cyt f). The 34- and 33 kDa polypeptides were later found to represent heterogeneous forms of Cyt f(Hurt and Hauska‚ 1982). The isolation procedure entails NaBr treatment of thylakoid membranes to remove extrinsic proteins‚ selective solubilization of the Cyt complex with an cholate detergent mixture‚ and purification by ammonium sulfate fractionation and sucrose density gradient centrifugation in Triton X-100. Triton X-100 was found to be inhibitory and turnover rates of were later reported (Hauska et al.‚ 1983; Hauska‚ 1986). Modified procedures include a larger scale one by Black et al. (1987) which uses the less expensive MEGA-9 detergent and Ca-phosphate chromatography in place of the sucrose-gradient centrifugation step. This method yielded a preparation that contains four peptides (masses of 33‚ 23.5‚ 20‚ and 17 kDa); 2 mol 2 non-heme Fe atoms‚ per mol Cyt f; and showed a turnover number of Krinner et al. (1982) used essentially the Hurt and Hauska (1981) method (see also Malkin‚ 1988a) to isolate an active Cyt complex from the cyanobacterium Anabaena variabilis strain ATCC 29413. The latter contained Cyt f (31 or 38 kDa depending on sample preparation for gel electrophoresis)‚ Cyt (22.5 kDa)‚ a 22 kDa species (probably the Rieske)‚ a 16.5 kDa species (subunit IV)‚ two small polypeptides below 10 kDa‚ two Cyt hemes per Cyt f heme‚ and substoichiometric amounts of the Rieske Fe-S center. Rieske proteins are readily lost during purification of Cyt bc/bf complexes (Trumpower and Edwards‚ 1979; Hurt et al.‚ 1981) and the Anabaena variabilis preparation seemed particularly susceptible. Inclusion of Triton X-100 in the sucrose gradient resulted in almost complete inactivation of the Anabaena variabilis Cyt complex presumably because of loss of the Rieske protein (Krinner et al.‚ 1982). Cyt complexes isolated by these procedures contain 0.5–1.0 plastoquinone (PQ) molecules per complex (Hurt and Hauska‚ 1982a; Chain‚ 1985). Purified Cyt bc1 complexes contain similar amounts of quinone (e.g.‚ Gabellini et al.‚ 1982; Kriauciunas

Chapter 9 The

Complex

et al.‚ 1989; Robertson et al.‚ 1993). This presumptive ‘bound’ quinone was thought‚ by analogy to the tightly bound quinone (QA) in bacterial and PS II reaction centers (Deisenhofer and Michel‚ 1991)‚ to have an essential role in Cyt bc 1 and complexes (Hurt and Hauska‚ 1982a; Yu and Yu‚ 1982). The preparation of inactive‚ quinone-depleted Cyt complexes and restoration of activity by reconstitution with native (Hurt and Hauska‚ 1982a) or artificial quinones (Willms et al.‚ 1988) argued against special‚ tightly-bound quinones and instead supports the idea of one or more quinone-binding sites of broader specificity (see Section V B). In addition to quinones‚ isolated Cyt complexes typically contain lipid and chlorophyll in variable amounts depending on the conditions used for solubilization. The ~1.0–2.0 mol of chlorophyll per Cyt f in Cyt complexes purified from spinach (Hurt and Hauska‚ 1981) andAnabaena variabilis(Krinneretal.‚ 1982)wasgenerallybelieved to represent contamination (however‚ see below). Lipid (in particular the relatively minor thylakoid phospholipid phosphatidylcholine) may serve an essential role as suggested by its requirement together with PQ for restoration of activity to quinone-depleted Cyt complex (Chain‚ 1985; Willms et al.‚ 1988). The phospholipids phosphatidylcholine and Phosphatidylglycerol markedly stimulate binding of the quinone analog DBMIB to the quinol-oxidation site as detected by the effect of DBMIB on the Rieske center EPR signal (Malkin et al.‚ 1988). Stimulation by phospholipids of DBMIB binding (Gwak et al.‚ 1987) and electron transport activity (Schägger et al.‚ 1990) has also been observed in the mitochondrial Cyt complex and a tightly bound cardiolipin‚ detected by NMR spectroscopy‚ appears to be important for the integrity of this complex (Hayer-Hartl et al.‚ 1992). These results are consistent with a specific role for lipids in interaction of quinol molecules with their binding site in the vicinity of the Rieske Fe-S center (Malkin‚ 1988b; Malkin et al.‚ 1988). Recently‚ Koppenaal and Krab (1991) have modifiedthe Krinneretal. (1982) procedure to isolate from the thermophilic cyanobacterium Synechococcus sp. strain PCC 6716 a Cyt complex containing major heme-staining proteins of 30.2 kDa (Cyt f) and 23.4 (Cyt a third peptide of ~22 kDa (probably the Rieske)‚ peptides of 20 and 19 kDa (attributed to phycocyanin)‚ some smaller polypeptides‚ and some chlorophyll. Plastocyanin could be removed by gel exclusion chromatography. It is

265

not clear whether the Rieske Fe-S center has been stoichiometrically retained although the purified complex shows a respectable turnover number (mol Cyt c reduced per mol Cyt f) of In an approach that does not employ ammonium sulfate precipitation‚ Rögner et al. (1990) used 1% dodecyl extraction of thylakoids‚ sucrose density gradient centrifugation in 0.04% and HPLC anion-exchange chromatography (Mono Q HR 10/10‚ Pharmacia LKB Biotechnology‚ Inc.) to separate PS II and PS I complexes from thylakoids of Synechocystis sp. strain PCC 6803. A further HPLC-hydroxyapatite chromatography step allowed separation from PS I of a Cyt preparation containing subunits of 38‚ 24‚ 19‚ and 15 kDa and b- and c- cytochromes in a 2:1 molar ratio. This procedure has now been improved by use of alternative anion-exchange and hydroxyapatite HPLC matrices (HiLoad Q-Sepharose HP 16/10 and Superformance 75-5) to yield highly purified Cyt monomer particles of molecular mass (estimated by HPLC gel exclusion chromatography and electron microscopy) ~100 ± 30 kDa which contain immunologically defined peptides of 35 (Cyt f)‚ 20 (Cyt and 13.5 (SU IV) kDa‚ additional proteins of 29‚ 6.6‚ 4‚ and 3.3 kDa‚ and one tightly-bound chlorophyll (Bald et al.‚ 1992; Boekema et al.‚ 1994). The chlorophyll is not energetically coupled to other chlorophylls or cytochromes as determined by fluorescence measurements at 77 K but one molecule appears to be specifically associated with each Cyt monomer. In a related approach using sucrose density-gradient centrifugation and HPLC cationand anion-exchange chromatography‚ Tsiotis et al. (1992) have prepared membranes (also see Tsiotis et al.‚ 1993) and isolated from Synechocystis sp. strain PCC 6714 a highly purified Cyt complex containing four major peptides of 45‚ 40‚ 20‚ and 16 kDa‚ two small peptides below 10 kDa‚ and cytochromes and f in a 2:1 molar ratio. Immunoblotting and amino-terminal amino acid sequencing (of the 40- and 16 kDa peptides) identified the 40-‚ 20-‚ and 16 kDa subunits as Cyt f‚Cyt and SUIV‚respectively. Interestingly‚ both the Bald et al. (1992) andTsiotis et al. (1992) Cyt preparations appear to be depleted of Rieske Fe-S protein. In the case of the latter‚ no crossreactivity was detected to antibodies prepared against the spinach (Tsiotis et al.‚ 1992) or the Nostoc sp. strain PCC 7906 Rieske protein (G. Tsiotis‚ personal communication; Nostoc sp. antibody

Toivo Kallas

266

prepared by B. Holton‚ personal communication). The loss of the Rieske Fe-S protein from cyanobacterial cytochrome complexes during isolation seems to be a persistent problem that needs to be resolved before intact‚ highly active preparations can be obtained for further structure/function studies. The thermophilic complex from Synechococcus sp. strain PCC 6716 (Koppenaal and Krab‚ 1991)‚ like others from thermophilic‚ may offer the advantage of greater stability.

B. Monomer‚ Dimer‚ Supercomplex? Several lines of evidence now suggest that Cyt complexes from both mitochondrial (Nalecz et al.‚ 1985; Schmitt and Trumpower‚ 1990; Nieboer and Berden‚ 1992) and bacterial (Sone and Takagi‚ 1990; Fernandez-Velasco and Crofts‚ 1991) sources exist as functional dimers in vivo and in situ (in the membrane). Schmitt and Trumpower (1990) have presented genetic evidence indicating that subunit 6 of the yeast mitochondrial Cyt complex serves to maintain a fully active dimer state of the complex in vivo. They further postulate that subunit 6 serves in regulation ofelectron transfer (possibly mediated by the proton motive force) as a ‘silencer’ which by dissociation can inactivate half of the sites in the complex. Some evidence also supports a dimeric Cyt complex. Graan and Ort (1986) found that 0.5 DBMIB per Cyt complex completely inhibits PQ oxidation‚ suggesting a functional dimer. However‚ Rich et al (1991) present contrary data indicating that complete inhibition requires one DBMIB per complex suggesting a functional monomer. Chain and Malkin (1991) have shown that the Cyt complex isolated by the method of Hurt and Hauska (1981) contains both monomers and dimers. These could be resolved on a second sucrose-density gradient in the presence of Triton X-100 into separate dimer and monomer populations (Chain 1985; Chain and Malkin‚ 1991). Monomerization could be prevented by crosslinking and only the dimer fraction was active when supplemented with PQ and lipid. Recently gel exclusion chromatography has been used to isolate predominantly a dimer form of the spinach Cyt complex (Cramer et al.‚ 1992; Huang et al.‚ 1993). The dimer form is stabilized at high ionic strength and exhibits approximately five times the in vitro oxidoreductase activity of the monomer. At present the nature of the functional Cyt complex in vivo and the role of

possible monomer/dimer interconversions remain unknown. These questions have not yet been addressed in cyanobacteria. Evidence has also been presented for formation of ‘supercomplex’ assemblages between Cyt bc/bf and other protein complexes. Joliot et al. (1989) have interpreted kinetic data from Rhodobacter sphaeroides in terms ofa model for Cyt Cyt supercomplexes‚ but the issue remains controversial and other investigators (e.g.‚ FernandezVelasco et al.‚ 1991) favor models in which electron transfer among these centers is delocalized. Evidence for Cyt oxidase supercomplexes in Paracoccus denitrificans (Berry and Trumpower‚ 1985) and Bacillus sp. strain PS3 (Sone et al.‚ 1987) stems from the physical isolation of such supercomplexes. In spinach chloroplasts‚ the Cyt complex has been found functionally associated with a light harvesting chlorophyll-a/b protein (LHC II) kinase (Gal et al.‚ 1990) and in Chlamydomonas reinhardtii‚ a transient association with PS I complexes in stromal lamellae has been suggested (Vallon et al.‚ 1991). These possible ‘supercomplex’ associations of the Cyt complex are of great interest relative to the postulated central role of the Cyt complex in regulation of light energy distribution (state transitions) between the two photosystems (see Section V E and Anderson‚ 1992).

C. The Cytochrome

and Subunit IV Proteins

Heme-staining (Thomas et al.‚ 1976; Hurt and Hauska‚ 1981)‚ optical redox-difference spectroscopy‚ and redox titrations of isolated Cyt complexes and their constituent proteins have demonstrated two noncovalently bound protohemes (b-hemes) on the Cyt b and corresponding Cyt polypeptides (reviewed by Hauska et al.‚ 1983; Cramer and Knaff‚ 1990). Table 2 summarizes the redox properties of bhemes from representative membrane preparations and isolated Cyt andCyt complexes. The two b-hemes from both mitochondrial (Tsai and Palmer‚ 1983) and bacterial (Meinhardt and Crofts‚ 1983; Güner et al.‚ 1991; Robertson et al.‚ 1993) Cyt complexes exhibit midpoint redox potentials that differ by approximately 100–150 mV and that can be readily distinguished spectroscopically both in membranes and in the isolated complex because of separated‚ characteristic absorbance maxima. Accordingly‚ these centers have been named hemes (low-potential) and (high-potential). In

Chapter 9 The

Complex

contrast‚ the show separated maxima only at low temperature (77 K) and large midpoint potential differences (~120 mV) only in isolated Cyt complexes (Hurt and Hauska‚ 1983; Clark and Hind‚ 1983). Detection of any substantial midpoint potential difference between in chloroplasts or isolated thylakoids has been much more difficult (Girvin and Cramer‚ 1984; Furbacher et al.‚ 1989). This has been taken as one line of evidence to suggest that‚ in contrast to the Cyt complex‚ obligate interheme (transmembrane) electron transfer does not occur during steady-state conditions (of high in the Cyt complex (Cramer et al.‚ 1987; Furbacher et al.‚ 1989; Cramer et al.‚ 1991). Recently‚ this argument appears to have been weakened by the detection in isolated spinach thylakoids of distinctly different midpoint potentials of –150 and –45 mV for hemes and respectively (Kramer and Crofts‚ 1994). This issue is discussed further in Section V B. Analyses by EPR and magnetic circular dichroism (MCD) spectroscopy indicate that the two b-hemes are each axially liganded through bis-imidazole linkages to a pair of His residues on the Cyt b polypeptide (Carter et al.‚ 1981; Simpkin et al.‚ 1989). Amino acid sequences have been deduced

267

from the nucleotide sequences of well over 20 Cyt b (or Cyt IV) genes from diverse sources including mitochondria (Nobrega and Tzagoloff‚ 1980)‚ bacteria (Davidson and Daldal‚ 1987)‚ plants (Heinemeyer et al.‚ 1984)‚ algae (Büschlen et al.‚ 1991)‚ and cyanobacteria (Kallas et al.‚ 1988a; Brand et al.‚ 1992; Osiewacz‚ 1992). Alignments of these sequences (see Hauska et al.‚ 1988; Gabellini‚ 1988; Widger and Cramer‚ 1991) reveal four strictly conserved His residues‚ the presumptive heme ligands‚ located in the Cyt b and proteins (Widger et al.‚ 1984; Saraste‚ 1984). Recent site-directed mutagenesis of these residues in the Rhodobacter sphaeroides Cyt b protein strongly supports assignment of His (111 and 212) and His (97 and 198) as the axial ligands for the and hemes‚ respectively (Yun et al.‚ 1991). In the absence of known three-dimensional structures‚ hydropathy analyses (Kyle and Doolittle‚ 1982; Rao and Argos‚ 1986; von Heijne‚ 1992) have proven useful for prediction of hydrophobic‚ membrane-spanning alpha-helical regions in membrane proteins as judged by comparison of hydropathy predictions of such regions in the bacterial reaction center (Rao and Argos‚ 1986) with the threedimensional structure based on X-ray crystallography

268

(Deisenhofer et al.‚ 1985; Deisenhofer and Michel‚ 1991). Cyt b and the equivalent Cyt and subunit IV polypeptides are the most hydrophobic proteins in Cyt bc/bf complexes. Hydropathy analysis of the Cyt b protein initially led to a model which predicted nine membrane-spanning alpha helices (Widger et al.‚ 1984; Saraste 1984). Most current models (Crofts et al.‚ 1987; di Rago and Colson‚ 1988; Robertson et al.‚ 1990) predict eight helices. Helices I–IV and V– VII in the 8-span Cyt b model correspond‚ respectively‚ to the four helices of Cyt and the three helices in subunit IV (Fig. 3). Subunit IV from

Toivo Kallas chloroplasts (Heinemeyer et al.‚ 1984) and cyanobacteria (Kallas et al.‚ 1988a) is truncated relative to the carboxyl-terminus of Cyt b and therefore does not have a region corresponding to helix VIII of Cyt b. An additional‚ amphipathic‚ membrane-parallel helix (IIIb in Fig. 3) is now thought to lie between membrane-spanning helices III and IV (see below). The and hemes in this model are located on opposite sides of the membrane and are liganded to conserved His residues [101 and 202] and [86 and 187]‚ respectively‚ as shown for the Nostoc sp. strain PCC 7906 Cyt protein in Fig. 3 . This eight-span

Chapter 9 The

Complex

Cyt b (or seven-span Cyt IV) model is consistent with the mapping in Cyt b of mutations that confer resistance to quinol-oxidation and quinone-reduction inhibitors (di Rago and Colson‚ 1988; Howell and Gilbert‚ 1988; Daldal et al.‚ 1989;). These mutations are expected to map within respective and sites on opposite sides of the membrane (Sections V A‚ and VII B and C). Additional support for the eight-helix Cyt b (and four-helix Cyt rests on the following: 1) The hydrophobic moment (a measure of the amphiphilicity or asymmetry of hydrophobicity of helices‚ Eisenberg‚ 1984) and other secondary structure algorithms predict that amphipathic helix IV should not lie within the membrane (as depicted in the ninespan model) but rather parallel to it with polar residues facing the aqueous environment (Rao and Argos‚ 1986; Crofts et al.‚ 1987; Brasseur‚ 1988; Crofts et al.‚ 1992) as shown in Fig. 3 (helix IIIb). Conserved amphipathic regions are also thought to occur at the amino-terminus and between helices I-II and V-VI (Crofts et al.‚ 1992). 2) Fusions of the Rhodobacter sphaeroides fbcB gene for Cyt b to the phoA ‘proteinexport’ reporter gene are consistent with the eighthelix topology (Yun et al.‚ 1991). The ‘leaderless’ alkaline phosphatase enzyme encoded by the phoA gene shows activity only when fused to a protein domain that results in its export to the periplasm and thus serves as a probe for periplasmic domains of membrane proteins (Manoil and Beckwith‚ 1985; McGovern‚ 1991). Fusions of phoA to the L subunit of the Rhodobacter sphaeroides reaction center predicted the correct topology of this subunit based on its known X-ray crystallographic structure (Allen et al.‚ 1987). 3) The amino- and carboxyl-terminal epitopes of the spinach Cyt (detected by specific antibodies) in intact thylakoids are both more sensitive to trypsin cleavage than the lumenal Cyt f protein implying an even number of membrane helices and a stromal (cytoplasmic in cyanobacteria) location for both termini of Cyt (Szczepaniak and Cramer‚ 1990). The definitive location of quinone binding sites awaits three-dimensional structure determination‚ but evidence has begun to accumulate for involvement of the Cyt b (or Cyt IV) and Rieske (see Section IV D) proteins at a quinol-oxidation site and for Cyt b (or Cyt IV) at a quinone-reduction site. Studies with quinone photoaffinity labels suggest that the SU IV‚ and not the Cyt subunit of the

269 spinach Cyt complex contains an actual binding site for plastoquinone (Doyle et al.‚ 1989). Mapping in Cyt b of mutations that confer resistance to quinoloxidation and quinone-reduction inhibitors (di Rago and Colson‚ 1988; Howell and Gilbert‚ 1988; Daldal et al.‚ 1989) provides strong evidence that the affected residues (and the corresponding ones in Cyt and SU IV) are also involved in formation of two distinct binding sites: one for quinol-oxidation and one for quinonereduction (see Sections V A‚ and VII B and C). An important precedent for this argument comes from location within the three-dimensional structure of the bacterial reaction center of overlapping binding sites for ubiquinone and inhibitors of ubiquinone binding (Sinning‚ 1992). Residues corresponding to resistance mutations in Cyt b are localized in the amphipathic helix IIIb of Cyt and the loop between helices V and VI (loop V–VI) of SU IV (shown as bolded circles in Fig. 3). Those corresponding to resistance mutations are located at the amino- and carboxyl-termini of Cyt and the amino-terminus of SU IV (darkened circles in Fig. 3). Accordingly‚ these domains are thought to have roles in formation of the binding sites for quinol-oxidation and quinone-reduction‚ respectively. Additional evidence for quinone binding to Cyt b comes from azido-quinone (azido-Q) photoaffinity labeling of this polypeptide in the three subunit Cyt complex from Rhodospirillum rubrum. (Kriauciunas et al.‚ 1989). Azido-Q labeling of the Cyt complex from the closely related purple Photosynthetic bacterium‚ Rhodobacter sphaeroides‚ labelled Cyt b as well as the apparently essential (Yu and Yu 1991) 14.4 kDa subunit IV (not related to the ~17.5 kDa SU IV from Cyt complexes) peptide of the four-subunit complex from this organism (Yu and Yu‚ 1987; Usui and Yu‚ 1991). A small 9.5 kDa subunit of the mitochondrial Cyt complex has also been identified as a quinone-binding protein (Usui et al.‚ 1990). These results suggest that in some cases additional subunits may participate in formation of quinone-binding sites and that significant differences may exist in this regard between Cyt proteins from different sources and between Cyt and complexes. In addition to the four heme-binding His residues mentioned above‚ several other residues have been strictly conserved in the more than twenty Cyt b (Cyt IV) sequences thus far determined. These

Toivo Kallas

270 positions are shaded in the Nostoc sp. strain PCC 7906 Cyt IV sequence shown in Fig. 3‚ and many occur in the putative quinol binding regions (helix IIIb and loop V–VI). These and other highly conserved residues are likely to serve important structural or catalytic roles and many are now under investigation by site-directed mutagenesis (Sections VIIB and C). Overall‚ chloroplasts and cyanobacteria show strikingly high 84–90% and 72–85% identity among Cyt and SU IV amino acid sequences‚ respectively. Sequence identity values for Cyt b from complexes with Cyt IV are in the range 25– 40% (Hauska et al.‚ 1988; Kallas et al‚ 1988a). Cyt b from a trypanosome thus far shows the least identity (20–25%) with any other Cyt b or Cyt IV sequence (possibly because of a high mutation rate in this mitochondrial genome) and was thus extremely useful for identification of conserved residues that may be important for function (Hauska et al.‚ 1988). From cyanobacteria‚ Cyt (PetB) sequences are available from Nostoc sp. strain PCC 7906 (Kallas et al.‚ 1988a) and Synechococcus sp. strain PCC 7002 (Brand et al.‚ 1992) and SU IV (PetD) from Nostoc sp. strain PCC 7906 (Kallas et al.‚ 1988a)‚ Synechococcus sp. strain PCC 7002 (Brand et al.‚ 1992)‚ and Synechocystis sp. strain PCC 6803 (Osiewacz‚ 1992). The cyanobacterial Cyt and SU IV sequences are approximately as similar to sequences from chloroplasts as they are to ones from other cyanobacteria: Identity values for Cyt are: 86.5% for Nostoc-sp. strain PCC 7906-Synechococcus sp. strain PCC 7002; 83.6% for Nostoc-sp. strain PCC 7906-spinach (Heinemeyer et al.‚ 1984); 83.2% Synechococcus sp. strain PCC 7002-spinach; and 86% for Nostoc-sp. strain PCC 7906-Chlamydomonas reinhardtii (Büschlen et al.‚ 1991). Values for SU IV are 75.6% for Nostoc-sp. strain PCC 7906Synechococcus sp. strain PCC 7002; 78% for Nostocsp. strain PCC 7906-Synechocystis sp. strain PCC 6803; 85% for Synechococcus sp. strain PCC 7002Synechocystis sp. strain PCC 6803; 80.6% for Nostocsp. strain PCC 7906-spinach; 76% for Nostoc-sp. strain PCC 7906-Chlamydomonas reinhardtii‚ 72% for Synechococcus sp. strain PCC 7002-spinach; and 76% for Synechocystis sp. strain PCC 6803-spinach. These data are consistent with 16S ribosomal RNA sequence data showing the origin of the chloroplast lineage within the cyanobacterial phylogenetic grouping (Giovannoni et al.‚ 1988).

D. The Rieske Iron-Sulfur Protein All cytochrome bc/bf complexes contain a distinctive high-potential ‘Rieske’ iron-sulfur protein. These proteins have largely uninformative UV-visible absorption spectra (Hurt et al.‚ 1981) but at liquid helium temperature the reduced‚ paramagnetic Rieske centers show highly characteristic EPR spectra with g-values of 1.89–1.90 (Rieske et al.‚ 1964; Malkin and Aparicio‚ 1975; Prince et al.‚ 1975). These 2Fe2S centers are unusual because of their high redox midpoint potentials of ~280–320 mV in both bc1 (Prince et al.‚ 1975; Trumpower 1981) and (Malkin and Aparicio‚ 1975; Malkin‚ 1981b‚ 1986; Riedel et al.‚ 1991; Nitschke et al.‚ 1992) Cyt complexes (Table 3). Rieske centers from certain bacterial bc/bf complexes show somewhat lower midpoint potentials (Knaff and Malkin‚ 1976; Liebl et al.‚ 1990‚ 1992) but are still much higher than the ~–400 mV values typical for 2Fe-2S bacterial and plant ferredoxins (Tsukihara et al.‚ 1986). A common feature‚ apparently shared by all Rieske centers from Cyt bc/bf complexes‚ is the dependence of redox potential on pH at pH values greater than ~7–8 (Table 3). This may reflect dissociation of a proton from the oxidized Fe-S cluster that could be involved in the mechanism for oxidation of a semiquinone at the quinol-oxidation site (Prince and Dutton‚ 1976; Link et al.‚ 1992; Nitschke et al.‚ 1992). In addition to the EPR signal at 1.90‚ Rieske centers from complexes exhibit readily detectable and signals at ~1.78–1.81 and 2.01– 2.03‚ respectively (Trumpower et al.‚ 1980; Prince and Dutton‚ 1976; see Table II in Ding et al.‚ 1992 for a further compilation of Rieske g-values). The signal in particular has served as an extremely useful reporter of the quinol-oxidation site because its lineshape and location are sensitive to the redox state of the Q-pool (Siedow et al.‚ 1978; Matsuura et al.‚ 1983) and to inhibitors which bind to the site (Bowyer et al.‚ 1983; DeVries et al.‚ 1983; Ohnishi et al.‚ 1988; McCurley et al.‚ 1990). These features have been used in elegant characterizations of the quinol-oxidation domain in Rhodobacter capsulatus mutants affected at this site (Robertson et al.‚ 1990; Ding et al.‚ 1992). The EPR signal has not been readily detected (but see Salerno et al.‚ 1983) in Rieske centers from Cyt complexes (e.g.‚ Hurt et al.‚ 1981; Malkin‚ 1982). Recently‚ non-saturating EPR conditions were used to observe an inhibitor and oxidized-quinone sensitive

Chapter 9 The

Complex

signal from the spinach Cyt complex (Riedel et al.‚ 1991). This study concluded that the orientation of the Rieske center is the same as that in the Cyt complex and suggests that the feature‚ although flatter‚ may also be useful for investigations of the iron-sulfur center in Cyt complexes. ‘Rieske’-type iron-sulfur centers have also been detected‚ based on their characteristic EPR spectra‚ in proteins that are not components of Cyt bc/bf complexes. These include a ~20 kDa‚ two [2Fe-2S] iron-sulfur protein from the thermophilic archaebacterium Thermus thermophilus (Fee et al.‚ 1984‚ 1986) and a variety of bacterial ring-hydroxylating dioxygenases (Fee et al.‚ 1986; Mason and Cammack‚ 1992). A high-potential Rieske-type center has recently been found in another archaebacterium‚ the thermoacidophile‚ Sulfolobus acidocaldarius.

271

(Anemüller et al., 1993). No b- or c-type cytochromes have been detected in this organism and therefore its ‘Rieske’ protein is apparently not a component of a Cyt bc/bf complex. Important evidence for the involvement of the Rieske Fe-S center at the site has come from studies of the effects of quinol-oxidation inhibitors on the Rieske EPR signal. The quinone-analogs, DBMIB (Trebst et al., 1970), DNP-INT (Trebst et al., 1978), UHDBT (Bowyer et al., 1982), and stigmatellin (Oettmeier et al., 1985) block quinoloxidation in the Cyt complex and to differing degrees in the Cyt complex also (Section V A). Malkin (1981 a, 1981 b, 1982) has shown that DBMIB lowers the midpoint potential of the Rieske center and shifts its EPR signal, in both thylakoid membranes and the isolated spinach Cyt complex, from g =

272

1.90 to g= 1.94 in a redox-dependent reaction. These results suggested a strong‚ redox-dependent‚ electrostatic interaction of a DBMIB semiquinone with the unpaired electron on the reduced 2Fe-2S center consistent with the stabilization of a semiquinone at the Fe-S center as predicted by the Qand b-cycle models for electron transport (see Section V B). Similar DBMIB-induced displacements of the g = 1.90 signal have been observed in membranes from Nostoc muscorum (Malkin‚ 1981a); in the Cyt Rieske in Chromatium sp. chromatophores (Malkin‚ 198la)‚ mitochondria (Degli Esposti et al.‚ 1984)‚ and the isolated Rhodobacter sphaeroides complex (McCurley et al.‚ 1990); but to a much lesser extent in membranes from Heliobacterium chlorum (Liebl et al.‚ 1990); and not at all from the Rieske protein in membranes of Sulfolobus acidocaldarius (Anemüller et al.‚ 1993). UHDBT‚ DNPINT‚ and stigmatellin have different effects on the Cyt Rieske center but can displace DBMIB‚ suggesting different but overlapping binding sites at the (Malkin‚ 1982‚ 1986). Stigmatellin and DNP-INT alter the EPR spectrum but‚ in contrast to their effect on the Cyt complex (Von Jagow and Ohnishi‚ 1985; Andrews et al.‚ 1990; Güner et al.‚ 1991)‚ do not raise the of the Rieske center either in spinach (Malkin‚ 1986) or in the complex from Bacillus sp. strain PS3 (Liebl et al.‚ 1992). These findings point to an overall similarity of Rieske Fe-S centers in and cytochrome complexes but also to significant differences. The following results provide further support for Rieske interaction with the site: 1) Crosslinking studies suggest an interaction between the Rieske and Cyt subunits in spinach (Vater et al.‚ 1992). 2) Stigmatellin does not bind to the bovine mitochondrial Cyt complex from which the Rieske Fe-S protein has been removed (Brandt et al.‚ 1991). 3) Several yeast Rieske mutants show increased resistance to myxothiazol‚ an inhibitor of the site in Cyt complexes (Ljungdahl et al.‚ 1989). 4) A point mutation‚ N256Y (Nostoc sp. strain PCC 7906 SUIV: T62)‚ in the yeast Cyt b results in both myxothiazol resistance and dissociation of the Rieske protein during purification of the complex (Geier et al.‚ 1992). At least 19 Rieske protein sequences have been determined from phylogenetically diverse sources including Neurospora crassa (Harnisch et al.‚ 1985)‚ bovine mitochondrion (Schägger et al.‚ 1987)‚ yeast mitochondrion (Beckman et al.‚ 1987)‚ and plant mitochondrion (Huang et al.‚ 1991); the bacteria

Toivo Kallas Paracoccus denitrificans (Kurowski and Ludwig‚ 1987)‚ Bradyrhizobium japonicum (Thony-Mayer et al.‚ 1989)‚ Rhodobacter sphaeroides strain GA (Gabellini et al‚ 1985)‚ Rhodobacter capsulatus (Davidson and Daldal‚ 1987a)‚ Rhodopseudomonas viridis (Verbist et al.‚ 1990)‚ Rhodospirillum rubrum (Majewski and Trebst‚ 1990)‚ and Rhodobacter sphaeroides (Yun et al.‚ 1990); spinach chloroplast (Steppuhn et al.‚ 1987) pea chloroplast (Salter et al.‚ 1992)‚ and tobacco chloroplast (Madueño et al.‚ 1992); and from the cyanobacteria Nostoc sp. strain PCC 7906 (Kallas et al.‚ 1988a)‚ Synechococcus sp. strain PCC 7002 (Widger‚ 1991)‚ and Synechocystis sp. strain PCC 6803 (Mayes and Barber‚ 1991). A striking feature of these sequences (see Davidson et al.‚ 1992 and Graham et al.‚ 1993 for recent alignments) is the strict conservation of the two hexapeptides‚ Cys-Thr-His-Leu-Gly-Cys (box I) and Cys-Pro-Cys-His-Gly-Ser (box II)‚ located near the carboxyl-terminus. In typical 2Fe-2S proteins (e.g.‚ plant ferredoxins)‚ the two iron atoms are each liganded to the protein by two cysteinyl sulfur atoms with the spacing: Cys–4 residues–Cys–2 residues– Cys–~28 residues–Cys (Tsukihara et al.‚ 1986). The arrangement of cysteines in the Rieske is clearly different. Detection in the Thermus thermophilus Rieske-like protein of only four cysteines indicated that at least half of the ligands for the two [2Fe-2S] clusters in this protein must come from residues other than Cys (Fee et al.‚ 1984). Spectroscopic analysis of Rieske centers from the Thermus thermophilus protein‚ the Pseudomonas sp. phthalate dioxygenase(Fee et al.‚ 1984‚ 1986)‚ and yeast (Telser et al.‚ 1987) suggested that both S-atoms from Cys and N-atoms from His act as ligands for these centers. Recent electron spin echo modulation (ESEEM) studies of the bacterial phthalate dioxygenase (Gurbiel et al.‚ 1989)‚ resonance Raman spectroscopy of the dioxygenase and the Thermus thermophilus Rieske protein (Kuila et al.‚ 1992)‚ ESEEM of purified Cyt bc/bf complexes from spinach‚ Rhodospirillum rubrum‚ Rhodobacter sphaeroides‚ and bovine mitochondria (Britt et al.‚ 1991)‚ and electron nuclear double resonance (ENDOR) of the Cyt complex from Rhodobacter capsulatus (Gurbiel et al.‚ 1991)‚ all strongly support the involvement of two His and two Cys residues in coordination of the Rieske 2Fe2S cluster. An Fe-S center liganded to the two more electronegative histidines should itself be more electropositive and thus be a better electron acceptor. This is thought to account for the high redox potential

Chapter 9 The

Complex

of the Rieske center (Fee et al.‚ 1986; Telser et al.‚ 1987). Ionization of the histidine imidazole ring may explain the observed pH-dependent redox properties of the Rieske protein (Fee et al.‚ 1986). Recent site-directed mutagenesis of yeast (Graham and Trumpower‚ 1991) and Rhodobacter capsulatus (Davidson et al.‚ 1992a) Rieske proteins confirms the essentiality of all of the Cys and His residues in the conserved box I and II regions. Both studies showed that the His immediately following box II (H184 yeast‚ H159 Rhodobacter capsulatus ) is not a ligand‚ consistent with the non-conservation of this residue in plant‚ cyanobacterial (Q132 occurs at this position in Nostoc sp. strain PCC 7906 as shown in Fig. 4)‚ and some bacterial sequences (see Fig. 1‚ Davidson et al.‚ 1992a). Cys 155 (C128 in Nostoc sp.

273 strain PCC 7906) was judged not to be a ligand because a Cys 155 to Ser mutation resulted in the synthesis of a small amount of the Rieske protein with an altered 2Fe-2S cluster (Davidson et al.‚ 1992a). These authors suggest‚ based on strict sequence conservation relative to the putative Fe-S binding domains from the ‘Rieske-type’ subunits of benzene‚ naphthalene‚ benzoate‚ and toluate dioxygenases‚ as well as the ‘ferredoxin-like’ subunits of benzene and naphthalene dioxygenases (references in Davidson et al.‚ 1992a; see also Mason and Cammack‚ 1992)‚ that Cys 133‚His 135‚Cys 153‚ and His 156 (equivalent to C108‚ H110‚ C126‚ and H129 in Nostoc sp. strain PCC 7906‚ Fig. 4) serve as ligands for the Rieske 2Fe-2S center. Spectroscopic evidence indicates that two His residues are ligated

274 to the same Fe atom (Gurbiel et al.‚ 1989‚ 1991; Britt et al.‚ 1991;Kuila et al.‚ 1992) and that in the Cyt bc/ bf complexes‚ the Fe-Fe axis of the Rieske center is in the plane of the membrane (Salerno et al.‚ 1979; Prince‚ 1983; Riedel et al.‚ 1991). Six published sequences are currently available for Rieske proteins from oxygenic phototrophs (see Salter et al.‚ 1992 for an alignment). Rieske proteins encoded in plant nuclear genomes have cleavable presequences of 49–69 residues for transport into chloroplasts. Apparently the information fortargeting to thylakoid membranes resides within the mature polypeptide (Bartling et al.‚ 1990). Accordingly‚ cyanobacterial Rieske proteins seem to lack presequences‚ although Mayes and Barber (1991) have identified in Synechocystis sp. strain PCC 6803 a possible ATG translation start codon 36 bases upstream of the start site identified in Nostoc sp. strain PCC 7906 (Kallas et al.‚ 1988a) and Synechococcus sp. strain PCC 7002 (Widger‚ 1991). The mature Rieske proteins from oxygenic phototrophs are all ~180 amino acids long (Salter et al.‚ 1992) and those from Cyt complexes are similar or slightly longer: e.g.‚ 179‚ 184‚ 191‚ and 199 residues in beef (Schägger et al.‚ 1987)‚ yeast (Beckmann et al.‚ 1987)‚ Rhodobacter capsulatus (Davidson and Daldal‚ 1987)‚ and Neurospora crassa (Harnisch et al.‚ 1985)‚ respectively. In addition to the two conserved Fe-S binding domains‚ the carboxyl-terminal regions of Rieske proteins are highly conserved as illustrated by the shaded or blackened residues between lines d2 and d3 in Fig. 4 of the Nostoc sp. strain PCC 7906 protein. Sequence identity in this region (residues 106–164 in Nostoc sp.) is relatively high (~40%) even between such distantly related sources as Nostoc sp. strain PCC 7906 and yeast or Nostoc sp. strain PCC 7906 and Rhodobacter capsulatus. Outside of this region‚ alignment of Cyt and sequences (e.g.‚ Kallas et al.‚ 1988a; Widger and Cramer‚ 1991; Graham et al.‚ 1993) becomes difficult. Nonetheless‚ there appear to be several discontinuities (deletions or insertions) in relative to Rieske proteins. All Cyt Rieske proteins clearly have 12–17residue carboxyl-terminal extensions (that extend past the point marked d3 in Fig. 4) relative to Cyt Rieske proteins. Two other possible discontinuities are marked d l and d2 in Fig. 4. The structural and/or functional significance of these discontinuities is not clear at present. Comparisons of Rieske proteins from chloroplasts

Toivo Kallas and cyanobacteria also reveal much higher conservation in the carboxyl- (e.g.‚ residues 103–179 in Nostoc sp. strain PCC 7906) than in the aminoterminal (e.g.‚ residues 1–102) segment. Amino acid sequence identities in the carboxyl-terminus are 84‚ 78‚ 79‚ 76‚ 73‚ and 74%‚ and in the amino-terminus 66‚ 69‚ 69‚ 55‚ 45‚ and 43%‚ respectively‚ between Nostoc sp. strain PCC 7906-Synechococcus sp. strain PCC 7002‚ Nostoc sp. strainPCC 7906-Synechocystis sp. strain PCC 6803‚ Synechococcus sp. strain PCC 7002-Synechocystis sp. strain PCC 6803‚ Nostoc sp. strain PCC 7906- spinach‚ Nostoc sp. strain PCC 7906-pea‚ and Synechocystis sp. strain PCC 6803spinach. Comparisons of the Rieske carboxylterminal domain are consistent with the idea of recent evolutionary divergence of chloroplasts and cyanobacteria; but the amino-terminus might suggest a more distant relationship. One obvious interpretation is that Rieske protein amino-terminal domains have fewer structural constraints and are therefore more tolerant of mutation. Huang et al. (1991) recently showed that plasmids expressing Rieske fusion proteins comprised of the aminoterminal third of the yeast protein and carboxylterminal two-thirds of the maize or tobacco mitochondrial proteins could restore respiratory growth to yeast bearing Rieske-deletion mutations. Expression ofthe intact plant mitochondrial proteins‚ or fusions of their amino-termini with the yeast carboxyl-terminal domain did not result in complementation. These results provide direct evidence for functional conservation in the Rieske carboxylterminal region and suggest that the non-interchangeable amino-termini serve species-specific functions. Hydropathy profiles for Rieske subunits from sources including mitochondria (Harnisch et al.‚ 1985; Schägger et al.‚ 1987)‚ bacteria (Gabellini and Sebald‚ 1986; Davidson and Daldal‚ 1987; Yun et al.‚ 1990)‚ plastids (Steppuhn et al.‚ 1987)‚ and cyanobacteria (Kallas et al.‚ 1988a; Malkin et al.‚ 1988) show similarities but the topology of these proteins in the membrane is still highly controversial. Most Rieske proteins have a long hydrophobic region near the amino-terminus (residues 21–47 in Nostoc sp. strain PCC 7906)‚ a shorter one in the box I region (103– 118 in Nostoc sp. strain PCC 7906)‚ and often a third short stretch near the carboxyl-terminus (142–153 in Nostoc sp. strain PCC 7906). The first (region I) has generally been interpreted to represent one (Harnisch et al.‚ 1985; Steppuhn et al.‚ 1987; Yun et al.‚ 1990)or

Chapter 9 The

Complex

two (Schägger et al.‚ 1987) membrane-spanning alpha-helices but zero-span models have also been proposed (Hartl et al.‚ 1989; González-Halphen‚ 1991). Trypsinization of beef mitochondrial Cyt complex III or the isolated Rieske protein yields a water-soluble‚ Fe-S containing peptide which‚ in contrast to the intact protein‚ cannot reconstitute oxidoreductase activity with Rieske-depleted complex (González-Halphen et al.‚ 1988‚ 1991). A 20-amino acid synthetic peptide from region I competes with the Fe-S protein in this reconstitution. González-Halphen et al. (1991) argue on this basis and on secondary structure analysis that region I represents an exposed‚ non-spanning amphipathic stretch of ~18 residues required for hydrophobic binding of the Rieske to the Cyt complex. Other studies demonstrating production of a water-soluble Rieske Fe-S domain by proteolytic separation from region I (Li et al.‚ 1981; Cocco et al.‚ 1991; Link et al.‚ 1992;Van Doren et al.‚ 1993) have been interpreted differently. Van Doren et al. (1993) showed that expression of the Rhodobacter sphaeroides fbcF gene in Escherichia coli or Rhodobacter sphaeroides‚ lacking Cyt b or proteins‚ results in assembly of the Rieske protein into membranes‚ suggesting that the Rieske subunit itself must have a transmembrane anchor. Szczepaniak and Cramer (1991) demonstrated that extraction at pH 11.3 releases half of the total Rieske protein from spinach membranes. Previously this has been taken as evidence for peripheral membrane association (Singer‚ 1971)‚ but surprisingly the more hydrophobic Cyt and SU IV components were released at lower pH values of 10.7 and 11.1‚ respectively. These data therefore suggested an important role for electrostatic stabilization between membrane-spanning helices in Cyt SU IV‚ and hydrophobic‚ possibly membrane-spanning domains of the Rieske protein. The issue of membrane domains for the Rieske remains unresolved and the recently elucidated structure for Cyt f (Section IV E) suggests that most‚ if not all‚ of the Fe-S protein could lie in the aqueous phase between Cyt f and the P-side of the membrane. The second region near box I is generally thought to form a partially hydrophobic pocket for the Fe-S center at or near the lipid/water interface on the P(positive)-side of the membrane in proximity to the site. This location of the Rieske center is consistent with EPR spin-relaxation studies (Prince‚ 1983; Ohnishi et al.‚ 1989) and places the Fe-S center in the vicinity of heme and Cyt (or Cyt f). A recent

275 proteolytic study of oriented membrane vesicles has placed the Rhodobacter sphaeroides Fe-S center on the N(neutral)-side of the membrane in proximity to the site (Theiler and Niederman‚ 1991). This conclusion is incompatible with the consensus of current evidence in support of electron transport models in which the Fe-S center oxidizes a quinol molecule at the site and in turn passes an electron to Cyt (or Cyt f; see Section V B). Strong support for Rieske center involvement at the site on the Pside comes from the finding that a Rhodobacter capsulatus mutant lacking the Rieske protein forms a stable Cyt subcomplex that retains an active but not a (Davidson et al 1992b). Fig. 4 shows a model for the Nostoc sp. strain PCC 7906 Rieske protein with one membrane-spanning helix and other hydrophobic domains‚ including one surrounding the Fe-S center‚ partially buried within the membrane. This representation is compatible with most current data but should be considered highly speculative for the reasons discussed above. Possible alpha-helical regions‚ shown as rectangles‚ are based on hydropathy (Kyte and Doolittle‚ 1982)‚ secondary structure predictions (Garnier et al.‚ 1978)‚ and on elements of previous models for the yeast Rieske protein (Beckmann et al 1989; Trumpower‚ 1990). Additional findings based on mutational analysis are discussed in Section VII D.

E. The Cytochrome f Protein Hill and coworkers first described (Hill and Scarisbrick‚ 1951) and subsequently purified (Davenport and Hill‚ 1952) Cyt f from the leaves of various plants. An excellent‚ recent review discusses the structure‚ function and biogenesis of this subunit (Gray‚ 1992). Important features of Cyt f, aspects peculiar to cyanobacteria‚ and more recent developments will be summarized here. Amino acid sequences for Cyt f have been deduced from the petA gene sequences from 13 different plant or algal species (see Gray‚ 1992) including the cyanobacteria Nostoc sp. strain PCC 7906 (Kallas et al.‚ 1988a)‚ Synechococcus sp. strain PCC 7002 (Widger 1991)‚ and Synechocystis sp. strain PCC 6803 (Mayes and Barber‚ 1991). Direct amino-terminal protein sequences have been determined for Cyt f purified from membranes of spinach‚ Spirulina maxima‚ and Aphanizomenon flos-aquae (Ho and Krogmann‚ 1980) and from wheat (Willey et al.‚ 1984b). All 13 Cyt f species are synthesized as precursor proteins

276 with cleavable‚ amino-terminal presequences of 31– 44 residues for membrane targeting. The role of this precursor as a procaryotic signal sequence has been established by expression in Escherichia coli of gene fusions (5' end of the pea petA fused to lacZ) which direct the encoded in a SecAdependent manner into the plasma membrane (Rothstein et al.‚ 1985). Interestingly‚ the Escherichia coli plasma-membrane leader peptidase correctly cleaves the pea Cyt f presequence (Anderson and Gray‚ 1991). In the mature protein‚ 36% of residues are invariant among all 13 Cyt f sequences (Gray‚ 1992). Identity among mature Cyt f sequences from all higher plants is ~80%; between Nostoc sp. strain PCC 7906 and spinach‚ 60%; and 71.5% and 69% between Nostoc sp. strain PCC 7906-Synechococcus sp. strain PCC 7002 and Nostoc sp. strain PCC 7906Synechocystis sp. strain PCC 6803‚ respectively. Overall similarity between Cyt f and proteins is extremely low‚ about 15–17% (allowing conservative replacements) with no large blocks of homology. The heme binding residues‚ C-X-X-C-H‚ located near the amino-terminus in all Cyt f and proteins‚ are among the few residues which are invariant in all species (see Widger and Cramer‚ 1991 for an alignment). Hydropathy analyses (see Malkin et al.‚ 1988‚ for data from Nostoc sp. strain PCC 7906) show two long hydrophobic stretches common to all Cyt f and proteins. One is in the distal portion of the presequence (positions –28 to –1 in Nostoc sp. strain PCC 7906). The other near the carboxyl-terminus (positions 255–275 in Nostoc sp. strain PCC 7906) is thought to form a lone membrane-spanning helix anchored by positively charged Arg or Lys residues on either side of the membrane (Willey et al.‚ 1984a). Proteolytic studies of intact thylakoids and oriented vesicles have largely confirmed the topological configuration for Cyt f shown in Fig. 2 (Willey et al.‚ 1984a; Ortiz and Malkin‚ 1985; Szczepaniak et al.‚ 1989). In such models ~ 15 carboxyl-terminal residues are exposed on the stromal (cytoplasmic) side whereas the amino-terminus‚ including the heme-binding domain‚ protrudes into the lumenal (intrathylakoidal) space. This transmembrane topology of Cyt f is consistent with its extreme resistance to extraction from membranes at high pH (Szczepaniak et al.‚ 1991) and with the role of Cyt f as an electron acceptor for the Rieske Fe-S center and an electron donor to PC or Cyt on the lumenal (P) side of the membrane (see Section V B). Water-soluble forms

Toivo Kallas have been obtained by proteolytic removal of the carboxyl-terminal hydrophobic domain of Cyt f from Cruciferae such as turnips‚ radish‚ and oil-seed rape (Gray‚ 1978; Tanaka et al.‚ 1978; Martinez et al.‚ 1992) or by genetic insertion of a stop codon at Gln228 in Cyt from Rhodobacter sphaeroides (Konishi et al.‚ 1991). Apparently the activation of an endogenous protease during purification of Cyt f from Cruciferae results in trimming of 33 residues (~3-4 kDa) from the carboxyl-terminus (Tanaka et al.‚ 1978; Gray‚ 1992). In the absence of such proteolysis‚ purified‚ full-length Cyt f proteins form insoluble aggregates‚ often octamers‚ in aqueous solution (see Gray‚ 1992). Protein sequencing and X-ray crystallography have established the structure of several c-type cytochromes (reviewed by Meyer and Kamen‚ 1982). In all cases the heme is attached to the protein by covalent thioether linkages to two Cys residues and by one or more axial ligands (the 5th and 6th nonheme ligands) to its Fe atom. The 5th ligand is always His‚ and this residue and two Cys residues thus comprise the distinguishing heme-binding motif‚ C-X-X-CH‚ found in c-type cytochrome proteins. The 6th ligand is usually Met but other combinations are possible: for example‚ dual His ligands (bis-His) for one of the four hemes of the Rhodopseudomonas viridis reaction center cytochrome (Weyer et al.‚ 1987). These axial ligands are thought to have an important role in determining the heme redox potential (Moore and Williams‚ 1977; Meyer and Kamen‚ 1982; Moore and Pettigrew‚ 1990). Because nitrogen is a better electron donor and sulfur a better a Met ligand should stabilize the reduced form of iron and thereby raise the redox potential relative to His or Lys ligands. Accumulated evidence suggested His-Met and His-Lys axial ligands for Cyt and Cyt f‚ respectively. Firstly‚ a 690 nm absorbance band‚ characteristic of a Met ligand‚ is observed in bovine Cyt (Kaminsky et al.‚ 1975) but not in Cyt f (Siedow et al.‚ 1980). Secondly‚ EPR and magnetic circular dichroism studies of Cyt (Simpkin et al.‚ 1989) and Cyt f (Siedowetal.‚ 1980; Rigby et al.‚ 1988; Simpkin et al.‚ 1989)‚ and Raman (Davis et al.‚ 1988; Hobbs et al.‚ 1991) or NMR (Rigby et al.‚ 1988) spectroscopy of Cyt f support the above assignments. Thirdly‚ there are no conserved Met residues among Cyt f sequences (with the exception of Met 287‚ numbered as in Nostoc sp. strain PCC 7906‚ at the extreme carboxyl-terminus) but two invariant Lys residues (29 and 145) occur in

Chapter 9 The

Complex

the putative lumenal domain (Gray, 1992). Because of proximity in alignments to the conserved Met 164 in Cyt (yeast numbering), Lys 145 was thought to be the likely corresponding 6th ligand in Cytf(Davis et al., 1988). The axial ligand role for Met164 (equivalent to Met 183 in Rhodobacter capsulatus ) is supported by recent site-directed mutagenesis studies of this residue in yeast (Nakai, 1990) and Rhodobacter capsulatus (Gray et al., 1992). On the contrary, normal photoautotrophic growth in Chlamydomonas reinhardtii bearing a site-directed Lys 145 to Val mutation in Cyt f argues strongly against Lys 145 as an axial ligand (J. Zhou and R. Malkin, personal communication). The recently elucidated 2.8Å X-ray crystal structure of turnip Cyt f reveals an unexpected and novel solution to this problem. The 6th axial ligand of Cytf is not an amino acid side chain at all, but rather the amino-terminal amino group of the protein (Cramer et al., 1994; and S. E. Martinez, J. L. Smith, D. Huang, A. Szczepaniak, and W. A. Cramer, personal communication; see Fig. 6, discussed in Section VI A below). Based on these findings, Cytf and structures appear to be radically different with respect to heme axial ligation. Possibly because of the unusual axial ligation, Cyt f species have the highest midpoint potentials known

277 for c-type cytochromes (+320 to +365 mV)‚ ~50– 100 mV higher than those for Cyt (see Table 4 for examples). The midpoint potentials of both Cyt f and are constant in the physiological pH range but have been shown in several species to fall by about 60 mV/pH at pH values above pH 8. The pKa values for these dissociations are approximately 8.5–9.0 (Bendall‚ 1982). The characteristic absorbance peak (552–556 nm range) of Cyt f or can be detected by redox difference spectroscopy at room temperature (Hauska et al.‚ 1983). Cyt f transfers electrons from the Rieske Fe-S center to the water-soluble copper-protein plastocyanin (PC) in chloroplasts and in algae and cyanobacteria to either PC or Cyt depending on the species and on copper availability (Ho and Krogmann‚ 1984; Sandmann‚ 1986; Merchant and Bogorad‚ 1986; see Chapter 12). The Rieske Fe-S– Cyt f and Cytf–PC reactions occur extremely rapidly with rate constants (Haehnel‚ 1980; Whitmarsh et al.‚ 1982; He et al.‚ 1991) relative to the ca. rate constant for oxidation by the Rieske center (Selak and Whitmarsh‚ 1982). A marked decrease in the rate of Cyt f oxidation is observed as a function of increasing ionic strength‚ suggesting protein-protein electrostatic interactions

278 between Cyt f and its acceptors (Takabe et al.‚ 1980; Niwa et al.‚ 1980). A number of more recent kinetic investigations of Cyt f/acceptor (e.g.‚ Qin and Kostic‚ 1992) as well as Cyt (e.g.‚ Güner et al.‚ 1993) reactions strongly support this idea. Examination of the Cyt f protein sequences reveals an unusually high proportion (20–22%) of charged amino acids for an integral membrane protein (Gray‚ 1992). Cross-linking and peptide-mapping studies have identified two positively charged regions on the chloroplast Cyt f that interact with acidic residues on PC (Takabe and Ishikawa‚ 1989; Morand et al.‚ 1989). At one site Lysl87 of Cyt f from spinach is covalently linked to Asp44 of PC and at another site‚ Glu59 and/or Glu60 of PC to residues in the aminoterminus of Cyt f (Morand et al.‚ 1989)‚ possibly Lys50 or the invariant Lys29 (Gray‚ 1992). Affinitybinding studies likewise indicate an interaction of the amino-terminus of Cyt f with PC and suggest involvement of the invariant Arg88 and Argl54 residues (Adam and Malkin‚ 1989). Recent sitedirected mutagenesis of PC has established roles in binding and electron transport for the surface-exposed Tyr83 (He et al.‚ 1991; Modi et al.‚ 1992a‚ b) and in binding for Asp42 (Modi et al.‚ 1992b) both in the ‘eastern patch’ of PC for which three-dimensional structures are available (Guss et al.‚ 1986). In contrast to considerable knowledge of Cyt f interactions with its acceptors‚ little is known about Rieske/Cyt f interactions. The latter clearly must occur and a Cytf/Rieske subcomplex has been isolated from spinach (El-Demerdesh et al.‚ 1988). Gross et al. (1991) have identified an epitope on turnip Cyt f (Leu223–Arg250) that is exposed in the isolated Cyt f protein but buried in the Cyt complex‚ suggesting a possible interface with the Rieske protein. Widger (1991) observed that Cyt f from Nostoc sp. strain PCC 7906 and Synechococcus sp. strain PCC 7002 are considerably more acidic (net charge –13 and –14 and pI 4.59 and 4.47‚ respectively) than Cyt f proteins from plant and algal chloroplasts (net charge –1 to 3 and pI 6.56 to 9.06). Cyt f from Synechocystis sp. strain PCC 6803 also appears to be acidic (Mayes and Barber‚ 1991). Ho and Krogmann (1984) had previously noted that the soluble electron carriers from filamentous cyanobacteria‚ Anabaena flos-aquae +5‚ pI 10.10)‚ Anabaena variabilis +5‚ pI 10.06; PC‚ charge 0‚ pI 7.55)‚ Plectonema boryanum +3‚ pI 9.32)‚ and Spirulina maxima charge 0‚ pI 7.17)‚ are generally more basic

Toivo Kallas than those from the unicellular cyanobacteria‚ Microcystis aeruginosa charge 0‚ pI 7.18)‚ Synechococcus lividus –2‚ pI 5.65)‚ algae charge –3 to –7‚ pI 4.14 to 4.68)‚ or higher plants (PC‚ charge –6 to –10‚ pI 4.04 to 4.41). These observations suggest that in some‚ but not all‚ cyanobacterial groups‚ predominantly basic residues on Cyt f interact with complementary‚ predominantly acidic residues on Cyt or PC. Widger (1991) has thereby postulated that chloroplasts may have arisen from a cyanobacterial progenitor that already possessed the ‘plant-type’ basic Cyt f and acidic PC. Based on currently available sequences‚ the residues Lys29‚ Lys50‚ Arg88‚ andArgl54‚ implicated in PCbinding (Takabe and Ishikawa‚ 1989; Morand et al.‚ 1989; Adam and Malkin‚ 1989; Gray 1992)‚ have all been conserved in cyanobacteria; however‚ Lys187 has not. The corresponding residues areAsp in Nostoc sp. strain PCC 7906 and Ala in Synechococcus sp. strain PCC 7002 and Synechocystis sp. strain PCC 6803. It is also clear that cyanobacterial Cyt and PC proteins are often poor acceptors and do not bind well to the weakly basic chloroplast Cytf(Morand et al.‚ 1989). In contrast‚ the basic‚ horse-heart cytochrome c is a good acceptor for Cyt f from some cyanobacteria (e.g.‚ Anabaena variabilis; Krinner et al.‚ 1982)‚ but not for spinach Cyt f(Hurt and Hauska‚ 1981). In contrast to chloroplast PC‚ the electron acceptor for Cyt f in cyanobacteria probably needs to interact with a terminal oxidase as well as with PS I (Scherer‚ 1990; Scherer and Böger‚ 1988; Sections III and V D‚ and Chapter 13). This in turn might place constraints on the allowable structure of PC or Cyt for interaction with Cyt f Such constraints may be different in different cyanobacterial groups because of the existence of possible alternative electron carriers. Cyt and PC are well known‚ alternate acceptors for Cyt f in some cyanobacteria (Sandmann‚ 1986). Recently‚ Laudenbach et al. (1990) raised the possibility of additional alternative acceptors by showing that inactivation of the gene encoding Cyt does not impair the growth of Synechococcus sp. strain PCC 7942‚ a strain presumably incapable of synthesizing PC. Further evidence is provided by growth in copper-depleted medium ofa Synechocystis sp. strain PCC 6803 mutant lacking Cyt (Zhang et al.‚ 1994). This strain does not express the PC gene in low copper (20–30 nm) medium (Zhang et al.‚ 1992) and thus electron transfer from the Cyt complex can occur through an alternative‚ as yet

Chapter 9 The

Complex

undescribed pathway. It should be noted that Rhodobacter capsulatus is capable of Photosynthetic growth in the absence of a functional gene for Cyt the mobile acceptor for Cyt in purple bacteria (Daldal et al.‚ 1986). An alternative‚ membraneassociated cytochrome‚ capable of transferring electrons from the Cyt complex to the reaction center‚ has recently been described in this organism (Jenny and Daldal‚ 1993). Interestingly‚ the closely related Rhodobacter sphaeroides apparently does not have Cyt and is not immediately capable of Photosynthetic growth in the absence of Cyt but Photosynthetic pseudorevertants overproduce yet another alternative cytochrome‚ an ordinarily minor‚ soluble‚ iso-Cyt (Donohue et al.‚ 1988; Fitch et al.‚ 1989).

F. Additional Low Molecular Mass Subunits? The number of subunits in Cyt complexes varies from the three‚ core catalytic subunits (Rieske‚ Cyt b‚ and Cyt in bacteria such as Paracoccus denitrificans (Yang and Trumpower‚ 1986)‚ Rhodospirillum rubrum (Kriauciunas et al.‚ 1989)‚ and Rhodobacter capsulatus (Robertson et al.‚ 1993) to four in Rhodobacter sphaeroides (Andrews et al.‚ 1990; Yu and Yu‚ 1991) to at least eleven in mitochondria (González-Halphen et al.‚ 1988). Evidence has begun to accumulate‚ as discussed above‚ that some of these extra subunits serve important roles in quinone binding (Rhodobacter sphaeroides subunit IV; Usui andYu‚ 1991)‚ electron transfer activity at the quinol-oxidation site (yeast subunit 9; Graham et al.‚ 1992)‚ and regulation of monomer-dimer transitions (yeast subunit 6; Schmitt and Trumpower‚ 1990). Cyt complexes require four subunits for activity in vitro (Cyt subunitIV‚ Rieske‚ and Cyt f)‚ but additional low-molecularmass polypeptides have consistently appeared in preparations from chloroplasts (Hurt and Hauska‚ 1982b; Haley and Bogorad‚ 1989) as well as cyanobacteria (Krinner et al.‚ 1982; Koppenaal and Krab‚ 1991; Bald et al.‚ 1992; Tsiotis et al.‚ 1992). Do these bands represent small subunits with roles in the Cyt complex? The best evidence in favor comes from the identification of a maize chloroplast gene for a 4 kDa peptide that copurifies with the maize Cyt complex (Haley and Bogorad‚ 1989). Not surprisingly‚ this gene‚ originally designated petE but since renamed petG (Stirewalt and Bryant‚ 1989)‚ also occurs in spinach‚ tobacco‚ pea‚ wheat‚

279 rice (references in Haley and Bogorad‚ 1989)‚ Chlamydomonas reinhardtii (R. Malkin‚ personal communication)‚ probably in all other plastid genomes‚ and it has been detected in the cyanelle genome of Cyanophora paradoxa (Stirewalt and Bryant‚ 1989). The petG gene encodes a 37 aminoacid peptide predicted by hydropathy and proteolytic analysis to span the membrane once with its carboxylterminus protruding into the stromal (or cytoplasmic‚ N-side) space. Antibodies against a synthetic decapeptide from the carboxyl-terminus of the maize PetG protein crossreacted specifically against polypeptides in thylakoids from various higher plant chloroplasts but failed to do so with thylakoids from Chlamydomonas reinhardtii or Synechocystis sp. strain PCC 6803 (Haley and Bogorad‚ 1989). The petG gene‚ however‚ appears to exist in cyanobacteria as suggested by use of primers‚ synthesized to conserved regions within PetG‚ for PCR amplification of a fragment of the expected size from Synechococcus sp. PCC 7002 DNA (D. London andT. Kallas‚ unpublished results). The availability of the petG gene from a transformable cyanobacterium should help to establish the role of the PetG protein in vivo and its relationship to the Cyt complex.

G. In Vitro Reconstitution Depletion of an essential component followed by restoration of an activity upon reconstitution of the component into a protein complex provides one unequivocal way to establish function. In this way‚ as discussed above‚ a role for lipids has been established in the Cyt complex (Chain‚ 1985; Willms et al.‚ 1988) and the necessity for a specific‚ tightly bound quinone has been excluded (Hurt and Hauska‚ 1982a; Willms et al.‚ 1988). As shown for the Cyt complex from yeast‚ extraction of the Rieske Fe-S protein with 0.7% cholate and 1.5 M guanidine hydrochloride results in loss of activity and the g = 1 .90 Rieske EPR signal (Trumpower and Edwards‚ 1979; Trumpower et al.‚ 1980). A purified Rieske preparation can be simply mixed with the Rieske-depleted Cyt complex in the presence ofa phosphate buffer‚ soybean phospholipids‚ and ubiquinone-10‚ to restore activity and the g = 1.90 EPR signal. These experiments clearly established the role of the Rieske protein in the Cyt complex (Trumpower and Edwards‚ 1979; Trumpower et al.‚ 1980). The Rieske protein can be resolved from the

280 spinach Cyt complex by chromatography on hydroxyapatite in the presence of 0.5% Triton X-100 but reconstitution by the simple mixing procedure did not succeed (Hurt et al‚ 1981) possibly because of the large Triton micelles associated with both the Rieske protein and the depleted complex (Adam and Malkin‚ 1987). Subsequently‚ Adam and Malkin (1987) demonstrated reconstitution of the Rieske protein into Rieske-depleted Cyt complex by a procedure that involved replacement of Triton with lipids. The success of this procedure should permit structure/function analysis by reconstitution with overproduced‚ mutant forms of the Rieske protein. This strategy has been used‚ with great success‚ for analysis of PS I components from cyanobacteria (Parrett et al.‚ 1990; Zhao et al.‚ 1990; Mehari et al.‚ 1991; Zhao et al.‚ 1992; see Chapter 10). Toward use of this approach for analysis in the cytochrome complex‚ Holton et al. (1992) have recently overproduced the full-length‚ Nostoc sp. strain PCC 7906 Rieske protein in Escherichia coli. Preliminary results indicate that the Rieske 2Fe-2S center with its unique Cys and His ligands can be reestablished in this protein in vitro by reconstitution with iron and sulfide (B. Holton‚ R. Malkin‚ T. Kallas‚ unpublished results) as shown some time ago for soluble ferredoxins (Malkin and Rabinowitz‚ 1966) and more recently for iron-sulfur proteins from PS I (Parrett et al.‚ 1990). Optimal activity in vitro may require interaction with other complexes and lipids as recently suggested by co-reconstitution into proteoliposomes of isolated Cyt and ATP synthase complexes from the thermophile Synechococcus sp. strain PCC 6716 (Krenn et al.‚ 1993). Co-reconstitution stimulated the activity of both complexes and native cyanobacterial lipids were required for high oxidoreductase activity (turnover number

V. Electron and Proton Transfer Pathways

A. Inhibitor Specificities Much of the evidence for two discrete quinonebinding sites located on opposite sides of the membrane and involvement of the Rieske Fe-S center at the site comes from inhibitor studies. Inhibitor effects on the Rieske EPR signal were discussed in Section IV D. Inhibitors of Cyt bc/bf complexes fall into two general classes: 1) those that prevent quinol-

Toivo Kallas oxidation or Cyt b reduction at the site‚ and 2) those that prevent Cyt b oxidation or quinonereduction at the site. The chemistry and modes of action of these inhibitors‚ quinone-analogs or antibiotics with affinities for quinone-sites‚ have been reviewed previously (Hauska et al.‚ 1983; Rich‚ 1984; Von Jagow and Link‚ 1986; Cramer et al.‚ 1987). Many have strikingly different actions on Cyt and complexes suggesting important structural differences between these complexes. Myxothiazol and mucidin (strobilurin A; Von Jagow et al.‚ 1986) effectively inhibit quinol-oxidation in Cyt complexes but have virtually no effect on Cyt complexes. Conversely‚ DBMIB (Trebst et al.‚ 1970) and DNP-INT (Trebst et al.‚ 1978) are potent inhibitors of quinol-oxidation in the Cyt complex but react only weakly with Cyt complexes (Hauska et al.‚ 1983; Degli Esposti et al.‚ 1984). UHDBT (Malkin et al.‚ 198 la‚ 1982) and stigmatellin (Oettmeier et al.‚ 1985) prevent quinol-oxidation in both. Binding of the latter does not occur in the absence of the Rieske subunit (Brandt et al.‚ 1991) suggesting a stigmatellin binding site on a conserved region of this protein. Differences in the primary structures of Cyt b and Cyt IV polypeptides that might account for differences in sensitivities are discussed in Section VII B. Antimycin-A is a classical and potent inhibitor of Cyt but not Cyt complexes (Hauska et al.‚ 1983). In the former it prevents Cyt b reoxidation at the site and thereby blocks further electron transfer and proton translocation through the complex (Snozzi and Crofts‚ 1984). Antimycin-A has been reported to inhibit cyclic electron flow in chloroplasts (Schürman et al.‚ 1971) but its site of action may be on a separate protein‚ possibly a ferredoxin quinonereductase (Moss and Bendall‚ 1984; Cleland and Bendall‚ 1992). Inherent antimycin-A resistance in the Cyt complex may be related to the split between Cyt and SU IV polypeptides which occurs in a region of the Cyt b protein to which several Qreduction inhibitor-resistance mutations have been mapped (Fig. 3 and Section VII C). However‚ a single amino acid substitution in this region could also govern inhibitor sensitivity/resistance as shown in characterizations of several antimycin-A-resistant mutants from mitochondria (di Rago and Colson‚ 1988; Howell and Gilbert‚ 1988) and Rhodospirilium rubrum (Park and Daldal‚ 1992). Several other inhibitors (diuron: di Rago et al.‚ 1986; HQNO: Howell et al.‚ 1987; and funiculosin: Colson et al.‚

Chapter 9 The

Complex

1988) also act at the site in Cyt but not complexes. NQNO (or HQNO‚ Selak and Whitmarsh‚ 1982; Jones and Whitmarsh‚ 1988) binds to the Cyt to prevent Cyt b oxidation but NQNO does not inhibit steady-state turnover of the enzyme (Jones and Whitmarsh‚ 1988) nor proton translocation (Hope and Rich‚ 1989). The methoxyacrylate‚ MOAstilbene behaves similarly (Rich et al.‚ 1992). It shows greater specificity for the Cyt complex but still only partially inhibits steady-state turnover. Consequently a inhibitor has not yet been found that can completely block electron transfer through the Cyt complex. As mentioned earlier and discussed in Section V B‚ this has complicated measurements of interheme electron transfer and led to proposals questioning obligate activity during noncyclic operation of the Cyt complex (Cramer et al.‚ 1987‚ 1991). Interestingly‚ MOA-stilbene‚ an acrylate inhibitor closely related myxothiazol‚ in contrast to its effect on the in the complex‚ is an inhibitor of the in complexes (Von Jagow and Link‚ 1986; Brandt et al.‚ 1988). Rich et al. (1992) have offered several possible explanations for this paradox including the speculation that part of the Cyt b structure used to form the in the complex has been shifted to form a part of the in the Cyt complex. This seems unlikely based on the bulk of evidence for similar topologies (see Sections IV C and VI A); however‚ there are some data compatible with this idea (see Li et al.‚ 1991; Section VIA).

B. Is There a Q-Cycle in the Cytochrome Complex? Mitchell’s quinone (Q)-cycle hypothesis (Mitchell‚ 1976) was formulated to explain two phenomena observed initially in the mitochondrial Cyt complex: 1) oxidant-induced reduction of Cyt b; and 2) a ratio of translocated across the membrane per electron transferred through the complex. This topic has been extensively studied and reviewed (Hauska et al.‚ 1983; Crofts et al.‚ 1983; Rich‚ 1986; Cramer et al.‚ 1987; Trumpower‚ I990b; Rich‚ 1991). The so-called ‘modified Q-cycle’ model (Crofts et al.‚ 1983; Dutton‚ 1986; Rich‚ 1986) is summarized here as are points of controversy in regard to the Cyt complex. The model requires two sites‚ one for quinol oxidation or located on the more positive (P) side of the membrane‚ and another for

281 quinone reduction or located on the opposite‚ more negative (N) side of the membrane as shown in Fig. 5. According to this model: 1) A quinol (plastoquinol‚ in the case of the Cyt complex) binds to the and is oxidized by the Rieske Fe-S center (from where the electron passes to the f heme and then to plastocyanin or Cyt 2) This oxidation by the Rieske center generates a more reactive (lower potential) semiquinone (PQH·) which immediately reduces the nearby low-potential b heme 3) Heme rapidly reduces the second‚ higherpotential b heme located near the opposite side of the membrane. 4) Heme reduces a quinone (PQ) to a stable semiquinone (PQH·) at the nearby 5) A second turnover of the complex (steps 1–4) reduces the semiquinone (PQH·) at the to the fully reduced quinol which dissociates from the site to complete the cycle. Proton translocation occurs as the result of deprotonation of on the P-side of the membrane‚ protonation of PQ on the N-side‚ and diffusion from the N to the P side.

282 Steps 1 and 2 above account for oxidant-induced reduction and one cycle (two turnovers) of the complex achieves the ratio. Other models such as the b- (Wikström and Krab‚ 1980) and semiquinone- (Wikström and Krab‚ 1986) cycles have also been formulated to account for these phenomena but have to some degree converged with the Q-cycle model (Wikström and Krab‚ 1986; Rich‚ 1986; Trumpower‚ 1990). These models differ in their fundamental explanations for establishment of the electrochemical charge gradient across the membrane. The Q-cycle operates electrogenically‚ i.e. by the physical movement of electrons across the membrane. In the b- and semiquinone cycles‚ the charge gradient is established by transmembrane movement of protons or charged semiquinones‚ respectively. The Q-cycle model‚ or modifications thereof‚ are widely accepted and account for much of the experimental data on electron and proton movements in the Cyt complex. Supporting evidence includes the following: 1) Site-specific inhibitors (Section V A) have led to characterization of separate binding sites for quinol-oxidation and quinone-reduction reactions. 2) Sequence determinations and topological models (Section IV C) show two pairs of conserved histidines‚ the likely heme ligands‚ each on separate sides of the membrane. 3) Numerous and inhibitor resistance mutations have been mapped to separate P- and N-side domains‚ respectively‚ of Cyt b (see Fig. 3 of Cyt IV‚ and Sections IV C‚ VII B‚ and VII C). 4) EPR measurements locate heme and the Rieske Fe-S center near the lipid/water interface on the P side‚ heme near the center of the membrane (Ohnishi et al.‚ 1989)‚ and a stable semiquinone anion within 6–10 Å of the N side (Meinhardt and Ohnishi‚ 1992). 5) EPR analyses of the Rieske center shows proximity and close interaction with the site (Section IV D); 6) Rapid electron transfer has been demonstrated between hemes and (Crofts and Wang‚ 1989). 7) Analysis of carotenoid bandshifts (sensitive indicators of membrane potential) in response to flash-activated‚ partial reactions in Rhodobacter sphaeroides chromatophores confirms the above localization of redox centers and provides direct evidence for electrogenic turnover of the Cyt complex (Glaser et al.‚ 1984; Robertson and Dutton‚ 1988). Electron transfer from heme to accounts for approximately 60% of the charge separation and electron transfer from to the for the remainder (Robertson and Dutton‚ 1988). 8) Recent studies of

Toivo Kallas mutants strongly support discrete‚ independently acting and sites. Mutants with a nonfunctional site (Robertson et al.‚ 1986; Brandt et al.‚ 1991; Davidson et al.‚ 1992b) have been shown to retain quinone-reduction activity and vice versa (Yun et al.‚ 1991; Hacker et al.‚ 1993). Acceptance of the Q-cycle as an explanation for turnover in the Cyt complex has been more controversial. There is general agreement as to a similar juxtaposition of redox centers (Sections IV C‚ D‚ E) and operation of the high potential chain At issue is transmembrane electron transfer through the low potential chain Does this electrogenic step occur obligatory‚ as demanded by the Q-cycle model‚ or only facultatively as proposed by Cramer et al. (1987‚ 1991) and others (Joliot and Joliot‚ 1985‚1992; Furbacher et al.‚ 1989; Votz and Rumberg‚ 1990). Firstly‚ detection of interheme electron transfer in the Cyt complex has been difficult because the two b-hemes typically show similar absorbance maxima (Section IV C)‚ and absence of an effective inhibitor to block rapid b-heme reoxidation (Section V A). As mentioned above‚ NQNO (Jones and Whitmarsh‚ 1988) and recently MOA-stilbene (Rich et al.‚ 1992) do prevent reoxidation but not turnover of the high-potential chain. If these inhibitors indeed bind specifically to the Q-reduction site (this too is controversial; Jones and Whitmarsh‚ 1985‚ 1988; Furbacher et al.‚ 1989; Rich et al.‚ 1991‚ 1992)‚ then this lends support to the idea that the Cyt complex can function non-electrogenically. Secondly‚ detection of a significant redox potential difference between the two Cyt hemes in membranes and a ratio of per transferred has been difficult‚ suggesting that interheme transfer is not favored thermodynamically and that the complex does not operate electrogenically under steady-state conditions of high membrane potential (Cramer et al.‚ 1987; Furbacher et al.‚ 1989). Despite these limitations‚ it appears that under appropriate conditions the two hemes can be distinguished spectroscopically and that a potential difference of ~ 100 mV exists between and even in thylakoids (Kramer and Crofts‚ 1994). Rapid electron transfer between them has been reported (Nitschke et al.‚ 1988; Kramer and Crofts‚ 1994) and the results of several studies support a concerted reduction of high and low potential chains by and associated‚ obligatory electrogenic step in the Cyt complex under most conditions (Rich‚ 1986‚

Chapter 9 The

Complex

1988; Rich et al.‚ 1992; Kramer and Crofts‚ 1992‚ 1993). Translocation of has also been observed (Hangarter et al.‚ 1987). The bulk ofcurrent evidence favors the Q-cycle but allows that alternative mechanisms may prevail under special conditions such as when the low-potential chain is reduced and cannot be readily oxidized. Proposed alternative mechanisms include versions of semiquinone (Joliot and Joliot‚ 1986‚ 1992; Rich et al.‚ 1992) and ‘bypass’ (Cramer et al.‚ 1987; Votz and Rumberg‚ 1990) schemes. The former was mentioned above; the latter allows a pair of electrons from to simultaneously reduce a pair of Fe-S centers in high potential chains‚ possibly in aCyt dimer (Cramer et al.‚ 1987). The resolution ofthese issues should be aided greatly by the determination of threedimensional structures upon which to base directed mutagenesis experiments. Flash-induced absorbance changes of Cyt (unresolved and in cyanobacteria were undetectable in a photoautotrophically grown thermophilic Synechococcus sp. (Nanba and Katoh‚ 1983‚ 1985a) or very small in aerobic‚ photoheterotrophically grownSynechocystis sp. strain PCC 6714 (Matsuura et al.‚ 1988). However‚ a rapid reduction of followed by HQNO-sensitive reoxidation could be clearly seen when the respiratory chain was inhibited by KCN (Matsuura et al.‚ 1988). Other characteristic‚ flash-activated Cyt reactions consistent with a Q-cycle interpretation were observed under more reduced conditions. These authors suggest that the Q-cycle occurs obligatorily in these cyanobacteria but that interheme electron transfer is too rapid to be detected under oxidative conditions. It should be noted that oxidation by the Cyt complex is the overall rate-limiting step in Photosynthetic electron transport (Witt‚ 1971; Joliot and Joliot‚ 1992). Recent evidence indicates that this rate limitation results neither from PQ diffusion nor binding of to the site but rather from processes within the Cyt complex (Joliot and Joliot‚ 1992). The forward rate constants for transfers between Rieske–Cyt f‚ and Cyt f–PC in chloroplasts are approximately 200‚ and respectively (Selak and Whitmarsh‚ 1982; Whitmarsh et al.‚ 1982; Hope et al.‚ 1992). Halftimes for these reactions measured from intact cells of a thermophilic Synechococcus sp. strain are 2 ms‚ and respectively (Nanba and Katoh‚ 1983).

283

C. Role in Cyclic Electron Transport Ferredoxin-mediated cyclic electron transfer involving PS I and the Cyt complex has no counterpart in the Cyt complex and therefore specific structural features are expected to reflect this unique role. Several major questions remain unanswered as to the protein subunits and pathway(s) used for these reactions. Three alternatives are illustrated in Fig. 1‚ namely: 1) transfer from ferredoxin (fd) into the PQ pool mediated by a ferredoxin-quinone reductase (FQR)‚ 2) indirect transfer from fd into the PQ pool involving anNADH dehydrogenase (NDH)‚ and 3) direct transfer from fd into the Cyt complex. The existence of an FQR enzyme in chloroplasts has been postulated based on antimycin-A sensitivity of cyclic electron transport and binding of antimycin to a quinone-reduction site outside of the Cyt complex (Moss and Bendall‚ 1984; Davies and Bendall‚ 1987). As yet no FQR protein has been isolated‚ although it has been suggested that ferredoxin: oxidoreductase (FNR) has a role in cyclic flow (Shahak et al.‚ 1981; Hosler andYocum‚ 1987) possibly as a quinone-reductase (Cleland and Bendall‚ 1992). Ferredoxin reduction of mediated by FNR has been well documented (Knaff and Hirasawa‚ 1991) but suggested roles in cyanobacterial respiration (Scherer et al.‚ 1988a) and cyclic electron transport are more controversial. Several studies have reported association of forms of FNR with the stromal (cytoplasmic) side ofthylakoid membranes and various binding-proteins have been implicated including the 17 kDa subunit IV of the Cyt complex (Clark et al.‚ 1984)‚ the PSI-E (PsaE) subunit (Andersen et al.‚ 1992)‚ and peripheral phycobilisome rods in Synechococcus sp. strain PCC 7002 (Schluchter and Bryant‚ 1992). This topic remains controversial although it now seems clear that FNR does not bind to subunit IV of the Cyt complex (Coughlan et al.‚ 1985‚ Matthijs et al.‚ 1986). The FNR protein has been isolated from a number of cyanobacteria (e.g.‚ Scherer et al.‚ 1988b) and the encoding petH gene cloned fromAnabaena PCC 7119 (Fillat et al.‚ 1990; partial sequence only) and Synechococcus sp. strain PCC 7002 (Schluchter and Bryant‚ 1992). The latter study revealed an amino-terminal extension‚ a phycobilisome-binding domain related in sequence to the phycocyaninassociated linker protein CpcD (see Chapter 7)‚ previously undetected in isolated FNR proteins.

284 Structurally related domains have been identified in the FNR proteins of Synechococcus sp. strain PCC 7942 (Schluchter and Bryant‚ 1992) and Anabaena sp. strain PCC 7119 (W. M. Schluchter and D. A. Bryant‚ personal communication). Genetic inactivation of FNR in Synechococcus sp. strain PCC 7002 did not succeed‚ suggesting an essential function in photoautotrophic growth (Schluchter and Bryant‚ 1992); consequently‚ it has not been possible to examine cyclic electron transport in mutants lacking FNR. Mi et al. (1992a) have presented data showing antimycin-A insensitive but cyclic electron transport in Synechococcus sp. strain PCC 7002 and suggested the involvement of NADH dehydrogenase (NDH). A role for the NDH enzyme in cyclic transport is further supported by the demonstration that Synechocystis sp. strain PCC 6803 ndhB and ndhL mutants have lost or impaired abilities‚ respectively‚ for PS I reduction in the presence of the PS II inhibitor DCMU (Mi et al.‚ 1992b). An ndhF interposon mutant of Synechococcus sp. strain PCC 7002 (Schluchter et al.‚ 1993) also showed impaired reduction in the presence of DCMU (Yu et al.‚ 1992) consistent with an NDH pathway for return of electrons to the P700 reaction center. Yu et al. (1992‚ 1993) further demonstrated a requirement for the PS I PsaE protein (psaE gene product) in another‚ as yet uncharacterized‚ pathway for cyclic electron return. FNR binding to the PsaE protein (Andersen et al.‚ 1992) may be significant in this context‚ although such an interaction has only been observed in higher plants in which FNR is not localized in phycobilisomes but may be bound directly to PS I complexes. Schluchter et al. (1993) question whether NDH is in the major pathway of cyclic flow from PS I because this would presumably require NADPH/NADH transhydrogenase activity which has not yet been detected at high levels in Synechococcus sp. strain PCC 7002 or other cyanobacteria. These accumulated findings suggest that there may be different routes for cyclic electron transport in cyanobacteria possibly to meet strain specific physiological requirements. Additional details concerning cyclic electron transport may be found in Chapter 10. Cyt reduction by ferredoxin has been demonstrated in chloroplasts in the presence of NADPH‚ fd‚ and DBMIB (Furbacher et al.‚ 1989) and in the isolated Cyt complex in the presence of NADPH‚

Toivo Kallas fd‚ FNR‚ and DNP-INT (Lam and Malkin‚ 1982). Inclusion of these inhibitors suggests reduction of heme on the N-side of the membrane. Cramer and coworkers (Cramer et al.‚ 1987; Furbacher et al.‚ 1989; Cramer et al.‚ 1991) have proposed a cooperative mechanism involving hemes and for generation of in the Cyt complex during cyclic electron transport. They propose: 1) oxidation of at the by oxidant-induced reduction to generate reduced P700 and reduced heme 2) concomitant reduction of heme by PS I-generated reduced ferredoxin; and 3) cooperative two-electron oxidation of both hemes by PQ at the center of the membrane to complete the cycle. They further envision that monomer/dimer transitions of the Cyt complex might regulate electron flow between the noncyclic (high NADPH/ATP) and cyclic (high ATP/NADPH) pathways as needed to meet metabolic needs. The formation of semiquinone anion (PQH) required for oxidant-induced reduction of in the cyclic scheme is thought to be blocked in the dimeric form of the Cyt complex which allows simultaneous 2 oxidation of PQH2 through the high-potential chain (i.e. via two centers Fe-S). This hypothesis is of considerable interest but controversial in that many current studies support an obligate Qcycle mechanism under most conditions (Section VB).

D. Are There Alternative Electron Transport Pathways in Cyanobacteria? As mentioned earlier‚ cyanobacteria possess several mechanisms for reduction of the PQ pool but only one known mechanism‚ the Cyt complex‚ for its oxidation (Fig. 1). Accordingly‚ the Cyt complex appears to be indispensable for both Photosynthetic and respiratory growth of cyanobacteria (Sandmann and Malkin‚ 1984; Peschek‚ 1987; Scherer et al.‚ 1988a; Scherer‚ 1990). In striking contrast‚ many other bacteria have several pathways that can bypass the corresponding Cyt complex. Examples include alternative cytochromes (Zannoni et al.‚ 1992)‚ quinol oxidase (Zannoni et al.‚ 1986)‚ and nitric oxide reductase (Bell et al.‚ 1992) in purple bacteria; bdtype and o-type quinol oxidases in Escherichia coli (Anraku and Gennis‚ 1987); and b-‚ and d-type terminal oxidases in Paracoccus denitrificans (Ludwig‚ 1992). Alternative‚ cyanide-insensitive oxidases also exist in plants (Siedow and Berthold‚

Chapter 9 The

Complex

1986). A recent example is the cloned Arabidopsis thaliana non-heme‚ single 31 kDa subunit‚ oxidase which can restore respiratory growth to hemedeficient Escherichia coli (Kumar and Söll‚ 1992). A phenomenon termed ‘chlororespiration’ has been described in Chlamydomonas reinhardtii (Bennoun‚ 1982) and other algae (see Scherer‚ 1990) in which a cyanide-insensitive‚ oxidase-inhibitor sensitive‚ but otherwise largely uncharacterized‚ oxidase can oxidize the chloroplast PQ pool. Significantly‚ chlororespiration has also been shown in Chlamydomonas reinhardtii mutants deficient in subunits of the Cyt complex (Bennoun‚ 1983). The common ancestry of cyanobacteria and chloroplasts suggests that the ‘chlororespiratory’ alternative oxidase might also exist in cyanobacteria. Low levels of cyanideinsensitive oxygen-uptake (~ 10–20% of cyanidesensitive respiration) have been observed in several different cyanobacterial strains‚ but there is no evidence currently to support coupling of this pathway to ATP synthesis (Scherer et al.‚ 1988c). Fermentation has been shown as a mode of maintenance metabolism in Oscillatoria limnetica (Oren and Shilo‚ 1979) and Oscillatoria terebriformis (Richardson and Castenholz‚ 1987) but there is no evidence for fermentative growth of any cyanobacterium. Other findings suggesting possible alternative pathways include: 1) the ‘light-activatedheterotrophic-growth’ phenomenon recently observed in Synechocystis sp. strain PCC 6803 which had been thought to be a nonheterotroph (Anderson and McIntosh‚ 1991); and 2) the recently described‚ low potential‚ soluble cytochrome proposed to be part of an anaerobic‚ energy-yielding pathway from pyruvate to hydrogenase (Krogmann‚ 1991; see Chapter 12). These pathways have not been well characterized but the former may require the usual cytochrome oxidase of this strain (see Chapter 13). An alternative which has apparently not been explored in cyanobacteria is the use of artificial acceptors to support anaerobic growth. Purple bacteria (Yen and Marrs‚ 1977; Madigan et al.‚ 1980) and Escherichia coli (Rothery and Weiner‚ 1991)‚ for example‚ can use acceptors such as dimethyl sulfoxide (DMSO) or trimethylamine-N-oxide (TMAO) because they possess the appropriate reductases. Consistent with physiological data‚ it has not yet been possible to genetically inactivate any of the pet genes for the Cyt complex in cyanobacteria.

285 Interposon mutagenesis of petD in Synechocystis sp. strain PCC 6803 resulted in merodiploids bearing both inactivated and wild-type gene copies (Osiewacz‚ 1992). Similar results have been obtained with interposon insertions in the petB and petD genes of Synechococcus sp. strain PCC 7002 (B. Holton‚ R. Alsaadi‚ T. Kallas‚ unpublished results) and Nostoc sp. strain PCC 7121 (D. Zarka and T. Kallas‚ unpublished results). In Synechococcus sp. strain PCC 7002‚ interposon insertion at a site upstream of petB readily allowed recovery of segregants lacking the corresponding wild-type region. In Nostoc sp. strain PCC 7121‚ segregation has not occurred even after prolonged heterotrophic growth of merodiploids in darkness nor after treatment with fluorodeoxyuridine‚ an agent used to lower chromosome copy number in chloroplasts (Newman et al.‚ 1990). One must conclude that conditions have not yet been found for cyanobacterial growth in the absence of the Cyt complex. As a possible alternative to endogenous Cyt bypass pathways‚ artificial electron carriers have been used in this capacity in other systems. A dramatic example comes from treatment of a particular mitochondrial myopathy‚ a rare disease resulting from deletion of Cyt b and other subunits of the Cyt complex (Capaldi‚ 1988). Treatment with menadione (which is reduced by ubiquinone) and ascorbate (which can transfer electrons from menadione to Cyt c) resulted in a remarkable reversal of the disease symptoms. Reduced menadione has also been shown to restore respiration to yeast mitochondria blocked by antimycin-A (Nosoh et al.‚ 1968). Nanba and Katoh (1985) showed that N‚N‚N'N'-tetramethyl-p-phenylenediamine(TMPD) restores Photosynthetic electron transport to a DBMIB-blocked‚ thermophilic Synechococcus sp. strain. These data indicate that TMPD bypasses the Cyt complex by accepting electrons from the PQ pool and donating them directly to P700. It is not clear whether electron flow through TMPD could support cyanobacterial growth in the absence of the energy transducing Cyt complex. If the complex normally translocates only one across the membrane per as suggested by Cramer et al. (1987‚1991) and others‚ then perhaps this would be possible. Some cyanobacteria possess a type-I‚ NADH dehydrogenase (Berger et al.‚ 1991) which might partially compensate for loss of energy transduction capacity in Cyt mutants.

286

E. Role in Redox-Sensing and Mediation of State Transitions Another function for which there is no parallel in the Cyt complex‚ is the emerging role of the Cyt complex as a redox sensor and mediator of state transitions. State transition refers to a short-term response to changes in light quality (wavelength) resulting in excitation energy redistribution between the two photosystems (Fork and Sato‚ 1986; Biggins and Bruce‚ 1989; Allen‚ 1992; see also Chapter 7 and Chapter 22). The mechanism of energy redistribution is controversial and beyond the scope of this chapter‚ but evidence from chloroplasts indicates that the sensor for imbalances in photon flux (manifested as changes in the redox state of the PQ pool) resides in the Cyt complex (reviewed by Knaff‚ 1991; Anderson‚ 1992; Allen‚ 1992). In brief‚ overexcitation of PS II in chloroplasts results in reduction of the PQ pool and activation of a kinase (LHC II kinase; Bennett‚ 1991) which is sometimes associated with the Cyt complex (Gal et al.‚ 1990). The activated kinase phosphorylates itself‚ LHC II (the peripheral chlorophyll a/b light-harvesting complex of PS II)‚ and also the Cyt peptide of the Cyt complex (Gal et al.‚ 1992). According to one model‚ LHC II is thought to detach from PS II. Recent evidence also indicates migration of the Cyt complex from the appressed granal (PS II-enriched) to the stromal (PS I enriched) thylakoid region (Vallon et al.‚ 1991). These changes shift excitation energy to PS I and electron flow from linear toward cyclic thereby raising the ratio of ATP to NADPH. Overexcitation of PS I has the converse effect. Evidence for the central role of the Cyt complex in mediating these profound changes comes from Lemna gibba (Gal et al.‚ 1987)‚ Chlamydomonas reinhardtii (Wollman and Lemaire‚ 1988)‚ and maize (Coughlan‚ 1988; Bennett et al.‚ 1988) Cyt mutants that can no longer phosphorylate LHC II nor undergo state transitions‚ and from deactivation of the LHC II kinase by quinoneanalog inhibitors of the Cyt complex (Frid et al.‚ 1992). Clearly‚ cyanobacteria do not have granal and stromal thylakoid lamellae and the mechanism for state transitions must be somewhat different from that in chloroplasts. Studies with phycobilisome-less mutants of Synechococcus sp. strain PCC 7002 (Bruce et al.‚ 1989) and Synechocystis sp. strain PCC 6803 (Vernotte et al.‚ 1990) support ‘spillover’ models for energy transfer between the photosystems but other

Toivo Kallas models have advocates also (e.g.‚ Mullineaux and Holzwarth‚ 1990). The Cyt complex has also been implicated in cyanobacteria as the primary sensor of redox state and mediator in the statetransition signal transduction pathway (Vernotte et al.‚ 1990; Fujita et al.‚ 1990; Murakami and Fujita‚ 1991; see Chapter 22). The role of phosphorylation in this process in cyanobacteria has apparently not been investigated. It would be of interest to determine whether a Cyt associated kinase and related mechanisms for regulation of state transitions in chloroplasts have been conserved in cyanobacteria.

VI. Three-Dimensional Structure and Biogenesis

A. Structure Determination and Overall Topography Three-dimensional structures based on X-ray crystallography have been immensely useful for elucidation of functional aspects of the bacterial reaction center (Deisenhofer et al.‚ 1985; Deisenhofer and Michel 1991) and by analogy PS II. Recently crystals of PS I from a thermophilic Synechococcus sp. diffracting to ~4 Å resolution (not yet sufficient for atomic resolution) have allowed the calculation of a 6 Å-electron density map and detection of overall features including transmembrane helices (Witt et al.‚ 1992; Krauß et al.‚ 1993; see Chapter 10). Such structures have not yet been developed for entire Cyt nor complexes‚ although several groups are pursuing this objective and crystals of the bovine mitochondrial complex have been obtained which diffract to 5–7 Å (Yue et al.‚ 1991; Kubota et al.‚ 1991; Berry et al.‚ 1992). Very recently‚ the first structure for a component from a Cyt bc/bf complex has been solved for the water-soluble‚ prostheticgroup-bearing‚ lumen-side domain (amino-terminus to residue 252) of turnip Cyt f (Section IV E). Crystals of this protein diffract to 1.8–2.5 Å resolution (Martinez et al.‚ 1992) and a 2.8 Å structure has been determined (Cramer et al.‚ 1994; and S. E. Martinez‚ J. L. Smith‚ D. Huang‚ A. Szczepaniak‚ and W. A. Cramer‚ personal communication). Fig. 6 shows one representation of the Cyt f model which reveals two flat domains (~15 Å thick)‚ one large and one small‚ both comprised largely of with a bend between the two domains. The large domain is a sheet sandwich and is thought to lie roughly parallel

Chapter 9 The

Complex

to the membrane surface with the small domain angled toward the membrane. Heme f lies within the plane of the large domain‚ close to the junction with the small domain‚ with the plane of the heme tilted approximately 30° from the plane of the membrane. As discussed in Section IV E‚ Lysl87 and Arg250 (shown in Fig. 6) form probable contacts with PC and the Rieske protein‚ respectively. The bulk of the Rieske protein and its Fe-S center may be sandwiched in a pocket between the roughly flat Cyt f and the membrane surface. This Rieske/Cyt f topology could presumably accommodate Rieske models that are either entirely lumenal or largely lumenal but with a single membrane-spanning helix. Crystallization of

287 the water-soluble‚ prosthetic-group-bearing domain of the Rieske protein (Section IV D) should also be possible and of obvious interest for elucidation of the Fe-S binding domain and contacts between the Rieske and Cyt f subunits. A model for the overall topography and relations of subunits within the Cyt complex was presented in Figs. 2 and 5 and discussed in Sections IV B‚ C‚ D‚ and E. Demonstration of crosslinking between the Cyt f—subunit IV‚ Cyt IV‚ Cyt (Shallan et al.‚ 1991)‚ Rieske—subunit IV (Lam‚ 1986)‚ and polypeptides (Vater et al.‚ 1992) from isolated complexes is consistent with the subunit interactions shown. Other aspects‚ such as the number of membrane-spanning helices in the Rieske (Section IV D) and the topology of subunit IV are more controversial. One would expect subunit IV to have the same topology as that widely accepted for the homologous region in Cyt b‚ as shown in Fig. 3. However‚ Li et al. (1991) recently presented evidence for trypsin cleavage of spinach subunit IV at Lys 119 or Arg 125 or 126 (conserved in Nostoc sp. strain PCC 7906; Fig. 3). Proteolysis data from oriented membranes suggested that this site protrudes from the lumenal (P) side of the membrane which would place the amino- and carboxyl-termini of subunit IV on the lumenal (P)- and stromal (N)sides‚ respectively‚ opposite from that shown in Fig. 3. Although this seems unlikely‚ it might offer an explanation for the unusual action of MOA-stilbene on the Cyt complex (Rich et al.‚ 1992) discussed in Section V A. Barring structural constraints‚ this kind of ‘flip’ in orientation may be readily possible during evolution based on recent results with Escherichia coli leader peptidase. Addition or removal of as few as one positively charged residue from a critical exposed region resulted in a reversal of transmembrane orientation from amino- and carboxyl-termini outside to amino- and carboxyltermini inside the cytoplasm (Von Heijne‚ 1989; Nilsson and Von Heijne‚ 1990). However‚ the topology as shown in Fig. 3 for subunit IV is consistent with von Heijne’s ‘positive-inside rule’(von Heijne‚ 1992; see also Cramer et al.‚ 1992). According to this rule‚ which correctly predicted the topology of 23 of 24 known bacterial inner membrane proteins‚ positively charged residues occur more frequently on loops exposed to the cytoplasm than to the periplasm. In the case of the Nostoc sp. strain PCC 7906 subunit IV as shown in Fig. 3‚ eight positive residues are located on the cytoplasmic (N) side and only three on

288 the intrathylakoidal (P) side (equivalent to the periplasmic side). For Cyt 8 and 5 positive charges occur on the N and P sides‚ respectively. Further work will be required for full resolution of this issue.

B. Localization in Cyanobacteria and Biogenesis The Photosynthetic apparatus in typical cyanobacteria is located on thylakoid membranes. In Gloeobacter violaceus (Rippka et al.‚ 1974)‚ which lacks internal membranes‚ it is clearly on the cytoplasmic membrane (CM). The cellular location of the respiratory chain is more controversial and possibly strain-specific. Evidence based on separation of thylakoid‚ CM‚ and outer membrane (OM) fractions has indicated respiratory components on the thylakoid‚ CM‚ or both (Omata and Murata‚ 1984‚ 1985; Peschek‚ 1987; Scherer et al.‚ 1988a; Nicholls et al.‚ 1992; see Chapter 13). The continuity of thylakoid and cytoplasmic membranes is clearly established in a few species (see Chapter 13)‚ has been suggested in other cases (e.g.‚ Peschek et al.‚ 1989) but remains unresolved for other species. Kraushaar et al. (1990) have presented immunological evidence for occurrence of Cyt subunits in both thylakoid and CM fractions of Synechococcus sp. strain PCC 6301‚ Synechocystis sp. strain PCC 6714‚ Anabaena variabilis strain ATCC 29413‚ and Nostoc sp. strain PCC 8009. This is expected based on the role of the Cyt complex in both photosynthesis and respiration. However‚ very surprisingly‚ NADH or PQ-9 to horse heart Cyt c electron transfer activities in the CM‚ but not the thylakoid‚ fractions were sensitive to antimycin-A. Kraushaar et al. (1990) suggested that the cyanobacterial CM might possess a separate‚ mitochondrial-like‚ antimycin-A sensitive cytochrome complex (i.e.‚ a Cyt complex). This idea is intriguing but difficult to reconcile with evidence for only one copy each of the petB‚ D‚ C‚ and A or related genes from Nostoc sp. strain PCC 7906 (Kallas et al.‚ 1988b) and Synechococcus sp. strain PCC 7002 (Brand et al.‚ 1992) and the similarity of corresponding Cyt proteins (suggested by immunological crossreactivity) detected in the thylakoid and CM (Kraushaar et al.‚ 1990). Unless lateral gene transfer has taken place‚ the occurrence of Cyt (chloroplast-type) and (mitochondrialtype) genes in the same procaryote also seems unlikely because of the clear common ancestry but considerable evolutionary separation of these sequences. An explanation for the data of Kraushaar et al. (1990) might be association of the CM Cyt

Toivo Kallas complex with additional subunits that confer antimycin sensitivity. The probable occurrence of the Cyt complex in thylakoid and cytoplasmic membranes of at least some cyanobacterial strains raises the question of how the product of a single gene is targeted to more than one cellular location (although this is perhaps a moot point if the two membrane systems are connected). Electron microscopic immunolocalization studies would be useful for confirming the presence of Cyt complexes in the CM. In higher plants and algae‚ the Cyt f‚ Cyt and subunit IV polypeptides are encoded by single-copy‚ chloroplast petA‚ B‚ and D genes‚ respectively‚ whereas petC encoding the Rieske protein resides in the nucleus (Steppuhn et al.‚ 1987). Most plant species have one petC gene but tobacco has two (Madueño et al.‚ 1992). In higher plants Cyt f is the only known organellar protein encoded with a cleavable precursor for membrane (thylakoid) targeting (Willey et al.‚ 1984a; Bartling et al.‚ 1990). Cyt f polypeptides from cyanobacteria have a comparable presequence (Section IV E). There is no evidence‚ either from chloroplasts or cyanobacteria‚ for precursor forms of Cyt and subunit IV suggesting that the mature proteins have internal domains for thylakoid insertion. All plant and algal Rieske proteins have cleavable presequences of variable length for transfer into chloroplasts. A presequence with two cleavable domains‚ one for plastid entry and one for thylakoid targeting‚ had been suggested (Steppuhn et al.‚ 1987) but more recent evidence (Bartling et al.‚ 1990) supports a single cleavable presequence and a thylakoid targeting domain within the mature polypeptide. This is consistent with the apparent absence of any presequence in cyanobacterial Rieske proteins (Section IV D). Import of the Rieske protein into mitochondria requires two post translational processing steps in lower eucaryotes (Hartl et al.‚ 1986; Cheng et al.‚ 1989) but apparently only a single processing step in mammalian cells (see Graham et al.‚ 1993). Assembly of the Cyt complex in higher plant chloroplasts requires additional nuclear gene products for roles such as mRNA processing (Barkan‚ 1993)‚ heme biosynthesis and insertion (Howe and Merchant‚ 1992)‚ and probably specific chaperonins and other as yet uncharacterized functions. No higher plant mutant has been obtained that lacks only the Rieske protein and the Rieske has emerged as the ‘key assembly protein’ required for assembly and stability of the other components in the Cyt complex

Chapter 9 The

Complex

(Metz et al.‚ 1983; Barkan et al.‚ 1986; Willey and Gray‚ 1988). This has been clearly demonstrated in a Lemna purpusilla mutant that lacks all components of the Cyt complex (Lam and Malkin‚ 1985) because of a mutation resulting in 100-fold lower level of Rieske mRNA (Bruce and Malkin‚ 1991). Genes for plastid-encoded subunits in this mutant are transcribed normally and the mRNAs are translated‚ but the proteins degrade rapidly in the absence of the Rieske polypeptide. In algae‚ the central role of the Rieske subunit is not as clear. Certain algal mutants possess some of the chloroplast-encoded subunits‚ even in the absence of Rieske protein (Bendall et al.‚ 1986; Lemaire et al.‚ 1986). Chlamydomonas reinhardtii mutants defective in c-heme attachment to soluble Cyt are also deficient in Cyt f‚ and the Rieske protein (Howe and Merchant‚ 1992). These results suggest that heme attachment is required for stability of mature Cyt and f proteins and that the latter in turn stabilizes the Cyt complex. Very recent site-directed mutagenesis of the Cyt heme and ligands in Chlamydomonas reinhardtii further supports this conclusion. Any one of the mutations‚ H86A‚ H187G‚ H187S‚ H100L‚ and H202Q results in loss of Photosynthetic growth and in nondetectable levels (less than 5–10% of wildtype) of the Cyt complex (F.-A. Wollman‚ personal communication). In contrast‚ in the purple bacterium Rhodobacter sphaeroides‚ mutagenesis of the His ligands resulted in loss of both b-hemes and much reduced levels of protein but changes in ligands resulted only in selective loss of heme and retention of (Yun et al.‚ 1991b). Assembly of mitochondrial and bacterial Cyt complexes does not depend on the Rieske protein as shown in yeast (Crivellone et al.‚ 1988) and Rhodobacter capsulatus (Davidson et al.‚ 1992b). Insertion ofthe Fe-S center into the Rieske protein is not necessary for assembly of the apoprotein into the complex (Gatti et al.‚ 1989; Graham et al.‚ 1991). In Rhodobacter sphaeroides‚ genetic deletion of the carboxyl-terminal membrane anchor from the Cyt protein prevented the proper assembly of the Cyt complex suggesting a key stabilizing role for the subunit (Konishi et al.‚ 1991). These findings suggest the evolution of different mechanisms for biogenesis of cytochrome bc/bf complexes in divergent phylogenetic groups. The assembly ofthe Cyt complex in cyanobacteria has apparently not yet been investigated.

289

VII. Genetics and Mutational Analysis

A. The pet Genes for Photosynthetic Electron Transport In plants and algae‚ the Cyt f‚ Cyt subunit IV and the putative 4.0 kDa PetG subunit are encoded by the chloroplast petA‚ B‚ D‚ and G genes‚ respectively (Wiley et al.‚ 1984a; Heinemeyer et al.‚ 1984; Haley and Bogorad‚ 1989); whereas the petC gene for the Rieske Fe-S protein resides in the nucleus (Steppuhn et al.‚ 1987). These pet genes are so named because their products have roles in photosynthetic electron transport. Other pet genes encode nonmembrane polypeptides such as plastocyanin (petE)‚ Cyt (petJ)‚ Cyt (petK)‚ ferredoxins (petF)‚ and FNR (petH) (see Hallick‚ 1989). The petB and petD genes of land-plant chloroplasts contain introns and are part of a complex operon (psbB-psbN-psbH-petBpetD) that includes PS II (psb ) genes (Heinemeyer and Herrmann‚ 1984; Rock et al.‚ 1987; Kohchi et al.‚ 1988; Ikeuchi et al.‚ 1989; also see Chapters 4 and 5). A series of mRNA processing and posttranslational events involving nuclear gene products are required for translation into protein of both monoand polycistronic pet mRNAs from this region (Rock et al.‚ 1987; Barkan‚ 1988). The petA gene in these chloroplasts is located approximately 15 kb upstream of psbB and is cotranscribed with unidentified ORFs; the primary transcript also undergoes complex processing events (Gray‚ 1992). The petG gene is located between the petA and psbB regions. A radically different organization with no semblance of a psbBNH-petBD operon is found in Chlamydomonas reinhardtii. The petA gene lies ~3 kbp upstream of petD‚petB at least 5 kbp downstream of petD‚ and all three separately transcribed pet genes are far from psbB (Büschlen et al.‚ 1991; Matsumoto et al.‚ 1991). Cyanobacterial pet gene organization shows some similarity to that in land plants although introns are lacking and the transcription pattern considerably simpler. In Nostoc sp. strain PCC 7906 (Kallas et al.‚ 1988a)‚ Synechococcus sp. strain PCC 7002 (Widger‚ 1991)‚ and Synechocystis sp. strain PCC 6803 (Mayes and Barber‚ 1991; Mayes et al.‚ 1993) the petC and petA genes are closely linked and cotranscribed as 1.6–2.0-kb mRNAs. In Synechocystis sp. strain PCC 6803 the psbN and psbH genes (part of the land plant psbB-psbN-psbH-petB-petD operon) are located directly upstream of petCA but transcribed separately (Mayes et al.‚ 1993). The psbNH genes have not yet

290 been identified in either Nostoc sp. strain PCC 7906 or Synechococcus sp. strain PCC 7002‚ but little sequence information is available upstream of petC. The petB and petD genes have been sequenced from Nostoc sp. strain PCC 7906 (Kallas et al.‚ 1988b) and Synechococcus sp. strain PCC 7002 (Brand et al.‚ 1992) in which they are cotranscribed but not part of an operon containing PS II genes. The longest major transcript in Nostoc sp. is 1.8 kb and the petD probe also anneals uniquely to an abundant 0.8 kb transcript in Northern blots. An unidentified 1.2-kbp ORF has been reported upstream of petB in Synechococcus sp. strain PCC 7002 (Brand et al.‚ 1992). In the Anabaena sp. strain PCC 7120 genome‚ thepetA and petB map far apart‚ approximately 1.4 Mbp or onequarter of the genome (Bancroft et al.‚ 1989; Kuritz et al.‚ 1993). The psbB gene has not been accurately assigned to this map but lies no closer than 10 kbp to the 5' end of petB in Nostoc sp. strain PCC 7906 (Kallas‚ 1988b). In several species of purple Photosynthetic bacteria (Gabellini and Sebald‚ 1986; Davidson and Daldal‚ 1987; Verbist et al.‚ 1989; Majewski and Trebst‚ 1990; Yun et al.‚ 1990; Shanker et al.‚ 1992) and Paracoccus denitrificans (Kurowski and Ludwig‚ 1987) the genes encoding the Rieske‚ Cyt b‚ and Cyt polypeptides are cotranscribed and designated either fbcFBC (Gabellini and Sebald‚ 1986) or petABC (Davidson and Daldal‚ 1987)‚ respectively. Bradyrhizobium japonicum shows the same order of genes except that the fbcB and JbcC genes are fused and form a singlefbcH gene which encodes a precursor polyprotein that is cleaved posttranslationally to yield mature Cyt b and Cyt subunits (Thony-Meyer et al.‚ 1991). The ancestral genes for the cytochrome bc/bf complex‚ as suggested by Gray (1992)‚ may have been part of a single operon as in purple bacteria or Bradyrhizobium sp. During the course ofevolution these genes have become separated‚ split‚ extensively rearranged‚ and even transferred into the host nuclear genome in the case ofplant and algalpetC genes. The chloroplast psbBNH-petBD operon may reflect gratuitous operon fusion or a requirement for coordinate regulation of these genes. In the prochlorophyte‚ Prochlorothrix hollandica‚ the petBD operon and the psbH gene are unlinked as in cyanobacteria‚ but a strictly conserved 93-bp element occurs upstream of each and may represent an alternative mechanism for achieving coordinate regulation (Greer and Golden‚ 1992). Other than in

Toivo Kallas Synechocystis sp. strain PCC 6803‚ in which the monocistronic psbH and dicistronic petCA transcripts appear to be initiated separately (Mayes et al.‚ 1993)‚ the possible co-regulation of psbH and pet genes has not been examined in cyanobacteria. Transcription start sites have been mapped 333 and 125 bp upstream of the Prochlorothrix hollandica petB translation start codon consistent with two major transcripts of 2.1 and 1.9 kb detected on RNA blots (Greer and Golden‚ 1992). Nostoc sp. strain PCC 7906 appears to have three transcription start sites approximately 420‚219‚ and 187 bp upstream of the petB ATG start codon (D. Zarka and T. Kallas‚ unpublished results)‚ compatible with transcript sizes of 1.8 and 1.5 kb determined by blot hybridization (Kallas et al.‚ 1988b). The –420 start site is located the appropriate distance downstream from sequences bearing some similarity to Escherichia coli ‘–35’ and ‘–10’ promoter motifs. Transcript sizes and start sites have not been reported for the Synechococcus sp. strain PCC 7002 petBD operon but their steady-state level‚ as well as that of petCA transcripts‚ is regulated by light (Brand et al.‚ 1992). In cells shifted to darkness‚ the petBD mRNA falls to undetectable levels within two hours but rises rapidly following resumption of illumination to three-fold the steady-state level. A single transcription start site has been mapped ~130 bp upstream of the start codon for the Synechocystis sp. strain PCC 6803 petC gene‚ consistent with a major hybridization band of ~2.0 kb on RNA blots (Mayes et al.‚ 1993). Motifs resembling the Escherichia coli promoter lie upstream of this position. Two other pet genes‚ petP and petR‚ have recently been identified immediately upstream of the petABC(fbcFBC) operon in Rhodobacter capsulatus (Tokito and Daldal‚ 1992). Specific functions have not been determined for the petP and R products but petR shares homology with bacterial response regulators and is essential for both Photosynthetic and respiratory growth. The sequence of petR is different from RegA (involved in oxygen sensing and regulation ofphotosynthesis gene expression; Bauer et al.‚ 1993) as are the phenotypes of mutants suggesting involvement in different signal transduction processes. Sequencing of regions surrounding cyanobacterial pet genes for the Cyt complex may similarly lead to identification of genes with important roles in regulation of photosynthesis or assembly of membrane protein complexes.

Chapter 9 The

Complex

B. The Quinol-Oxidation Site As mentioned in Section IV C and shown in Fig. 3‚ the mapping of inhibitor resistance mutations to the Cyt b sequence has been instrumental in defining separate domains‚ one for quinol-oxidation and one for quinone-reduction‚ located on the P- and Nsides‚ respectively‚ of the membrane spanning model for Cyt b. The interactions of residues within these domains and their specific functions have been further probed by analysis of revertants and site-directed mutations. Based on these data and by analogy to quinone-binding sites in the bacterial reaction center‚ refined models for the quinol-oxidation site have been constructed. Despite these considerable advances‚ it should be noted that many questions remain unanswered‚ that these models have not been confirmed by three-dimensional structural information‚ and that corresponding mutations in the Qbinding domains of the Cyt complex (where important functional differences exist) are as yet completely lacking. Table 5 shows a compilation of quinol-oxidation‚ inhibitor resistance mutations mapped to Cyt b proteins from Rhodobacter capsulatus‚ yeast‚ mouse‚ and Chlamydomonas reinhardtii mitochondria (references in Table 5). The corresponding sites in the Cyt and subunit IV proteins from Nostoc sp. strain PCC 7906 are shown as bolded circles in Fig. 3. With the possible exception of the Rhodobacter capsulatus L106P mutation (Daldal et al.‚ 1989) corresponding to Cyt L95 in Nostoc sp. strain PCC 7906‚ these mutations map to the P-side of the membrane spanning model and are localized predominantly in the amphipathic helix IIIb‚ the loop V–VI (between helices V and VI in subunit IV)‚ and adjacent regions on the transmembrane helices III‚ V‚ and VI. In addition‚ mutations at some of these sites‚ e.g. F144S‚ F144L‚ and F144G in Rhodobacter capsulatus (V133 in Nostoc sp. strain PCC 7906) result in weakened binding and slower turnover (Robertson et al.‚ 1990; Ding et al.‚ 1992). These data strongly support the involvement of residues in these regions of Cyt b (and the corresponding ones in Cyt and subunit IV) in quinone-binding and catalysis of quinol-oxidation. The mapping of null mutations to sites in these regions confirms their essentiality for respiration in yeast and photosynthesis in Rhodobacter capsulatus (Table 6 and references therein). In yeast‚ several respiratory revertants have

291

been obtained through back-mutation at the site of the original lesion (Table 6). These ‘pseudorevertants’ provide insights into the characteristics of residues tolerated at these critical sites and into the possible basis for functional differences between the Cyt and complexes. For example‚ reversion of the yeast C133 Y mutation in helix III fromTyr to Ser‚ Asp‚ Asn‚ or Phe results in restoration of respiratory growth to 55%–95% of the wild-type rate (di Rago et al.‚ 1990a). The corresponding residue in Cyt (position 135) is already Ser suggesting that genetic engineering of this or other sites could lead to Cyt complexes with higher turnover at the rate-limiting step. Further evidence for interaction of the P-side helix IIIb and loop V–VI (Fig. 3) in quinol-oxidation is provided by characterization of second-site mutations that restore respiration to yeast respiratory-defective‚ ‘petite’ mutants (Table 7 and references therein). In two of these‚ G131S/G260A and G137E/N256K (corresponding to Nostoc sp. strain PCC 7906 IV:A66 and SU IV:T62‚ respectively)‚ a second mutation in the V–VI loop (a region within SU IV ofthe Cyt complex) partially complements the primary mutation at the P-side end of helix III (di Rago et al 1990a‚ 1990b). It is also of interest that an acidic residue (Glu) at position 137 in yeast results in loss of respiration whereas the corresponding wild-type residue in Cyt (Asp 141) is acidic already. Nearby second-site mutations C133S or H141 Y‚ which make the defective Cyt b more like Cyt (the corresponding residues in Cyt are S13 7 and Y145‚ respectively)‚ restore respiration to a low level (di Rago et al.‚ 1990a‚ 1990b). These data indicate that compensatory mutations can result in slightly different structural configurations that still permit quinol-oxidation activity although at lower rates. Other second-site revertants in this same region additionally show resistance to inhibitors‚ e.g. yeast Y133N/A126T and Y133D/A126T (corresponding to Nostoc sp. strain PCC 7906 S137/S130)‚ consistent with the idea of overlapping quinol and inhibitor binding sites (Tron et al.‚ 1991‚ and Table 7). Table 8 shows a compilation of site-directed mutations in the Cyt b quinol-oxidation domain of purple bacteria. Mutations of the His97 and His 198 (Nostoc sp. strain PCC 7906 H86 and H187) ligands for heme resulted in loss of all heme signals as well as the Cyt b and proteins (Yun et al.‚ 1991b). Nonconservative substitutions in the P-E-W-Y region

292

(positions 294–297 in Rhodobacter sphaeroides and subunit IV: 77–80 in Nostoc sp. strain PCC 7906)‚ thought to form part of the quinol-oxidation site (Hauska et al.‚ 1988)‚ significantly lowered activity consistent with important roles for these residues in catalysis (Crofts et al.‚ 1992a‚ b; Table 8). That activity was not completely abolished is surprising in light of the universal conservation of P-E-W and nearly universal conservation of Y at these positions (Section IV C). Conservative mutations E295D‚ W296F‚ and Y297F at these sites in Rhodobacter sphaeroides showed oxidation kinetics largely similar to those ofthe wild-type (Crofts et al.‚ 1992a‚

Toivo Kallas

b). The nonphotosynthetic Rhodobacter capsulatus G158D mutant displays impaired quinol-oxidation but a functional quinone-reduction site (Robertson et al.‚ 1986‚ 1990). Atta-Asafo-Adjei and Daldal (1991) have made 15 amino-acid substitutions by directed mutagenesis of this site A147 in Nostoc sp. strain PCC 7906). Of these only G158A and G158S‚ and to a much lesser extent G158C and G158P permitted Photosynthetic growth (Table 8). These data demonstrate that only residues with small molar volumes (less than 90 function well atthis position in the Q-oxidation site structure. Ala and Ser (molar volumes approximately 89 relative to

Chapter 9 The

Complex

293

294

60 for the native Gly 158) allow activity at lowered rates and also confer resistance to myxothiazol (Myx) presumably by blocking access to the site. The corresponding G143A mutation in mouse had previously been characterized from spontaneous Myx-resistant mutants (Howell and Gilbert‚ 1988). Ala occurs at this position in the wild-type Cyt complex suggesting a basis for the natural myxothiazol-resistance of the complex and another target for engineering of the quinol-oxidation site. In addition to the Rhodobacter capsulatus G158 (Nostoc sp. strain PCC 7906 residue discussed above‚ two others have been implicated as determinants for inhibitor specificities of Cyt and complexes. F144V and G152D mutations in Rhodobacter capsulatus (Daldal et al.‚ 1989) confer myxothiazol resistance. The corresponding residues in the naturally myxothiazol-resistant Cyt protein are already Val and Asp (V133 and D141 in Nostoc sp. strain PCC 7906)‚ respectively. Substitutions of the aromatic Rhodobacter capsulatus F144 (Daldal et al.‚ 1989) or the corresponding mitochondrial F129 (di Rago et al.‚ 1989; Bennoun et al.‚ 1991) for smaller‚ aliphatic residues also confer myxothiazol resistance and in some cases significantly impair

Toivo Kallas

quinol-binding and oxidation (Robertson et al.‚ 1990; Ding et al.‚ 1992) as outlined in Table 5. Similarly‚ conversions from the aliphatic Gly 152 (G137 in yeast) to charged or hydroxylated residues result in resistances to myxothiazol or mucidin (Table 5). In contrast‚ T163 (Rhodobacter capsulatus‚ T148 in mouse Cyt b‚ S152 in Cyt is a conserved polar residue that appears to be a determinant for stigmatellin-sensitivity of both Cyt and complexes. T163A and T148M mutations confer stigmatellin resistance in Rhodobacter capsulatus and mouse. It is not yet clear whether single amino acid changes can confer particular characteristics to the Cyt complex‚ or whether multiple‚ more subtle modifications are required. Based on analyses of Rhodobacter capsulatus mutants such as those mentioned above (Tables 5 and 8) and by analogy to the and binding sites of the bacterial reaction center‚ Robertson et al. (1990) have proposed a model for the ubiquinolbinding/oxidation site of the Cyt complex. Key features as illustrated in Fig. 7 of Robertson et al. (1990) include: 1) a carbonyl oxygen of Q (or the corresponding phenolic oxygen of suggested to interact through hydrogen bonding to the imidazole

Chapter 9 The

Complex

295

Toivo Kallas

296 ring of one of the invariant heme-liganding His residues (Rhodobacter capsulatus H97 or H198, Nostoc sp. strain PCC 7906 and H187); 2) an aromatic residue, F144 (in Nostoc sp. strain PCC 7906 in the aliphatic V133), oriented cofacially with and providing van der Waals contacts to the quinone molecule; 3) a small, aliphatic residue, I162 or V333 (Nostoc sp. strain PCC and SUIV:V98, respectively) on the opposite side of the quinone ring; 4) an invariant, nonionic, polar residue T163 (Nostoc sp. strain PCC that may hydrogen bond to the quinone; and 5) a small, aliphatic residue, G152 or G158 (Nostoc sp. strain PCC and A147) within the binding site. Table 9 lists essential residues at the reaction center and sites (Deisenhofer et al., 1985; Komiya et al., 1988; Sinning, 1992), residues thought to correspond to these in the Cyt b site (Robertson et al., 1990), and the corresponding ones in the Cyt complex as described above. With the exception of F144 and G152 (if the latter is the key aliphatic residue), the corresponding sites in the complex contain identical amino acids or conservative replacements. F144 has not been conserved (V133 in Cyt Thus, an aromatic residue may not be essential at the site, or perhaps other nearby residues (e.g., F131 or the invariant Y136) serve this role. It should be noted again that other residues (e.g., S, L, G, V) function at position 144 in Rhodobacter capsulatus (Table 5) but with lowered catalytic efficiency. By comparison to the Cyt complex (Table 9) and consistent with site-directed mutagenesis results (Atta-Asafo-Adjei and Daldal, 1991), Rhodobacter capsulatus G158 (A147 in Nostoc sp. strain PCC 7906) might be the essential, small, aliphatic residue rather than G152 (D141 in Nostoc sp. strain PCC 7906). Based on these considerations and the models of Robertson et al. (1990), Ding et al. (1992), and Crofts et al. (1992a, 1992b), a possible structure is shown for the quinol-oxidation site of the Cyt complex (Fig. 7A, 7B, and 7C). Loop V-VI ofsubunit IV is also likely to be involved but not shown. Fig. 7A shows helices II and IV with the heme-liganding H86–H187 and H100–H202 pairs close to the respective and sites together with helices III and IIIb which carry several inhibitor-resistance mutation sites. Fig. 7B represents an expanded view illustrating possible H-boding interactions of the molecule with H187 and S152. V133 corresponds to the F144 thought to have a role in quinone binding in Rhodobacter capsulatus and the strictly conserved Y136 is shown in a position where

it could interact with Fig. 7C shows the model viewed from the ‘top’ (N-side) along the axes of the membrane-spanning helices. Helix IV contains a proline and therefore a bend. One should further imagine that the Rieske Fe-S center resides in close proximity to the pocket‚ perhaps with one of the His ligands for the Fe-S center capable ofhydrogenbonding to the molecule as depicted in Fig. 7 of Robertson et al. (1990). Recently‚ Ding et al. (1992) have extended the earlier model and proposed a ‘double-occupancy’ site structure in which two quinone molecules are each hydrogen-bonded (possibly in a dimer) to the imidazole ring of a His ligand for the Rieske Fe-S center (see Section IV D). As observed by Widger et al. (1984)‚ fourteen residues lie between the two heme-liganding His in helix IV of Cyt rather than thirteen as found in Cyt b. The extra residue (Thr at position 188 in Nostoc sp. strain PCC 7906) shifts the orientation of the His residues in the Cyt complex such that the planes ofthe imidazole rings are all in the same orientation‚ i.e. all parallel to the membrane according to the model of Cramer et al. (1987) or perhaps all perpendicular as shown in Fig. 7A and 7C. In Cyt b of the complex‚ one pair ofimidazoles is predicted to be parallel and the other perpendicular to the membrane (Cramer et al.‚ 1987). The functional ramification of the extra Thr in Cyt is currently under investigation by insertions of Thr‚ Ile‚ and Asp at the corresponding position in Cyt b from Rhodobacter capsulatus to create a more ‘Cyt like’ structure. All of these insertion mutants assemble the complex but grow at lower rates consistent with the importance to structure/function ofa residue at this site (F. Daldal‚ personal communication).

C. The Quinone-Reduction Site Table 10 shows quinone-reduction inhibitor resistance mutations that have been mapped to Cyt b (references in Table 10). The darkened circles in Fig. 3 mark the corresponding sites in the Cyt IV model. These mutation sites are all located on the Nside of the membrane at the amino- and carboxyltermini of Cyt and at the amino-terminus of subunit IV. In the Cyt complex the latter two regions are joined to form a contiguous loop, IV–V, (see Sections IV C and V A) which appears to be at the heart of the site. As discussed above, the lack of this connection in the Cyt complex could explain the altered catalytic properties and inhibitor

Chapter 9 The

Complex

insensitivity of the relative to that in the complex. Alternatively, the Cyt IV split may be relatively unimportant in relation to the roles of specific residues in this region. Bruce and Malkin (1991) noted lower turnover of Cyt than of other subunits in the Lemna Cyt complex, and postulated that the Cyt IV split may be an adaptation to allow, by analogy to high turnover in the protein of PS II (Ohad et al., 1985), high turnover of the quinone-binding subunit IV Until recently, genetic analysis of the site had been confined to characterization of mouse (Howell et al., 1987; Howell and Gilbert 1988) and yeast (di Rago et al., 1986; di Rago and Colson, 1988) Cyt b mutants resistant to the inhibitors diuron, antimycinA, funiculosin, and HQNO (Table 10). Similar

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resistance mutations could not be obtained in the purple bacteria Rhodobacter capsulatus or sphaeroides because of insensitivity to these Q-reduction inhibitors in vivo. Rhodospirillum rubrum‚ however‚ has been found to be antimycin-A sensitive (Shanker et al.‚ 1992) and a subtle D252E change (Nostoc sp. strain PCC 7906 SU IV:D35) confers antimycin-A resistance in one mutant (Park and Daldal‚ 1992). As in the case of the reaction center and Cyt site inhibitor-resistance mutations (Sections IV C and VII B)‚ the mutations in Table 10 are thought to be located within the domain for quinone binding and reduction. Recently the roles of amino acids at these and highly conserved residues in the domain of purple bacteria have been probed in greater detail by site-

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directed mutagenesis coupled with biophysical characterizations of mutants (Table 11). Some of these, e.g. Rhodobacter sphaeroides A52V (Nostoc sp. strain PCC 7906 and K251M (Nostoc sp. strain PCC 7906 SU IV:N34) reproduce in purple bacteria the antimycin-A resistance previously seen in spontaneous yeast and mouse mutants (Tables 10 and 11). These mutants show weaker antimycin-A binding and lower turnover at the site (Crofts et al., 1992b; Hacker et al., 1993). Mutants H217A (Nostoc sp. strain PCC 7906 D252A, and D252N (Nostoc sp. strain PCC 7906 SU IV:D35) are of particular interest. In these, the reoxidation of Cyt is inhibited even in the absence of antimycin demonstrating the importance of these residues for electron transfer from to quinone. H217A and D252A cannot grow photosynthetically but surprisingly D252N still can despite severe impairment of transmembrane electron transfer as indicated by a markedly decreased electrochromic absorption change at 503 nm (Crofts et al., 1992b; Hacker et al., 1993). Although D252 is conserved as SU IV:D35 in the Cyt complex, the behavior of the D252N mutant resembles that proposed by Cramer et al. (1987, 1991) for the Cyt complex operating under conditions of high membrane potential; namely, that the Q cycle is inoperative and that electron flow occurs primarily through the high potential chain > Fe-S > Cyt f). These mutational studies provide strong evidence that A52, H217, K251, and D252 in Rhodobacter sphaeroides have roles at the site and helices I, IV, and V of Cyt b can be positioned in models such that these residues are in juxtaposition to form this site (see the helical wheel diagram, Fig. 8 of Hacker et al., 1993). Very recently the role of H217 has been investigated further in Rhodobacter capsulatus and suggested to be essential for stabilization of the semiquinone at the site (Gray et al., 1994). An H217L mutant shows assembly of the complex but no growth whereas H217D and R mutants grow slowly but are impaired in to quinone electron transfer. The Leu and Asp substitutions appear to destabilize and Arg apparently overstabilizes the semiquinone at the site. Destabilization would impair the ‘gated,’ two electron reduction of Q to and overstabilization would impede dissociation of from the site, both required for turnover of the complex by the Q cycle mechanism (Section V B). Interestingly, Arg occurs at the corresponding 207 position in Cyt (Nostoc ). Substitution of a basic Arg for a weakly acidic His residue at this position may be a key

determinant for different site characteristics in Cyt and complexes. However‚ relatively stable semiquinones have long been observed in Cyt complexes (Siedow et al.‚ 1978; Ohnishi and Trumpower‚ 1980; de Vries et al.‚ 1983) but only recently in the complex (Pace et al.‚ 1992) and there is currently no evidence for a more highly stabilized semiquinone in the than the complex. Thus additional elements probably have roles in determining the overall properties of the sites in these complexes. Yun et al. (1992) have mutagenized five other highly conserved residues‚ Rhodobacter sphaeroides G48‚ Q58‚ S102‚ F104‚ and P202 (Nostoc sp. strain PCC 7906 G37‚ Q47‚ S91‚ M93‚ and P192‚ respectively)‚ with likely roles in structure or catalysis (Table 11). The small aliphatic residues G48/G62 in helix I and G132/G146 in helix III (Nostoc sp. strain PCC 7906 and respectively‚ see Fig. 3) are postulated to allow these helices to pack close to the heme-liganding helices II and IV (Tron et al.‚ 1991b). Consistent with this idea‚ G48A is indistinguishable from wild-type‚ substitution of the considerably larger Val (G48V) prevents Cyt oxidation‚ and in G48D the complex is not assembled (Yun et al.‚ 1992). The S102A‚ F104I‚ and P202L mutations‚ surprisingly‚ had little effect. Alteration of the strictly conserved P202 presumably removed the bend from within helix IV Phe 104 was of interest because it lies in helix II equidistant between hemes and and has been suggested as an electron carrier between them. The absence of Phe at this position (Met93 in Cyt has been suggested as a basis for restricted interheme electron transfer in the Cyt complex (Cramer et al.‚ 1987; Furbacher et al.‚ 1989). The lack of any major effect in the F104I mutant argues against an electron transport role for Phe 104 (Yun et al.‚ 1992). It may be significant‚ however‚ that F105 lies next to F104 in Cyt b. The former could possibly serve in place of F104 as the electron carrier between the b-hemes in the complex.

D. The Rieske Iron-Sulfur Protein Numerous mutations have now been mapped to or generated within genes for the Rieske proteins from yeast and Rhodobacter capsulatus (Tables 12 and 13). These organisms offer the clear advantage of growth in the absence of the Cyt complex and mutants have been obtained in three ways. Firstly‚ respiratory defective (null or petite)‚ yeast mutants

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have been generated by chemical mutagenesis and identified as Rieske mutants by noncomplementation in crosses with a tester strain carrying an inactivated Rieske gene (Gatti et al.‚ 1989). These are listed in Table 12 and the altered residues marked with dots in the corresponding Nostoc sp. strain PCC 7906 Rieske structure shown in Fig. 4. Secondly‚ a yeast Rieske deletion strain has been constructed (Beckmann et al.‚ 1989) and respiratory growth restored by complementation with in vitro‚ randomly mutagenized plasmids carrying the Rieske gene. This has

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permitted screening for both null (Graham et al.‚ 1993) and temperature-sensitive (Beckmann et al.‚ 1989; Ljungdahl et al.‚ 1989) mutations. These are listed in Table 12 and marked in Fig. 4 with dots or bolded‚ respectively. Thirdly‚ site-directed mutations in Rieske genes have been introduced into yeast (Graham and Trumpower‚ 1991; Graham et al.‚ 1993) and Rhodobacter capsulatus (Davidson et al.‚ 1992a‚ 1992b) Rieske deletion strains. These are listed in Table 13 and marked in Fig. 4 with arrows. It should be noted that several mutations (listed in Table 12 or

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its footnote) map to the nonconserved N-terminal domain and therefore corresponding sites cannot be located on the Nostoc sp. strain PCC 7906 Rieske model (Fig. 4). These mutations (e.g.‚ the temperaturesensitive T85I‚ I101T‚ and P102L in the amino-

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terminal hydrophobic domain‚ Table 12) generally have less severe effects on activity than those in carboxyl-terminal regions and some show increased resistance to myxothiazol suggesting interaction of this region with the site in Cyt b (Ljungdahl et al.‚

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1989; Graham et al.‚ 1993). I101T and P102L additionally block post-translational processing of the intermediate to the mature form of the yeast Rieske protein (Graham et al.‚ 1993; Section VI B). These results are consistent with ‘domain-swapping’ (Huang et al.‚ 1991) and other experiments (e.g.‚

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González-Halphen et al.‚ 1991) discussed in Section IV Dsuggestingimportant‚species-specificfunctions for the amino-terminal region ofthe Rieske‚ probably in assembly and intersubunit interactions. Most of the null‚ and temperature-sensitive‚ respiratory-defective mutations in the Rieske occur

302 in strictly or highly conserved residues within the carboxyl-terminal Fe-S binding domain or in the adjacent distal region (Fig. 4). The latter (approximately positions 133 to 151 inNostoc sp. strain PCC 7906) is suggested‚ because it contains Gly and Pro residues‚ to serve as a flexible domain which together with more distal residues may ‘cap’ the Fe-S pocket (Graham et al.‚ 1993). A number of mutations in this domain as well as in the ‘box II’ Fe-S binding region result in loss of all catalytic activity and lower amounts of the Rieske protein and the Fe-S center‚ consistent with important roles for residues in these regions. For example‚ any mutation in the Cys or His residues of box II causes loss of function (Graham et al.‚ 1993) and loss of the Rieske protein in all cases tested (e.g.‚ C178Y‚ Gatti et al.‚ 1989). S183L‚ D186N‚ and G189D (Nostoc sp. strain PCC 7906 S131‚ D134‚ and G137‚ respectively) in the adjacent ‘flexible’ domain also caused loss ofthe Fe-S center and activity but allowed retention of small amounts of the Rieske protein (Gatti et al.‚ 1989). These mutations demonstrate the importance of residues other than the immediate Fe-S center ligands for integration and or stability of the Fe-S center. Temperature-sensitive mutations in other strictly conserved residues (e.g. T160A‚ V165A‚ A170T corresponding to Nostoc sp. strain PCC 7906 T109‚ V114‚ and V119‚ respectively) have only minor effects although activity at the permissive temperature is somewhat lower in each case (Graham et al.‚ 1993). The G175S (Nostoc sp. strain PCC 7906 K123) mutation permits 20% of wild-type activity at the permissive temperature (Ljungdahl et al.‚ 1989) but G175D (to the negatively charged Asp) is null (Graham et al.‚ 1993). The Cyt Rieske protein has a positively charged Lys (K) at this important site in the loop between box I and box II suggesting one possible basis for differences in the properties of Rieske Fe-S centers from Cyt and complexes. Other mutations point to stringent requirements for particular residues or charged groups at specific positions. For example‚ aliphatic to negatively charged‚ G189D (Nostoc sp. strain PCC 7906 G137) mentioned above; positively charged to aliphatic at the adjacent position R190G (Nostoc sp. strain PCC 7906 K138); or small volume to larger aliphatic‚ A196V (Nostoc sp. strain PCC 7906 A144)‚ all resulted in complete loss of activity (Table 12). All site-directed mutations in the Rieske (listed in Table 13) have been aimed at identification of Cys and His ligands for the 2Fe-2S center as discussed in

Toivo Kallas Section IV D. Substitutions for any of the strictly conserved Cys or His residues in box I and II ofthe yeast Rieske resulted in loss of the Fe-S center but did not prevent processing and assembly of the Rieske protein into the complex (Graham and Trumpower‚ 1991). In a somewhat different result‚ mutagenesis of any of the conserved Cys or His (except C155 corresponding to Nostoc sp. strain PCC 7906 C128) in Rhodobacter capsulatus caused loss of the Fe-S center and also greatly decreased the amount of Rieske protein in the membrane (Davidson et al.‚ 1992a). The C155S mutation permitted synthesis of a small quantity of the Rieske protein which contained an altered 2Fe-2S cluster with a midpoint redox potential 180 mV lower than that of the wild-type. A His residue has been conserved in the position distal to box II (H184 in yeast and H159 in Rhodobacter capsulatus) in the Rieske from but not complexes where Gln (Nostoc sp. strain PCC 7906 Q132) occupies this site. H184R in yeast (Graham and Trumpower‚ 1991) and H159A and S mutants of Rhodobacter capsulatus (Davidson et al.‚ 1992a) all had properties similar to the wild-type. This position appears to tolerate diverse residues suggesting that the Gln/His difference in Cyt versus complexes may not be of functional significance.

E. Prospects for Genetic Analysis of the Cytochrome Complex from Cyanobacteria Because of the similarity of the Photosynthetic apparatus to that in chloroplasts‚ and the relative ease of genetic manipulation in several strains‚ cyanobacteria have proven enormously useful for mutational analysis ofPS II and PS I (see Chapters 8 and 10). Genetic studies of the cytochrome complex have lagged far behind for the principal reason that this complex is required‚ based on all current indications‚ for Photosynthetic as well as all other known modes of growth in cyanobacteria (Section V D). Accordingly‚ one cannot make lethal mutations nor perform reverse genetics by the proven‚ ‘straightforward’ approach of complementation of deletion strains with in vitro mutated copies of the corresponding genes. Continued refinements in chloroplast transformation and gene replacement by particle bombardment (e.g. Roffey et al.‚ 1991; Takahashi et al.‚ 1991b; Weeks‚ 1992) have now made site-directed mutagenesis of Chlamydomonas reinhardtii chloroplast genes an almost routine‚ if still time-consuming

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and sometimes difficult process. As discussed above‚ the plastid petA‚ B and D genes have been cloned and sequenced from Chlamydomonas reinhardtii (Büschlen et al.‚ 1991; Bertsch and Malkin‚ 1991) and very recently‚ site-directed mutations in Cyt (F. A. Wollman‚ personal communication) and Cyt f (J. Zhou and R. Malkin‚ personal communication) obtained and characterized from homoplasmic segregants lacking the wild-type gene copies. These are the first characterized mutations in any structural gene for the Cyt complex and Chlamydomonas currently appears to be the organism best amenable for studies of the chloroplast encoded Cyt subunits. It offers the advantage of growth in the absence of the Cyt complex and chloroplast gene replacements are possible. The advantages of Chlamydomonas reinhardtii for studies of the nuclear encoded Rieske protein are not as apparent. Nuclear gene replacements in Chlamydomonas reinhardtii are highly uncertain at best and there is currently no stable mutant lacking the Rieske protein. Because of greater ease and rapidity of genetic manipulations‚ cyanobacteria should still be useful for in vivo analysis of nonlethal mutations in the Cyt complex‚ particularly in the case of the Rieske subunit. It should be possible to obtain segregants bearing such mutations by linking mutations to selectable markers inserted into neutral sites flanking cyanobacterial pet genes. Discovery or engineering of a cyanobacterium with an active quinol oxidase would solve the problem of an alternative pathway for mutant propagation and such a strain would rapidly become the organism of choice for genetics of the Cyt complex.

VIII. Unresolved Questions and Perspective As discussed above‚ the cytochrome complex plays a central role in electron transport in cyanobacteria. Issues specific to cyanobacteria include the nature and extent of this role in different strains. Where is the complex localized within cyanobacterial cells and with what other complexes and carriers does it interact? Are there alternative acceptors to plastocyanin and Cyt as suggested in Section IV E? Are there alternative electron transport pathways that might bypass the Cyt complex altogether? Does the complex act as a redox sensor to mediate state transitions or balance noncyclic versus cyclic electron transport pathways?

303 If so‚ what is the molecular basis for this signal transduction function? Issues of more general significance include the pathways for electrons and protons. How do electrons re-enter the Cyt complex during cyclic transport and what domains and other proteins are involved? Does inter-heme electron transfer and the Q-cycle occur obligatory during turnover or does transfer through the high potential chain > Fe-S > Cyt f) or by another mechanism such as the semiquinone cycle predominate under certain circumstances? Is the active form of the complex in vivo a monomer or dimer and do transitions between these configurations regulate pathways of electron transfer? Are there roles for low molecular weight subunits? What are the mechanisms for attachment of hemes and the Fe-S center and for assembly of the complex? A central issue of broad significance remains the extent and basis for structural and functional similarity/dissimilarity between the Cyt and complexes. Dissimilarities include inhibitor specificities‚ biophysical properties of redox centers‚ differences in turnover rates‚ and possible differences in electron transfer pathways. Elucidation of structure/ function in the Cyt complex would be greatly advanced by further progress in two areas: firstly‚ by mutational analysis of the Cyt complex coupled with biochemical and spectroscopic analysis of mutants; secondly‚ by solution of the three dimensional structures of Cyt and complexes or component proteins. The recent solution of the Cyt f structure at high resolution (Cramer et al.‚ 1994; and S.E. Martinez‚ D. Huang‚ A. Szczepaniak‚ J.L. Smith‚ and W.A. Cramer‚ personal communication) underscores the importance of this type of information. Spectroscopic evidence had suggested Lys145 as the sixth axial ligand for heme f. Recent site directed mutagenesis experiments in Chlamydomonas reinhardtii showed Lys 145 not to be important (J. Zhou and R. Malkin‚ personal communication) but the Cyt f structure ultimately revealed that the amino-terminal amino acid of the protein serves as the axial ligand. This unexpected and novel outcome could not readily have been predicted by any other means of analysis. Three dimensional structures provide the rational framework for subsequent high resolution mutational and biochemical studies and we can look forward to future insights provided by these combined approaches‚ some of which will undoubtedly be performed with cyanobacteria.

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Acknowledgments I would like to thank a great number of colleagues for responding to requests for reprints and particularly Drs D. A. Bryant‚ W. A. Cramer‚ A. R. Crofts‚ F. Daldal‚ R. B. Gennis‚ J. H. Golbeck‚ K. A. Gray‚ D. M. Kramer‚ U. Liebl‚ R. Malkin‚ S. R. Mayes‚ M. Rögner‚ Y. Shahak‚ B. L. Trumpower‚ G. Tsiotis‚ J. Whitmarsh‚ F. A. Wollman‚ and J. Zhou for sharing data prior to publication. The quinol-oxidation site model was constructed by use of the ‘Hyperchem’ molecular modeling program in collaboration with Dr Robert Moore‚ UW-Oshkosh Chemistry Department. Work from my laboratory was supported by RUI Grant DMB-8902695 from the National Science Foundation and by UW-Oshkosh Faculty Development Award R128.

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PsaC and PsaD proteins expressed in Escherichia coli. FEBS Lett 276: 175–180 Zhao J‚ Li N‚ Warren PV‚ Golbeck JH and Bryant DA (1992) Site-directed conversion of a cysteine to aspartate leads to the assembly of a [3Fe-4S] cluster in PsaC of Photosystem I. The photoreduction of is independent of Biochemistry 31: 5093–5099

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Chapter 10 Photosystem I in Cyanobacteria John H. Golbeck Department of Biochemistry, Center for Biological Chemistry, University of Nebraska, Lincoln, NE 68583-0718, USA Summary I. Introduction II. Unifying Principles III. Architecture of Photosystem I A. Nomenclature B. Polypeptide Composition C. Electron Microscopy D. X-Ray Crystal Structure 1. Polypeptides 2. Cofactors IV. Integral Polypeptides A. PsaA/PsaB Heterodimer 1. Structure 2. Function B. PsaF 1. Structure 2. Function C. Psal 1. Structure 2. Function D. PsaJ 1. Structure 2. Function E. PsaK 1. Structure 2. Function F. PsaL 1. Structure 2. Function G. PsaM 1. Structure 2. Function V. Peripheral Polypeptides A. PsaC 1. Structure 2. Function B. PsaD 1. Structure 2. Function C. PsaE 1. Structure 2. Function Acknowledgments References D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 319–360. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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John H. Golbeck

Summary The Photosystem I (PS I) complex in cyanobacteria functions most typically as a light-driven, cytochrome oxidoreductase. The adaptability of cyanobacteria to conditions of nutrient availability allows cytochrome to be replaced by plastocyanin when copper is plentiful, and ferredoxin to be replaced by flavodoxin when iron is limiting. These changes, however, do not lead to any known alterations in the polypeptide composition ofthe membrane-bound PS I complex. This multiprotein complex incorporates all of the biochemical machinery required to produce efficient charge separation across the thylakoid membrane in a process that culminates in the conversion of a red photon to chemical free energy. The membrane-bound components which comprise the complex include an array of~ 110 antenna chlorophyll a molecules to provide a large optical cross-section to incoming photons, a series of inorganic and organic cofactors to carry out the acts of charge separation and charge stabilization, and a matrix of eleven polypeptides to provide ligands to the photoactive components. These components are arranged in a motifbelieved to be shared by all photochemical reaction centers: in PS I a chlorophyll (a) dimer serves as the primary electron donor; a chlorophyll (a) monomer serves as the primary electron acceptor; and a quinone (phylloquinone) serves as the intermediate electron acceptor. Other common features include the presence ofa protein (hetero)dimer (PsaA and PsaB), which binds the antenna chlorophylls, the electron donor and acceptor chlorophylls, and the two quinone molecules. This shared photochemical motif is broken by the inter-polypeptide iron-sulfur cluster which occupies the same relative position as the non-heme iron in Type-II (quinone-type) reaction centers, but which is redox active in Type I (iron-sulfur type) reaction centers. The addition of two iron-sulfur clusters, and located on a separate polypeptide, PsaC, provides a path for the electrons out ofthe membrane phase and to the stromal phase, allowing ferredoxin to be reduced with high quantum efficiency. The other PS I polypeptides serve ancillary roles in stabilizing PsaC and docking ferredoxin or flavodoxin (PsaD), in enhancing ferredoxin reduction and allowing for cyclic electron flow (PsaE), and in forming trimers of the PS I complex in the membrane and facilitating state transitions (PsaL). The functions of the remaining polypeptides, PsaF, Psal, PsaJ, PsaK, and PsaM, are unclear; however it is increasingly unlikely that they participate directly in the primary processes of photochemical energy conversion.

I. Introduction The PS I reaction center is a membrane-bound, multisubunit enzyme found in all naturally-occurring photosynthetic cyanobacteria and plants. It is classified according to IUPAC rules as a light-driven plastocyanin:ferredoxin oxidoreductase. Its function, implicit in the name, is to transform part of the energy of a photon into chemical free energy by oxidizing plastocyanin and reducing ferredoxin against a steep thermodynamic gradient. Decades of detailed study have shown that the photoconversion process is remarkably efficient: the quantumyield (the number of charge separation events per absorbed photon) is 1.00 ± 0.02 (Kok, 1973), and the efficiency of photoconversion of a red photon (equivalent to Abbreviations: Chl – chlorophyll; DCMU – 3-(3,4-dichlorophenyl)-1,1-dimethylurea; EDC – N-ethyl-3-[3-(dimethylamino)propyl] carbodiimide; ESP – electron spin polarization; EPR – electron paramagnetic resonance; LHC – light-harvesting chlorophyll protein complex; NMR– nuclear magnetic resonance ; PMS – N-methylphenazonium 3-sulfonate; rms – root mean square; SDS – sodium dodecylsulfate; zfs – zero-field splitting

1.76 eV) is about 43%. The detailed mechanism by which this photochemical conversion process is carried out in cyanobacteria is the topic of this chapter. It is probably accurate to state that while the functional aspects of PS I have been the primary focus of research in the 1970s and early-to-mid 1980s, structural aspects have attracted the most attention in the past several years. This renewed emphasis on structure is due, in part, to the successful determination of the 3-dimensional structures of the soluble electron transfer proteins which function as donors and acceptors to the membrane-bound PS I complex: plastocyanin, ferredoxin, flavodoxin, and oxidoreductase (see Chapter 12). More recently, the sustained efforts of several laboratories to crystallize the cyanobacterial PS I complex has led to a major success in the determination of its 3-dimensional structure at a current resolution of 6 Å (Krauß et al., 1993). Additionally, attempts are now underway to solve the 3-dimensional structures of the peripheral polypeptides PsaC, PsaD, and PsaE at a higher resolution, and an NMR solution structure of PsaE

Chapter 10 Photosystem I Reaction Center has recently become available (see Section V C below). The emphasis on structure is also a natural consequence of the successful use of molecular genetics, including deletion mutagenesis of the PS I polypeptides and site-specific mutagenesis of the ligands to the inorganic and organic cofactors. The biochemical resolution of the PS I cofactors and peripheral stromal polypeptides, and the successful rebuilding of a functioning reaction center (see Golbeck, 1993c) has permitted reconstitution experiments with non-native and geneticallymodified proteins. This technique has the potential to uncover the functions of the hydrophilic stromal proteins, and to unravel the route of non-cyclic as well as cyclic electron flow. Finally, the promise is that deletion mutagenesis will continue to uncover the functions ofthe low molecular mass hydrophobic polypeptides. It is perhaps significant that many of the above advances have occurred using cyanobacteria as the target organisms. This purpose of this chapter is to summarize and review the cyanobacterial PS I literature of the last 5 years. It is primarily an update to the large body of knowledge that already exists on the structure and function of PS I in cyanobacteria, green algae and plants. For a more comprehensive examination of the PS I architecture, the reader is directed to general review articles by Malkin (1982, 1987), Nelson (1987), Mathis and Rutherford (1987), Golbeck (1987), Golbeck and Bryant (1991) and Guikema et al. (1993). Specialized articles concerning Photosystem I polypeptides are provided by Reilly and Nelson (1988), Margulies (1989), Scheller and Møller (1990), Chitnis and Nelson, 1991, Almog et al., (1992), Ikeuchi (1992), and Bryant (1992); and articles dealing primarily with electron transfer components are covered by Lagoutte and Mathis (1989), Evans and Bredenkamp (1990), Sétif (1992), and Golbeck (1992, 1993a). An analog of PS I exists in the anaerobic green sulfur bacteria including Chlorobium limicola f. thiosulfatophilum, Chlorobium phaeobacteroides, Chlorobium vibriforme, (Nitschke et al. 1990a; Miller et al., 1992; Feiler et al., 1992; Kjær et al., 1994), and in anaerobic, gram-positive bacteria such as Heliobacterium chlorum, Heliobacterium gestii, and Heliobacillus mobilis (Trost and Blankenship, 1989; Nitschke et al., 1990b). The degree to which these reaction centers are homologous to the cyanobacterial and plant PS I reaction centers is still under active investigation. At this point, the bacterial analogs

321 serve to reinforce findings on the cyanobacterial and plant PS I core, and to stimulate interest in the evolutionary precursors of the two photosystems. For further information, the reader is directed to three recent review articles on the evolutionary relationship between the bacterial reaction centers and Photosystem I (Nitschke and Rutherford, 1992; Golbeck, 1993b; Lockau and Nitschke, 1993).

II. Unifying Principles As a discipline of biochemistry, the field photosynthesis became unified at the point when the ‘ironsulfur’ type photosystems (PS I, the green sulfur bacterial reaction center and the heliobacterial reaction center) and the ‘quinone-type’ photosystems (PS II and the purple non-sulfur bacterial reaction center) were recognized to share common structural and functional features in the early events of charge separation. These features include a (hetero or homo) dimeric protein core which contains antenna chlorophylls and a rigidly-spaced series of cofactors that function to produce and stabilize the photochemically-generated charge-separated species. In PS I, these initial events can be depicted in the following two step sequences:

where P700 is primary electron donor, a chlorophyll a dimer; is the electron acceptor, a chlorophyll a monomer; and is the secondary electron acceptor, a bound molecule of phylloquinone (2-methyl-3phytyl-1,4-naphthoquinone). According to this scheme, the absorption of a photon by one of the resident 100 chlorophyll antenna molecules results in transient charge separation between P700 and The electron on is rapidly transferred to the bound phylloquinone, which stabilizes the separated charges against the rapid, and inevitable, charge recombination that would occur between the primary electron donor and acceptor pair. This threecomponent photochemical motif is present in all known photochemical reaction centers; the differences in the reaction centers lie in the precise identity of the cofactors, in the redox potentials of the components, and in the function of the secondary electron donors and acceptors on the stromal and lumenal phases of the membrane. Where the ‘ironsulfur type’ and ‘quinone-type’ reaction centers specifically differ is the point at which the electron is

322 transferred from the primary quinone to either a secondary quinone or an iron-sulfur cluster. The non-heme iron in the protein boundary between the heterodimeric core may be the key component which distinguishes the two reaction centers in terms of function. In the ‘quinone-type’ reaction center, a single non-heme iron is ligated with histidine ligands provided by each of the reaction center polypeptides (D1 and D2), and the iron is redox-inactive. The electron is transferred from the primary plastoquinone across the heterodimeric protein boundary to the secondary plastoquinone which functions as a two electron (and two proton) accumulator [for details, see Chapter 8]. One consequence of this architecture is that in the ‘quinone-type’ reaction centers, the electron remains located within the boundaries of the lipid bilayer by virtue of the hydrophobic nature of plastohydroquinone. In Photosystem I, and most

John H. Golbeck likely in the green bacterial and the heliobacterial reaction centers, an interpolypeptide [4Fe-4S] cluster is positioned roughly analogous to the non-heme iron in Photosystem II, and it is redox-active under physiologically-relevant conditions (Fig. 1). This cluster, known as may have evolved with relatively high midpoint potential to preclude the electron from being transferred to the second phylloquinone (at this writing, the function of the second phylloquinone in Photosystem I is unknown). The electron is subsequently transferred from to the and iron-sulfur clusters as depicted below.

The sequence of electron flow between the three

Chapter 10 Photosystem I Reaction Center iron-sulfur clusters is not known with precision; neither is it known whether they function in cyclic flow. Nevertheless, thispathway enables the electron to be transferred over a sufficiently long distance to reduce ferredoxin and flavodoxin, soluble electron carriers located in the stroma. According to this line of thinking, the mechanism that governs the fate of the electron on the reducing side of the reaction center is the salient feature that separates the ‘quinone-type’ from the ‘iron-sulfurtype’ reaction centers. The more obvious differences between the reaction centers lie in the structure ofthe proteins which serve to modulate the redox potentials of the photoactive components, and in the identities ofthe secondary electron donors and acceptors which have evolved to provide the specialized functions of water oxidation in PS II and reduction in PS I. Nevertheless, this close correspondence in function does not necessarily imply an evolutionary link between the two types of reaction centers. Attempts to find a significant homology between the sequences of PsaA/B and D1/D2 or the L/M polypeptides have failed (Nugent et al., 1993), but it is possible that the constraints on transmembrane proteins that function in vectorial electron transport have led to a common motif in terms ofcofactors and geometry. The large majority of spectroscopic studies have been performed on PS I complexes isolated from higher plants rather than cyanobacteria. It might be assumed from the previous discussion that the cyanobacterial and higher plant PS I complexes would be so much alike that the source of the biological material would be of little relevance for most studies. However, results on the electron transfer events surrounding have not been totally consistent between plants and cyanobacteria, and the kinetics of electron transfer have been found to depend on parameters such the detergent used to isolate the complex, the number of chlorophylls per P700, and the presence of secondary electron acceptors. Further, many of the spectra are differences between oxidized and reduced species in which inherent assumptions limit the degree of accuracy. The reader is cautioned to take note ofthe species and experimental conditions when electron transfer events are discussed among the components P700, and The majority of this article focuses on the architecture and polypeptide composition of the cyanobacterial PS I complex.

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III. Architecture of Photosystem I

A. Nomenclature Although the nomenclature used for the genes encoding PS I components has been standardized (psaA through psaM) (Hallick, 1989), the nomenclature for the PS I proteins themselves has not. As a consequence, the literature contains a variety of names for the polypeptides, which currently range from the form ‘PsaX’ and ‘PsI-X’, where X is derived from the locus designation of the gene, to ‘Subunit X’, where X is a Roman numeral identifying a polypeptide with some relative on an SDSpolyacrylamide gel. In this article, the names of the PS I polypeptides are derived from the cyanobacterial gene loci according to a bacterial convention established by Demerec et al. (1966); hence, the psaA gene encodes the PsaA protein (see also Dure, 1991). It should be noted that the same cyanobacterial species can be represented by different names; this is probably true here of Synechococcus sp., Synechococcus vulcanus and Synechococcus elongatus, that minimally represent very closely related, if not identical, species.

B. Polypeptide Composition A compilation of the published amino acid sequences of PS I polypeptides from Synechococcus sp. strain PCC 7002, Synechococcus sp. strain PCC 6301, Synechococcus sp., Synechococcus vulcanus, Synechococcus elongatus, Synechocystis sp. strain PCC 6803, Nostoc sp. strain PCC 8009, Anabaena sp. strain ATCC 29413, and one or two additional species has resulted in a consensus that the cyanobacterial reaction center consists of 11 discrete proteins labeled PsaA through PsaF and PsaI through PsaM (a recently discovered 4.8 kDa protein in cyanobacterial PS I complexes, ‘PsaO’ (formerly referred to as PsaN by Bryant, (1992) may represent the twelfth polypeptide (Koike et al., 1989; Nyhus et al., 1992). Table 1 lists the number of residues in each polypeptide, the mass, the presence ofcofactors, a function, and whether or not the protein is predicted to have transmembrane helices in PS I from Synechococcus sp. strain PCC 7002. One important distinction from higher plants and algae is that the latter may not contain PsaM; although a psaM homolog exists in the chloroplast genome of

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Marchantia polypmorpha (Ohyama et al., 1986), no such homolog exists in the tobacco chloroplast genome (Shinozaki et al., 1986). Moreover, the protein has not yet been identified in isolated PS I complexes from higher plants. However, higher plants and algae contain PsaG, PsaH, and PsaN, a newlydiscovered 9 kDa lumenal protein (Knoetzel and Simpson, 1992; He and Malkin, 1992). Those proteins exclusive to higher plants and algae will not be discussed further.

C. Electron Microscopy High-resolution electron microscopy of PS I complexes has provided information on the presence of trimers and monomers, and more recently, on the location of the extrinsic polypeptides PsaC, PsaD, PsaE and PsaF. Recent studies in cyanobacteria have involved examination of both negatively stained monomers and trimers isolated from Phormidium laminosum by detergent solubilization (Ford, 1987; Ford et al., 1987). The monomers appear as pearshaped, ellipsoidal objects about 14.8 × 9.1 nm, and the trimers appear as disk-shaped, triangular objects about 18.5 nm (edge length) × 6.2 nm (thickness).

John H. Golbeck

The ‘bottom’ face of the trimer has no obvious structural features, but the ‘top’ face exhibits a chiral organization with a protrusion from each monomer projecting inward toward the central stain cavity (Ford and Holzenberg, 1988). In addition, the PS I trimers form long stacks of disks, which are the result of face-to-face pairing of the individual trimers. The dimensions of Synechococcus sp. PS I are 15.3 × 10.6 × 6.4 nm and the shape is that of a prolate ellipsoid (Boekema et al., 1989; Rögner et al., 1990). These dimensions are roughly compatible with a molecular mass of 300 to 400 kDa. The cyanobacterial PS I complex contains 260 kDa of protein (PsaA through PsaF and PsaI through PsaM), 98 kDa of chlorophyll (110 molecules of Chl a), 1.0 kDa of iron and sulfur (three [4Fe-4S] clusters), and an unspecified amount of and this corresponds to a total mass of 359 kDa. Hence, the particles examined were PS I monomers rather than trimers. The dimensions of the Synechococcus sp. trimer are similar to those described for Phormidium laminosum with a diameter of 19 nm and a thickness of 6.5 nm. Trimers of similar size have also been isolated by isoelectric focusing from the marine cyanobacterium Synechococcus sp. strain PCC 7002

Chapter 10 Photosystem I Reaction Center

(Tsiotis et al., 1993). Finally, van der Staay et al. (1993) have characterized the structure and polypeptide composition of PS I complexes from Prochlorothrix hollandica (1993; also see Chapter 3). The PS I complexes of this prochlorophyte were found to form trimers that were similar in size, appearance in the electron microscope, and polypeptide composition to those of cyanobacteria, and no evidence for the association of the LHC a/b complexes with the PS I complex was observed. Boekema and colleagues have carried out a sophisticated image analysis which reveals distinctions between the two surfaces of the PS I complex (Boekema et al., 1989). A low-resolution 3-D model was constructed from a tilt series of 2-D electron

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micrographs in which the images were processed by a combination of Fourier-peak-filtering and correlation averaging (Böttcher et al., 1992). At a resolution of 15 to 18 Å, one side (purportedly the stromal side) shows a 30 Å-high ridge which most likely represents the PsaC, PsaD and PsaE polypeptides (Fig. 2). The other side is flat, but in the center there is a 30 Å-deep indentation, which may partially separate the PsaA and PsaB subunits. This could represent the ‘docking’ site for plastocyanin and/or cytochrome on the PS I core. A comparative structural analysis was carried out on the trimeric complex that contains the full complement of polypeptides, and the trimeric core that was selectively depleted of the PsaC, PsaD and PsaE polypeptides (Kruip et al., 1993). A contour difference analysis of the two preparations indicates the relative location of these three proteins on the stromal surface of the Photosystem I complex (Fig. 3). Most notably, the height of the complex is reduced 25 to 33 Å upon the removal of the stromal polypeptides, indicating that

326 the stromal-exposed mass is composed exclusively of these three proteins. This agrees very well with earlier biochemical resolution and reconstitution experiments, that showed that only PsaC, PsaD and PsaE are removed with chaotropic agents (Li et al., 199la). When PsaE is selectively removed, the distance between stacked trimers of PS I decreases about 10 Å, and a contour difference analysis of the two preparations shows a slight change at a location near the connecting domain (M. Rögner and E. Boekema, unpublished results). The implication is that PsaE overlays PsaC and possibly part of PsaD, a conclusion which is also in agreement with crosslinking experiments (Oh-Oka et al. 1989) and biochemical resolution and reconstitution studies (Parrett et al., 1989, 1990; Zhao et al., 1993). The three-dimensional NMR solution structure of PsaE indicates that it is a rather flat molecule composed of mostly and hence it would not contribute more than 15 to 20 Å to the height ofthe complex (C. Falzone, J. Lecomte, J. Zhao, and D. A. Bryant, personal communication). There is still considerable uncertainty whether PS I trimers exist in the thylakoid membrane or are simply artifacts of detergent solubilization (Hladik and Sofrova, 1991). Studies from electron microscopy have shown that when cyanobacterial thylakoid membranes are incubated with high concentrations of the PS I complexes form monomers (Hefti et al., 1992). When the arrays are reconstituted in a lipid membrane (Ford et al., 1990), the PS I complexes remain in the form ofmonomers under all conditions studied. In contrast, deletion mutagenesis experiments and functional studies have pointed to the existence ofPS I trimers in the membrane. For example, PsaLless mutants of Synechocystis sp. strain PCC 6803 (Chitnis et al., 1993) and Synechococcus sp. strain PCC 7002 (W. M. Schluchter, J. Zhao and D. Bryant, personal communication) fail to form PS I trimers when solubilized with dodecyl maltoside or Triton X-100. Cross-linking studies performed with thylakoids of the wild-type strain and the psaL mutant exhibit differences that are consistent with the interpretation that trimers are present in the thylakoids (J. Zhao and D. A. Bryant, personal communication). The Synechococcus sp. strain PCC 7002 psaL mutant also fails to perform state transitions normally, implying that at least in cyanobacteria, trimers are necessary to accommodate an important physiological function. As expected,thepsaL mutant grows substantially slower than the wild-type strain

John H. Golbeck when grown in green light. (W. M. Schluchter and D. A. Bryant, unpublished results).

D. X-Ray Crystal Structure Due to the greater stability of proteins from thermophilic organisms, crystallization has been attempted with PS I complexes isolated from the following cyanobacteria: Phormidium laminosum (grown at 42 °C, Ford et al., 1987); Mastigocladus laminosus (grown at 50 °C; Shoham et al., 1990); and Synechococcus sp. (grown at 55 °C; Witt et al., 1990). Crystals of trimeric PS I complexes from the mesophilic, marine cyanobacterium Synechococcus sp. strain PCC 7002 have also been reported (Tsiotis et al., 1993). The high diffraction quality of the PS I crystals from the thermophilic Synechococcus sp. has led to the determination of the 3-dimensional structure of the trimeric form at a resolution of 6 Å (Krauß et al., 1993). The 1–2 mm long and 0.5 mm wide hexagonal crystals contain a large hexagonal unit cell with a = b = 287 Å and c = 167 Å; the space group is with two trimers in the unit cell and with one trimer in the asymmetric unit.

1. Polypeptides Amino-terminal amino acid sequencing of the polypeptides in the Synechococcus sp. PS I crystals has shown the presence of all but the PsaI and PsaL proteins (Witt et al., 1992). However, the psaI and psaL genes have been located in the Synechococcus sp. genome (Mühlenhoff et al., 1993), and it is fully expected that the corresponding proteins are present in the crystallized PS I complex. The electron density map shows that the overall architecture of PS I monomer is represented by two distinct domains. The smaller is a so-called ‘connecting domain’, which does not extend above or below the 40 Å thick membrane and which apparently links the PS I monomers into trimers. The larger is the ‘catalytic domain’, that has protrusions extending 35 Å and 15 Å into the stromal and lumenal phases, and that contains the components involved in photochemical charge separation. The stromal-protruding region most probably represents the relatively hydrophilic polypeptides PsaC, psaD and PsaE. The cavity on the lumenal side may represent the ‘docking site’ for plastocyanin and/or cytochrome near P700 (Krauß et al., 1993). This

Chapter 10 Photosystem I Reaction Center

cleft is also found in the image analysis of electron micrographsofthe2-D crystalsoftheSynechococcus sp. PS I complex (Böttscher P. et al., 1992). By localizing among the three electron-rich regions (12 r.m.s. deviations above the mean density) that represent the iron-sulfur clusters, the remaining electron density (approximately 1.2 r.m.s. deviations above the mean density) canbe fitted withpolyalanine helices and disks (representing the tetrapyrrole Chls) to show a 2-fold rotation axis running through the cofactors P700 and iron-sulfur center A total of 28 can be mapped onto the electron density, 21 of which are transmembrane helices oriented at angles between 3° and 30°to the membrane-normal (Fig. 4 A). Eight of the transmembrane helices (labeled a to h) are symmetry-related with eight other helices (labeled a’ to h’), and are likely to be derived from PsaA and PsaB subunits (Fig. 4B). This analysis of the catalytic domain in the crystal structure confirms that PS I is a protein heterodimer, as had been inferred earlier from consideration of the protein composition and cofactor stoichiometry, from the expected dimeric nature of P700, and from the known inter-polypeptide location of In addition, two helices run parallel to the membrane on the lumenal side ofthe reaction center, and these are also probably also derived from PsaA and PsaB (Fig. 4A). Five additional helices (labeled

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i, j, k. l, m) have no symmetry equivalents; however, tentative assignments can be made. Helices j and k are close to the three-fold axis, implying a role in connecting the monomers to trimers. The electron density indicates that they may be linked on the stromal side, which implies a protein large enough to pass twice through the membrane. The only low molecular mass polypeptides which are large enough to contain two transmembrane helices are PsaK. and PsaL. A PsaK-less mutant of Synechococcus sp. strain PCC 7002 forms predominantly trimers (and the same percentage as the wild-type strain) of PS I in solution (W. M. Schluchter, J. Zhao, L. Yu, J. H. Golbeck and D. A. Bryant, unpublished results). However, PsaL-less mutants of Synechocystis sp. strain PCC 6803 (Chitnis. et al. 1993) and Synechococcus sp. strain PCC 7002 (W. M. Schluchter, J. Zhao, and D. A. Bryant, personal communication) form only monomers in solution. Hence, it is likely that PsaL represents the protein containing the j and k helices close to the 3-fold axis, that organizes the PS I monomers into trimers. This assignment is also consistent with protease studies which show that PsaL has stromal-exposed regions (Zilber and Malkin, 1992). As shown in Fig. 4A, helix i is rather long and protrudes through the membrane to the stromal side. There are two probable candidates for this protein:

328 PsaJ and PsaF. PsaJ has a hydrophilic region at the carboxyl-terminal end of the predicted but otherwise no other information is available. Hydropathy profiles of PsaF suggest that it also contains one transmembrane located near the middle of the protein (similar to PsaJ, there is also a hydrophilic region at the carboxyl-terminus). It was recently found that PsaF can be cross-linked with PsaE, and that PsaF is easily removed with detergents in a PsaE-less mutant (U. Mühlenhoff, J. Zhao and D. A. Bryant, personal communication); this requires that PsaF has a stromal-protruding segment on the transmembrane The protrusion of the i helix on the stromal side near the connecting domain also agrees with the location of PsaE deduced from electron microscopic studies; hence, the data suggest that this helix could represent the transmembrane portion of PsaF. The large, broken (Fig. 4) on the stromal side that appears to come in close contact with the iron-sulfur clusters may be part of an extended structure of PsaD. While there is yet no 3-dimensional structure, the close contact with the electron density that represents PsaC, and the location on the stromal side distal to the connecting domain, is consistent with the identity of the stromal helix as PsaD. A cross-linking product of PsaD to a small, unidentified protein of approximately 8–10 kDa could suggest that the m helix is derived from PsaK; this suggestion can be tested with the PsaK-less mutant of Synechococcus sp. strain PCC 7002 (U. Mühlenhoff and D. A. Bryant, personal communication). It should also be noted that the PsaE protein contains only and loop structures and no segments (see below and Fig. 8). The results from electron microscopy also make it unlikely that the three broken helices on the stromal side represent any protein other than PsaD (Kruip et al., 1993). If one assumes that PsaA and PsaB each contain between 9 and 11 that PsaL and PsaK each have two transmembrane and that PsaF, PsaI, PsaJ and PsaM each have one transmembrane the electron density map of the PS I complex should reveal a total of 26 to 30 helices. As indicated above, 28 are found in the crystal structure, but this includes the three broken on the stromal side that may be part of PsaD; hence, a maximum of 25 are likely to be represented by the hydrophobia PS I polypeptides. The variance between the number of helices found in the crystal structure and the number expected indicates either

John H. Golbeck that one to five remain to be found in a more refined structure or that the hydropathy profiles have overestimated the number of represented by the hydrophobic PS I polypeptides.

2. Cofactors Of the 90 to 110 Chl a molecules present in the cyanobacterial PS I complex, 45 have been assigned to the electron density map. Most of the Chls appear to be oriented similar to P700 (Fig. 5A), with their dihydroporphyrin planes perpendicular to the membrane. Even with only one-half identified, it is still possible to observe chains of chlorophylls, separated by 8 to 15 Å, that run across the thylakoid membrane in a manner reminiscent of the Chls of LHC II (Kühlbrandt and Wang, 1991). Several of the Chl molecules have a catalytic rather than lightharvesting role. The primary donor Chl is likely to be close to the 2-fold rotation axis and near the lumenal depression, allowing it to accept an electron from either plastocyanin or cytochrome Its depiction as two dihydroporphyrin planes separated by 9 Å (Fig. 5B) and normal to the membrane is consistent with existing spectroscopic evidence for the orientation of P700. Two additional Chl molecules, that have not been found spectroscopically but which may be analogous to the bridging, monomeric bacteriochlorophylls of the purple bacterial RC, are found 11 Å from P700. The primary acceptor Chl, is thought to correspond to the electron density 12 Å distant from one of the bridging Chl molecules. Although this is a quite reasonable assignment, the electron density in a symmetry-related position appears insufficient to map the expected Chl molecule on the proposed second chain of cofactors on the PS I complex (see Fig. 1). This position instead has been assigned to the naphthoquinone head group of one of the two resident molecules of phylloquinone (Krauß et al., 1993). If this assignment bears out at higher resolution (the best crystals diffract to 3.9 Å; P. Fromme, personal communication), the symmetry will have been broken early in the electron transport pathway. The two white dots in Fig. 5A show an alternative location in the electron density map for phylloquinone. The distance from the center of the left dot to is 14 Å and the distance from the center of the right dot to is 16 Å (N. Krauß, personal communication). This properly takes into account the stoichiometry of phylloquinone (two per P700)

Chapter 10 Photosystem I Reaction Center

and renders the quinones more similar to and of the purple bacterial reaction center. However, even at this alternate location, the phylloquinones in PS I are buried more deeply in the membrane than the corresponding quinones in the bacterial RC (relative to the non-heme iron). It must be kept in mind that the assignment of small organic molecules such as phylloquinone is very difficult at 6 Å resolution, and that the phase error in the current electron density map is not known. The most striking feature revealed by the crystal structure is the arrangement of the and ironsulfur clusters in the form of an irregular triangle, with one cluster 15 Å, and the other, 22 Å from (Figs. 4A, 5A, and 5B). Although the crystal structure gives no hint as to which cluster represents and spectroscopic data detailed below implies that the proximal cluster is and the distal cluster is The distance of 12 Å from (the purported) to (the distance from the alternative phylloquinone locations is probably closer to 15 Å) strongly suggests that functions as an electron transfer agent from

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to This is quite unlike PS II, in which the iron can removed or replaced with a non-redox active metal with little effect on the efficiency of forward electron transfer. This distinction in roles is particularly striking the electron density map of PS I is superimposed on the Rhodopseudomonas viridis reaction center. is only 3 to 4 Å ‘higher’ in the membrane than the bacterial non-heme iron when P700 and the acceptor chlorophyll are overlaid with the special pair and acceptor chlorophyll of the bacterial reaction center (N. Krauß, personal communication), yet their functions appear quite different.

IV. Integral Polypeptides

A. PsaA/PsaB Heterodimer 1. Structure The psaA and psaB genes that encode the reaction

330 center polypeptides were first identified and sequenced from the chloroplast genomes of higher plants (Fish et al., 1985; Fish and Bogorad, 1986; Kirsch et al., 1986), but sequences are now available for Synechococcus sp. strain PCC 7002 (Cantrell and Bryant, 1987), Synechocystis sp. strain PCC 6803 (Smart and McIntosh, 1991), and the thermophilic species Synechococcus sp. (probably identical to S. vulcanus; Shimizu et al., 1992; Mühlenhoff et al., 1993). The deduced amino acid sequences of the PsaA and PsaB proteins are tabulated and compared in Tables 2 and 3. Because of the larger number of sequenced higher plant and algal genes, inferences of structure and function based on sequence comparisons are understandably weighted more heavily by the eucaryotic data. In all known cases, the cyanobacterial psaA and psaB genes are cotranscribed, and in Synechococcus sp. strain PCC 7002 the genes encode polypeptides of 740 and 734 amino acids (81383 and 81683 kDa) which are ~45% identical. The sequence similarity between the cyanobacterial and eucaryotic PsaA polypeptides is about 95%, and the Synechocystis sp. strain PCC 6803 and Synechococcus sp. (as well as eucaryotic) sequences contain a short amino-terminal extension that is absent in the Synechococcus sp. strain PCC 7002 sequence. The sequence similarity between the cyanobacterial and eucaryotic PsaB polypeptides is slightly greater than 95%. As a point of comparison, the Chlorobium limicola f. thiosulfatophilum reaction center gene encodes a protein of 730 amino acids with a calculated mass of 82 kDa, a value similar to PsaB in PS I (Büttner et al., 1992). The predicted protein is also highly hydrophobic; hydropathy analyses suggest the presence of 11 transmembrane segments, a number identical to that calculated for PsaB (the actual number of transmembrane helices may be as low as 8). Yet, the sequence similarity of the reaction center in the green sulfur bacterium is only 15% with PsaA and 14% with PsaB, which is marginally above chance (especially when the restricted set of hydrophobic amino acids common in transmembrane is considered). The similarity of the external loop regions is especially low except for the single loop between putative spans VIII and IX which contains two cysteines separated by eight amino acids (five which are identical to those in PsaA and PsaB). This sequence occurs in the same relative position on the Chlorobium limicola f. thiosulfatophilum (Büttner et al., 1992) and Heliobacillus mobilis ( Liebl et al., 1992) reaction center proteins as in PS I and represents

John H. Golbeck the likely binding site for iron-sulfur center A high degree of primary amino acid sequence similarity is likely to translate to a high degree of similarity in the secondary and tertiary structure of the protein. The obvious consequence is that the 3dimensional structure determined for Synechococcus sp. will most likely represent a general foldingpattern for procaryotes and eucaryotes alike. Even in the Chlorobium sp. and heliobacterial reaction centers, there is the expectation that the overall protein folding pattern may match that of PS I. By way of example, the overall primary sequence homology of the higher plant D1 and D2 proteins with the L and M subunits ofthe purple bacterial RC is also only about 15%, yet the three-dimensional folding pattern of the five membrane-spanning is predicted to be very similar (Trebst, 1986). Topological studies involving antibody recognition domains have been initiated using higher plant PS I complexes. Immunogold labeling studies with antibodies prepared against synthetic peptides have identified residues of PsaA and PsaB in maize PS I complexes which are exposed to the stromal or lumenal surfaces (Vallon and Bogorad, 1993). Assuming that PsaA and PsaB have 11 membranespanning the labeling of residues 371– 379, 413–421, and 497–505 (numbering system in maize) are consistent with models that have the amino-terminus and the regions exposed to the stromal surface. Studies with polyclonal and domain-specific antibodies are also being used with higher plant PS I complexes to provide neighbor analysis of cross-linked or genetically modified materials (Andersen et al., 1990, 1992; Henry et al., 1992). The most interesting region in the sequence of PsaA and PsaB is that surrounding helices VIII and IX (nomenclature of Kirsch et al., 1986). The presence of appropriately spaced leucine residues in helix VIII of both proteins has led to the suggestion that the heterodimer is held together by a ‘leucine zipper’ similar to that found in DNA-binding proteins (Webber and Malkin, 1990; Kössel et al., 1990). That this mechanism would be needed to dimerize two highly hydrophobic membrane proteins appears a bit surprising, and there has been criticism of this proposal based on both theoretical and experimental grounds. Unfortunately, the experimental data are equivocal, since strains harboring conservative mutations in the proposed leucine zipper region L522V, L536M and L522V/L536M of PsaB in Synechocystis sp. strain PCC 6803 exhibited wild-

Chapter 10 Photosystem I Reaction Center

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Chapter 10 Photosystem I Reaction Center type characteristics, grewphotoheterotrophically and showed normal light-induced charge separation from P700 to and (Smart et al., 1993). The PS I analogs may again be helpful in establishing structural requirements for reaction center assembly, since neither the Chlorobium limicola nor Heliobacillus mobilis reaction center proteins appear to contain the leucine-zipper motif. The conserved cysteine residues in the extra stromal connecting region connecting helices VIII and IX most probably represent the binding site for ironsulfur center Indeed, the immediate region surrounding the binding site shows an astonishingly high sequence identity in PsaB among all sequenced species (the PS I analogs excepted). A total of 100 amino acids from residues 500 to 600 of PsaB are nearly invariant in eucaryotes and procaryotes, suggesting that an extraordinarily high degree of conservation is related to primary photoprocesses and hence to the binding of electron transport cofactors. The ligands to P700, and may be located on amino acids located on helices VIII and IX of PsaA and PsaB, which may form a 4membered bundle analogous to the LD, LE, MD and ME core of the bacterial reaction center (Deisenhofer et al., 1985). An antiparallel arrangement ofadjacent helices has been suggested to provide more stability in the bundle due to the contributions of the helix-dipole interactions (Rodday et al., 1993). The binding region is represented by the sequence CDGPGRGGTC and is identical in PsaA and PsaB ofall species sequenced thus far. The PsaA and PsaB polypeptides each contribute two cysteine ligands to bind the interpolypeptide [4Fe-4S] cluster (Petrouleas et al., 1989; McDermott et al., 1989; Scheller et al., 1989b). The preponderance of glycines and the presence of a proline in this region may be due to the need for the peptide to fold back on itself to place the cysteines in proper position to ligate the iron atoms in Alternately, they may allow the positively-charged arginine residue to participate in the binding of PsaC to the PS I core. To this end, the chemical modification of arginine residues on PS I with phenylglyoxal prevented the reconstitution of the cyanobacterial PsaC and PsaD proteins onto a modified spinach core. In this same work, an attempt at molecular modeling ofthis domain has led to the finding that a wide variety ofconformations can be generated as the result of the flexibility conferred by the glycines (Rodday et al., 1993). The loops could form a cavity ofsufficient size to contain

333 a protein of the size predicted for PsaC (Fig. 6). The positively charged arginines at the apices of the loops have been suggested to keep an open conformation and to function in the binding of the negatively-charged PsaC. It will be very interesting to test this proposal by deleting the arginines using site-specific mutagenesis. The chemical identity of P700 as a Chl a species has recently been questioned. Two molecules of Chl a' (the of Chl a) have been found in PS I complexes isolated from cyanobacteria (as well as higher plants and algae; Maeda et al., 1992); one Chl a' is extractable with acetone and the other with chloroform, implying that one is in a more hydrophobic site than the other. Although these findings have led to the speculation that P700 (or may consist ofa specialized Chl molecule(s), there is no other evidence to support this assignment. The ligands to the cofactors P700, and on the PsaA and PsaB polypeptides are not known. However, there are several conserved histidines in PS I (especially H524, 525 and 542 in PsaA and H520, 521 and 528 in PsaB of Synechococcus sp. strain PCC 7002) and in the Chlorobium limicola and Heliobacillus mobilis reaction centers (see Büttner et al., 1992; Liebl et al., 1992) that are excellent candidates for binding P700, the bridging chlorophyll, or

2. Function All reaction centers are designed to carry out three necessary roles in the process of photoconversion: light capture, charge separation, and charge stabilization. In cyanobacteria, the vast majority of the PsaA/PsaB heterodimer serves as a scaffold for the approximately 100 Chl a molecules and (an unspecified number of) molecules that function in light absorption serves a second role of photoprotection in all reaction centers (Mathis and Schenck, 1982)). Unlike higher plants and algae, the cyanobacterial PS I complex does not contain Chl b, nor does it contain additional lightharvesting Chl proteins similar to the LHC I complement (although such proteins have been identified in the chloroplasts ofred algae by Wolfe et al., 1992). Through a process of excitation energy transfer described first by Forster (1965), the energy of the Chl a excited singlet state is transferred to a low-lying photochemical trap, P700. It is now widely accepted that this trap is a Chl a dimer in the ground

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state that exhibits a bleaching centered at 698 to 700 nm when oxidized (hence the name P700). Studies of energy transfer and charge separation kinetics using fluorescence and transient absorption methods show that the charge separation kinetics in PS I from Synechococcus sp. is essentially trap-limited (Holzwarth et al., 1993). The major fluorescence lifetime components were found to be 12 and 36 ps; the first component is attributed to an energy transfer process between pigments in the main antenna Chl pool and a small long-wavelength Chl pool, and the second is assigned to the overall charge separation lifetime. The transient absorption lifetimes were 7–8 ps, 33 ps, and > 1 ns; the first and second components are assignments were the same as in the fluorescence studies, and the long-lived component is attributed to the lifetime of the oxidized primary donor Measurements of the spin-polarized EPR spectra of the triplet state of P700 in Synechocystis sp. strain

John H. Golbeck

PCC 6803 at 4.5 K yielded zfs parameters which are identical with those of the Chl monomer (Sieckmann et al., 1991b). In order to explain this behavior, it was proposed that the triplet excitation is delocalized over both halves of a Chl dimer at room temperature but appears localized on one Chl of the dimer at low temperature. The dimer/monomer controversy has further subsided with the realization that localization of the charge on one of the two Chl may be responsible for the optical properties of the cation (Mathis and Sétif, 1981) and the ESR properties of the P700 triplet state (Rutherford and Mullet, 1981). There is widespread agreement that the primary acceptor, is a Chl monomer with an absorption band in the red centered between 682 and 690 nm depending on the preparation (Mathis et al., 1988; Kim et al., 1989). The transfer of the exciton to the photochemical trap is followed by an initial rapid bleaching which is dominated by the formation of Charge separation follows rapidly, leading to a bleaching in

Chapter 10 Photosystem I Reaction Center the visible region which represents a contribution from both the cation and the anion (Mathis et al, 1988). The charge-separated state has a lifetime that is not yet known with certainty (values of< 10 ps are reported by Fenton et al. (1979) whereas values > 10 ps are reported by Wasielewski and co-workers (1987). The acceptor undergoes a redoxidation in 32 ± 5 ps (Shuvalov et al., 1986; Trissl et al., 1987), leading to the reduction of phylloquinone to the semiquinone anion radical. The identity of as phylloquinone is now well established after a decade ofuncertainty. A quinonelike molecule was suspected from early EPR electron spin polarization (ESP) studies ofPS I particles from Synechococcus sp. (Petersen et al., 1987). Two consecutive spectra are found after a laser flash; the time constant of the electron transfer steps is 260 ± 20 ns (Bock et al., 1989). The first spectrum is assigned to the pair, where is the second electron acceptor, probably a quinone. Compelling evidence for the identity of was also provided by reconstitution experiments and by ESP EPR of higher plant PS I complexes (Rustandi, et al., 1990). A further study showed that reconstitution is possible only with a limited number of quinone analogs (Biggins, 1990). Indeed, when a PS I complex from Synechocystis sp. strain PCC 6803 is extracted and then reconstituted with phylloquinone, the characteristic ESP signal is restored; however, when it is reconstituted with naphthoquinone and other phylloquinone analogs, subsequent electron transfer to the iron-sulfur cluster is blocked (Sieckmann et al, 1991a). The binding site therefore appears to be quite specific for phylloquinone. Using the loss and recovery of P430 as the criterion, the extraction and reconstitution of phylloquinone has also been demonstrated in a refined core isolated from Synechococcus sp. strain PCC 6301 (Ikegami et al, 1993). Studies on the orientation of the native and reconstituted quinones have been carried out in PS I particles from Synechocystis sp. strain PCC 6803 and Synechococcus lividus. In fully deuterated Synechococcus lividus, the principal x-axis ofthe gtensor of is roughly parallel to the symmetry axis of the pair-dipole coupling tensor (Gierer et al., 1991). Simulations of PS I ESP spectra using independently measured g-tensors confirm that the phylloquinone x-axis (along the C = O bonds) lies roughly parallel to the axis connecting and (Stehlik et al., 1989; Fuchsle et al., 1993). Studies by K-band EPR spectroscopy suggest that the orientation

335 of reconstituted naphthoquinone is different from that ofphylloquinone. Additional studies in spinach PS I complexes show that restoration of the ESP signal is possible only when the midpoint potential of the reconstituted acceptor is more positive than about 750 mV and structure contains either two aromatic rings, such as naphthoquinone, or a benzoquinone derivative substituted with an alkyl tail (Rustandi et al., 1992). Surprisingly, the phylloquinone that gives rise to the signal appears to be easily displaced; it can be exchanged with perdeuterated phylloquinone, and the replacement rate is strongly dependent upon temperature, and occurs on the time scale of hours to days at 4 °C and at 37 °C, and on the time scale of minutes when illuminated. Reconstitution of higher plant PS I complexes with 2-azido-9,10-anthraquinone followed by illumination with UV light selectively labels the PsaA and PsaB polypeptides (Iwaki et al, 1992). This makes it feasible, at least in principle, to identify the covalent attachment site on PsaA/PsaB by an analysis of endopeptidic digest ion products. In Synechocystis sp. strain PCC 6803, phylloquinone has been shown to become doubly reduced when the PS I complex is illuminated in the presence of a reductant (Bottin and Sétif, 1991). Forward electron flow is thereby thwarted until molecular oxygen can re-oxidize the quinone in a process which requires several minutes. Snyder et al. (1991) exploited this finding to show that the ESP signal in higher plant PS I complexes is not observed under conditions where the quinone is likely to have been doubly reduced. Heathcote and coworkers (1993) used a similar approach to show that doublereduction of the bound phylloquinone abolished the asymmetric EPR signal (g = 2.0048, mT) attributed to These results provide compelling biochemical evidence that the spectroscopic signal commonly known as is derived from phylloquinone. The back reaction between and is observed only when and are prereduced and when the iron-sulfur clusters are removed physically from the PS I complex. The back reaction pathway follows two different routes, depending on the conditions. When the iron-sulfur centers are prereduced in either cyanobacterial or higher plants, the 250 ns recombination between and populates the triplet state of P700 (Sétif and Bottin, 1989; Sétif and Brettel, 1990), which then relaxes with a half-time of When the iron-sulfur clusters

336 are removed biochemically from a Synechococcus sp. strain PCC 6301 PS I complex, the recombination between and populates the ground state of P700 with a half-time of about (Warren et al., 1990, 1993 a). The electrostatic charge on and appears to bias the pathway for back electron flow. The direct back reaction to the ground state of P700 is observed at cryogenic temperatures under both sets of conditions. When the oxidation of is determined by optical kinetic spectroscopy, a half-time of about 25 ns if found in a Synechococcus sp. PS I complex (Brettel, 1988). A similar study of the spinach PS I complex showed that is oxidized with a half-time of250 ns (Mathis and Sétif, 1988). This discrepancy was recently resolved when the experiments were repeated in spinach, Synechococcus sp. and Synechocystis sp. strain PCC 6803 at a higher S/N (Sétif and Brettel, 1993). It was found that forward electron transfer in all PS I complexes was biphasic; a 15 ns-phase and a 200 ns-phase are both present, the ratio of which is species-dependent as well as detergent-sensitive. The 200 ns-kinetic phase was found to be variable, and is considerably more prominent in Synechococcus sp. strain PCC 6301 than in Synechocystis sp. strain PCC 6803 (K. Brettel andJ. H. Golbeck, unpublished observations). When the oxidation of is determined from the decay ofthe ESR ESP signal, a halftime of 200 ns is found in both cyanobacteria and higher plant PS I complexes (Bock et al., 1989). The 15 ns-kinetic phase is likely to be present (the risetime of the EPR spectrometer is about 25 ns), and it can be inferred by the presence of a prominent ‘late signal’ in the ESP measurement. Further, there is no change in the 200 ns rate ofreoxidation of if PsaC is removed. This provides direct evidence that the electron acceptor following is (A. Van der Est, C. Bock, J. H. Golbeck, K. Brettel, P. Sétif and D. Stehlik, unpublished resultsi). is likely to participate as a redox-active component in forward electron transfer simply because of distance: it is 12 Å from the proposed location ofthe preceding acceptor (14 to 15 Å to the alternate position of the quinones; see Fig. 5B) and 15 Å distant from the nearest iron-sulfur cluster on PsaC. This assessment is supported by recent dynamic studies of forward electron flow in sitespecific mutants generated in Synechocystis sp. strain PCC 6803 (Smart et al., 1993). When cysteine 565 is changed to serine, the PS I complex assembles properly, as shown by the presence of P700 and the

John H. Golbeck low temperature photoreduction of and However, only 60% of and is capable ofbeing photoreduced relative the amountpresent by chemical reduction with sodium dithionite. When and are prereduced, a new set ofresonances appear upon illumination at g = 2.015, 1.941 and 1.811, which have been proposed to arise from a mixed-ligand cluster located in the site (Warren et al., 1993b). The missing 40% of the photochemistry is coupled to a population of clusters located in the site. The [3Fe-4S] clusters are reducible with dithionite at pH 6.5, which implies a midpoint potential considerably more oxidizing than that foreither or Hence, the [3Fe-4S] cluster in the site serves as a ‘redox trap’, disallowing forward electron flow to and on the basis of thermodynamics alone. Similar to the behavior of the wild-type the [3Fe-4S] cluster undergoes partial, reversible photoreduction when the complex is illuminated at 15 K. Therefore, even when the [3Fe-4S] cluster is reduced, and hence unavailable as an electron acceptor, there is no further increase in the amount of or photoreduction. This experiment shows that thenon-functioning site is not bypassed by the electron from ; the 27 to 30 Å from to the next-available iron-sulfur center probably cannot be spanned at 15 K. This behavior is compatible with the participation of as an intermediate in electron transfer from to A C556S site-directed mutant in the cysteine adjacent to the proline does not show the presence of a [3Fe-4S] cluster in isolated particles, but the g = 1.811 resonance of is in evidence when the sample is illuminated in the presence of reduced and (R. Schulz, Y.-S. Jung, J. H. Golbeck and L. McIntosh, unpublished results). This satisfying result indicates that the constraints on the iron-sulfur cluster that lead to the high broad linewidth, fast spin-relaxing cluster are relieved when either of the cysteine ligands on PsaB are replaced by serine. It is also intriguing that both cysteines on PsaB and one cysteine on PsaA are flanked with aspartic acid residues (the fourth flanking residue on PsaA is glutamine). The presence of the nearby carboxylate has been shown to exert a subtle effect on the electronic properties of the cluster. For example, the resonances of is shifted downfield from g = 2.05, 1.86 and 1.76 to g = 2.04, 1.87 and 1.80 when aspartate 566 is changed to alanine; when aspartate 557 is changed to alanine, the g-values remain relatively unaltered, but the linewidths are much

Chapter 10 Photosystem I Reaction Center greater. These changes in spectral properties do not prevent from functioning as an electron carrier between to (R. Schulz, Y.-S. Jung, J. H. Golbeck and L. McIntosh, unpublished results). It will be interesting to determine whether the midpoint potential of is altered by these changes. Site-specific mutants in PsaB have also been generated in the eucaryote Chlamydomonas reinhardtii (Webber et al., 1993). When cysteine 560 (the cysteine adjacent to the proline) in PsaB was changed to histidine, the cells were non-photosynthetic and there was no accumulation of PS I. It is interesting that replacement of the proline by alanine and leucine in Chlamydomonas reinhardtii leads to photosynthetic growth and accumulation of normal amounts of PS I. This result is borne out by recent studies of the homologous 2[4Fe-4S] protein from Clostridium pasteurianum, in which the two prolines adjacent to cysteines were changed to other amino acids (Gaillard et al., 1993). The EPR characteristics of the reduced iron-sulfur clusters were not significantly affected, which implies that there is little change in the folding of the polypeptide or in the electronic structure of the clusters. These results shows that the Cys-Pro motif, which is found in the vast majority of 2[4Fe-4S] ferredoxins, may not be a necessary structural element in binding the ironsulfur clusters. Further progress from site-directed mutagenesis of the PS I reaction center genes is expected, especially now that targeted mutagenesis has been accomplished in a second cyanobacterium, Anabaena variabilis strain ATCC 29413 (Nyhus et al., 1993).

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B. PsaF

indicated above, the psaF and psaJ genes are cotranscribed in all known instances in cyanobacteria (see section on PsaJ). PsaF is the only known peripheral polypeptide in cyanobacteria located on the lumenal side of the membrane in cyanobacteria. The protein has a transit sequence similar to those directing proteins such as plastocyanin and the extrinsic proteins of PS II to the thylakoid lumen. The transit sequence, which is cleaved after the protein has passed through the membrane, can be seen as the 23- to 27-amino acid extension at the amino-terminus when the gene andprotein sequences are compared (Table 4). In the three cyanobacterial proteins for which the complete sequences are known, the processed protein contains 141 amino acids, and the sequence shows a high degree ofconservation. In Synechococcus sp. strain PCC 7002 PsaF has a mass of 15,412 Da, and is negatively charged (the calculated pI is 5.5). As shown in Table 4, there is a clustering of positively-charged groups (KQRAKNFR) near the amino-terminus of the protein. Hydropathy analysis indicates that PsaF contains one hydrophobic region in the middle ofthe protein between residues 92 and 109. This implies that there exists a membrane-spanning helix which could serve to anchor the protein firmly to the PsaA/ PsaB heterodimer. The argument was made earlier that helix ‘i’ in the crystal structure (Figure 4) may be the derived from the carboxyl-terminus of PsaF. This firm attachment could account for the fact that the protein cannot be removed with chaotropic agents (Li et al., 1991a) but that it can be partially removed from the PS I complex with high concentrations of detergents, including 1% Triton X-100 (Bengis and Nelson, 1975).

1. Structure

2. Function

The primary amino acid sequence of PsaF has been deduced by sequencing the psaF gene in Synechococcus sp. PCC 7002 (W. M. Schluchter and D. A. Bryant, unpublished results), Synechococcus sp. (Mühlenhoff et al., 1993), and Synechocystis sp. strain PCC 6803 (Chitnis et al., 1991), and partial sequences are available by amino-terminal amino acid sequencing of the PsaF protein from Synechococcus sp. PCC 6301 (Li et al., 1991a), Synechococcus vulcanus (Koike et al., 1989), and Anabaena sp. strain ATCC 29413 (Nyhus et al., 1992). The deduced or actual amino acid sequences of the PsaF protein are tabulated and compared in Table 4. As

The function of PsaF in PS I is not well-understood. It is tacitly assumed that the patches of positive charge are instrumental in positioning the negativelycharged donor proteins, plastocyanin and cytochrome so that the cofactors are in the proper orientation to donate electrons efficiently to However, nearly all of the kinetic studies have been performed on higher plant PS I complexes, and it will be necessary to carry out studies in cyanobacteria to ensure that the results are universal. The reason is that in higher plants plastocyanin is the likely electron donor, whereas in most cyanobacteria cytochrome is the most abundant electron donor to (see

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Chapter 12). It is curious that in spite of the presence of two dissimilar electron carriers, there exists only one form of PsaF in which the primary amino acid sequence of is well-conserved between higher plants and cyanobacteria. If PsaF is responsible for docking these two proteins, then it must recognize and orient two soluble electron carriers with very different 3dimensional structures. In one of the first experimental indications of function, Nelson and Bengis (1977) reported that photoreduction was lost when PsaF (identified as Subunit III) was removed from the thylakoid membrane of spinach with Triton X-100. Since photoreduction mediated by Nmethylphenazonium 3-sulfonate (PMS) was relatively unaffected, the implication was that PsaF had a role in mediating plastocyanin function on the lumenal side of PS I. This assessment was supported, indirectly, by a report that plastocyanin could be cross-linked to PsaF in spinach Photosystem I

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complexes the zero-length cross-linking agent EDC (Wynn and Malkin, 1988). A kinetic study, that showed that cross-linked plastocyanin could support rapid electron donation of to in spinach PS I complexes, supported this assessment (Hippler et al., 1989). This confirmed an earlier study in which the fast-phase of plastocyanin donation to spinach complexes was lost when a polypeptide (presumed to be PsaF) was removed with Triton X100. In total, these studies led to the conclusion was that PsaF provided a positive charge at the highly negative oxidizing side of PS I to facilitate the fast electron transfer from negatively-charged plastocyanin to (Ratajczak et al., 1988). Wynn et al. (1989) closed the argument with the finding that cytochrome (previously cytochrome could be cross-linked with PsaF in Synechococcus sp. strains PCC 7002 and PCC 6301 PS I complexes in a manner analogous to that of plastocyanin in higher plants. From these studies, it appeared that PsaF

Chapter 10 Photosystem I Reaction Center supplied a necessary plastocyanin and/or cytochrome ‘docking’ site near on the lumenal side of PS I. A major difficulty with the older literature is that it was not known, or widely appreciated, that Triton X100 could remove PsaF from the PS I complex. Further complicating the issue is the report from Ikeuchi et al (1990) that Triton X-100 can deplete higher plant PS I complexes not only of LHC I and variable amounts of PsaF, but also significant amounts of PsaK and PsaJ. Hence, reports of photoreduction (or lack of photoreduction) must be interpreted with caution unless data on polypeptide composition is also available. Probably the most interesting result so far was provided by a PsaF deletion mutant of Synechocystis sp. strain PCC 6803, for which it was shown that the rate of cell growth was relatively unaffected by the absence of PsaF (Chitnis et al., 1991). Surprisingly, the mutant organism grew at a faster rate in the presence of high concentrations of than did the wild-type organism either in the absence of presence of Recent results using the thermophilic cyanobacterium Synechococcus elongatus indicate that the PsaF polypeptide is not required for the bimolecular reaction between P700 and cytochrome (Hatanaka et al., 1993). Interestingly, had a moderate stimulating effect on the rate of P700 reduction whether PS I complexes were associated with the PsaF or not. The whole issue is, of course, complicated by the known participation of cations in the interaction of plastocyanin and the PS I complex (see Gross, 1993), and by the facile interchangeability of cytochrome and plastocyanin in cyanobacteria (see Chapter 12). It will take a careful sorting out of these factors using the full range of tools available in molecular biology, biochemistry and spectroscopy before the function of PsaF is understood.

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C. Psal 1. Structure The psaI gene has been cloned and sequenced in Synechococcus sp. strain PCC 7002 (W. M. Schluchter and D. A. Bryant, personal communication), Synechococcus sp. (Mühlenhoff et al., 1993), Synechocystis sp. strain PCC 6803 (Q. Xu, L. Yu, V. P. Chitnis and P. R. Chitnis, personal communication), and Anabaena variabilis strain ATCC (Sonoike et al., 1992). The deduced amino acid sequences of the PsaI proteins are tabulated and compared in Table 5. In Synechococcus sp. and Synechococcus sp. strain PCC 6803 the psaI gene is located downstream from psaL , but it is not known whether the two genes are co-transcribed. In Synechococcus sp. strain PCC 7002, the psaI gene is approximately 200 bp upstream from psaL and transcribed from the opposite DNA strand. The carboxyl-terminus of PsaI in procaryotes is acidic (2–3 aspartic/glutamic acid residues) whereas in higher plants it is basic (2 to 3 lysines; see Bryant, 1992). In Anabaena variabilis strain ATCC 29413, the psaI open reading frame could encode a polypeptide of 46 amino acids, whereas amino-terminal amino acid sequencing of the mature protein shows that it contains only 35 residues (Ikeuchi et al, 1991). The protein is clearly cleaved at the amino-terminus, and there is some resemblance within the 11 missing residues for a signal processing site (PxILA). Also, the last eight amino acids in the presequence have some similarity to the signal peptide of plastocyanin in Anabaena variabilis. It has been suggested that the presequence targets the amino-terminus of the PsaI protein to the thylakoid lumen, thereby allowing the carboxyl-terminus to be exposed to the stroma

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340 (Sonoike et al., 1992). However, the significance of the this putative presequence is not clear, since the psaI genes of Synechococcus sp. strain PCC 7002 and Synechococcus sp. do not appear to encode such presequences. Although the PsaI protein has not been detected in the PS I preparations from Synechococcus sp. (Ikeuchi et al., 1992), the psaI gene was detected downstream from the psaL gene (Mühlenhoff et al., 1993). PsaI has been detected in the PS I complexes of Synechococcus sp. strain PCC 7002 and Synechocystis sp. strain PCC 6803, although the protein comigrates upon SDS-polyacrylamide gels with PsaJ in both cases (Ikeuchi et al., 1993). The amino-termmal sequences for PsaI and PsaJ from the Synechococcus sp. strain PCC 7002 precisely matched the sequences predicted by the genes (M. Ikeuchi, Y. Inoue, W. M. Schluchter, and D. A. Bryant, personal communication). The hydropathy profile of PsaI from is consistent with an extremely hydrophobic protein that most probably forms a single transmembrane This finding is consistent with the fact that PsaI can be extracted from the PS I complex with a mixture of chloroform/ methanol (Ikeuchi et al., 1990).

2. Function No function is known for PsaI, nor have any deletion or interruption mutants of PsaI become available for study. There has been one proposed role for PsaI based on weak sequence similarity to helix E of the PsbD (D2) reaction center protein in PS II. Helix E participates in binding the quinone and iron; hence, the suggestion was made that PsaI may contain a partial a quinone-binding site (Scheller et al., 1989a). It is intriguing that both PsaI and PsaJ are encoded in the chloroplast genome of higher plants (Ohyama et al., 1986; Shinozaki et al., 1986; Bryant, 1992),

which is where most of the catalytic components of the photosynthetic reaction centers are encoded. Ikeuchi et al. (1990) have found that PsaI is lost when a higher plant PS I complex is prepared with SDS. While this may appear to weaken the argument that PsaI has a role in primary processes, it has not been shown that the SDS preparation is able to support electron transport beyond

D. PsaJ

1. Structure The psaJ gene is located downstream from and is cotranscribed with the psaF gene in Synechococcus sp. strain PCC 7002 (W. M. Schluchter and D. A. Bryant, personal communication), Synechocystis sp. strain PCC 6803 (Xu et al., 1993), and Synechococcus sp. (Mühlenhoff et al., 1993). PsaJ has also been identified on the basis of amino acid sequencing in Synechococcus vulcanus (Koike et al., 1989), and in Anabaena variabilis strain ATCC 29413 (Ikeuchi et al., 1991), and it is present in the cyanelle genome of Cyanaphora paradoxa, where it is located similarly and also apparently co-transcribed with the psaF gene (D. A. Bryant and V. L. Stirewalt, unpublished observations; see Chapter 4). The actual or deduced amino acid sequences of the PsaJ proteins are tabulated and compared in Table 6. Because the psaF and psaJ genes are on the same operon, interposon mutagenesis of psaF in Synechocystis sp. strain PCC 6803 may affect the expression of PsaJ due to polarity effects. The PsaJ proteins have a mass of about 4 kDa and the protein in Synechococcus vulcanus is blocked at the amino-terminus (Koike et al., 1989). The hydropathy profile of PsaJ is consistent with an extremely hydrophobic protein that may form a single transmembrane The degree of

Chapter 10 Photosystem I Reaction Center homology between species is not as high as for other cyanobacterial proteins, but this may be expected of a transmembrane protein which has little tertiary structure.

2. Function The phenotype of the psaF psaJ double deletion mutant of Synechocystis sp. strain PCC 6803 appears identical to that of the wild-type. The organism grows at wild-type rates in high and low light, and in high and low concentrations in the absence of both PsaF and PsaJ (P. R. Chitnis, personal communication). Accordingly, PsaJ may have little or no role in cofactor binding or in PS I assembly.

E. PsaK 1. Structure The amino acid sequence for PsaK has been deduced from the sequence of the psaK gene in Synechococcus sp. strain PCC 7002 (W. M. Schluchter and D. A. Bryant, unpublished results) and Synechococcus sp. (Mühlenhoff et al., 1993), and partial sequences have been obtained directly from the protein in Synechococcus sp. strain PCC 7002 (M. Ikeuchi and Y. Inoue, personal communication), Synechococcus vulcanus (Koike et al., 1989) and Anabaena variabilis (Ikeuchi et al., 1991). The actual or deduced amino acid sequences of the PsaK protein are tabulated and compared in Table 7. Eight amino acids are cleaved from the precursor of the PsaK protein in Synechococcus sp. strain PCC 7002 (M. Ikeuchi and Y. Inoue, personal communication). This presequence

341 is identical in length and resembles the presequences of the Synechocystis sp. strain PCC 6803 and Cyanophora paradoxa PsbK proteins, components of PS II which are similarly processed (Ikeuchi et al., 1991; D. Bryant, personal communication). Ikeuchi et al. (1991) have suggested that the PsbK presequence targets the N-terminus of that protein to the thylakoid lumen, and this could also be true for the PsaK protein. The hydropathy profile of PsaK is consistent with a membrane-intrinsic protein containing two transmembrane, regions, and, indeed, the 79 amino acid protein in Synechococcus sp. strain PCC 7002 has enough mass to comfortably thread twice through the membrane. Along with PsaA and PsaB, PsaK has been found as a component of spinach CP 1, a minimal PS I particle isolated from the PS I reaction center with the use of the strong ionic detergent SDS (Wynn and Malkin, 1990). CP 1 shows charge separation between P700 and but all secondary electron transfer to the quinone and iron-sulfur clusters is lost. Ikeuchi et al. (1990), however, report that PsaK polypeptide is released, along with LHC I and the PsaJ protein by treating the spinach PS I-LHC I holocomplex with 1% Triton-X-100. More work will be needed to determine whether PsaK is more tightly associated with the PS I core than any of the other low molecular mass polypeptides.

2. Function Interposon mutagenesis of the psaK gene in Synechococcus sp. strain PCC 7002 has not yet provided any useful information on its function in

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PS I. The PsaK-less mutant grows normally at high and low light intensities, and the PS I complexes isolated from the mutant are trimeric complexes that do not appear to be deficient in any other subunits (W. M. Schluchter and D. A. Bryant, personal communication). Moreover, the mutant shows normal charge separation between P700 and and is competent in both linear and cyclic electron flow (L. Yu and J. Golbeck, unpublished results).

F. PsaL

John H. Golbeck (Li et al., 1991a), Synechococcus vulcanus (Koike et al., 1989), and Anabaena variabilis strain ATCC 29413 (Nyhus et al., 1992). The actual or deduced amino acid sequences of the PsaL protein are tabulated and compared in Table 8. PsaL shows more diversity in its amino acid sequence than any other PS I polypeptide. Nevertheless, hydropathy plots suggest the presence of two hydrophobic regions, and the PsaL protein is large enough to contain two complete membrane-spanning The amino-terminus of the Synechococcus sp. strain PCC 7002 protein is not processed.

1. Structure 2. Function The amino acid sequence of PsaL has been deduced from the sequence of the psaL gene in Synechococcus sp. strain PCC 7002 (W. M. Schluchter and D. A. Bryant, personal communication), Synechococcus sp. (Mühlenhoff et al., 1993), and Synechocystis sp. strain PCC 6803 (Chitnis et al., 1993), and partial sequences are available from N-terminal protein sequencing in Synechococcus sp. strain PCC 6301

Interposon mutagenesis of the psaL gene in Synechocystis sp. strain PCC 6803 (Chitnis et al., 1993) and Synechococcus sp. strain PCC 7002 (W. M. Schluchter and D. A. Bryant, personal communication) has provided significant information on its function in PS I. In both instances, the psaL mutants grow normally at high and low intensities of white

Chapter 10 Photosystem I Reaction Center

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light, show normal charge separation between P700 and and are competent in both linear and cyclic electron flow. However, trimers of the PS I reaction center are not found in the mutant after detergent solubilization with dodecyl maltoside and sucrose gradient ultracentrifugation. No other PS I polypeptide is missing from the monomeric complexes. It is therefore likely that PsaL represents the j and/or k helices depicted on the model of the 3dimensional crystal structure (Fig. 4A). The functional significance of the PS I trimers, and whether they exist in vivo, remains unclear. However, cross-linking experiments performed with thylakoids from the wild-type and psaL mutant strains of Synechococcus sp. strain PCC 7002 show clear differences that are consistent with the presence of trimers in the wild-type but not in the mutant (J. Zhao and D. A. Bryant, personal communication). In addition to the inability to form trimers, the PsaLless mutant of Synechococcus sp. strain PCC 7002 appears to be permanently locked in State I (high fluorescence condition), which suggests that either trimers, or Chls associated with PsaL, or both, have some role in facilitating state transitions and/or allowing energy transfer from phycobilisomes to PS I (J. Zhao and D. A. Bryant, personal communication) . Consistent with these observations, the psaL mutant strain grows much more slowly than the wild-type strain in green light, when energy transfer from the phycobilisome to PS I is most important (W. M. Schluchter and D. A. Bryant, personal communication).

Bryant, personal communication), Synechococcus sp. (Haehnel et al., 1992) and Synechocystis sp. strain PCC 6803 (Ikeuchi et al., 1992), and from amino-terminal protein sequencing of PsaM in Synechococcus vulcanus (Ikeuchi et al., 1992). The actual or deduced amino acid sequences of the PsaM protein are tabulated and compared in Table 9. Although PsaM has not yet been found in the PS I complexes of higher plants, the sequence is highly similar to the presumed product of ORF 32 in the liverwort Marchantia polymorpha (Ohyama et al., 1986). It is noteworthy that no homologous ORF is present in the chloroplast DNA of tobacco or rice (Shinozaki et al., 1986). The PsaM protein is highly hydrophobic, and due to its small size, the protein is expected to span the membrane once.

G. PsaM

The primary amino acid sequences of PsaC proteins have been deduced from the sequences of the psaC genes in Synechococcus sp. strain PCC 7002 (Rhiel et al., 1992), Nostoc sp. strain PCC 8009 (Bryant et al., 1990), Synechocystis sp. strain PCC 6803 (psaC1: Anderson and McIntosh, 1991b; psaC2: Steinmuller, 1992), Synechococcus sp. (Mühlenhoff et al., 1993);

1. Structure The amino acid sequence of PsaM has been deduced from the sequence of the psaM gene in Synechococcus sp. strain PCC 7002 (W. M. Schluchter and D. A.

2. Function Nothing is known about the function of PsaM in PS I. No interruption or deletion mutants of PsaM are yet available for study, although the availability of the gene from transformable organism Synechococcus sp. strain PCC 7002 should allow the creation of such mutants in the near future.

V. Peripheral Polypeptides

A. PsaC 1. Structure

344 Synechococcus sp. strain PCC 6301 (Herman et al., 1993), Anabaena variabilis strain ATCC 29413 (Mannan, and Pakrasi, 1991), Anabaena sp. strain PCC 7120 (Mulligan and Jackman, 1992), and/or by amino acid sequencing the PsaC protein in Synechococcus vulcanus (Koike et al., 1989; Shimizu et al., 1990), Synechococcus elongatus Naegeli (Kotani et al., 1992), Calothrix sp. strain PCC 7601 (Mann et al., 1991). The actual or deduced amino acid sequences of the PsaC proteins are tabulated and compared in Table 10. The primary amino acid sequences of PsaC are conserved to an extraordinary degree among cyanobacterial species, and the only differences are conservative changes at the aminoterminus and near the center of the protein. These changes are not likely to lead to any differences in the 3-dimensional structures of PsaC. The higher plant PsaC sequences differ significantly from the cyanobacterial sequences in only two regions. At position 37, higher plants contain a lysine residue

John H. Golbeck while cyanobacteria contain a neutral amino acid (glycine or alanine), and at positions 70 and 71, dicots contain a tryptophan-histidine pair, monocots contain a glycine-proline pair and cyanobacteria contain a glycine-alanine pair. Additional conservative changes,particularly in the carboxyl-terminal region of the protein, are seen in higher plant sequences when compared to cyanobacteria. These changes are also not expected to lead to significant differences in either structure or function ofthe PsaC protein. One interesting anomaly is that Synechocystis sp. strain PCC 6803 may harbor two psaC genes; psaC1 codes for a protein with an amino acid sequence identical to PsaC of tobacco (Anderson and McIntosh, 1991b), while psaC2 codes for a protein with an amino acid sequence typical of PsaC in cyanobacteria (Steinmüller, 1992). The presence of two psaC genes in Synechocystis sp. PCC 6803 begs the question offunction(s) for the two proteins. The g-values of the and iron-

Chapter 10 Photosystem I Reaction Center sulfur clusters in Synechocystis sp. strain PCC 6803 are identical to those of other cyanobacteria and slightly different from those ofhigher plants (Mehari et al., 1991), which would indicate that the psaC2 rather than psaC1 gene product is expressed. This is consistent with the finding that the amount of psaC1 mRNA transcript is low under conditions where the PS I polypeptides are expressed (Anderson and McIntosh, 1991b). Interposon mutagenesis of the psaC1 gene with a kanamycin cartridge leads to a phenotype that is lethalunder LAHG (light-activated heterotrophic growth) as well as photoautotrophic growth conditions (Anderson and McIntosh, 1991a). Since the organism is capable of LAHG growth under conditions where the entire PS I complex is deleted, it is not expected that the product of the psaC1 gene would be present in the PS I complex. This conclusion is supported by more recent findings which show that when the psaC2 gene has been interrupted, the organism grows under LAHG conditions, the PsaC, PsaD and PsaE proteins are missing, and the EPR spectrum shows reversible electron donation to in the light (J. Yu, L. Smart, Y.-S. Jung, J. H. Golbeck and L. McIntosh, unpublished results). The function of PsaC1 is unknown, but it clearly serves an essential function in the overall metabolism of the cyanobacterium. ThepsaC gene encodes a slightly acidic (apparent pI of 5.5) hydrophilic protein containing 81 amino acids (8682 Da in Synechococcus sp. strain PCC 7002) including the amino-terminal methionine, which is proteolytically removed from the mature protein. The PsaC protein contains nine cysteine residues, eight of which are clustered in two distinct CxxCxxCxxxCP motifs that are characteristic of iron-sulfur proteins that contain two distinct [4Fe4S] clusters. PsaC has some sequence homology with proteins such as the 54-amino acid ferredoxin of Peptococcus aerogenes and the first 58 residues of the 106-amino acid Azotobacter vinelandii ferredoxin. There are also regions ofsignificant homology with ferredoxins from Desulfovibrio gigas (Kissinger et al., 1989) and Bacillus thermoproteolyticus (Fukuyama et al., 1988) especially in the area around the single [4Fe-4S] cluster. The 3-dimensional structures ofthese proteins have been solved, and the overall folding pattern is remarkably similar for all four, especially near the iron-sulfur clusters. The Peptococcus aerogenes ferredoxin has a twofold rotational symmetry axis related to the two ironsulfur clusters (see Carter, 1977). When the protein

345 is rotated 180 degrees around this axis, the main chain atoms and the iron-sulfur clusters approximately overlay. The protein can therefore be divided into two symmetrical domains, each of which assumes nearly the same main chain conformation, and each of which contains a partial iron-sulfur binding site. The first three cysteines of each CxxCxxCxxxCP binding motif provide three ligands to the two [4Fe-4S] clusters. The fourth cysteine of each ironsulfur binding motif provides the fourth ligand to the other [4Fe-4S] cluster. Because ofthis geometry, the two half-chains do not form separate globular ironsulfur domains, but each half chain cooperates in the same manner to form an approximate intramolecular diad axis. The protein can be visualized as a dimer of two connected polypeptides containing shared [4Fe4S] clusters. This structure may represents the most frugal manner in which a small 2[4Fe-4S] protein can fold to create the internal iron-sulfur binding pockets. The extremely small size of these 2[4Fe-4S] ferredoxins has led several groups to suggest that, overall, PsaC will exhibits a similar folding pattern to the Peptococcus aerogenes ferredoxin (Oh-oka et al., 1988; Dunn and Gray, 1988; Zhao et al., 1992). The only significant differences between the Peptococcus aerogenes ferredoxin and PsaC are an additional ten amino acids between the two ironsulfur binding sites, which may have the effect of enlarging the internal loop structure as the polypeptide chain folds back on itself, and an additional fourteen amino acids at the carboxyl-terminus. The internal addition should not translate to a greater distance between the iron-sulfur clusters since this is largely pre-determined by the spacing between the third and fourth cysteine in the motif, CxxCxxCxxxCP. The availability of large quantities of PsaC apoprotein from an Escherichia coli expression system, combined with an efficient iron-sulfur insertion protocol (Mehari et al., 1991), should allow structural analyses of the protein. A crystal structure or NMR solution structure of PsaC is obviously needed to test these predictions, and efforts to crystallize the protein after reinsertion of the and iron-sulfur clusters are in progress (J. H. Golbeck, unpublished results). The locations of the and clusters are known relative to their cysteine ligands. These assignments could be made because of several experimental advances, including the isolation of a core from Synechococcus sp. strain PCC 6301 which was stripped of PsaC (Golbeck et al., 1988a; Parrett et al,

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John H. Golbeck

1989); the isolation of the psaC gene from Synechococcus sp. strain PCC 7002 which coded for a protein nearly identical in amino acid sequence to Synechococcus sp. strain PCC 6301 (Bryant et al., 1990); the generation of an expression plasmid in which mutant forms ofPsaC could be synthesized in Escherichia coli (Zhao et al., 1990; Li et al., 1991b); and the development of a reconstitution protocol which produced a functional PS I complex from the core and added PsaC (Golbeck et al., 1988b; Parrett et al. 1990; Li et al., 1991b). These experiments showed that the substitution ofaspartate for cysteine in position 14 of PsaC led to the formation of a [3Fe4S] cluster at the site-modified site, and that the unmodified [4Fe-4S] cluster had the g-values and characteristics of (Zhao et al., 1992; Yu et al., 1993a). Similarly, the substitution of aspartate for cysteine in position 51 of PsaC led to the formation of a [3Fe-4S] cluster at the site-modified site, and that the unmodified [4Fe-4S] cluster had the g-values and characteristics of Since the pattern ofcysteine ligation in PsaC is expected to be identical to that of other 2[4Fe-4S] ferredoxins, it was concluded that is ligated by cysteines 11,14,17, and 58 and is ligated by cysteines 21, 48, 51, and 54 (Fig. 7). Further studies with site-specific mutants has hinted that the ninth cysteine at position 34 does not participate in ligating either or (Yu et al., 1993a).

2. Function PsaC contains two [4Fe-4S] clusters, and which function in forward electron transfer from to ferredoxin or flavodoxin. Their role is to provide a pathway for electrons to move out of the hydrophobic reaction center core and into the hydrophilic stromal phase. The participation of the iron-sulfur clusters in forward electron flow was demonstrated conclusively in an experiment which showed that the photoreduction of was lost after PsaC, PsaD and PsaE were removed from a cyanobacterial PS I complex (Y.-S. Jung, V. Chitnis, P. Chitnis and J. H. Golbeck, unpublished results). The photoreduction of was recovered upon rebinding PsaC and PsaD to a core. This experiment further demonstrates that cannot donate electrons directly to ferredoxin (orflavodoxin). A similar result had been shown earlier using higher plant PS I complexes (Hanley et al., 1992). Hence, or (or

both) participate in forward electron flow from to ferredoxin. In spinach PS I complexes, is selectively destroyed with (Sakurai et al., 1991; Inoue et al., 1992), and photoreduction is lost (Hanley et al., 1992). The crystal structure at 6 Å-resolution of the PS I complex from Synechococcus sp. (Krauß et al., 1993), as well as a recent EPR study of oriented PS I complexes from Synechocystis sp. strain PCC 6803 (Guigliarelli et al., 1993), indicate that the axis is tilted away from the membrane plane (see Fig. 5B). This geometry carries with it the implication that electron flow is serial through the three ironsulfur clusters and Also, the absolute orientation of the principal axes of relative to the crystal axes have been determined through an EPR study of single crystals of PS I from Synechococcus sp. (Brettel et al., 1992). However, in none of the

Chapter 10 Photosystem I Reaction Center above work was it possible to identify clusters and nor was it possible to determine if the pathway of electron flow is or An additional complication is that one of the two iron-sulfur clusters may be silent (i.e., not used). Yet, this now seems unlikely as the distancefrom to the distal iron-sulfur cluster is 22 Å; moreover, from the X-ray crystallographic studies, the amount of electron density surrounding the ironsulfur cluster indicates the presence of protein structure that would preclude close contact with a stromal protein such as ferredoxin (N. Krauß and P. Fromme, personal communication). The linear pathway may have a different route than the cyclic pathway; however, so little is known about cyclic flow that this almost seems an unwarranted speculation. Unfortunately, the room-temperature optical properties of and are identical, which makes it difficult to study the dynamics of electron flow through the two clusters. Since the of is approximately–580 mV and the of is about –520 mV (a relatively restricted range ofpotentials have been reported; see Golbeck, 1987 for a tabulation), the sequence of electron flow might be expected to be on thermodynamic grounds alone. There is some spectroscopic evidence, however, which supports the alternative sequence It should be possible to approach the issue ofwhether or is proximal to through the use of site-specific mutants in combination with the measurement of the lowtemperature photoreduction of or The premise behind these experiments is that [3Fe-4S] clusters have relatively high midpoint potentials; if they were engineered into PsaC, there should exist a thermodynamic sink in the electron transfer chain. For example, the midpoint potential of the [4Fe-4S] cluster in the site in the C14D mutant protein was measured to be –520 mV, and that of the [3Fe-4S] cluster in the site was –90 mV, and the midpoint potential of the [4Fe-4S] cluster in the site in the C51D mutant was measured to be –580 mV, and that of the [3Fe-4S] cluster in the sitewas–98mV(Yu et al., 1993a). The redox potentials of the [4Fe-4S] clusters in the two mutant proteins did not change after rebinding to the PS I core (L. Yu, J. Zhao, D. A. Bryant and J. H. Golbeck, unpublished results). Hence, not only are the reduction potentials of the [4Fe-4S] clusters independent of the [3Fe-4S] clusters, but they are identical to the [4Fe-4S] clusters in the wild-type PS I complex. This indicates that the

347 primary amino acid sequence of the PsaC protein is solely responsible for conferring the midpoint potentials of the two iron-sulfur clusters. It also indicates that the midpoint potential of is not driven more electronegative due to the electrostatic influence of The [3Fe-4S] clusters have not been studied in the reconstituted PS I complex. However, if the [3Fe-4S] clusters remain intact when the protein is rebound to the PS I core, and iftheir electrochemical properties remain unchanged from those of the free protein, then the complex reconstituted with C14D will contain a high-potential redox trap in the site, and the complex reconstituted with C51D will contain a high-potential redox trap in the site. As an extension of this idea, it should be possible to insert other elements of the first transition series in the unoccupied site to create a mixed-ligand, heterometal cluster with modified properties. Indeed, the [3Fecluster in the unbound C14D mutant protein has been found to take up a ion, creating a cluster on further reduction with a resulting spin state (L. Yu and J. H. Golbeck, unpublished results). It will be very interesting to study the electron transfer properties of rebound mutant proteins when the redox potentials of and have been modulated by the presence of a heterometal cluster. Since the geometry of the and clusters places one ofthem 15 Å from and the other 22 Å from the mutant which allows low-temperature photoreduction ofthe native [4Fe-4S] cluster should identify or as the cluster. The logic of these experiments can also be extended to the next electron transport reaction—the transfer ofthe electron from one of the [4Fe-4S] clusters to soluble ferredoxin or flavodoxin. One of the two mutants should be significantly or completely impaired in the ability to support ferredoxin photoreduction.

B. PsaD 1. Structure The amino acid sequence of PsaD has been deduced from the gene sequence in Synechococcus sp. strain PCC 7002 (W. M. Schluchter and D. A. Bryant, unpublished results),Synechococcus sp. (Mühlenhoff et al., 1993), Nostoc sp. strain PCC 8009 (Bryant et al., 1990), Synechocystis sp. strain PCC 6803 (Reilly et al., 1988) and Synechococcus sp. strain PCC 6301 (Wynn et al., 1989) and whole or partial sequences are available by amino acid sequencing of the PsaD

348

proteins from Synechococcus vulcanus (Koike et al., 1989) Synechococcus elongatus (Enami et al., 1990), Calothrix sp. strain PCC 7601 (Mann et al., 1991) and Anabaena variabilis ATCC 29413 (Nyhus et al., 1992). The actual and deduced amino acid sequences of PsaD are tabulated and compared in Table 11. The cyanobacterial PsaD proteins range in size from 139 amino acids to 144 amino acids (15.4 to 15.9 kDa) and contain large numbers of lysines and arginines. PsaD in higher plants is somewhat larger, containing 158 to 162 amino acids (17.5 to 18 kDa). The sequence identity is about 65% among cyanobacterial sequences, which are in turn about 55% similar to the higher plant sequences. In most cyanobacteria PsaD has a highly basic isoelectric point (calculated pI of 9.0 to 9.6), but there are exceptions, such as a

John H. Golbeck

near-neutral isoelectric point (calculated pI of 6.7) for PsaD from Synechococcus sp. strain PCC 7002. The hydropathy profile indicates that PsaD is a highly hydrophilic protein which does not span the thylakoid membrane. This conclusion is supported by the finding that PsaD can be removed from the PS I complex with chaotropic agents (Parrett et al., 1989, 1990; Li et al., 1991a), and that free PsaD remains soluble up to concentrations of Chemical modification and proteolysis studies have shown that a large portion of PsaD is exposed to the solvent on the stromal surface of the PS I complex (Ortiz et al., 1985; Zilber and Malkin, 1992), where it may form a ‘cap’ over much of PsaC. Secondary structure predictions based on Chou-Fasman and Robson-Garnier criteria indicate significant

Chapter 10 Photosystem I Reaction Center structure, especially at the amino-terminal end between residues 20 and 45, but also between residues 48 and 53 and residues 60 and 70. These regions could represent the three broken stromal helices visualized in the X-ray structure on the stromal side of the reaction center (see Fig. 4 A). The predominance of lysine and arginine residues on the carboxylterminal end of the protein make it likely that this portion is in contact with PsaC and/or PsaE (the charged residues may be involved in ‘docking’ ferredoxin and flavodoxin on the stromal side of the reaction center), and that the amino-terminus is in contact with the PsaA/PsaB heterodimer. The expression of large quantities of functional PsaD in Escherichia coli (Li et al., 1991b) should allow detailed structural analyses of the isolated protein.

2. Function The primary amino acid sequence of PsaD provides some insight into the mechanism of docking ferredoxin to the PS I reaction center. In most instances, PsaD carries a net positive charge due to the large number of clustered lysine and/or arginine residues in the center portion of the protein, and this may be related to binding negatively-charged ferredoxin (pI of about 4.0 to 4.5). This role is supported by nearest-neighbor analyses using the zero-length cross-linker N-ethyl-3-[3-(dimethylamino)propyl] carbodiimide (EDC), in which ferredoxin from Synechococcus sp. strain PCC 7002 was found to be cross-linked to PsaD in a PS I complex of Synechococcus sp. strain PCC 6301 (Wynn et al., 1989). Flavodoxin from Synechococcus sp. PCC 7002 has also been found to cross-link with PsaD and PsaC in PS I complexes ofSynechococcus sp. strain PCC 7002 (U. Mühlenhoff and D. A. Bryant, unpublished results). The role of PsaD as a participant in the reduction of soluble electron acceptors is also supported from studies of a PsaD-less mutant of Synechocystis sp strain PCC 6803. In this mutant, ferredoxin-mediated photoreduction with cytochrome is used as electron donor is observed for the wild-type control) is totally inhibited in isolated thylakoids or in dodecyl-maltoside PS I complexes (Y.-S. Jung, V. Chitnis, P. Chitnis and J. H. Golbeck, unpublished results). The addition of the highly basic PsaD from Synechococcus sp. strain PCC 6803, or the neutral PsaD from Synechococcus sp. PCC 7002 PsaD to the isolated thylakoids or PS I

349 complexes led to the recovery of about 25% of the wild-type rate. Surprisingly, flavodoxin reduction and flavodoxin-mediated photoreduction were both found to occur in the PsaD deletion mutant, albeit at a much reduced rate vs. This surprising finding shows that PsaD may not be needed for flavodoxin-mediated photoreduction, provided that the iron concentration in the medium is sufficiently low to induce the synthesis of flavodoxin. A second function for PsaD was deduced from in vitro reconstitution experiments, in which Synechococcus sp. strain PCC 7002 PsaC and Nostoc sp. strain PCC 8009 PsaD were differentially rebound to a Synechococcus sp. strain PCC core (Li et al., 1991 b). These experiments show that when PsaC is rebound to a core in the absence ofPsaD, rather than is the terminal acceptor predominantly photoreduced at 15 K. The EPR linewidths of and those of photoaccumulated and are significantly broader, and the g-values are slightly different, than and for the wild-type control. When PsaD is present, becomes the terminal acceptor predominantly photoreduced at 15 K, the linewidths of and those of photoaccumulated and are sharp, and the g-values are identical to the control. It was also shown that PsaC is bound loosely to the core in the absence of PsaD and can be removed with 0.1% Triton X-100, but not with 0.1% dodecyl maltoside. When PsaD is present, PsaC is stable to 1% Triton X-100 (Zhao et al. 1990; Li et al. 1991b). Neither protein is rebound unless iron-sulfurcenter is intact and unless the and clusters are present in PsaC (Li et al., 1991a). These experiments show that PsaD stabilizes (and possibly orients) PsaC on the PS I heterodimer. Identical results were found using isolated thylakoids membranes from the PsaD-less mutant of Synechocystis sp. strain PCC 6803 (Y.-S. Jung, V. Chitnis, P. Chitnis and J. H. Golbeck, unpublished results); hence, this action of PsaD is not an artifact of the in vitro reconstitution procedure. Interposon mutagenesis of psaD in Synechocystis sp. strain PCC 6803 with a gene conferring kanamycin resistance has provided some information on its role in PS I (Chitnis et al. 1989a). The most significant result was that the PS I polypeptides were found to be turned over rapidly in the absence of PsaD. It was also shown that several low-molecularmass polypeptides (including PsaE) were missing in a Triton-isolated PS I complex. If these proteins are

350 also missing in vivo, the implication would be that PsaD is required for the efficient assembly of the low-molecular-mass subunits on the stromal side of PS I. However, based on the stabilizing role of PsaD, it is more likely that Triton X-100 simply removed the loosely-bound PsaC and PsaE during the isolation of the PS I complex. It was also reported that while the organism grew at rates equivalent to the wild type under photoheterotrophic conditions in the presence of 5 mM glucose and DCMU, the mutant was also capable of slow photoautotrophic growth. At the time, this remained unexplained, especially in light of the fact that ferredoxin does not accept electrons from in the absence of PsaD. However, it is now known that flavodoxin can accept electrons in the absence of PsaD, and it is possible, although unproven, that the flavodoxin gene can become induced in the absence of PsaD. This finding is idiosyncratic to Synechocystis sp. strain PCC 6803. PsaD is required in Synechococcus sp. strain PCC 7002, which is unable to grow photoheterotrophically in its absence even at iron concentrations which led to the full induction of flavodoxin formation (W. M. Schluchter and D. A. Bryant, personal communication). A more recent study of a psaD deletion mutant of Synechocystis sp. strain PCC 6803 has shed light on some of these uncertainties (Y.-S. Jung, V. Chitnis, P. Chitnis and J. H. Golbeck, unpublished results). Firstly, it was found that isolated thylakoids did contain wild-type levels of PsaC and PsaE. This shows that Synechocystis sp. strain PCC 6803 is able to assemble a functional PS I complex in the absence of PsaD. It also indicates that PsaE can assemble on the PS I complex in the absence of PsaD; this result differs from a conclusion reached by Chitnis and Nelson (1992), who reported that stable assembly of PsaE on the PS I complex required the presence of PsaD. Secondly, it was found that PsaC (assayed by immunoblotting) and the iron-sulfur clusters (assayed by EPR spectroscopy) were lost when the thylakoids were treated with 0.1 % Triton X-100. When recombinant Nostoc sp. strain PCC 8009 PsaD was rebound to the membranes, PsaC and were retained in the presence of 1% Triton X-100. The mutant strain did not grow photoautotrophically but did grow photoheterotrophically in the presence of glucose and 3-(3,4-dichlorophenyl)-1,1 -dimethylurea (DCMU). This implies that cyclic electron flow was unimpaired, even though there was no measurable reduction of ferredoxin (measured by FNR-mediated photoreduction) in isolated thylakoids.

John H. Golbeck

C. PsaE 1. Structure The primary amino acid sequence of PsaE has been deduced from the gene sequence in Synechococcus sp. strain PCC 7002 (Bryant et al., 1990; Zhao et al., 1993), Synechococcus sp. (Mühlenhoff et al., 1993), Synechococcus sp. strain PCC 6301 (Rhiel and Bryant, 1993), Nostoc sp. strain PCC 8009 (Bryant et al., 1990), and Synechocystis sp. strain PCC 6803 (Chitnis et al., 1989b; Rousseau and Lagoutte, 1990), and partial or whole sequences are available from amino acid sequencing in Synechococcus vulcanus (Koike et al., 1989), Calothrix sp. strain PCC 7601 (Mann et al., 1991) and Anabaena variabilis strain ATCC 29413 (Nyhus et al., 1991). The actual or deduced amino acid sequences of the PsaE protein are tabulated and compared in Table 12. The sequences predict a basic protein with a mass between 7.7 and 8.1 kDa containing 70 to 75 amino acids. There is a high degree of sequence identity among cyanobacteria (about 73%); the only significant difference is that PsaE proteins from the filamentous, heterocystous cyanobacteria Nostoc sp. strain PCC 8009, Calothrix sp. strain PCC 7601 and Anabaena variabilis strain ATCC 19413 (as well as the plastidencoded PsaE protein of the red alga Porphyra umbilicalis and the nuclear-encoded proteins of higher plants) have a seven-residue deletion near the carboxyl-terminus (see Table 12). This deletion occurs in a large, external loop region of the protein (see below), and it should not alter the dominant sheet structure of PsaE (see below). Higher plant PsaE proteins are somewhat larger than the cyanobacterial homologs; these proteins, containing 91 to 101 amino acids, have the additional amino acids located at the amino-terminal ends of the proteins. The PsaE protein is hydrophilic except for a small stretch of hydrophobic amino acids (SGIIYPVIV) at positions 31 to 39 and it is positively charged at pH 7.0. In most cyanobacteria, the carboxyl-terminal sequence is entirely composed of alanine, proline, and lysine. The exception is Synechococcus sp. strain PCC 7002, in which the carboxyl-terminus is truncated prior to this region. There are between seven or eight conserved aromatic residues, depending on whether the external loop is truncated. The location of PsaE on the stromal side of PS I is inferred from the absence of a transit peptide and from the fact that it can be cross-linked to PsaD,

Chapter 10 Photosystem I Reaction Center

which in turn, can be cross-linked to soluble ferredoxin (Oh-oka et al., 1989; Andersen et al., 1990). PsaE, along with PsaD and PsaC, can be removed from the PS I complex with chaotropic agents, and it is rebound only in the presence of refolded or native PsaC (Li et al., 199 la; Zhao et al., 1993). Certain cationic detergents, such as dodecyltrimethylammonium bromide, when used in combination with 2.5 M NaCl, extract PsaE selectively. PsaE can be rebound by simple addition to the depleted complex (Sonoike et al., 1993). Secondary structure predictions based on ChouFasman and Robson-Garnier criteria indicate that the protein should be largely between residues 15 and 55, but there may be some structure between residues 60 and 65 (but see below). Limited proteolysis ofhigher plant PS I complexes show that the amino-terminal region of PsaE up to residue 15 is surface exposed (Zilber and Malkin, 1992; Lagoutte and Vallon, 1992). This region is totally missing in the cyanobacterial PsaE protein. Internal regions near Met39 in PsaE and between Met74 and Met 140, including most ofthe basic amino acid clusters, may be surface exposed in the spinach PsaE protein (Lagoutte and Vallon, 1992). Comparable data is unavailable for the cyanobacterial protein, although it is expected to be quite similar (except for the 7

351

amino acid deletion), especially in the carboxylterminal region. An NMR solution structure of PsaE has been recently determined (see Fig. 8; Falzone et al., 1994a,b). As the structural algorithm predicted, the secondary structure ofthe protein includes a relatively high content of the NOE constraints give rise to a well-defined structure composed of five antiparallel segments of The topology is represented by (+1, +1, +1, –4x), which brings the first and last strands (and consequently the aminoand carboxyl-termini) together. Surprisingly, the structure is quite similar to Src homology 3 domains found in eucaryotic proteins of signal transduction pathways (Falzone et al., 1994). There is a large, poorly constrained loop connecting strands three and four, that calculations indicate can adopt many energy-equivalent conformations. This loop has an rms deviation that is much greater than for the remainder of the structure (for regions of the structure not found in disordered loops, the rms deviations are less than 0.3 Å). The distance from the aminoterminus to the farthest residue in the loop is between 26 and 44 Å; it is possible that the loop is mobile in solution, although this cannot be yet be concluded with certainty from the NMR data. Perpendicular to that longer axis of the protein, the widest dimension

352 is about 30 Å. The large unconstrained loop is seven amino acids shorter in several cyanobacteria, including Nostoc sp. strain PCC 8009, and in all higher plants, but this is not expected to change the overall appearance ofthe protein and suggests that it is not the external loop but the compact domain of the protein that is functionally important. It is interesting to note that the charged residues are located almost exclusively on two ‘edges’ of the protein (see Fig. 8), and these faces might represent the two binding sites for PsaC (or the PsaA/PsaB heterodimer) and for a soluble electron carrier.

2. Function There are two suggested roles for PsaE in PS I. One proposed function is to stimulate the reduction of soluble electron acceptors such as ferredoxin and/or flavodoxin. Weber and Strotmann (1993) showed that the rate of ferredoxin-mediated photoreduction declined in spinach PS I as the PsaE protein was removed with low concentrations of chaotropic agents. At the point ofcomplete removal, the rate of photoreduction was about onehalf of the control rate. The addition of recombinant PsaE from Synechococcus sp. strain PCC 7002 led to the recovery of about one-half of the missing rate. Flavodoxin reduction was shown to be enhanced when Synechococcus sp. strain PCC 7002 PsaC and Nostoc sp. strain PCC 8009 PsaD were reconstituted onto a core isolated from Synechococcus sp. strain PCC 6301 (see Golbeck, 1993b). The rate of flavodoxin photoreduction was enhanced by a factor of two when Synechococcus sp. strain PCC 7002 PsaE was included in the reconstitution protocol. Sonoike et al. (1993) showed that the addition of PsaE to a depleted PS I complex from Synechococcus sp. promoted the interaction between and ferredoxin after a single-turnover flash. Rousseau et al (1993) found that PS I lacking PsaE through deletion mutagenesis exhibited a 25-fold slower rate of ferredoxin reduction relative to the wild-type after a saturating flash. When PsaE was added back to the deficient membranes, the original rate of ferredoxin reduction was recovered (second-order rate constant of approximately These rate experiments imply that PsaE functions as a structural element in facilitating interaction between ferredoxin and flavodoxin and the PS I complex. Yet, this cannot be the major role for PsaE since no aberrant phenotype was initially found when

John H. Golbeck psaE was inactivated by interposon mutagenesis in Synechocystis sp. strain PCC 6803 (Chitnis et al., 1989b) or Synechococcus sp. strain PCC 7002 (Bryant et al., 1990; Zhao et al., 1993). There is no effect in either organism on the rate of photoautotrophic growth or oxygen evolution in saturating light, and there is no effect on non-cyclic electron transport (Bryant et al., 1990; Zhao et al., 1993). Why then, does there appear to be a mismatch ofthe biochemical and physiological data? Analysis of overall electron transport rates in whole cells support the idea that faster PS I turnover does not necessarily lead to higher rates of overall electron transport activity. To illustrate this point, the throughput of electrons through PS I (and PS II) in plants is limited by a rate bottleneck imposed on plastoquinone oxidation at the cytochrome complex. The 200 eq. throughput of electrons in the electron transport chain (Kok, 1973) implies that PS I needs to turn over only once every 5 ms. However, full sunlight excites a chlorophyll molecule 10 times per second, and with 100 chlorophylls per trap, PS I should be minimally capable of turning over once every 1 ms (many cyanobacteria rarely experience full sunlight, however). The rate of ferredoxin reduction implied by the single-turnover experiments (a half-time of at ferredoxin and 2.2 – PS I) is capable of supporting a throughput of 10,000 eq. It is reasonable to suppose that the rapid half-time for the interaction of ferredoxin with PS I ensures that the 30 ms back reaction between P700+ and is completely suppressed (no more than 0.3% of the electrons would backreact under these conditions). It is also possible that a rapid donation of electrons from to ferredoxin resets the reaction center sufficiently rapidly to accommodate the statistically-uneven flux of photons that strike the PS I reaction center. Yet these must be relatively minor issues, since all of these arguments fail to account for the equal rates of oxygen evolution and equal growth rates of cyanobacteria in the presence and absence of the PsaE polypeptide. The data might be explained if the overall rate limiting step in photosynthesis and growth lies elsewhere than the throughput of electrons in PS I. A second function for the PsaE protein was discovered after it was found that a psaE mutant strain of Synechococcus sp. strain PCC 7002 was more sensitive to increased growth temperature than the wild-type (Golbeck and Bryant, 1991). This phenotype suggested that a lesion might exist in the

Chapter 10 Photosystem I Reaction Center

PS I complex that becomes apparent at elevated temperatures. One possibility was that this function might be associated with the need for additional ATP to cope with the increased demands of ion pumping and metabolism at elevated temperatures.

353

Zhao et al. (1993) then showed that the psaE mutant of Synechococcus sp. strain PCC 7002 grew much more slowly than the wild type strains at low light intensities and at air levels of (compared to the normal growth conditions of 1% routinely

John H. Golbeck

354 employed in the laboratory). Moreover, the Synechococcus sp. strain PCC 7002 psaE mutant did not grow at all under photoheterotrophic conditions (glycerol + DCMU), and the Synechocystis sp. PCC 6803 psaE deletion mutant only grew extremely slowly (if at all) under these conditions (Zhao et al., 1993). To check for the absence of cyclic electron flow, a protocol was developed determine the relative number of PS I electrons returned to P700. The reduction rates of were measured in whole cells in the presence of mutations to inhibit the input of electrons from the respiratory complex and inhibitors to block input of electrons from PS II(DCMU) or the efflux of electrons to the respiratory oxidases (cyanide). The presence of a cycle around PS I could then be inferred from the rate of reduction in the presence and absence of methyl viologen. In the absence of methyl viologen, the psaE deletion mutant and ndhF/PsaE double deletion mutant showed reduction kinetics identical to the wild-type in the presence of methyl viologen, indicating that cyclic electron flow around PS I was absent (Yu et al., 1992; Yu et al., 1993b). A quantitative analysis of the throughput showed that under saturating light conditions, cyclic electron flow in wild-type Synechococcus sp. strain PCC 7002 accounts for 2.6% of the electron flux, and the NADH dehydrogenase contributes about 5%. Maxwell and Biggins (1976, 1977) concluded from the reduction kinetics in various cyanobacteria that cyclic electron transport does not contribute appreciably to photosynthesis in oxygen-evolving autotrophs. Myers (1986, 1987) also sought evidence of cyclic flow from analysis of the steady-state fraction of P700 reduced as a function of the intensity of light, and found that the total return electron flow, which is defined as cyclic plus respiration in the light, cannot be much larger than dark respiration. All three studies agree that cyclic electron flow cannot represent a significant process in cyanobacteria at high light intensities. However, the growth experiments indicate that cyclic electron transport must be much more important at low light intensity or under conditions of limitation (Zhao et al., 1993). The electron transport agent(s) involved in the cyclic process is not known; certainly, the low percentage ofelectron fluxthrough the cyclic pathway may make the search for cofactors much more difficult.

Acknowledgments Unpublished work from the author’s laboratory was supported by grants from the National Science Foundation (DMB-904333 and MCB-9205756) and the Layman Fund of the University ofNebraska. The author thanks Don Bryant, Parag Chitnis, Chris Falzone, Juliette Lecomte, Wolfgang Nitschke, and Matthias Rögner for providing preprints of unpublished work; Wendy Schluchter, Jindong Zhao, and Uli Mühlenhoff, for communicating unpublished results; John Biggins, Egbert Boekema, Norbert Krauß, Matthias Rögner, and Wolfram Saenger for generously supplying figures; Paul Scott and YeanSung Jung for proofreading the manuscript; and Marion O’Leary for the generous use ofhis secluded cabin on Washington Island, WI.

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Chapter 10 Photosystem I Reaction Center 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 Rousseau F and Lagoutte B (1990) Amino acid sequence of Photosystem I subunit IV from the cyanobacterium Synechocystis sp. PCC 6803. FEBS Lett 260: 241–244 Rousseau F, Sétif P and Lagoutte B (1993) Evidence for the involvement of PS I-E in the reduction of ferredoxin by Photosystem I. EMBO J 12: 1755–1765 Rustandi RR, Snyder SW, Biggins J, Norris JR and Thurnauer MC (1992) Reconstitution and exchange of quinones in the site of Photosystem I - An electron spin polarization electron paramagnetic resonance study. Biochim Biophys Acta 1101: 311–320 Rustandi RR, Snyder SW, Feezel LL, Michalski TJ, Norris JR, Thurnauer MC and Biggins J (1990) Contribution of vitamin to the electron spin polarization in spinach Photosystem I. Biochemistry 29: 8030–8032 Rutherford AW and Mullet JE (1981) Reaction center triplet states in Photosystem I and Photosystem II. Biochim Biophys Acta 635: 225–235 Sakurai H, Inoue K, Fujii T and Mathis P (1991) Effects of selective destruction of iron-sulfur center B on electron transfer and charge recombination in Photosystem I. Photosynth Res 27: 65–71 Scheller HV and Møller, BL (1990) Photosystem I polypeptides. Physiol Plant 78: 484–494 Scheller HV, Okkels JS, Høj PB, Svendsen I, Roepstorff P and Møller BL (1989a) The primary structure of a 4.0 kDa Photosystem I polypeptide encoded by the chloroplast psaI gene. J Biol Chem 264: 18402–18406 Scheller HV, Svendsen I and Møller BL (1989b) Subunit composition of Photosystem I and identification of center X as a [4Fe-4S] iron-sulfur cluster. J Biol Chem 264: 6929–6934 Sétif P (1992) Energy transfer and trapping in Photosystem I. In: Barber J (ed) Current Topics in Photosynthesis, Vol 11, The Photosystems: Structure, Function, and Molecular Biology, pp 471–499. Elsevier, Amsterdam Sétif P and Bottin H (1989) Identification of electron-transfer reactions involving the acceptor of Photosystem I at room temperature. Biochemistry 28: 2689–2697 Sétif P and Brettel K (1990) Photosystem I photochemistry under highly reducing conditions: study of the P700 triplet state formation from the secondary radical pair Biochim Biophys Acta 1020: 232–238 Sétif P and Brettel K (1993) Forward electron transfer from phylloquinone–A(l) to iron sulfur centers in spinach Photosystem-I. Biochemistry 31: 7846–7854 Shimizu T, Hiyama T, Ikeuchi M, Koike H. and Inoue Y (1990) Nucleotide sequence of the psaC gene of the cyanobacterium Synechococcus vulcanus. Nucl Acids Res 18: 3644–3644 Shimizu T, Hiyama T, Ikeuchi M and Inoue Y (1992) Nucleotide sequences of the psaA and psaB genes encoding the Photosystem I core proteins from the thermophilic cyanobacterium Synechococcus vulcanus. Plant Mol Biol 18: 785–791 Shinozaki K, Ohme M, Tanaka M, Wakasugi T, Hayashida N, Matsubayashi T, Zaita N, Chunwongse J, Obokata J, Yamaguchi-Shinozaki K, Ohto C, Torazawa K, Meng BY, Sugita M, Deno H, Kamogashira T, Yamada K, Kusuda J, Takaiwa F, Kato A, Tohdoh N, Shimada H, and Sugiura M (1986) The complete nucleotide sequence of tobacco chloroplast

359 genome: its gene organization and expression. EMBO J 5: 2043–2049 Shoham G, Michaeli D and Nechushtai R (1990) Photosystem I reaction center of Mastigocladus laminosus: Structural and functional aspects. In: Baltscheffsky M (ed.) Current Research in Photosynthesis, Vol II, pp 555–562. Kluwer, Dordrecht Shuvalov VA, Nuijs AM, van Gorkom HJ, Smit JWJ and Duysens LNM (1986) Picosecond absorption changes upon selective excitation of the primary electron donor P-700 in Photosystem I. Biochim Biophys Acta 850: 319–323 Sieckmann I, Brettel K, Bock C, Van der Est A and Stehlik D (199la) Transient electron paramagnetic resonance of the triplet state of P(700) in photosystem .1. Evidence for triplet delocalization at room temperature. Biochemistry 32: 4842– 4847 Sieckmann I, Van der Est A, Bottin H, Sétif P and Stehlik D (1991 b) Nanosecond electron transfer kinetics in Photosystem I following substitution of quinones for vitamin as studied by time resolved EPR. FEBS Lett 284: 98–102 Smart LB and McIntosh L (1991) Expression of photosynthesis genes in the cyanobacterium Synechocystis sp. PCC 6803: psaA-psaB and psbA transcripts accumulate in dark-grown cells. Plant Mol Biol 17: 959–971 Smart L, Warren PV, Golbeck JH and McIntosh L (1993) Mutational analysis of the structure and biogenesis of the Photosystem I reaction center in the cyanobacterium Synechocystis sp. PCC 6803. Proc Natl Acad Sci USA 90: 1132–1136 Snyder SW, Rustandi RR, Biggins J, Norris JR and Thurnauer MC (1991) Direct assignment of as the secondary in Photosystem I. Proc Natl Acad Sci USA 88: 9895–9896 Sonoike K, Ikeuchi M and Pakrasi H (1992) Presence of an Nterminal presequence in the PsaI protein of the Photosystem I complex in the filamentous cyanobacterium Anabaena variabilis ATCC 29413. Plant Mol Biol 20: 987–990 Sonoike K, Hatanaka H and Katoh S (1993) Small subunits of Photosystem I reaction center complexes from Synechococcuselongatus .2. The psaE gene product has a role to promote interaction between the terminal electron acceptor and ferredoxin. Biochim Biophys Acta 1141: 52–57 Stehlik D, Bock C and Petersen J (1989) Anisotropic electron spin polarization of correlated spin pairs in photosynthetic reaction centers. J Phys Chem 93: 1612–1619 Steinmuller K (1992) Identification of a 2nd psaC gene in the cyanobacterium Synechocystis sp PCC 6803. Plant Mol Biol 20: 997–1001 Trebst A (1986) The topology of the plastoquinone and herbicide binding peptides of Photosystem II in the thylakoid membrane. Z Naturforsch 41C: 240–245 Trissl H.-W, Leibl W, Deprez J, Dobek A and Breton J (1987) Trapping and annihilation in the antenna system of Photosystem I. Biochim Biophys Acta 893: 320–332 Trost JT and Blankenship RE (1989) Isolation of a photoactive photosynthetic reaction center-core antenna complex from Heliobacillus mobilis Biochemistry 28: 9898–9904 Tsiotis G, Nitschke W, Haase W and Michel H (1993) Purification and crystallization of Photosystem I complex from a phycobilisome-less mutant of the cyanobacterium Synechococcus PCC 7002. Photosynth Res 35: 285–297 Vallon O and Bogorad L (1993) Topological study of PS I-A and

360 PS I-B, the large subunits of the Photosystem-I reaction center. Eur J Biochem 214: 907–915 van der Staay, GWM, Boekema E, Dekker JP, and Matthijs HCP (1993) Characterization of trimeric Photosystem I particles from the prochlorophyte Prochlorothrix hollandica by electron microscopy and image analysis. Biochim Biophys Acta 1142: 189–193 Warren PV, Parrett KG and Golbeck JH (1990) Characterization of a Photosystem I core containing P700 and intermediate electron acceptor Biochemistry 29: 6545–6550 Warren PV, Golbeck JH and Warden JT (1993a) Charge recombination between and occurs directly to the ground state of P700 in the absence of and Biochemistry 32: 849–857 Warren PV, Smart LB, McIntosh L and Golbeck JH (1993b) Site-directed conversion of cysteine 565 to serine in PsaB of Photosystem I results in the assembly of [3Fe-4S] and [4Fe4S} clusters in A mixed-ligand [4Fe-4S] cluster is capable of electron transfer to and Biochemistry 32:4411–4419 Wasielewski MR, Fenton JM and Govindjee (1987) The rate of formation of in Photosystem I particles from spinach as measured by picosecond transient absorption spectroscopy. Photosynth Res 12: 181–190 Webber AN and Malkin R (1990) Photosystem I reaction-center proteins contain leucine zipper motifs. A proposed role in dimer formation. FEBS Lett 264: 1–4 Webber AN, Gibbs PB, Ward JB and Bingham SE (1993) Sitedirected mutagenesis of the Photosystem-I reaction center in chloroplasts—the cysteine-proline motif. J Biol Chem 268: 12990–12995 Weber N and Strotmann H (1993) On the function of subunit PsaE in chloroplast Photosystem I. Biochim Biophys Acta 1143: 204–210 Witt, HT, Rögner M, Mühlenhoff U, Witt I, Hinrichs W, Saenger W, Betzel C, Dauter Z and Boekema, EJ (1990) On isolated complexes of reaction center I and X-ray characterization of single crystals. In: Baltscheffsky M (ed) Current Research in Photosynthesis, Vol II, pp 547–554. Kluwer, Dordrecht Witt HT, Krauß N, Hinrichs W, Witt I, Fromme P and Saenger, W (1992) Three-dimensional crystals of Photosystem I from Synechococcus sp and X-ray structure analysis at 6 Å resolution. In: Murata N (ed) Research in Photosynthesis, Vol I pp 521– 528. Kluwer, Dordrecht Wolfe GR, Cunningham FX Jr and Gantt E (1992) In the red alga Porphyridium cruentum Photosystem I is associated with a putative LHC complex. In: Murata N (ed) Research in

John H. Golbeck Photosynthesis, Vol I, pp 315–318. Kluwer, Dordrecht Wynn RM and Malkin R (1988) Interaction of plastocyanin with Photosystem I: a chemical cross-linking study of the polypeptide that binds plastocyanin. Biochemistry 27: 5863–5869 Wynn RM and Malkin R (1990) The Photosystem I 5.5 kDa subunit (the psaK gene product). An intrinsic subunit of the PS I reaction center complex. FEBS Lett 262: 45–48 Wynn RM, Omaha J and Malkin R (1989) Structural and functional properties of the cyanobacterial Photosystem I complex. Biochemistry 28: 5554–5560 Xu Q, Yu L, Chitnis VP and Chitnis PR (1993) Function and organization of Photosystem I in a cyanobacterial mutant strain lacking PsaF and PsaJ subunits. J Biol Chem (in press). Yu L, Golbeck JH, Zhao JD, Schluchter W, Mühlenhoff U and Bryant D (1992) The PsaE protein is required for cyclic electron flow around Photosystem I in Synechococcus sp. PCC 6301. In: Murata N (ed) Current Research in Photosynthesis, Vol I, pp 565–568. Kluwer, Dordrecht Yu L, Zhao J, Lu W, Bryant DA and Golbeck JH (1993a) Characterization of the [3Fe-4S] and [4Fe-4S] clusters in unbound PsaC mutants CUD and C51D. The midpoint potentials of the single [4Fe-4S] clusters are identical to and in bound PsaC of Photosystem I. Biochemistry 32:8251– 8258 Yu L, Zhao JD, Mühlenhoff U, Bryant DA and Golbeck JH (1993b) PsaE is required for cyclic electron flow around Photosystem I in the cyanobacterium Synechococcus sp. PCC 7002. Plant Physiol 103: 171–180 Zhao JD Warren PV, Li N, Bryant DA and Golbeck JH (1990) Reconstitution of electron transport in Photosystem I with PsaC and PsaD Proteins expressed in Escherichia coli. FEBS Lett. 276: 175–180. Zhao JD, Li N, Warren PV, Golbeck JH and Bryant DA (1992) Site-directed conversion of a cysteine to an aspartate leads to the assembly of a [3Fe-4S] cluster in PsaC of Photosystem I. Photoreduction of is independent of Biochemistry 31: 5093–5099 Zhao JD, Snyder WB, Mühlenhoff U, Rhiel E, Warren PV, Golbeck JH and Bryant DA (1993) Cloning and characterization of the psaE gene of the cyanobacterium Synechococcus sp. PCC 7002: characterization of a psaE mutant and overproduction of the protein in Escherichia coli. Mol Microbiol 9: 183–194 Zilber AL and Malkin R (1992) Organization and topology of Photosystem I subunits. Plant Physiol 99: 901–911

Chapter 11 The F-type ATPase in Cyanobacteria: Pivotal Point in the Evolution of a Universal Enzyme Wayne D. Frasch Department of Botany, Arizona State University, Tempe, AZ 85287-1601, USA 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 A. Potential Phosphate Oxygen Binding Sites B. Potential Metal Ligands C. Potential Adenosine Ring Binding Sites VI. Location of the Noncatalytic Metal-Nucleotide Binding Site VII. Model of the Metal-Nucleotide Binding Sites of CF1 VIII. Regulation of Catalytic Activity A. Regulation by the and subunits B. Regulation via Noncatalytic Nucleotide Binding Sites Acknowledgments References

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Summary The structure and mechanism of the from cyanobacteria shares several characteristics with the chloroplast enzyme but also resembles a bacterial ATPase in many ways. The overall subunit composition and organization of the enzyme appear closely similar in all organisms. The substructure is composed of three heterodimers of and subunits in which the subunits are somewhat more distal from the membrane. Comparison of the organization ofthe genes encoding these subunits in cyanobacteria with that found in other species supports the hypothesis that two events occurred in the divergence of bacteria from cyanobacteria: (i) separation of the atp1 and atp2 operons and; (ii) the duplication and divergence of the gene for the b subunit into b and b'. The b, b' and subunits are each thought to bind to one of the three heterodimers to make an asymmetric complex. The subunit is also associated with the heterodimers in an asymmetric manner. Metals and nucleotides bind as complexes to the ATPase at a total of six sites of which three are catalytic. At the catalytic sites, all ofthe phosphates are coordinated to the metal in the active state and ATP synthesis occurs via an in-line transfer of the phosphoric residue between ADP and water. The equilibrium constant at each catalytic site (unisite catalysis) is about one which implies: (i) that the is not required to make ATP but to release the product and (ii) that the catalytic sites are coupled (binding-change mechanism) such that binding of substrate to one site facilitates release of product from a second site. Several independent lines of evidence suggest that induces sequential conformational changes that facilitate ATP release (indirect coupling). The catalytic and non-catalytic metal-nucleotide binding sites can be distinguished and selectively filled by their unique binding properties. The non-catalytic sites show higher specificity for adenine nucleotides and bind to the metal as a monodentate complex in the latent enzyme form. In the chloroplast the six sites (N1–N6) fill in order of increasing dissociation constants in which N1, N3 and N4 are catalytic sites. Both the and D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 361–380. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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subunits contain the P-loop sequence (GXXXXGKT) that is conserved in most enzymes that bind ATP. The Ploop of the subunit is directly involved with the catalytic site. The noncatalytic site is less well defined and is thought to bridge the and subunits. Several amino acid side chains on these subunits have been implicated in binding the metal-nucleotide complexes. The measured distances between the N1–N3 sites and the crystal structure of the suggests that the N2 and N3 sites comprise one heterodimer. The F-ATPase is a highly regulated enzyme that presumably throttles the supply of ATP to the cell and minimizes the ATPase reaction. The enzyme from cyanobacteria contains some but not all of the regulatory mechanisms present in the chloroplast enzyme. The required for activation is decreased significantly in by reduction ofthe disulfide in the subunit. Since activation ofthe oxidized form of the enzyme requires a higher than the enzyme needs to catalyze ATP synthesis, the hydrolysis of ATP rarely occurs. This disulfide is missing in the cyanobacterial ATPase and results in a low requirement for activation. High levels of ATP are thought to be maintained in the dark by the oxidative phosphorylation system in the cyanobacteria. Dissociation of the e subunit also activates the ATPase and eases disulfide reduction. Disulfide reduction also facilitates dissociation. The binding of metal-nucleotide complexes or free metal to the noncatalytic sites also induces an up- or down-regulation of the ATPase activity, respectively. The strongly cooperative binding of metals may be involved in regulating the activity by affecting the binding-change mechanism.

I. Introduction In cyanobacteria, as in all other life forms, energy is harvested by electron transfer reactions that create a transmembrane, electrochemical proton gradient The is then used to drive the synthesis of ATP from ADP and phosphate via an F-type ATPase, thereby converting the energy into a relatively stable form that is accessible by a wide variety ofenzymes. Despite the fact that the F-type ATPase has been highly conserved throughout evolution, subtle changes have occurred that distinguish the enzyme in bacteria from that found in animal mitochondria and plant chloroplasts. The universal nature of this ATPase has provided a means with which to follow the increased complexity due to the evolution from procaryotic to eucaryotic organisms. The increased complexity is evident in terms of gene organization, subunit composition and in the layers of regulatory mechanisms that govern the enzymatic activity of the ATPase. Under some circumstances, the F-type ATPases can hydrolyze ATP to pump protons in the reverse direction. In some cases the ATPase-generated is used to drive other cellular processes. However, Abbreviations: portion chloroplast coupling factor ATPase; DCCD – N,N'-dicyclohexylcarbodiimide; Escherichia coli ATPase; – mitochondrial ATPase; FRET– fluorescence resonance energy transfer; FSBA – 5'-pfluorosulfonylbenzoyladenosine; FSBI – 5'-p-fluorosulfonylbenzoylinosine; OGP – n-octyl

the F-type ATPases of most organisms, and particularly the higher-plant-chloroplast ATPase involved in photosynthetic ATP synthesis, have regulatory mechanisms within the enzyme that minimize the extent of ATPase activity. Reviews of the F-ATPases from bacteria, mitochondria and chloroplasts (Futai et al., 1989; Fillingame, 1990; Boyer, 1993; Peterson and Amzel, 1993) and their evolution with regards to other proton ATPases (Nelson, 1992) have recently appeared. The cyanobacterial F-ATPases have not been well characterized compared to the enzymes from some other organisms. Methods have now been developed to purify the from cyanobacterial sources that can be used to reconstitute photophosphorylation (Lubberding et al., 1983; Hicks and Yocum, 1986). These purified ATPases contain the same five subunits with similar molecular masses to those of other F-ATPases. The has also been purified and functionally reconstituted into proteoliposomes with the cyanobacterial cytochrome complex (Krenn et al., 1993). Thermodynamically, catalysis of ATP synthesis by the cyanobacterial enzymes is similar that of other F-ATPases (Peschek et al., 1986; Nitschmann and Peschek, 1986). However, the cyanobacterial F-ATPase is particularly interesting due to the presence of both oxidative phosphorylation complexes and a photosynthetic apparatus analogous to that of higher plant chloroplasts in the same cellular compartment (see Chapter 13). Just as cyanobacteria share traits with

Chapter 11 F-Type ATPase both chloroplasts and bacteria, the cyanobacterial Ftype ATPase shares some ofthe characteristics ofthe chloroplast enzyme but also resembles the enzyme of bacterial origin in many aspects. II. Organization of Subunits The F-type ATPase is composed of and portions. The portion is an integral membrane protein complex comprised of single copies of subunits a, b and b' subunits and about ten copies of subunit c. This complex serves as a protonpore in the membrane to couple the energy derived from electron transfer reactions to the portionoftheenzyme.Theextrinsic protein complex attached to has a subunit stoichiometry (Süss and Manteuffel,

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1982; Walker et al., 1985). A total ofsix nucleotide binding sites are known to exist on (Cross and Nalin, 1982; Senior, 1988; Girault et al., 1988). Three of these sites are catalytic, each residing primarily on a subunit, while the other three noncatalytic sites are primarily on each of the subunits. The chloroplast enzyme also has six metal binding sites (Haddy et al., 1985; Hiller and Carmeli, 1985). Coordination of the nucleotide phosphates by the bound metals facilitates catalysis (Frasch and Selman, 1981). Figure 1 shows the structure of the rat mitochondrial as it is understood at this time from protein crystallography at 3.6 Å resolution (Bianchet et al., 1991). The and subunits each exist as trimeric arrays that are organized in two offset layers along the three-fold symmetry axis. On the bottom

364 (proximal to and the membrane surface) the subunits are located close to the axis and are in contact with each other. The subunits are above the subunits and are farther from the three-fold symmetry axis. Each subunit interacts with an subunit. Due to the sequence homologies of the and subunits, these subunits can be considered to be heterodimers. Image-averaged electron micrographs of from spinach chloroplasts (Fig. 2) provide a lower resolution structure but resolve the location of the smaller subunits (Boekema et al., 1988; Lücken et al., 1990). The enzymes from a variety ofsources are closely similar in appearance as viewed by this technique (Akey et al., 1984; Ysern et al., 1988; Boekema et al., 1988). Comparison of imageaveraged micrographs of with those of subunitdepleted enzyme revealed that the and subunits are associated with one ofthe three heterodimers (Boekema et al., 1990). The structure ofthe bovine mitochondrial at 6.5 Å resolution has recently appeared as shown in Fig. 3 (Abrahams et al., 1993). Unlike the rat enzyme, this structure shows clear evidence of asymmetry. Viewed from the top, the bovine crystal structure has an asymmetric cleft similar to that observed by image-averaged electron microscopy. A 40 Å stem protrudes from the bottom ofthe which is thought to be part of the stalk that joins the protein to the membrane domain. Adjacent to the stem is a pit that may be the docking site of the b subunits from The central core of the bovine structure contains two helices 90 Å and 45 Å in length, oriented vertically from the bottom of the stem, that form a left-handed coiled coil. The longer helix contacts the and subunits at several points and is suggested to be part of the or subunits.

III. Gene Organization In cyanobacteria, the subunits for the ATPase are encoded on two operons. The gene products of the atp1 operon of Synechococcus sp. strain PCC 6301 are a,c,b',b, in order of transcription (Cozens and Walker, 1987). The atp2 operon contains the genes for the and subunits. In this operon, the gene for the subunit precedes the subunit with a 4-basepair overlap between the two genes. Anabaena sp. strain PCC 7120 and Synechocystis sp. strain PCC 6803 have a gene organization that resembles

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that of Synechococcus sp. strain PCC 6301 (McCarn et al., 1988; Lill and Nelson, 1991) This organization appears to be an intermediate evolutionary step between bacteria and chloroplasts (see Chapter 5). In E. coli, the genes for the all of the subunits of the ATPase are found in the unc operon, so named because deletion of this operon leads to a phenotype of uncoupled respiration (Butlin et al., 1971). The order of transcription of the genes is such that the products are formed in the order I, a, c, b, and (Futai and Kanazawa, 1983). The UncI protein is not a subunit ofthe enzyme but may be involved in the folding or assembly. It is noteworthy that the order of transcription produces the components first, followedbythe subunits which are transcribed roughly in the same order as their proximity to at the membrane surface in the assembled complex. An open reading frame of unknown function occurs at the 5' end of the atp1 operon in Synechocystis sp. strain PCC 6803, coding for a putative gene product similar to UncI in E. coli (Lill and Nelson, 1991). Two events have occurred in the divergence of E. coli from its common ancestor with cyanobacteria. Firstly, E. coli appears to have condensed the two operons into a single unit. This hypothesis is based on the observation that the green bacterium Chlorobium sp. and other eubacteria all contain a

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366 separate operon encodingthe and subunits.Based on this observation, it has been suggested that atp2 is the more primitive of the two operons (Nelson, 1992). In several chloroplast genomes, the atpB subunit) and atpE ( subunit) genes overlap by a few nucleotides (Palmer, 1983). While this is the case for most of the cyanobacterial species sequenced thus far, the spacing between these genes inAnabaena sp. strain PCC 7120 is more similar to E. coli in which a spacer region separates the two genes (Curtis, 1987). The second event is the presence of the genes for the b and b' subunits in Synechococcus sp. strain PCC 6301 andAnabaena sp. strainPCC 7120 (Cozens and Walker 1987; McCarn et al., 1988). Since the ratio of a to b subunits in E. coli is 1:2 (Foster and Fillingame, 1982), the presence ofthe closely related b and b' subunits in cyanobacteria appears to have resultedfroma gene duplication event (Nelson, 1992). The independent evolution of these two genes then led to the asymmetry of the complex in cyanobacteria. The subunit composition and gene organization of the cyanobacterial ATPase also likely represents an intermediate step in the formation of the symbiotic relationship that led to the chloroplast in higher plants. The atp2 operon in chloroplasts is still present with no major alterations. However, the atp1 operon only contains the genes encoding subunits in the order of IV (a), III (c), I (b) and The genes encoding subunits II (b'), and have migrated to the nucleus. In Anabaena sp. strain PCC 7120, the atp1 cluster is preceded by regions similar to the Shine-Dalgarno ribosome binding sequences in E. coli (McCarn et al. 1988). Despite low sequence homology, subunits I, II, III and IV have been correlated to b, b', c and a in bacteria (Herrmann et al., 1985; Henning and Herrmann, 1986). In fact, the chloroplast subunit I can successfully substitute for subunit b in E. coli (Schmidt et al., 1990). Thus, the cyanobacterial and chloroplast genes in the atp1 operon exist in the same transcriptional order with the gene duplication event occurring in the cyanobacteria. The asymmetry of the ATPase subunit organization in the chloroplast is likely to have resulted from divergence of the genes after the duplication event that occurred in cyanobacteria. Consistent with the intermediary position of cyanobacteria in the evolution between bacteria and chloroplasts, the gene that codes for chloroplast subunit I contains an intron that is lacking in Anabaena sp. strain PCC 7120, Synechococcus

Wayne D. Frasch sp. strain PCC 6301 and E. coli (Cozens and Walker, 1987; McCarn et al., 1988). Analysis of amino acid sequences for the ATPase from a wide variety oforganisms show that the different subunits have undergone variable rates of evolution. The most highly conserved sequences are in the subunit followed bythe subunit. The c and a subunits in cyanobacteria have greater sequence homology to the chloroplast subunits than to those from E. coli. Comparison of subunit c sequences from 38 organisms in bacteria, fungi, mammals and higher plants have been used to create phylogenetic maps (Recipon et al., 1992). This comparison places cyanobacteria together with chloroplasts in one lineage while other eubacteria except for purple bacteria are in a second lineage. A third lineage comprised of the purple bacteria, mitochondria and eucaryotic nuclei also exists in which the nuclear copies ofthe gene encoding subunit c are believed to originate from a mitochondrial transfer event.

IV. Mechanism of ATP Synthesis Figure 4 shows the mechanism ofATP synthesis at a single catalytic site on the The stereochemistry ofthe ribose ofthe nucleotide bound to the catalytic site of is believed to be in the anti-gauche-gauche conformation from experiments that compared the binding of nucleotide analogs in which the conformation of the ribose was restricted (Schlimme et al., 1979) and by NMR studies (Devlin and Grisham, 1990). NMR-nuclear Overhauser enhancement studies show that the nucleotide binds to the bovine mitochondrial enzyme in a similar manner (Garin et al., 1988). Divalent cations like and serve as cofactors of this reaction by forming coordination complexes with the phosphates of the bound nucleotides (Hochman et al., 1979; Bossard et al., 1980; Frasch, 1981). The stereochemistry of the reaction was determined using exchange-inert complexes of nucleotides with as inhibitors of or synthase as well as the ability of the complexes to serve as slow substrates (Frasch, 1981; Frasch and Selman, 1982). Synthesis of ATP proceeds in the chloroplast enzyme by the binding of the metalADP isomer and insertion of phosphate into the coordination sphere of the metal. The product of the reaction is a tridentate metal-ATP complex that is

Chapter 11 F-TypeATPase

probably either the or the epimer (shown). The coordination ofall ofthe phosphates of the nucleotides to the metal at the catalytic sites of has beenconfirmedby EXAFS analysis (Carmeli et al., 1986) and by NMR studies (Devlin and Grisham, 1990). However, ATP synthesis in bovine mitochondrial is thought to involve the conversion of monodentate metal-ADP to abidentate metal-ATP complex (Bossard et al., 1980). Thus, the stereochemistry of the reaction may not be the same in the from all organisms. Kinetic analyses of the dependence of the rate of ATP synthesis driven by a light-induced on the concentrations of phosphate and ADP supported a compulsory-ordered Bi-Uni mechanism in which the binding of ADP precedes the binding of phosphate (Selman and Selman-Reimer, 1981). However,

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alteration of the rate of phosphorylation by changing the concentrations of the substrates ADP and phosphate necessarily causes changes in the extent of The kinetics of ATP synthesis when was held constant support a random mechanism for the binding of phosphate and ADP (Kothen et al., 1992). The of bovine mitochondria (Webb et al., 1980) and of bacterial origins (Senter et al., 1983) have been shown to catalyze a direct, in-line transfer ofthe phosphoric residue between ADP and water(Fig. 5). Thiswasmeasuredbythe racemization of and on the phosphate produced by the enzyme during the exchange of with water. The high extent of inversion of configuration found in these experiments suggests that the catalyzes the synthesis of ATP

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via an reaction mechanism. For the to catalyze the synthesis of ATP, there must be two amino acid sidechains that can protonate an oxygen of the inorganic phosphate to make that oxygen a good leaving group and allow a nucleophilic attack by the oxygen of the of ADP (Fig. 4). Since the reaction occurs with inversion of configuration, these amino acids must be on the side ofthe inorganic phosphate that is distal from the ADP. When ATP concentrations are held sufficiently low to limit catalysis to only one of the three catalytic sites (unisite catalysis), the equilibrium constant of the reaction approaches one (O’Neal and Boyer, 1984). By measuring the extent of incorporation of into the formed by upon addition of and 40 nM ATP, it was estimated that the interconversion of ATP with ADP and phosphate occurred about 400 times prior to the release of product (O’Neal and Boyer, 1984). These and other related studies (Boyer, 1993) have supported two conclusions concerning the mechanism of ATP synthesis. Firstly, energy from the is not required to make ATP from ADP and phosphate but is required to release the product from the enzyme. Secondly, the catalytic sites are coupled to each other. This coupling of catalytic sites, known as the binding-change mechanism, proposes that the binding of substrates to one site influences the catalytic states ofthe other sites. Additional evidence to support the binding-change

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mechanism was obtained by examining the rate of ATP hydrolysis in the using the fluorescent nucleotideanalogTNP-ATP(Grubmeyer et al., 1982). When the enzyme is present in a threefold excess overATP as substrate (unisite conditions), ATP binds the enzymerapidlyand withhighaffinity, having a dissociation constant between picomolar and nanomolar. Under these conditions release of ADP and phosphate is very slow. However, the rate ofproduct release is accelerated by as much as fold when ATP is present at concentrations high enough to fill two of the three catalytic sites (Grubmeyer et al., 1982; Duncan and Senior, 1985; Noumi et al., 1986; Cunningham and Cross, 1988). Since the is not required to make ATP, the two protons needed for the synthesis of the gphosphate bond do not need to originate from the proton gradient (so-called direct coupling). Instead, proton conduction across the membrane may occur at a site that is distant from the catalytic site but is coupled to ATP synthesis by a mechano-chemical process in which the energy from the induces sequential conformational changes in the protein that facilitate release of ATP. Increasing evidence supports the indirect coupling hypothesis. Firstly,the complex from Propionigenium modestum has a subunit composition very similar to the E. coli enzyme but uses a to synthesize ATP (Laubinger and Dimroth, 1988). It is clear that ions would not be capable of directly participating in the formation of the phosphateanhydride bond. The close relationship between the

Chapter 11 F-TypeATPase ATPase of P. modestum and the ATPases was firmly established by the ability of a hybrid enzyme composed of E. coli and P. modestum to form a functional ATPase-driven pump (Laubinger et al., 1990). Secondly, the ability of the E. coli complex to catalyze ATPdriven transport and ATP synthesis at low has also been recently reported (Avetisyan et al., 1993). Finally, recent experiments examined the ratio of ADP: ATP bound to under unisite conditions to determine if the ratio changed with increasing (Labahn and Gräber, 1992). If the protons from the participatedirectly in the protonation of the phosphate oxygen to allow ATP bond formation, the equilibrium of bound ADP/ ATP should decrease at high The lack of change in bound ADP/ATP even at a was interpreted to indicate that the was used to make ATP via an indirect coupling mechanism (Labahn and Gräber, 1992).

V. Characteristics of the Metal-Nucleotide Binding Sites Conditions have been found which will fill specifically the catalytic or noncatalytic binding sites with metal-nucleotide complexes (Bruist and Hammes, 1981; Kironde and Cross, 1986). In these sites are distinguished by the ability of the catalytic sites to bind ATP, GTP or ITP which are hydrolyzed upon binding while the noncatalytic sites are specific for ATP. Dissociation of nucleotide from these sites can be prevented by covalent modification using the photoaffinity analog nucleotide. Each of the three catalytic and three noncatalytic sites can be distinguished to some extent by binding affinity which may be the result of the binding-change mechanism and/or the regulatory mechanisms of catalytic activity. The differences in binding affinities between sites varies with the source of the enzyme and are most pronounced in the chloroplast In the metal-nucleotide binding sites have been numbered N1-N6 in order of decreasing binding affinity. Sites Nl, N3 and N4 are believed to be catalytic whereas N2, N5 and N6 are noncatalytic (Bruist and Hammes, 1981; Leckband and Hammes, 1987; Xue et al., 1987; Shapiro et al., 1991a). Each ofthe first three sites can be easily distinguished and selectively filled due to its unique properties (Bruist

369 and Hammes, 1981). Resonance energy transfer measurements of fluorescent nucleotide analogs bound at these sites determined the distances between these site and other specific sites on the enzyme (Shapiro et al., 1991a,b). As shown in Fig. 6, the N1–N2, N2–N3 and N3–N1 distances are 44.3 Å, 36.7 Å and 48.4 Å, respectively. The location ofthe two cysteines (L and D) and the disulfide (SS) on the subunit as well as a unique labeling site on the subunit suggest that these subunits are associated with the heterodimer that contains the N3 catalytic site. Substitution of vanadyl for in the N2 site of chloroplast has provided a means by which to characterize the ligands to the metal at this noncatalytic site (Roskelley et al., 1992). In the latent enzyme, the N2 site binds a monodentate metal-ATP complex. Binding of the metal in the absence of nucleotide to this site occurs via cooperative binding to a second site. Analysis of the bound to the N2 site by ESEEM and ENDOR spectroscopies suggests that the most probable set of equatorial ligands to the metal consists of one water, two carboxyl oxygens and one lysine nitrogen (Houseman et al., 1994a). When the single phosphate oxygen of ATP is inserted into the coordination sphere of the bound at this site, the lysine becomes displaced from the coordination sphere but remains very close (Houseman et al., 1994b). Covalent modification studies indicate that the lysine that coordinates the metal at this site is on the subunit. Conversion of the ligands to the metal at the noncatalytic N2 site from the ATP-coordinate form to the lysine-coordinated form occurs upon the binding of nucleotide to catalytic sites at pH 8. Since it is the unprotonated amine of the lysine that ligates the metal, conversion of the noncatalytic site to the amine-coordinated form at this pH requires the forced deprotonation of the lysine. This forced deprotonation provides a feasible proton pump mechanism if coupled to the ATPase reaction at the catalytic site in the heterodimer (Houseman et al., 1994b).

VI. Location of the Metal-Nucleotide Binding Site on the subunit-the Catalytic Site The regions of the subunit sequence shown in Fig. 7 are thought to be the site of metal-nucleotide binding involved with catalysis. This structure is

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Chapter 11 F-TypeATPase based on covalent modification, on site-directed mutagenesis studies and by analogy to the structures of adenylate kinase, ras p21 and elongation factor Tu for which the crystal structures are known. The majority of site-directed mutants constructed to identifyresiduespotentiallyinvolvedwithnucleotide binding were made with the E. coli ATPase. For consistency between various studies, all residues discussed below will be described according to their location in the sequences ofthe subunits.

A. Potential Phosphate Oxygen Binding Sites The region of the subunit bounded by residues 134–155 is known as the P-loop. This sequence GGAGVGKT (149–156) which ends this glycine rich loop contains the consensus sequence GXXXXGKT found in several nucleotide binding enzymes including adenylate kinase, ras p21 and elongation factor Tu (Walker et al., 1982; Fry et al., 1985, 1986; Duncan et al., 1986) and has now been extended to include a large family of enzymes known as the adenine- nucleotide binding cassett (ABC) family. In adenylate kinase, the glycine-rich flexible loop is thought to fold over the nucleotide upon binding. A synthetic peptide of this loop (residues 134–182) binds ATP and TNP-ATP (Garboczi et al., 1988). Site-directed mutations in the E. coli subunit ofG146,G149,G150,A151,G154,K155andT156 have been made of which all but mutations of G150 or G154 had large effects on enzyme function (Parsonage et al., 1987a,b; Hsu et al., 1987; Iwamoto et al., 1991). The lysine-155 in the consensus sequence is thought to interact directly with a phosphate of the nucleotide at the catalytic site probably via a hydrogen bond to the phosphate oxygen. This amino acid becomes covalently modified by 4-chloro-7nitrobenzofurazan chloride (NBD-Cl) in beef heart at alkaline pH, thereby inactivating ATPase activity (Andrews et al., 1984a,b). Covalent derivation of by 3'-o-(5-fluoro-2,4-dinitrophenyl)-ATP also indicates that this residue is proximal to the nucleotide ribose (Chuan and Wang, 1988). In analogy to the crystal structures of adenylate kinase and elongation factor Tu, Garboczi et al. (1988) suggested that bound to the phosphate of the nucleotide. Alternatively, may bind the because or Q mutations greatly decreased ATP hydrolysis and the

371 MgATP binding affinity but had little effect on MgADP binding (Parsonage et al., 1988a). Moreover, adenosine triphosphopyridoxal modifies in the E. coli (Ida et al., 1991). Another potential binding residue is which can become modified by if the analog binds in the presence of (Ida et al., 1991).

B. Potential Metal Ligands Dicyclohexylcarbodiimide (DCCD) reacts with a carboxy-residue ofthe subunit from mitochondria, bacteria and chloroplast to inactivate the enzyme and cause the loss of a nucleotide binding site (Pougeouis et al., 1979; Shoshan and Selman, 1980; Yoshida et al., 1981). The presence of interferes with inactivation by DCCD suggesting that the modified residue is involved with metal binding (Vallejos, 1981). However, other work shows that although the nucleotide is unable to bind after DCCD modification, the binding of metal is unaffected (Dagget et al., 1985). DCCD modifies in the subunit of the thermophile PS3 but modifies in E. coli (Yoshida et al., 1981; Esch et al., 1981). The appears to be the more essential residue in E. coli since the mutation decreased MgATP binding by 1000-fold and catalytic activity by 100fold while the mutation only reduced activity by 6-fold (Parsonage et al., 1988b). However, mutations of either of these carboxyl residue in the thermophilic PS3 subunit abolished the ability to reconstitute activity upon mixing with and subunits (Ohtsubo et al., 1987). Mutations of or of or C in the E. coli subunit severely impair catalytic activity (Takeyama et al., 1990; Iwamoto et al., 1991). Since these mutations affect metal specificity, these hydroxyl-containing residues have been postulated to be essential for metal binding. It is ofnote that the ligands to the metal of the metal-nucleotide binding site of cH-ras p21 are a serine and a threonine (Pai et al., 1989).

C. Potential Adenosine Ring Binding Sites The adenine of the nucleotide binds to the subunit near positions 287–296 and 328–332 as determined by photoaffinity labeling (Hollemans et al., 1983; Garin et al., 1986; Cross et al., 1987). As shown in

372 Fig. 7, conserved tyrosines and may bind the adenosine via interactions. In , 5'-pfluorosulfonylbenzoylinosine (FSBI) covalently modifies ,whileFSB-adenosine labels which is now thought to lie outside the catalytic site (Bullough and Allison, 1986; Lunardi et al., 1987; Cross et al., 1987). This is consistent with the observation that the catalytic sites can bind a variety of nucleotides while the noncatalytic sites show strong specificity for adenine nucleotides. The nucleotide analogs and have also been used to fill catalytic and noncatalytic sites which are distinguished by the ability of the noncatalytic sites to retain the analog after several catalytic cycles. Covalent incorporation of these analogs has identified the at the catalytic site in the from bovine mitochondria (Garin et al., 1986), spinach chloroplasts (Xue et al., 1987) and E. coli (Wise et al., 1987). Results obtained using the photoaffinity analog led Zhou et al. (1992) to propose that only binds the adenosine ring of the nucleotide at the catalytic site.

VI. Location of the Noncatalytic MetalNucleotide Binding Site The noncatalytic site was suggested to straddle the and subunits from the observation that the both subunits can become covalently modified when the photoaffinity analog is bound specifically to the N2 site (Bruist and Hammes, 1981). Modification of the noncatalytic site with 2has identified as the adenosine binding region in the enzyme from bovine mitochondria (Garin et al., 1986), spinach chloroplasts (Xue et al., 1987) and E. coli (Wise et al., 1987). This site specificity was also confirmed when the analogs were incorporated into the noncatalytic (N2) and catalytic (N3) sites of using the methods of Bruist and Hammes (1981). Zhou et al. (1992) have proposed that the adenosine ring of the nucleotide at the noncatalytic site is bound to and The proximity of the latter tyrosine with strongly suggests that the adenosine rings of the catalytic and noncatalytic sites are proximal while the phosphate groups of the two nucleotides are distal. This is supported by measurements using an EPR spin labeled photoaffinity ATP analog which estimated the spin labels

Wayne D. Frasch attached to the ribose ring to be 15 Å apart (Vogel et al., 1992). Evidence indicates that the noncatalytic site is primarily associated with the subunit. The subunit has relatively high sequence homology to the subunit, including the GXXXXGKT consensus sequence such that presumably binds a nucleotide phosphate. Since evolution often conserves the tertiary structure of proteins, the homology of the and subunits suggest that the nucleotide binding siteofthe subunit has a structure similar to the subunit. In order to allow the adenosine rings of the catalytic and noncatalytic nucleotides to face each other, the subunit is probably rotated about 180° relative to the subunit which is consistent with the most recent crystal structure of the rat mitochrondrial (M. Bianchet, M. Amzel and P. Pedersen, personal communication). In the subunit, the carboxyl group thought to coordinate themetal is 26 amino acids from the GXXXXGKT lysine. In the subunit, is also 26 amino acids from the analogous lysine. This lysine has been implicated in binding nucleotide phosphates from studies that show covalent modification of by when nucleotide binds to in the absence of metal (Ida et al., 1991). Ida et al., (1991) did not specify which nucleotide binding site was filled in this experiment, but proposed that it was a catalytic site. This supports the suggestion that there is overlap between the metal binding sites of the catalytic site of one heterodimer and the noncatalytic site of an adjacent heterodimer. Such an association was originally proposed from the observation that the inhibitor diadenosine tetraphosphate, which is able to span the two nucleotide binding sites of adenylate kinase, is an effective competitive inhibitor of the (Vogel and Cross, 1991). Onthe subunit, there is also a sequence homology to adenylate kinase but not other ATP binding enzymes, known as the Walker B consensus region. This sequence of LL-F-D (238–242) is close to the AMP (or second nucleotide) binding site ofadenylate kinase. A weaker homology can also be found on the subunit. The role ofthis region of these proteins in binding catalytic or noncatalytic nucleotides is, at present, undetermined. It does leave open the remote possibility, however, that as many as twelve nucleotides may be able to bind to the ATPase even though no more than six have ever been observed.

Chapter 11 F-TypeATPase VII. Model of the Metal-Nucleotide Binding Sites of The crystal structure of rat mitrochrondrial has now been resolved to the point that the distance and orientation of the P-loop regions are defined. These results are in general agreement with the distances determined by fluorescence resonance energy transfer (FRET) measurements (Shapiro and McCarty, 1991) with the exception of the N1-N4 distance. The N2 site has been postulated to be noncatalytic and is slightly closer to the catalytic N3 than the catalytic N1 site. The position ofN2 is also consistent with the relative location of the subunit P-loops to two adjacent subunit P-loops. These consistencies suggest that N2 and N3 are nucleotide binding sites on the same heterodimer.

VIII. Regulation of Catalytic Activity Through the synthesis ofATP, the serves as the throttle for the energy supply to all living cells. As would be expected from an enzyme with such a role, the catalytic activity is highly regulated and capable of interconversion between slow and rapid rates ofcatalytic turnover. Presumably, this increases the supply of ATP during increased energy demand and decreases ATP production to conserve energy resources. The from all sources share some, but not all, ofthese regulatory mechanisms. For example, the ATPase inhibitor protein is unique to the mitochondrial (Schwerzmann and Pedersen, 1986). The extent of regulation is less apparent in the E. coli enzyme and most dramatic in the chloroplast ATPase. In the latter case, the interconverts between the active state and a latent state that has almost no activity. Since the from all sources efficiently catalyze ATP hydrolysis, the chloroplast ATPase probably requires the increased regulation to prevent hydrolysis of ATP during long periods of darkness when the photosynthetic light reactions are unable to operate. When associated with the membrane, activation of the enzyme is dependent on the generation of a for both synthesis and hydrolysis activity. In chloroplasts, can be induced in isolated thylakoids by light-driven electron transfer reactions, acid-base transition or a valinomycin-induced diffusion potential. The threshold level of

373 required for activation is dependent on the state of reduction of a disulfide bond in the enzyme (Nalin and McCarty, 1984). Reduction of this bond is mediated in vivo by thioredoxin which obtains reducing equivalents from the reducing side of Photosystem I (Shahak, 1982; Mills and Mitchell, 1982). To achieve 50 % activation, the oxidized and reduced forms of the ATPase require a of 19 and , respectively (Jünesch and Gräber, 1987). Since activation of the oxidized form of the enzyme requires a higher than the enzyme needs to catalyze ATP synthesis, the hydrolysis of ATP rarely occurs. Several other treatments can induce the activation of hydrolysis activity of the chloroplast ATPase in vitro. These include sulfhydryl reducing agents such as dithiothreitol to reduce the disulfide bond as well as trypsin, heat, alcohols, anions such as bicarbonate and sulfite, organic acids like maleate and detergents such as octyl glucoside. Characterization ofthe means by which these treatments activate the enzyme have revealed that the enzyme contains multiple layers of interrelated regulatory mechanisms.

A. Regulation by the and subunits Dissociation of the subunit has been shown to activate the ATPase activity ofpurified . When removed in a manner that does not destroy the protein, reconstitution of the subunit with inhibits ATPase activity of the chloroplast (Richter et al., 1984) and E. coli enzymes (Klionski et al., 1984). Activation by heat, alcohols and detergents have all been attributed to the dissociation of the subunit induced by these treatments (Richter et al., 1984; Patrie and McCarty, 1984; Yu and McCarty, 1985). The disulfide bond that is reduced upon activation of by dithiothreitol (or thioredoxin in is on the subunit Nalin and McCarty, 1984). Of the four sulfhydryls in this protein, C199 (S1) and C205 (S2) comprise the disulfide responsible for activation (Moroney et al., 1984). These cysteines lie between the S3 and S4 sulfhydryls which are located near the N and C-termini, respectively. In the S4 is easily modified by N-ethyl maleimide (NEM) and is therefore thought to be relatively exposed to solvent. However, S3 only becomes susceptible to NEM modification upon illumination of thylakoids, suggesting a possible rearrangement of the subunit upon light-induced activation.

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Activation of the ATPase of oxidized by trypsin is now known to be the result of cleavage of the subunit at a specific site (Schumann et al., 1985). The subunit, as it is bound to the enzyme, contains three sites that are susceptible to trypsin cleavage numbered in order of decreasing susceptibility to proteolysis. Site I yields 27 kDa and 7 kDa fragments with the latter, carboxyl-terminal fragment containing only the S4 sulfhydryl. In both and purified the subunit is resistant to attack at any site by trypsin until the subunit is removed by methanol or heat treatment. In the absence of the subunit, there is rapid proteolysis at Site I. In thylakoids, trypsin sensitivity of oxidized is increased in a similar manner by illumination of the thylakoids. Thus,the formedbyillumination induces a conformation of that causes the disulfide to become accessible to trypsin digestion. These changes also influence the ease of disulfide reduction. These functional interactions between the and subunits in regulating the ATPase activity are complemented by the apparent physical proximity of these subunits on the enzyme as shown by electron microscopy and fluorescence resonance energy transfer measurements (Figs. 2 and 6). Reduction of the disulfide in the subunit increases the susceptibility to cleavage at Site II (Schumann et al., 1985). This site causes the release of a 1.3 kDa peptide from the C-terminus ofthe 27 kDa fragment that contains the S2 cysteine residue ofthe disulfide. The 25 kDa fragment of the subunit that remains is then slowly digested at Site III into a 14 kDa (S3containing) and an 11 kDa (S1 -containing) fragment. Activation of the latent ATPase of occurs, then, via a conformational change that shifts the subunit (or dissociates this subunit completely) to expose the subunit to the solvent. Activation is also achieved by reduction of the disulfide bond on the subunit. Disulfide reduction lowers the required for complete activation. Conversely, the disulfide is more easily reduced if the has changed the conformation of and subunits or if the subunit is removed altogether. Trypsin cleavage allows the and subunits to form the active conformation and, if the disulfide is reduced, can remove the S2 cysteine-containing peptide thereby preventing the reformation of the disulfide. As with other aspects of the from cyanobacteria, the subunit appears to represent an intermediate evolutionary step between bacteria and

Wayne D. Frasch

chloroplasts. From sequence comparisons, the chloroplast protein contains a highly conserved region that spans residues 195–233 which is largely missing in bacteria and mitochondria (Cozens and Walker, 1987; McCaryetal., 1988; Werner et al., 1990). The S1 (C199) and S2 (C205) cysteines that comprise the disulfide and the trypsin proteolitic Sites I (R215) and II (K204) occur within this region. Although the subunits from several cyanobacteria including Synechococcus sp. strain PCC 6716 contains a portion of this sequence (residues 205–233) in a highly conserved manner to the chloroplast protein, the sequence from 197–205 is missing. Thus, the cyanobacterial subunit is missing both sulfhydryls required for disulfide bond formation andthe trypsin cleavage Site II. The S4 cysteine is also unique to the chloroplast subunit. It was not a surprise to find, then, that the ATPase activity of the cyanobacterial enzyme is not activated by dithiothreitol (Bakels et al., 1991). Similar results have been observed with the isolated from Mastigocladus laminosus (Binder and Bachofen, 1979). This also explains the observation that the in Nostoc sp. strain MAC (PCC 8009) appears to be fully activated in vivo in cells adapted to either light or dark at physiological temperatures (Austin et al., 1992). Despite the absence ofthe disulfide and the trypsin cleavage Site II in the subunit, the from cyanobacteria does show latent behavior (Bakels et al., 1991). Trypsin treatment increased the ATPase activity in the cyanobacterial ATPase by more than 5-fold while the spinach enzyme treated comparably only showed a 1.5-fold increase. This suggests that the trypsin Site I is primarily responsible for activation. It is noteworthy that the ATPase of Spirulina platensis is activated by dithiothreitol (Hicks and Yocum, 1986a,b). Unfortunately, the sequence ofthe subunit ofthis species has not been determined. Heat or methanol treatments were also ineffective in activating the Synechococcus sp. strain PCC 6716 enzyme and, like the thiol-reagent treatment, had a slightly inhibitory effect on activity (Bakels et al., 1991). The relative ineffectiveness of these treatments and the increased accessibility of trypsin to the cyanobacterial ATPase also suggest that the esubunit is less able to restrict access to the trypsin site (and inhibit the ATPase) than its chloroplast counterpart. This different conformation of and subunits may explain why the cyanobacterial ATPase always

Chapter 11 F-TypeATPase appears to be partially activated. The ATP synthase from Synechococcus sp. strain PCC 6716 can also be activated by a inasimilar manner to the chloroplast enzyme (Bakels et al., 1991). However, the threshold value of from this ATP synthase is which is significantly lower than the measuredforthereduced form of the chloroplast enzyme. This is consistent with the lack ofa disulfide bond in the cyanobacterial enzyme such that this ATPase is always in the ‘reduced’ form. The ATP synthase from Rhodospirillum rubrum, that also lacks the disulfide in the subunit (Falk et al., 1985), has been determined to have a threshold similar to Synechoccus sp. strain PCC 6716 (Slooten and Vandenbranden, 1989). According to Jünesch and Gräber (1987), such a low threshold for the ATP synthase should allow significant rates of ATPase activity upon transition from the light to the dark. However, the cellular ATP level of Synechoccus sp. strain PCC 6716 in the dark is about 75 % of the light level when grown aerobically (Lubberding and Schroten, 1984). These high levels of ATP are thought to be maintained by oxidative phosphorylation in the dark. If correct, this would eliminate the need for much ofthe regulation of the ATPase in cyanobacteria required by the chloroplast enzyme. The Synechococcus sp. strain PCC 6716 ATPase is also stimulated significantly by glucopyranoside (also known as octyl glucoside or OGP) or sodium sulfite Bakels et al., 1991). The OGP-induced activity in chloroplast has been attributed to the solubilization of the subunit from the enzyme (Yu and McCarty, 1985). This conclusion is consistent with the observation that the enzyme only needs to be incubated in OGP prior to, but not during, the assay to exhibit ATPase activity. However, higher rates of ATPase activity can be achieved if OGP is also present during the assay (Pick and Bassilian, 1981). This latter effect does not appear to involve the or subunits but is more directly involved with the catalytic process.

B. Regulation via Noncatalytic Nucleotide Binding Sites Activating anions and organic acids appear to share the ability of OGP to induce ATPase activity via a process that does not involve the or subunits. Sulfite is unable to replace the initial required for activation of ATP hydrolysis nor does it affect

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the ease of reduction of the disulfide bond on the subunit by thiol reducing agents Larsen and Jagendorf, 1989a). However, after the initial activation, sulfite can substitute for the needed to maintain the ATPase activity. In the presence of this activator, rates of ATP hydrolysis as high as 3 mmol mg havebeen achieved which is about five-fold faster than observed in the presence of a The conversion of from the latent to the active form has been correlated with a decrease in affinity of one tightly bound ADP that induces the exchangeability or loss of nucleotides at this site (Larsen and Jagendorf, 1989b). Reversion to the latent state is also accelerated after illumination by the addition of ADP. Based on these observations it has been proposed that this ADP binds tightly to a site that regulates the rate ofATPase at the catalytic site. Alternatively, the tightly bound ADP in the latent enzyme has been proposed to bind to a catalytic site (Milgrom and Boyer, 1990). Latent does contain an ADP bound at a catalytic site too tightly to be removed without irreversibly denaturing the enzyme. Consistent with the binding-change mechanism, a is required to remove this tight binding ADP from its catalytic site to allow ATP hydrolysis to occur at the next catalytic site. Once activated, the rate of ATP hydrolysis is greatest when the concentrations of and ATP are set to maximize the amount of metal-nucleotide complex and minimize the concentrations of uncomplexed metal (Carmeli and Lifshitz, 1972). The uncomplexed metal will inhibit catalysis as will the presence of significant concentrations of uncomplexed ADP (Nelson et al., 1972). The inhibition by ADP may be the result ofsimple product inhibition in which the presence of high concentrations ofADP interfere with the binding ofATP at the catalytic sites. This may also be the case for the metal, if there is a compulsory metal release step as part of the catalytic cycle. However, it is also possible that this inhibition results from the binding of or ADP to the putative regulatory site. The chloroplast enzyme binds free to two sites in a cooperative manner with high affinity Haddy et al., 1985; Hochman and Carmeli, 1985). Recent experiments indicate that these metal binding sites are noncatalytic sites (Roskelley et al., 1992). When the ATPase activity is examined over the course of the first minute at low free concentrations, an initial burst ofactivity is observed

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in the first 20 seconds which gradually converts to a slower steady-state rate that continues for several minutes. Both of these rates are inhibited by increasing concentrations of free which results from the binding of the metal to the high affinity noncatalytic binding sites. Bicarbonate, which activates ATPase activity in a manner similar to sulfite, only increases the rate of the free -inhibited form of the enzyme (Guerrero etal., 1990). Kinetic analysis indicates a competitive relationship between sulfite activation and inhibition by ADP (Larsen and Jagendorf, 1989b). Sulfite also stimulates the release/exchange of the tightly bound ADP from the putative regulatory site. However, the rate of ADP release induced by sulfite was three orders of magnitude lower than the rate of ATP hydrolysis. This observation appears to be more difficult to reconcile with the ADP originating from an obligatory alternating catalytic site than at a regulatory site. However, activation by sulfite is not totally independent from regulation by the subunit in This is based on the observation that modification of the reduced disulfide by NEM not only results in permanent activation of ATPase but also abolishes the stimulation of the ATPase by sulfite (Umbach et al., 1990). Thus, the regulation of catalysis originating with the and subunits may be mediated via the tightly bound ADP. In the the state of metal and nucleotide bound to the noncatalytic sites also influences the rate of ATP hydrolysis (Jault and Allison, 1993). Anions like sulfite also activate the ATPase activity of and thus regulation at this level is common to most ATPases. The rate of ATPase activity in has three phases. Upon addition of ATP there is an initial burst of ADP produced. The rate of ATPase then slows for several seconds (intermediate phase) but then gradually accelerates to its final steady-state rate. The initial burst of ADP presumably originates from the ATP-induced release of catalytic site-bound ADP formed at the end of the previous catalytic event. When the noncatalytic sites of nucleotide-depleted are covalently modified with FSBA prior to the assay, the ATPase rate never accelerates from the intermediate phase but remains slow (Jault and Allison, 1993). Conversely, when the noncatalytic sites of nucleotide-depleted are filled with MgATP or Mg-pyrophosphate prior to the assay, the slower (intermediate) phase is eliminated. These

results suggest that the binding of a metal-nucleotide to the noncatalytic site activates the ATPase activity. Presumably, the binding of ATP to the noncatalytic sites promotes the release of tightly bound ADP from the catalytic sites. However, these apparent regulatory aspects of the noncatalytic sites of the purified can also be explained if the metal and nucleotide at each noncatalytic site serves as a proton pump for the catalytic site in the same heterodimer(Houseman et al., 1994b). The conformational changes induced by the ATPase reaction would then be expected to be more efficient when metal and nucleotide are bound to the noncatalytic sites even if no is generated as a result. The from cyanobacteria appears to be more closely related to the chloroplast enzyme than to that of mitochondrial origin despite the presence of both photosynthetic and oxidative phosphorylation electron transfer complexes in these cells. However, even for the limited number of studies performed thus far, it is evident that the ATP synthases from some cyanobacteria have more chloroplast-like traits while others have more characteristics of a bacterial enzyme. Further characterization of the cyanobacterial enzyme should provide considerable insight into the evolutionary consequences that led to the divergence in the regulatory mechanisms of this essential enzyme. Acknowledgments This is publication number 188 of the Center for the Study of Early Events in Photosynthesis. The Center is supported by US Department of Energy grant DEFG02-88ER13969. References Abrahams JP, Lutter R, Todd RJ, van Raaij MJ, Leslie AGW and Walker JE (1993) Inherent asymmetry of the structure of ATPase from bovine heart mitochondria at 6.5 Å resolution. EMBOJ 12: 1775–1780 Akey C, Crepeau R, Dunn S, McCarty R and Edelstein S (1983) Electron microscopy of single molecule molecular averaging of subunit-deficient from E. coli and spinach chloroplasts. EMBO J 2: 1409–1415 Andrews W, Hill F. and Allison W (1984a) Identification of the essential tyrosine residue in the subunit of bovine heart mitochondrial that is modified by 7-chloro-4benzofurazan. J Biol Chem 259: 8219–8225

Chapter 11 F-TypeATPase Andrews W, Hill F and Allison W (1984b) Identification of the lysine residue to which the 4-citrobenzofurazan group migrates after the bovine mitochondrial is inactivated with 7-chloro-4-nitro benzofurazan. J Biol Chem 259: 14378– 14382 Austin P, Ross IS and Mills JD (1992) Light/dark regulation of photosynthetic enzymes within intact cells of the cyanobacterium Nostoc sp. Mac. Biochim Biophys Acta 1099: 226– 232 Avetisyan A, Bogachev A, Murtasina R and Shulachev V (1993) ATP-driven transport and ATP synthesis in Escherichia coli grown at low FEBS Lett 317: 267– 270. Bakels RHA, van Walraven HS, Scholts MJC, Krab K and Kraayenhof R (1991) Activation ofthe synthases of a thermophilic cyanobacterium and chloroplasts—a comparative study. Biochim Biophys Acta 1058: 225–234 Bianchet M, Ysern X, Hullihen J, Pedersen PL and Amzel LM (1991) Mitochondrial ATP synthase. J Biol Chem 266:21197– 21201 Binder A and Bachofen R (1979) Isolation and characterization of a coupling factor I ATPase of the thermophilic blue-green alga (cyanobacterium) Mastigocladus laminosus. FEBS Lett 104: 66–70 Boekema E, van Heel M and Gräber P (1988) Structure of the ATP synthase from chloroplasts studied by electron microscopy and image processing. Biochim Biophys Acta 933: 365–371 Boekema E, Xiao J and McCarty RE (1990) Structure ofthe ATP synthase from chloroplasts studied by electron microscopy. Localization of the small subunits. Biochim Biophys Acta 1020:49–56 Bossard M, Vik T and Schuster SM (1980) Beef heart mitochondrial adenosine triphosphatase—catalyzed formation ofa transition state analog in ATP synthesis. J Biol Chem 255: 5342–5346 Boyer PD (1993) The binding change mechanism for ATP synthase— some probabilities and possibilities. Biochim Biophys Acta 1140: 215–250 Bruist M and Hammes GG (1981) Further characterization of nucleotide binding sites on chloroplasts coupling factor one. Biochemistry 20: 6298–6305 Bullough D and Allison W (1986) Inactivation of the bovine heart mitochondrial by 5'-r-fluorosulfonylbenzoyl inosine is accompanied by modification of tyrosine345 in a single subunit. J Biol Chem 261: 14171– 14177 Butlin JD, Cox GB and Gibson F. (1971) Oxidative phosphorylation in Escherichia coli K-12. Mutations affecting magnesium ion or calcium stimulated ATP. Biochem J 124: 75–81 Carmeli C and Lifshitz Y (1972) Effects of orthophosphate and ADP on ATPase activity in chloroplasts. Biochim Biophys Acta 267: 86–95 Carmeli C, Huang J, Mills D, Jagendorf AT and Lewis A (1986) Extended X-ray absorption fine structure of and ATP complex bound to coupling factor 1 of the H+-ATPase from chloroplasts. J Biol Chem 261: 16969–16975 Carmeli C, Lewis A and Jagendorf AT (1989) EXAFS analysis ofthe structure of Mn-nucleotide bound to latent and activated In: Baltscheffsky M (ed) Current Research in Photosynthesis, Vol. III, pp 29–32. Kluwer, Dordrecht

377 Chuan H and Wang J (1988) 3'-o-(5-fluoro-2,4-dinitrophenyl )ADP ether and ATP ether. Affinity reagents for labeling ATPases. J Biol Chem 263: 13003–13006 Cozens AL and Walker JE (1987)The organization and sequence of the genes for ATP synthase subunits in the cyanobacterium Synechococcus 6301. J Mol Biol 194: 359–383 Cross RL and Nalin CM (1982) Adenine nucleotide binding sites on beef heart J Biol Chem 257: 2874–2881 Cross RL, Cunningham D, Miller C, Xue Z, Zhou J-M and Boyer P (1987) Adenine nucleotide binding sites of beef heart ATPase: photoaffinity labeling of Tyr-368 at a noncatalytic site and Tyr 245 at a catalytic site. Proc Natl Acad Sci USA 84: 5715–5719 Cunningham D and Cross RL (1988) Catalytic site occupancy during ATP hydrolysis by J Biol Chem 263: 18850–18856 Curtis S (1987) Genes encoding the beta and epsilon subunits of the proton-translocating ATPase from Anabaena sp. PCC 7120. J Bacterioi 169: 80–86 Dagget SG, Gruys KJ and Schuster SM (1985) Metal interactions with beef heart mitochondrial ATPase. J Biol Chem 260: 6213–6218 Devlin C and Grisham C (1990) and nuclear magnetic resonance and kinetic studies of the active site structure of chloroplast ATP synthase. Biochemistry 29: 6192–6203 Duncan T and Senior AE (1985) The defective proton-ATPase of uncD mutants ofEscherichia coli. Two mutations which affect the catalytic mechanism. J Biol Chem 260: 4901–4907 Duncan T, Parsonage D and Senior AE (1986) Structure of the nucleotide-binding domain in the subunit ofEscherichia coli . FEBS Lett 208: 1–6 Esch F, Bohlen P, Otsuka A, Yoshida M and Allison WS (1981) Inactivation of the bovine mitochondrial with Dicyclohexyl carbodiimide leads to the modification of a specific glutamic acid residue in the subunit. J Biol Chem 256: 9084–9089 Falk G, Hampe A and Walker JE (1985) Nucleotide sequence of the Rhodospirillum rubrum ATP operon. Biochem J. 228, 391–407 Fillingame RH (1990) Molecular mechanics of ATP synthesis by type ATP synthases. The Bacteria XII: 345–391 Foster D and Fillingame RH (1982) Stoichiometry ofsubunits in the complex of Escherichia coli. J Biol Chem 257: 2009–2015 Frasch WD (1981) Stereochemistry and substrate specificity of coupling factor 1. In: Selman BR and Selman-Reimer S (eds) Energy Coupling in Photosynthesis, pp 157–168. Elsevier, Amsterdam Frasch WD and Selman BR (1982) Mechanism of phosphorylation catalyzed by chloroplast coupling factor 1: stereochemistry. Biochemistry 21: 3636–3643 Frasch WD and Sharp RR (1985) Nucleotide binding to chloroplast ATP synthase: effect on the proton spin-echo NMR spectrum. Biochemistry 24: 5454–5458 Fry DC, Kuby S and Mildvan AS (1985) NMR studies of the MgATP binding site of adenylate kinase and of a 45-residue peptide fragment of the enzyme. Biochemistry 24:4680–4694 Fry DC, Kuby S and Mildvan AS (1986) ATP-binding site of adenylate kinase: mechanistic implications of its homology with ras-ecoded p21, , and other nucleotide-binding

378 proteins. Proc Natl Acad Sci USA 83: 907–911 Futai M and Kanazawa H (1983) Structure and function of proton-translocating adenosine triphosphatase biochemical and molecular biological approaches. Microbiol Rev 47: 285–312 Futai M, Noumi T and Maeda M (1989) ATP synthase ( ATPase): results by combined biochemical and molecular biological approaches. Annu Rev Biochem 58: 111–136 Garboczi D, Shenbagamurthi P, Kirk W, Hullihen J and Pedersen PL (1988) Mitochondrial ATP synthase. Interaction of a synthetic 50-amino acid, peptide with ATP. J Biol Chem 263: 812–816 Garin J Boulay F, Issartel J-P and Vignais P (1986) Identification of amino acid residues photolabeled with 2-azido adenosine diphosphate in the subunit of beef heart mitochondrial Biochemistry 25: 4431–4437 Garin J, Vignais P, Gronenborn A, Clore G, Gao Z and Baeuerlein E (1988) studies on nucleotide binding to the catalytic sites of bovine mitochondrial FEBS Lett 242: 178–182 Girault G, Berger G, Galmiche, J-M, and Andre F-A (1988) Characterization of six nucleotide-binding sites on chloroplast coupling factor 1 and one site on its purified subunit. J Biol Chem 263: 14690–14695 Gresser M, Myers J and Boyer PD (1982) Catalytic site cooperativity of beef heart mitochondrial adenosine triphosphatase. J Biol Chem 257: 12030–12038 Grubmeyer C, Cross RL and Penefsky HS (1982) Mechanism of ATP hydrolysis by beef heart mitochondrial ATPase. Rate constants for elementary steps in catalytic at a single site. J Biol Chem 257: 12092–12100 Guerrero K, Xue Z and Boyer PD (1990) Active/ inactive state transitions of the chloroplast are induced by a slow binding and release of J Biol Chem 265:16280–16287 Haddy A, Frasch WD and Sharp RR (1985) Cooperative binding of manganese(II) to chloroplast coupling factor 1 detected by NMR proton relaxation enhancement. Biochemistry 24:7926– 7930 Haddy A, Frasch WD and Sharp RR (1989) Properties of the manganese(II) binding site in ternary complexes of Mn-ADP and Mn-ATP with chloroplast coupling factor 1: magnetic field dependence of solvent and NMR relaxation rates. Biochemistry 28: 3664–3669 Henning J. and Herrmann RG (1986) Chloroplast ATP synthase of spinach contains nine non-identical subunit species, six of which are encoded by plastid chromosomes in two operons in a phylogenetically conserved arrangement. Mol Gen Genet 203: 117–128 Herrmann RG, WesthoffP, Alt J, Tittgen J and Nelson N (1985) In: Van Vlooten-Doting L, Groot G, and Halls T (eds) Molecular Form and Function ofthe Plant Genome, pp 233–256. Plenum, New York Hicks D and Yocum CF (1986a) Properties ofthe cyanobacterial coupling factor ATPase from Spirulina platensis. I. Electrophoretic characterization and reconstitution of photophosphorylation. Arch Biochem Biophys 245:220–229 Hicks D and Yocum, CF (1986b) Properties of the cyanobacterial coupling factor ATPase from Spirulina platensis, II. Activity of the purified and membrane-bound enzymes. Arch Biochem Biophys 245: 230–237 Hiller R and Carmeli C (1985) Cooperativity among manganese-

Wayne D. Frasch binding sites in the of chloroplasts. J Biol Chem 260: 1614–1617 Hiller R and Carmeli C (1990) Kinetic analysis of cooperative Interactions Induced by binding to the chloroplast ATPase. Biochemistry 29: 6186–6192 Hochman Y and Carmeli C (1981) Correlation between the kinetics of activation and inhibition of adenosine triphosphatase activity by divalent metal ions and the binding ofmanganese to chloroplast coupling factor 1. Biochemistry 20: 6287–6292 Hochman Y, Lanir A, Werber M and Carmeli C (1979) The effect of binding of cobalt(III)-nucleotide complexes on the kinetic properties of adenosine triphosphastase activity in coupling factor 1 from chloroplasts. Arch Biochem Biophys 192: 138–147 Hollemans M, Runswick M, Fearnley, I and Walker JE (1983) The sites of labeling of the subunit of bovine mitochondrial with 8-azido-ATP. J Biol Chem 258: 9307–9313 Houseman ALP, Morgan L, LoBrutto R and Frasch WD (1994a) Characterization of ligands of a high affinity metal-binding site in the latent chloroplast by EPR spectroscopy of bound Biochemistry 33: 4910–4917 Houseman ALP, LoBrutto R and Frasch WD (1994b) The coordination of nucleotides to the metals at the M2 and M3 metal-binding sites of the spinach chloroplast Biochemistry, in press Hsu S-Y, Nuomi T, Takeyam M, Maeda M, Ishibashi S and Futai M (1987) of Escherichia coli An amino acid replacement within a construed sequence(G-X-X-X-XG-K-T/S) of nucleotide-binding proteins. FEBS Lett 218: 222–226 Ida K, Noumi T, Maeda M, Fukui T and Futai M (1991) Catalytic site of of Escherichia coli. J Biol Chem 266:5424– 5429 Iwamoto A, Omote H, Hanada H, Tomioka N, Itai A, Maeda M and Futai, M. (1991) Mutations in and the glycine-rich sequence and in the subunit of Escherichia coli J Biol Chem 266:16350–16355 Jagendorf AT and Nathan GE (1985) In: Steinback KE, Bonitz S, Arntzen CJ and Bogorad L (eds) Molecular Biology of the Photosynthetic Apparatus, pp 121–140. Cold Spring Harbor Laboratory Press, Cold Spring Harbor Jault J-M and Allison W S (1993) Slow binding of ATP to noncatalytic nucleotide binding sites which accelerates catalysis is responsible for apparent negative cooperativity exhibited by the bovine mitochondrial J Biol Chem 268:1558– 1566 Jünesch U and Gräber P (1987) Influence of the redox state and the activation of the chloroplast ATP synthase on protontransport-coupled ATP synthesis-hydrolysis. Biochim Biophys Acta 893: 275–288 Kironde FAS and Cross RL (1986) Adenine nucleotide-binding sites ofbeef heart J Biol Chem 261:12544–12549 Klionski D, Brusilow W and Simoni R (1984) In vivo evidence for the role of the subunit as an inhibitor of the protontranslocating ATPase of Escherichia coli. J Bacteriol 160: 1055–1060 Kohlbrenner WE and Boyer PD (1983) Probes of catalytic site cooperativity during catalysis by the chloroplast adenosine triphosphatase and the adenosine triphosphate synthase. J Biol Chem 258: 10881–10886 Kothen G. Schwartz O and Strotmann H (1992) Enzyme kinetic

Chapter 11 F-TypeATPase studies on chloroplast by a clamptechnique. In: Murata N (ed) Research in Photosynthesis, Vol II, pp 661– 668. Kluwer, Dordrecht Krab K, Bakels R, Scholts M and van Walraven H (1993) Activation of the synthase in thylakoid vesicles from the cyanobacterium Synechococcus 6716 by Includinga comparison with chloroplasts and introducing a new method to calibrate light-induced Biochim Biophys Acta 1141: 197–205 Krenn B, Koppenaal F., Van Walraven H, Krab K and Kraayenhof R (1993) Co-reconstitution of the synthase and cytochrome b-563/c-554 complex from a thermophilic cyanobacterium. High ATP yield and mutual effects on the enzymatic activities. Biochim Biophys Acta 114: 271–281 Labahn A and Gräber P (1992) Transport portons do not participate in ATP synthesis/hydrolysis at the nucleotide binding site of the from chloroplasts. FEBS Lett 313: 177–180 Larson E and Jagendorf AT (1989) Sulfite stimulation of chloroplast coupling factor ATPase. Biochim Biophys Acta 973: 67–77 Larson E, Umbach A and Jagendorf, AT (1989) Sulfite-stimulated release of ADP bound to chloroplast thylakoid ATPase. Biochim Biophys Acta 973: 78–85 Laubinger W and Dimroth P (1988) Characterization ofthe ATP synthase of Propionigenium modestum as a primary sodium pump. Biochemistry 27: 7531–7537 Laubinger W, Deckers-Hebestreit G, Altendorf K and Dimroth P (1990) A hybrid adenosine triphosphate composed of of Escherichia coli and of Proprionigenium modestum is a functional sodium ion pump. Biochemistry 29: 5458–5463 Leckband D and Hammes GG (1987) Interactions between nucleotide binding sites on chloroplast coupling factor 1 during ATP hydrolysis. Biochemistry 26: 2306–2312 Lill H and Nelson N (1991) The atp1 and atp2 operons of the cyanobacterium Synechocystis sp. PCC 6803. Plant Mol Biol 17:641–652 Lubberding HJ and Schroten W (1984) The ATP level in the thermophilic cyanobacterium Synechococcus 6716 during lightdark transition and in the presence of some specific inhibitors. FEMS Microbiol Lett 22: 93–96 Lubberding HJ, Zimmer G, Van Walraven HS, Schrickx J and Kraayenhof R (1983)Isolation,purificationandcharacterization of the ATPase complex from the thermophilic cyanobacterium Synechococcus 6716. Eur J Biochem 137: 95–99 Lücken U, Gogol E and Capaldi R (1990) Structure of the ATP synthase complex of Escherichia coli from cryoelectron microscopy. Biochemistry 29: 5339–5343 Lunardi J, Garin J, Issartel J-P and Vignais P (1987) Mapping of nucleotide-depleted mitochondrial with 2-azidoadenosine diphosphate. Evidence for two nucleotide binding sites in the subunit. J Biol Chem 262:15172–15181 McCarn D, Whitaker R, Alam J, Vrba J and Curtis SE (1988) Genes encoding the alpha, gamma, delta, and four subunits of ATP synthase constitute an operon in the cyanobacterium Anabaena sp. strain PCC 7120. J Bacteriol 170: 3448–3458 Milgrom Y and Boyer PD (1990) The ADP that binds tightly to nucleotide-depleted mitochondrial and inhibits catalysis is bound at a catalytic site. Biochim Biophys Acta 1020:43–48 Mills JD and Mitchell P (1982) Modulation of coupling factor ATPase activity in chloroplasts. Reversal of thiol modulation

379 in the dark. Biochim Biophys Acta 679: 75–83 Moroney J, Fullmer C and McCarty RE (1984) Characterization of the cysteinyl-containing peptides of the subunit ofcoupling factor 1. J Biol Chem 259: 7281–7285 Murataliev M Milgrom Y and Boyer PD (1991) Characteristics of the combination of inhibitory and azide with the ATPase from chloroplasts. Biochemistry 30: 8305–8310 Nalin CM and McCarty RE (1984) Role ofa disulfide bond in the subunit inactivation of the ATPase of chloroplast coupling factor 1. J Biol Chem 259: 7275–7280 Nelson N (1992) Evolution of organellar proton-ATPases. Biochim Biophys Acta 1100: 109–124 Nelson N, Nelson H and Racker E (1972) Partial resolution ofthe enzymes catalyzing photophosphorylation. XI. Magnesioumadenosine triphosphatase properties ofheat-activated coupling factor 1 from chloroplasts. J Biol Chem 247, 6506–6510 Nitschmann WH and Peschek GA (1986) Oxidative Phosphorylation and energy buffering in cyanobacteria. J Bacteriol 168:1205–1211 Nuomi T, Kanazawa H and Futai M (1986) Replacement of arginine 246 by histidine in the subunit of Escherichia coli resulted in loss of multi-site ATPase activity. J Biol Chem 261:9196–9201 O’Neal C and Boyer PD (1984) Assessment of the rate of bound substrate intercon version and of ATP acceleration of product release during catalysis by mitochondrial adenosine triphosphatase. J Biol Chem 259: 5716–5767 Ohtsubo M, Yoshida M, Ohta S, Kagawa Y, Yohda M and Date T(1987)In vitro mutated subunits from the of the thermophilic bacterium, PS3, containing glutamine in place of glutamic acid in positions 190 or 201 assembles with the and subunits to produce inactive complexes. Biochem Biophys Res Comm 146:705–710 Pai E, Kabsch W, Krengel V, Holmes K, John J, and Wittinghofer A (1989) Structure ofthe guanine-nucleotide-binding domain of the Ha-ras oncogene product p21 in the triphosphate conformation. Nature 341: 209–214 Palmer JD (1983) Comparative organization of chloroplast genomes. Ann Rev Genet 19: 325–354 Parsonage D, Al-Shawi M and Senior AE (1988a) Directed mutagenesis of the strongly conserved lysine 155 in the catalytic nucleotide-binding domain of of from Escherichia coli. J Biol Chem 263: 4740–4744 Parsonage D, Duncan T, Wilke-Mounts S, Kironde F, Hatch L and Senior AE (1987a) The defective proton-ATPase of uncD mutants of Escherichia coli. Identification by DNA sequencing of residues in the which are essential for catalysis or normal assembly. J Biol Chem 262: 6301–6307 Parsonage D, Wilke-Mounts S and Senior, AE (1988b) E. coli site-directed mutagenesis of the FEBS Lett 232: 111–114 Parsonage D, Wilke-Mounts S and Senior AE (1987b) Directed mutagenesis of the of from Escherichia coli. J Biol Chem 262: 8022–8026 Patrie WJ and McCarty RE (1984) Specific binding of coupling factor 1 lacking the delta and epsilon subunits to thylakoids. J Biol Chem 259: 11121–11128 Pedersen PL and Amzel LM (1993) ATP synthases. J Biol Chem 268:9937–9940 Peschek GA, Hinterstoisser B, Riedler M, Muchl R and Nitschmann WH (1986) Exogenous energy supply to the

380 plasma membrane ofdark anaerobic cyanobacterium Anacystis nidulans: Thermodynamic and kinetic characterization of the ATP synthesis effected by an artificial proton motive force. Arch Biochem Biophys 247: 40–48 Pick U and Bassilian S (1981) Octyl glucoside stimulates a ATPase activity in chloroplast In: Energy Coupling in Photosynthesis, Selman BR and Selman-ReimerS (eds), pp 251–260. Elsevier, Amsterdam Pougeois R, Satre M and Vignais P (1979) Reactivity of mitochondrial to dicyclohexylcarbodiimide. Inactivation andbindingstudies. Biochemistry 18:1408–1413 Recipon H, Perasso R, Adoute A and Quetier F (1992) ATP synthase subunit c/III/9 gene sequences as a tool for interkingdom and metaphytes molecular phylogenies. J Mol Evol 34: 292–303 Richter M, Patrie W and McCarty RE (1984) Preparation of the subunit and subunit-deficient chloroplast coupling factor 1 in reconstitutively active forms. J Biol Chem 259:7371–7373 Roskelley A, LoBrutto R and Frasch W (1992) Characterization of the metal ligands at nucleotide binding sites of CF1. In: Murata N (ed) Research in Photosynthesis, Vol II, pp 745– 748. Kluwer, Dordrecht Schlimme E, deGroot E, Schott E, Strotmann H and Edelman K (1979) Photophosphorylation of base-modified nucleotide analogs by spinach chloroplasts. FEBS Lett 106: 251–256 Schmidt G, Rodgers A, Howitt S, Munn A, Hudson G, Holten T, Whitfield P, Bottomley W, Gibson F and Cox G (1990) The chloroplast CF0 I subunit can replace the b-subunit of the ATPase in a mutant strain of Escherichia coli K12. Biochim Biophys Acta 1015: 195–199 Schumann J, Richter M and McCarty RE (1985) Partial proteolysis as a probe of the conformation of the subunit in activated soluble and membrane-bound chloroplast coupling factor 1. J Biol Chem 260: 11817–11823 Schwerzmann K and Pedersen PL (1986) Regulation of the mitochondrial ATP synthase/ATPase complex. Arch Biochem Biophys 250: 1–18 Selman BR and Selman-Reimer S (1981) The steady state kinetics of photophosphorylation. J Biol Chem 256: 1722–1726 Senior AE (1988) ATP synthesis by oxidative phosphorylation. Physiol Rev 68: 1177–1231 Senter P, Eckstein F and Kagawa Y (1983) Substrate metaladenosine 5'-triphosphate chelate structure and stereochemical course of reaction catalyzed by adenosinetriphosphatase from the thermophilic bacterium PS3. Biochemistry 22:5514–5518 Shahak Y (1982) Activation and deactivation of proton ATPase in intact chloroplasts. Plant Physiol 70: 87–91 Shapiro A, Huber A and McCarty RE (1991a) Four tight nucleotide binding sites of chloroplast coupling factor 1. J Biol Chem 266:4194–4200 Shapiro A, Gibson K, Sheraga H and McCarty RE (1991b) Fluorescence resonance energy transfer mapping of the fourth of six nucleotide-binding sites of chloroplast coupling factor 1. J Biol Chem 266: 17276–17285 Shoshan V and Selman BR (1980) The relationship between light-induced adenine nucleotide exchange and ATPase activity in chloroplast thylakoid membranes. J Biol Chem 255: 384– 389 Slooten L, and Vandenbranden S (1989) ATP synthesis by proteoliposomes incorporating Rhodospirillum rubrum as measured with firefly luciferase: dependence of delta-psi

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Chapter 12 Soluble Electron Transfer Catalysts of Cyanobacteria Larry Z. Morand, R. Holland Cheng and David W. Krogmann Department of Biochemistry, Purdue University, West Lafayette, IN 47907-1153, USA

Kwok Ki Ho Department of Botany, National University of Singapore, Lower Kent Ridge Road, Singapore 0511, Republic of Singapore

Summary I. Ferredoxin A. Genes B. Crystal Structures C. Multiple Ferredoxins II. Flavodoxin A. Genes B. Protein Chemistry C. Physiological Response to Iron Deprivation Reductase (FNR) III. Ferredoxin A. Genes B. ProteinChemistry C. Interaction of Ferredoxin with FNR D. Enzyme Activity IV. Plastocyanin A. Genes B. Protein Chemistry C. Physiological Response to Copper Deprivation V. Cytochrome A. Genes B. Protein Chemistry C. Physiology of Cytochrome VI. Low-Potential Cytochrome c A. Protein Chemistry and Location in the Cell B. Physiological Role VII. Hydrogenase A. Genes B. Protein Chemistry C. Physiology of Hydrogen Metabolism D. A Speculative Model of Hydrogen Metabolism in Cyanobacteria References

D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 381–407. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

382 382 382 383 383 387 387 388 388 389 389 390 391 392 392 392 393 394 394 394 395 396 396 396 397 398 398 398 399 401 402

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L. Z. Morand, R. H. Cheng, K. K. Ho and D. W. Krogmann

Summary This review covers only a fraction of the area of one written a dozen years ago on photosynthesis in cyanobacteria (Ho and Krogmann, 1982) yet it cites many more references than that earlier work. The power of reductionist laboratory science has increased immensely in the intervening years. The soluble electron transfer catalysts of photosynthesis have received disproportionate attention since soluble proteins are more easily dealt with by the techniques of protein chemistry. Research on each of the catalysts reviewed here has exposed a variety of insights through the tools of contemporary science. The many studies of different forms of ferredoxin in cyanobacteria promise new understanding of the regulation of electron transfer and its mechanism. The crystal structures of ferredoxin, oxidoreductase, flavodoxin and plastocyanin are elegant examples ofwhat our broader understanding will become. The powerful technique ofgene deletion used on cytochrome (cytochrome has done more than confirm preconceptions. It has given us the intriguing puzzle of why more than two routes of electron flow between carriers of very similar redox potential may be used. The low potential cytochrome c beckons for an explanation of its catalytic function and for an understanding of its role in the ancient origin of other cytochromes. Finally, hydrogenase, whose catalytic act is the simplest—the movement ofan electron to or from a proton— seems ready for understanding. Hydrogenase has a long history of fragility and activity loss during purification. One type of hydrogenase has now been purified and there are glimpses of its metabolic role.

I. Ferredoxin The discoveries of ferredoxin and oxidoreductase (FNR) opened a large field ofstudies ofsoluble electron transfer catalysts which participate in photosynthesis and related processes. There are multiple forms of ferredoxin but one of these— ferredoxin I—seems to be the essential catalyst of electron transfer from membrane-bound, iron-sulfur centers in Photosystem I to FNR which, in turn, reduces for fixation. The ferredoxin I of cyanobacteria and all other oxygenic photosynthetic organisms is a strongly acidic protein of approximately one hundred amino acid residues with a [2Fe-2S] center that transfers one electron. The structure, function, and evolution of the ferredoxins has been reviewed (Rogers, 1987; Knaff and Hirasawa, 1991) but the literature on them continues to increase rapidly. Ferredoxin I in cyanobacteria and other oxygenic photosynthetic organisms has evolved from a single gene. The major ferredoxin in anoxygenic procaryotic photosynthetic organisms is a [4Fe-4S] protein with a rather different amino acid sequence. Abbreviations: FAD – flavin adenine dinucleotide; FMN – flavin mononucleotide; FNR – ferredoxin oxidoreductase; NMR – nuclear magnetic resonance; SDS – sodium dodecylsulfate.

A. Genes The petFI gene for ferredoxin I has been cloned from Synechococcus sp. strains PCC 7942 and PCC 7002 and several strains of Anabaena sp. (Reith et al., 1986; Van der Plas et al., 1986; Alam et al., 1986; Van der Plas et al., 1988; Leonhardt and Straus, 1992). All authors report that there is only a single copy of the gene and that the mRNA is monocistronic. There is no evidence that ferredoxin is transcribed with any other gene. Iron deprivation of cyanobacteria causes the replacement of most of the ferredoxin I with the non-iron containing flavoprotein, flavodoxin (see Chapter 25). Van der Plas et al. (1988) found that iron deficiency caused a 30-fold decrease in ferredoxin protein but only a two- to three-fold decrease in the mRNA for synthesis of this protein. Thus it is likely that iron deficiency accelerates the degradation of the ferredoxin apoprotein. Van der Plas et al. (1988) attempted to delete the petFI gene by replacing part of the coding sequence with a kanamycin resistance marker. Transformation was attempted in the presence and absence of iron in which condition flavodoxin might be expected to replace ferredoxin. No deletion mutants were recovered indicating that cells lacking the petFI gene are not viable. The further implication is that flavodoxin may replace most but cannot replace all of the functions of ferredoxin in the cell.

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B. Crystal Structures

C. Multiple Ferredoxins

Hall et al. (1971) noticed the extraordinary stability of ferredoxin from the cyanobacterium Spirulina maxima. The enzyme from higher plants and eucaryotic algae is quite unstable at room temperature and seems to suffer decomposition ofthe Fe-S centers when stored at -20 °C. Ferredoxins I from many genera of cyanobacteria are stable for years at -20 °C and suffer no damage for weeks at room temperature if no proteases or bacteria are present (D. W. Krogmann, unpublished results). Coupled with their ease of purification, the stability of cyanobacterial ferredoxins makes them ideal proteins for physical analysis, Markley and coworkers have undertaken an NMR spectroscopic analysis of Anabaena sp. strain PCC 7120 ferredoxin which has already identified resonances of many of the individual atoms in the molecule (Skjeldal et al., 1990; Oh and Markley, 1990). This work promises to give exquisite details of the structure, changes during catalysis and the molecular interactions of ferredoxin with its reaction partners. Another extraordinary advantage of the cyanobacterial ferredoxins is their ease of crystallization. Mitsui and Arnon (1971) first described the crystallization of a ferredoxin from Nostoc sp. In 1981, Tsukihara et al. published a crystal structure of the ferredoxin from Spirulina platensis at 2.5 Å resolution. In 1990, Tsukihara et al. solved the structure of ferredoxin I from Aphanothece sacrum at 2.2 Å resolution. Both showed a central barrel with minor helical regions. The crystals from A. sacrum were of better quality and the resulting structure suggested that the surface around Tyr 23, Tyr 80 (Tyr 25 and Tyr 82 in the Anabaena sp. strain PCC 7120 ferredoxin; see Fig. 1) and the iron-sulfur center might interact with FNR. Ryniewski et al. in 1991 reported a structure from crystals of ferredoxin I from Anabaena sp. strain PCC 7120 which is similar in overall fold to the ferredoxins of S. platensis and A. sacrum (see Fig. 1). Böhme and Haselkorn (1989) have achieved expression of genes encoding ferredoxins from Anabaena sp. strain PCC 7120 in Escherichia coli; hence, the stage is well set for informed site-directed mutagenesis of the ferredoxin molecule guided by a high-resolution threedimensional structure.

Ferredoxin I receives electrons from PS I and the details of that process are covered in Chapter 10 of this volume. Reduced ferredoxin I must then pass electrons on to FNR. This process will be discussed in a later section of this chapter (see Section III). In a more general vein, one may wonder why ferredoxin I is so loosely held on the thylakoid surface. Its electron donor, an Fe-S center on the psaC gene product, is firmly fixed, and it now appears that ferredoxin I’s electron acceptor, FNR, may be anchored as well. Since ferredoxin I may serve other electron acceptors—e.g., nitrite reductase, sulfate reductase, thioredoxin reductase, etc.—free movement of this rather small, soluble protein may be an advantage. Immunogold labeling shows that ferredoxin is located throughout the cytoplasm of Synechococcus sp. strain PCC 7942 rather than in association with thylakoid or cytoplasmic membranes (Van der Plas et al. 1988). Perhaps loose binding facilitates replacement of ferredoxin by flavodoxin in times of iron depletion. There is a large body of literature demonstrating multiple forms of ferredoxin in higher plants and cyanobacteria. Unfortunately, there is some confusion about how to identify these forms and certainly much mystery about their functions. When two forms of ferredoxin have been separated by chromatography, Roman numerals I and II have been used to distinguish them but the assignment of the Roman numeral has not been done on a consistent basis. Characteristics used to distinguish multiple forms of ferredoxin are: photosynthetic reduction, chromatographic elution profiles, relative abundance in the cells, isoelectric points, redox potentials, and amino acid sequences. Of these, the amino acid sequences may prove to be the most reliable. Identification by photosynthetic reduction activity may not be good since ferredoxin specificity in this assay system is not highly selective. Hutber et al. (1978) compared the activities of ferredoxin I and II from Nostoc sp. strain MAC using illuminated Nostoc sp. membrane particles as electron donor and found ferredoxin II had about 66% as much activity as ferredoxin I in catalyzing photoreduction. They use ‘ferredoxin II’ to designate the first eluting, less abundant ferredoxin in the extract. Ferredoxin II was slightly more active than ferredoxin I in a pyruvate

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oxidase assay system using enzymes from Clostridium pasteurianum (Hutson et al., 1978). Wada et al. in 1981 found no difference in photoreductionwhen ferredoxins I and II fromNostoc muscorum were compared under similar conditions. Nostoc sp. strain MAC cells had three times as much ferredoxin I as ferredoxin II when grown either photoautotrophically or photoheterotrophically. The amino acid sequences of these two proteins give clear distinction to their identities (Hase et al., 1982). It seems extremely likelythat the amino acid sequence which is similar in all ferredoxins I that have been isolated from a wide variety of photosynthetic

organisms is the sequence of the catalyst for photosynthetic reduction. Unfortunately, not every laboratory can perform amino acid sequence analysis to establish the identity of ferredoxin. Ferredoxin II usually has a lower pI than ferredoxin I but conservation of charged residues is not so rigid that one can be sure that ion-exchange chromatography profiles or isoelectric focusing patterns will identify all ferredoxins I and II. One suspects this is the case in the observation of Sakihawa and Shin (1987) that 97% of the ferredoxin of Synechococcus vulcanus eluted from DEAE ahead ofa minor band equal to 3% ofthe total. In several cases the two

Chapter 12 Soluble Electron Transfer Catalysts of Cyanobacteria forms were found to be present in equal concentration (Shin et al., 1977; D.W. Krogmann, unpublished results). So, relative abundance is not a valid basis for distinction. There is no help in sorting from measurements of the redox potential. Spirulina maxima ferredoxins I and II have values of–390 and –310 mV respectively while Nostoc sp. strain Mac ferredoxins I and II have values of–350 and –455 mV (Cammack et al., 1977). Some of this data may indicate that ferredoxins II may have different functions in different organisms and so may be distinctly different proteins. Support for this possibility comes from an unpublished observations ofone ofthe authors (D. W. Krogmann, unpublished results). Two ferredoxins from Aphanizomenon flos-aquae were resolved by conventional DEAE chromatography and the more tightlybound form was identified as a ferredoxin Iby amino acid sequence determination (Lee et al., 1983). A slightly less-abundant ferredoxin was eluted at a lower salt concentration. When this fraction was purified by adsorption on Sephacryl S-200 equilibrated with 80% saturated and eluted with gradually decreasing concentrations of the ferredoxin was resolved into three different fractions. The best defined member of the ferredoxin II cluster is ferredoxin H which Schrautemeier and Böhme (1985) isolated from heterocysts of Anabaena variabilis. When compared to the ferredoxin from vegetative cells, ferredoxin H was equal to it in ability to reduce with illuminated thylakoids, and it catalyzed the reduction ofnitrogenase at twice the rate of the vegetative cell ferredoxin. Ferredoxin H was uniquely able to reduce nitrogenase using and hydrogenase or NADPH + FNR. Böhme and Schrautemier (1987) went on to purify and characterize the two ferredoxins. Ferredoxin H, which is in the ferredoxin II sequence cluster, adsorbed to DEAE-Sepharose more tightly and had a lower isoelectric point than the ferredoxin of vegetative cells which is in the ferredoxin I sequence cluster. This again illustrates that the isoelectric point is not a useful distinction between ferredoxins I and II. There were small differences in the absorption maxima, redox potentials and molecular weights but the EPR spectra were identical. This paper contains the important observation that ferredoxin H is less stable in dilute solution than the ferredoxin I of vegetative cells. If this lower stability is shared by other ferredoxins II, this may account, at least in

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part, for their low abundance and apparent absence from manypreparations.Next, Böhme andHaselkorn (1988) isolated the gene (fdxH) for ferredoxin H from Anabaena sp. strain PCC 7120 and, more recently Schrautemeier and Böhme (1992) cloned this gene from Calothrix sp strain PCC 7601 as well. The Anabaena sp. strain PCC 7120 fdxH is present in a single copy and is seven kilobases downstream from the nifHDK gene cluster which encodes the subunits ofdinitrogenase and dinitrogenase reductase. The deduced amino acid sequence differs in 47 ofthe 98 amino acid residues found in ferredoxin I of vegetative cells. In the region of residues 40 to 50 which is highly conserved in ferredoxins I and is part of a metal cluster loop involved in catalysis, the ferredoxins H have substitutions 43 and which could contribute to the differences in redox potential and specificity. There is joy in noting that Jacobson et al. (1992) have described the crystallization of ferredoxin H from Anabaena a sp. strain PCC 7120 and so we may expect a three dimensional structure of this molecule. A recent survey of ferredoxin sequences in the data banks showed sixteen ferredoxins I and four ferredoxins II. The ferredoxin IIs stand apart as a group. Of the one hundred amino acids in the molecule, fifty-nine positions are highly conserved in all ferredoxins I. Of these conserved residues, fifty-one show no change, five show one amino acid substitution and one shows two substitutions. Focusing on the four ferredoxin II sequences, one finds that in the fifty-nine conserved positions, there are ten and eleven substitutions in the ferredoxins II from N. muscorumand A. sacrum, respectively.There are fifteen substitutions in these positions in ferredoxin H (a ferredoxin II) from Anabaena sp. strain PCC 7120 and there are twenty-three substitutions in the ‘ferredoxin II’ from Synechococcus sp. strain PCC 6301 described by Cozens and Walker (1987). Note that this latter gene encoding a ferredoxin was found within a cluster of seven genes encoding subunits ofATP synthase. Location on the genome map can hardly be used in distinguishing genes, although more completed maps may give this meaning. This combination of location and the striking difference in amino acid sequence may indicate a unique function (which has not yet been determined) for the gene product. Gogotov and his colleagues have addressed the problem of multiple ferredoxins in nitrogen-fixing,

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heterocystous cyanobacteria and have made some importantcontributionstotheproblem.Yakunin et al. (1990), using elegant purification methods, isolated two ferredoxins fromAnabaena sphaerica. They use the designations ferredoxin I and ferredoxin II for their ferredoxin in a different sense than is usual and it would be best to await some primary structure data to sort out the identities. What is important is that they have found a more abundant ferredoxin with the conventional molecular weight of 10,600 and another ferredoxin at about one-tenth the concentration of the first with a molecular weight of 33,600. The larger molecule has six iron atoms and can be converted into a 10,600 molecular-weight form by heating in SDS. There is no evidence yet for reassembly of the trimer from monomeric subunits and no conversion ofthe ferredoxin isolated as a low molecular weight form into a trimer. There are small differences in the light absorption and EPR spectra of the two ferredoxins. The smaller ferredoxin was an effective electron donor to the nitrogenase of Rhodobacter sphaeroides and to the hydrogenase of Clostridium butylicum and could couple the phosphoroclastic reaction of pyruvate to hydrogen production. The trimeric ferredoxin did not work in these assays. Yakunin et al. (1991) found these ferredoxins present in cells grown on ammonia and precede the appearance of nitrogenase in cells adapting to as a nitrogen source. The smaller ferredoxin is also active in NADP reduction while the larger ferredoxin is not. A gene encoding a bacterial-type ferredoxin was found in the DNA from Anabaena sp. strain PCC 7120 by Mulligan et al. (1988). The gene encodes a sequence of 126 amino acids with an arrangement of four cysteines that is characteristic of binding sites for [4Fe-4S] centers. The deduced protein sequence gives a hydropathy plot consistent with the possibility of a membrane binding region. The gene is found between the nifB and nifS genes. The gene has been found in nitrogen-fixing bacteria as well but the gene product has thus far escaped detection. Yakunin et al. (1993) have found a bacterial type ferredoxin in cells from nitrogen-fixing cultures of A. variabilis. In addition to the typical cyanobacterial ferredoxin I and the heterocyst ferredoxin of Schrautemeier and Böhme (1985), a third ferredoxin was present in low concentration. The absorption spectrum of this protein is like that of bacterial ferredoxins and there are three to four iron atoms per protein of mass 6 kDa. Yakunin et al. designate this

protein ferredoxin III and report that it reduces bacterial nitrogenase and hydrogenase at faster rates than do the cyanobacterial ferredoxins. Ferredoxin III is not present in cells grown on fixed nitrogen which supports the suggestion of a role in nitrogen fixation. Yet another strange variant in ferredoxin structure has turned up in the work of Bottin and Lagoutte (1992) in their very thorough characterization of the ferredoxin I from Synechocystis sp. strain PCC 6803. In addition to the more abundant ferredoxin which precipitates with high concentrations of ammonium sulfate, they found a small amount of ferredoxin in theammoniumsulfate-saturatedsupernatant fraction. The saturated-ammonium sulfate soluble ferredoxin was found to react far less readily with antibody made against spinach ferredoxin than did the ammonium sulfate-precipitated ferredoxin. Both ferredoxins had identical amino acid sequences. The authors suggest that slight differences in conformation may be the basis for this saturated-ammonium sulfatesoluble form of ferredoxin. Consideration of the multiple forms of ferredoxin in cyanobacteria requires notice of the admirable work that Hase and his colleagues have done on ferredoxin isoproteins in maize. Kimata and Hase (1988) found four electrophoretically distinct ferredoxin fractions (I–IV) in etiolated seedlings. Only ferredoxin II reacted with a polyclonal antiserum prepared against ferredoxin I, and ferredoxins I and II were the only ones found present in leaves. Their levels were increased five-fold by light. Ferredoxin II was found only in bundle sheath cells while ferredoxin I was detected in both bundle sheath and mesophyll cells. Ferredoxin III was found in both mesocotyl and root cells and IV was found in mesocotyl cells. The concentrations of ferredoxins III and IV did not respond to light. Hase et al. (1990) next cloned the genes encoding the precursor proteins of I and III and found strikingly different transit peptides. Most recently, this laboratory (Hase et al., 1991) has recombined fragments of the genes of ferredoxin I and III to establish the site ofa difference in catalytic specificity. They had found that ferredoxin III catalyzed the reduction ofhorse heart cytochrome c far more rapidly than did ferredoxin I. When the carboxyl-terminal half of the gene for ferredoxin III was fused to the amino-terminal half of the gene for ferredoxin I, the protein produced from the recombinant gene had the specificity of ferredoxin III for cytochrome c reduction. Thus they have

Chapter 12 Soluble Electron Transfer Catalysts of Cyanobacteria demonstrated differences in tissue and cell specific expression and in catalytic activity and narrowed the latter to the carboxyl-terminal region of the protein. Shin et al. (1977) mentioned that the two ferredoxins they found in Nostoc verrucosum showed a difference in catalytic activity like that reported by Hase et al. (1991) for the maize isoproteins. One ferredoxin was more active in reduction and the other was more active in cytochrome c reduction. As noted at the beginning of this section, the many distinct enzymatic functions of ferredoxin have prompted speculations that the multiple forms ofthis protein might, in some instances, be specific for one or another enzymatic process. Recently, several more ferredoxin-requiring reactions have been added to the list. Marques et al. (1992) have purified a ferredoxin-glutamate synthase from Synechococcus sp. strain PCC 6301. Rhie and Beale (1992) have described a ferredoxin-heme oxygenase to generate the biliverdin precursor for phycobilin synthesis in Cyanidium caldarium, and a ferredoxin-dependent protochlorophyllide reductase has also been described (see Chapter 17). It seems likely that these enzymes will have a counterpart in cyanobacteria. II. Flavodoxin Flavodoxin replaces ferredoxin in some cyanobacteria, algae and heterotrophic bacteria when they suffer iron deficiency (see Chapter 25). This catalyst was first recognized by R. M. Smillie (1965) who called it phytoflavin. Soon after this the enzyme was found in Clostridium and Azotobacter sp. in which it was recognized as a flavoprotein that, during iron deficiency, replaced the ferredoxin used in nitrogen fixation. Flavodoxin is an acidic protein (pI = 3.5 to 3.8) of 18 kDa mass with a single FMN prosthetic group. Redox potentials of–50 mV to –244 mV for the reduction ofthe oxidized to the semiquinone and of –370 mV to –450 mV for the reduction of the semiquinone to the hydroquinone form have been reported. Thus flavodoxin resembles ferredoxin in net charge and redox properties.

A. Genes The first complete primary structure of a cyanobacterial flavodoxin was deduced from the sequence of its gene (denoted isiB) from Synechococcus sp. strain PCC 7942 by Laudenbach et al. (1988). The

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isiB gene was found as a single-copy gene whose transcription was induced by low-iron concentration (see Chapter 25). In contrast, petFI mRNA concentration was unaffected by the iron level. Iron deficiency prompts the appearance of two transcripts of 1.1 and 1.9kb with initiation sites in close proximity. The 1.1 -kb transcript terminates just upstream from the start of the flavodoxin coding sequence while the 1.9-kb transcript contains the entire isiB gene. The deduced amino acid sequence codes for a mature protein of 169 amino acids with a calculated molecular weight of 18,609 and an estimated isoelectric point of 4.78. Leonhardt and Straus (1989) next isolated the isiB gene from Anabaena sp. strain PCC 7120. More recently these authors have cloned and sequenced the isiB gene and its upstream neighbor (isiA), that encodes an ironstress-induced, PS II-associated, chlorophyll-binding protein from Synechococcus sp. strain PCC 7002 (Leonhardt and Straus, 1992). These two genes form a dicistronic operon activated by iron stress to produce an abundant monocistronic message containing isiA and a much less abundant dicistronic message that contains both isiA and isiB. They find the same transcriptional phenomena in Synechococcus sp. strain PCC 7942. The genes for the corresponding proteins in iron-sufficient cells, CP43 (PsbB) and ferredoxin, have also been cloned and sequenced and they are constitutively expressed (with orwithout iron). Fillat et al. (1991) isolated the flavodoxin gene from Anabaena sp. strain PCC 7119. These authors confirmed that iron regulates flavodoxin expression at transcription. They also find a single-copy gene and note that if heterocysts have a distinct flavodoxin, then it must have a distinctive sequence. It should be noted that the genes encoding flavodoxins from heterotrophic nitrogen-fixing bacteria encode a slightly larger polypeptide of approximately 19 kDa. The gene is called nifF and is said to encode a Nifspecific flavodoxin. Thus, there is a heterocyst-specific ferredoxin in cyanobacteria and a Nif-specific flavodoxin in nitrogen-fixing heterotrophs. Since these genes are single-copy, one must conclude that if isozymic forms of these proteins exist, their genes don’t readily cross hybridize. There is, as yet, no mention of isozymic forms of flavodoxin but some careful protein fractionation may be needed to check this. Alternatively, the gene encoding flavodoxin in heterocystous cyanobacteria could be differentially transcribed in vegetative cells (iron regulation) and heterocystous (developmental regulation).

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B. Protein Chemistry Flavodoxin is not difficult to purify ifone induces the enzyme by iron depletion. Flavodoxin is frequently found in cyanobacteria collected from dense natural blooms where both iron and copper deficiency are common occurrences. The fact that flavodoxin precipitates above 50% saturation with ammonium sulfate provides a great advantage in removing the phycobiliproteins. Puyeo and Gomez-Moreno (1991) have described an excellentprocedure forpurification of flavodoxin, FNR and ferredoxin from Anabaena sp. strain PCC 7119. Earlier their laboratory had characterized this flavodoxin (Fillat et al., 1988, 1990). Paulsey et al. (1990) have done similar redox and spectral measurements with the flavodoxin of Anabaena sp. strain PCC 7120. Bottin and Lagoutte (1992) have isolated the flavodoxin from Synechocystis sp. strain PCC 6803 and the enzyme from this unicellular organism seems quite like those of the two filamentous Anabaena sp. A crystal structure (see Fig. 2) for flavodoxin from Synechococcus sp. strain PCC 6301 was published by Smith et al. (1983). Fukuyama et al. (1990,1992) solved the crystal structure of this enzyme from the red alga Chondrus crispus, so one can have considerable confidence in the knowledge of the structure of these molecules. These structures will certainly assist the description of flavodoxin in solution by nuclear magnetic resonance methods; this work has been initiated by Stockman et al. (1990). Thus the stage is set for bravura performances in molecular biology. Two molecules, ferredoxin — an Fe-S protein with a mass of 10 kDa and flavodoxin — an FMN-containing protein with a mass of 20 kDa — perform the same catalytic transfer of electrons from PS I to FNR. It is likely that both receive electrons from the same site on the PS I complex (see Chapter 10), and it is clear that both donate electrons to FNR and perhaps many other electron acceptors. Since FNR is soluble and a portion of its structure is known, one may hope that the precise interactions of ferredoxin and flavodoxin with FNR will be revealed. Both ferredoxin and flavodoxin can easily be produced in large amounts in E. coli, so surely one will soon see some interesting site-directed mutagenesis studies ofthese molecules. On the flavodoxin side, there is some chemical work on its interaction with FNR. Walker et al. (1990) report elegant kinetic measurements of electron flow through complexes of FNR and flavodoxin. There is

an interesting suggestion from this work that a protein rearrangement may occur during electron transfer. Pueyo and Gomez-Moreno (1991) describe a carbodiimide cross-linked complex of the two flavoproteins in a one-to-one ratio. Complex formation hinders electron flow from flavodoxin to cytochrome c and blocks flavodoxin reduction by PS I. The complex is useful in studying electron transfer between the two flavoproteins. Medina et al. (1992) derivatized flavodoxin with carbodiimide and glycine ethyl ester. This inhibited the ability of the molecule to act as an electron donor more severely than its ability to act as an electron acceptor. The authors conclude that Asp 123, Asp 126, Asp 144 and Asp 146 are involved in binding flavodoxin to FNR (see Fig. 2).

C. Physiological Response to Iron Deprivation Flavodoxin was first found in iron deficient Synechococcus sp. and it has been found in a number of other unicellular genera of cyanobacteria (for a more complete discussion of the iron-deprivation response in cyanobacteria, see Chapter 25). In contrast, some of the filamentous Anabaena sp. lack the ability to express this enzyme in response to iron depletion. Pardo et al. (1989) found that Anabaena variabilis strain PCC 6309 showed a parallel decline of ferredoxins I and II but no flavodoxin appeared to replace them. Sandmann et al. (1990) tested more strains of Anabaena sp. with interesting results. Anabaena sp. strain ATCC 29413 replaced ferredoxin with flavodoxin at low-iron levels. Anabaena sp. strain ATCC 29211 did not synthesize flavodoxin. Anabaena sp. strain ATCC 29151 has a constitutive flavodoxin in its heterocysts. It should also be noted that the red alga Chondrus crispus has a constitutive flavodoxin (Fitzgerald et al., 1978). These observations show the loss offlavodoxinin some organisms while there is constitutive expression in others. More such observations and a better knowledge of the disposition ofthese organisms in nature might allow us to see why these genes are lost or retained. An Anabaena sp. whose habit is to dwell on the bottom of an aquatic environment might have a mechanism for gathering precipitated iron and so not need flavodoxin. A unicellular or planktonic species growing high in the water column might regularly experience iron deficiency and so be obliged to retain the flavodoxin option. A Chondrus sp. battered constantly by the (possibly) iron-deficient waters of

Chapter 12 Soluble Electron Transfer Catalysts of Cyanobacteria

the Irish Sea might be obliged to use flavodoxin to spare iron for cytochromes.

III. Ferredoxin

Reductase (FNR)

This enzyme catalyzes the transfer of electrons from reduced ferredoxin or reduced flavodoxin to This enzyme consists of a single polypeptide of 43 to 55 kDa and has one non-covalently held FAD as its coenzyme. The midpoint potential at pH 7 for a twoelectron change is –320 mV to –370 mV for FNR from various species. There have been several models to describe the binding of this rather loosely held

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enzyme to the photosynthetic membrane and now there is a new suggestion for a unique binding mechanism to the membrane in cyanobacteria (Schluchter and Bryant, 1992). Karplus et al. (1991) have published a crystal structure of FNR from spinach (see Fig. 1) and their paper is a compact review of a great deal of biochemical studies performed earlier.

A. Genes Fillat et al. (1990b) reported the first partial gene sequence for FNR from the cyanobacterium Anabaena sp. strain PCC 7119. Sancho et al. (1988)

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had published an N-terminal sequence for this protein that matched well the amino-terminal sequence of Spirulina sp. FNR, which protein had been sequenced in its entirety byYao et al. (1984). ThepetH gene was isolated using a cDNA ofthe gene for FNR from pea and an oligonucleotide to the N-terminal region of the Anabaena sp. gene. The gene so isolated encoded a sequence of 304 residues and was 65% identical to the 294 residue sequence from Spirulina sp. The Anabaena sp. petH gene product is 50% identical to the pea FNR and the molecular weight is close to that of a number of higher plant FNRs. Schluchter and Bryant (1992) used the FNR gene of pea to isolate the petH gene from Synechococcus sp. strain PCC 7002. They found a gene with an amino acid sequence more than one hundred residues longer than the earlier sequences from cyanobacteria. They isolated a larger FNR protein and confirmed the assigned translation start codon by amino-terminal amino acid sequencing. Mapping of the petH transcript by primer extension indicated that transcription begins 112–114 bp upstream of translation initiation site. Interposon mutagenesis was attempted to delete the single-copy gene. The results indicate thepetH gene is required for viability under photoautotrophic growth conditions. The extension of amino acid sequence is at the aminoterminus ofFNR and residues 1 through 85 are 78% similar to the CpcD protein of Synechococcus sp. strain PCC 7002. The CpcD protein is a phycocyaninassociated linker protein in the phycobilisome; its role is to limit peripheral rod-length variation by terminating rod elongation (de Lorimier et al., 1990; see Chapter 7). A reexamination of the petH gene sequence indicates that an even larger amino-terminal domain (~ 136 residues) exists in Anabaena sp. strain PCC 7119, although a portion of this sequence also has a high degree ofsequence similarity to CpcD (W. M. Schluchter and D. A. Bryant, personal communication). Phycobilisomes of Synechococcus sp. strain PCC 7002 (Schluchter and Bryant, 1992) and Synechococcus sp. strains PCC 6301 or 7942 (A. N. Glazer, J. Collier and A. Grossman, personal communication) were found to contain high levels of FNR. Immunoblots of whole-cell extracts from Synechococcus sp. strain PCC 7002 showed immunoreactive bands at 45 and 43 kDa. Five other cyanobacteria from four distinct genera gave comparable bands on immunoblots. The amino-terminal extension was suggested to anchor FNR to the phycobilisome, where

it would be reasonably close to the fixed PS I source of electrons which ferredoxin must bring to FNR. The 45 kDa FNR polypeptide fractionated as a cytosolic (hydrophilic) protein while the 43 kDa species pelleted with the membrane fraction and was more hydrophobic. Suggestive evidence indicated that the 43-kDa species might be a fatty-acylated derivative of the 45-kDa protein. Fatty acylation might target some of the FNR to the cytoplasmic membrane, where this enzyme is thought to serve as a dehydrogenase for the respiratory chain (Scherer et al., 1988). It is interesting to note that a pea cDNA encoding FNR has been expressed in E. coli , and it appears that chaperonins are required to fold or assemble a functional enzyme (Carillo et al., 1992). The catalytically active and full-length form of FNR of Synechococcus sp. strain PCC 7002 has also been overproduced in E. coli (W. M. Schluchter and D. A. Bryant, personal communication).

B. Protein Chemistry Purification of FNR from cyanobacteria is made difficult by the coprecipitation of FNR and phycocyanin in both acetone and ammonium sulfate. Next one finds that FNR and phycocyanin coelute from DEAE columns. These observations may possibly be related to the phycocyanin-binding domain at the ammo-terminus of the protein. Susor and Krogmann (1966) overcame this difficulty by selectively denaturing the phycocyanin by stirring in 75% acetone at room temperature. Wada et al. (1983) were able to remove most of the phycocyanin from their preparation by chromatography on Blue Sepharose. Sancho et al. (1988) purified the enzyme by repeated chromatography on both DEAE and Cibachron Blue-Sepharose columns. Serre et al. (1991) have describedcrystals of FNR from Anabaena sp. strain PCC 7119. All of these preparations are haunted by the specter of proteolysis and some effective protease inhibitors must be used to assure isolation of the ‘intact’ enzyme (‘intact’ in this sense has in the past meant a 32–34 kDa form ofthe protein that is catalytically active but that is in fact already degraded). It should be noted that the cyanobacterial FNR is quite stable and resistant to proteolytic attack when isolated along with intact phycobilisomes (W. M. Schluchter and D. A. Bryant, personal communication). However, disruption of cells at low ionic strength, under which conditions phycobil-

Chapter 12 Soluble Electron Transfer Catalysts of Cyanobacteria isomes rapidly dissociate, leads to rapid degradation of FNR to smaller but often enzymatically active species. These observations strongly suggest that FNR is protected from proteolysis as long as phycobilisomes are intact and this supports the proposed localization of the protein within the phycobilisomes. Jon Herriott has long sought the tertiary structure of a higher-plant FNR and this has culminated in the paper of Karplus et al. (1991) describing the three dimensional structure of the spinach enzyme (see Fig. 1C). The spinach enzyme is 312 residues long when shorn of its leader sequence, that bears no resemblance to the amino-terminal extension sequence of Schluchter and Bryant (1992). This spinach enzyme sequence corresponds to and closely resembles the shorter forms of cyanobacterial FNR that have been studied in detail. Thus, the structure of the spinach enzyme is likely to closely resemble the structures of the catalytic domains of the Anabaena sp. strain PCC 7119 and Spirulina sp. enzymes. The spinach FNR structure seems to confirm most of the inferences obtained by derivitization and crosslinking studies of the cyanobacterial enzymes.

C. Interaction of Ferredoxin with FNR Computer modeling of the interaction of ferredoxin and FNR with the derived crystal structures of these molecules from spinach has been reported by Karplus et al. (1991). The negatively charged region of ferredoxin, Glu92, Glu93 and Glu94, was placed near the positively charged region of FNR, positions 85–93: KNGKPHKLR (see Fig. 1).These two regions previously were shown to be the major sites of 1ethyl-3-[3-(dimethylamino)propyl]carbodiimide (EDC) cross-linking between ferredoxin and spinach FNR by Zanetti et al. (1988). Either Lys85 or Lys88 of FNR (see Fig. 1) was covalently linked to a glutamic acid residue of ferredoxin. Also, the Fe-S center of ferredoxin was positioned near the exposed edge of the flavin ring of FNR. This arrangement had ferredoxin filling the cleft between the two domains of FNR (i.e., the FAD and binding domains) without overlapping with the binding site. The modeling exercise agreed with the findings of other workers. Sancho et al. (1990) demonstrated that and ferredoxin have different binding sites by differentially protecting Anabaena sp. strain PCC 7119 FNR from phenylglyoxal modification. Ferredoxin protected Arg93 (Arg94 in Fig. 1) from

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modification and this residue is located within the positively charged region of FNR which has been shown above to interact with ferredoxin. Also, as discussed by Apley and Wagner (1988), several lines of evidence, taken collectively, suggest that ferredoxin binds near the center of FNR with the exposed portion of the flavin ring of FNR in close proximity to the Fe-S center of ferredoxin. Tryptophan residues were shown to be in or near the ferredoxin binding site of spinach FNR by the work of Davis (1990). The presence of ferredoxin decreased tryptophan fluorescence by 50%. There are six tryptophan residues in spinach FNR (see Schluchter and Bryant, 1992; Karplus et al., 1991 for the comparisons of derived primary structures of several FNR species relative to the tertiary structure of spinach FNR). Interestingly, the positions ofthree tryptophans in the crystal structure are within or adjacent to the ferredoxin binding site and the other three are in positions that would make them unlikely candidates for interaction with ferredoxin. Trp 182 is in the cleft on a helix which extends through the two domains of FNR. Thus, the filling of the cleft between the two domains of FNR by ferredoxin may be responsible for quenching the fluorescence of this residue. The other two tryptophan residues whose fluorescence may be quenched by the binding of ferredoxin are Trp296 and Trp309 (Fig. 1). The location of these residues are in the domain and adjacent to the cleft between the two domains of FNR where ferredoxin has been proposed to bind. The other three tryptophans, Trp57, Trp199 and Trp259, are on the opposite side ofthe FNR molecule from the proposed site of ferredoxin binding and therefore would not be directly affected by the presence of ferredoxin. Medina et al. (1992) investigated the involvement of lysine residues in the interactions between ferredoxin and FNR of Anabaena sp. strain PCC 7119 by chemical modification with pyridoxal phosphate. Lys53 of FNR was modified to the greatest extent relative to other lysines, and ferredoxin afforded protection from modification for this residue. Lys53 corresponds to Arg71 of spinach (Fig. 1). When Arg71 is examined on the crystal structure of FNR, its location is seen to be on an extended chain above the ferredoxin binding site of FNR, positions 85–93, and near Trp182 proposed here to be affected by the binding of ferredoxin. Also, by analogy with lysine residues affected by modification of Anabaena sp. strain PCC 7119, Lys304 and Lys305 of spinach

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would be considered in or near the binding site of ferredoxin. As seen in the crystal structure, these residues lie on a helix that is adjacent to the cleft between the two FNR domains. Lys304 and Lys 305 are also near Trp309, which is proposed here to be affected by the ferredoxin binding. Apley and Wagner (1988) also investigated the role of lysines on FNR in the interaction of this protein with ferredoxin. Chemical modification of spinach FNR with eosin isothiocyanate identified a singularly labeled lysine within the region involved in ferredoxin binding. Within the span of residues extending from 179 to 228, there are two lysine ‘rich’ regions: 179 to 192 (SFLWKMFFEKHDDYK) and 213 to 222 (KDDFEKMKEK). As seen from the crystal structure, the positions of these residues suggest that Lys 183 is the likely candidate for protection by ferredoxin. Lys 183 is located next to Trp l82 and near Arg71, both of which are proposed to be affected by ferredoxin binding (Fig. 1).

D. Enzyme Activity Pueyo et al. (1992) have made a covalently crosslinked complex of ferredoxin and FNR. Treatment of the two proteins with EDC gives a one-to-one complex. The complex shows activity in NADPHdependent, cytochrome c reductase activity indicating passage of electrons through FNR to ferredoxin and then to cytochrome c. No ternary complex is formed with flavodoxin, as would be expected if ferredoxin and flavodoxin bind at the same site. A light-dependent deactivation/reactivation of A. variabilis FNR has been described by Fillat et al. (1991). When cells reach the stationary phase of growth, the activity of FNR declines to 20% of its maximal level without a corresponding loss of the enzyme protein. If the cells are diluted and illuminated, reactivation occurs at a rate dependent upon light intensity. It appears that there is an alteration in the structure of FNR, possibly by an intramolecular thiol disulfide interchange, which lowers the affinity of FNR for ferredoxin and the for NADP and for ferredoxin in the diaphorase and cytochrome c reductase assays, respectively. There is an intriguing report by Hodges et al. (1990) that, when FNR is incubated with labelled ATP, the label was transferred to FNR in the presence of pea thylakoids. In darkness, the label goes to serine and in light to threonine. These observations may lead to the elucidation of a new

control mechanism in higher plants; it is not known if such a mechanism occurs in cyanobacteria. IV. Plastocyanin Plastocyanin is a protein of approximately 100 amino acid residues and contains a single atom of copper which is the redox-active component of electron transport. The protein is believed to transfer electrons from cytochrome f to P700 of PS I in the photosynthetic electron transfer chain. The redox potential of plastocyanin is about +360 mV. The isoelectric point of plastocyanin varies among cyanobacterial species and ranges from 3.8 in the unicellular Synechococcus sp. strain PCC 6301 to 9.3 in the filamentous, heterocystous Anabaena flosaquae. In eucaryotic algae and higher plants the isoelectric point is uniformly acidic—typically around pH 4. In the event of copper deprivation, which is common in aquatic habitats, plastocyanin may be replaced by one or more electron carriers.

A. Genes Van der Plas et al. (1989) isolated the first cyanobacterial petE gene encoding plastocyanin from Anabaena sp. strain PCC 7937. The gene sequence indicated a preprotein of 139 amino acids with an amino-terminal extension of 34 amino acids; this extension resembled other transit peptides and probably could facilitate passage ofthe plastocyanin across the thylakoid into the lumen. A monocistronic transcript of 740 bases was found, and this transcript was diminished by copper deficiency. The gene was transferred to Synechococcus sp. strain PCC 7942 which hasnoplastocyanin; inthese cells transcription occurred regardless of the copper concentration in the medium. Briggs et al. (1990) isolated the plastocyanin gene from Synechocystis sp. strain PCC 6803 and found it also encoded a precursor protein of 126 amino acids with a transit peptide of 29 amino acids. Zhang et al. (1992) examined the regulation of plastocyanin synthesis by copper in Synechocystis sp. strain 6803 and found that no plastocyanin could be detected at 30 nanomolar copper but the protein was present at one micromolar copper. Cytochrome which substitutes for plastocyanin in copper-deficient cells, was present in the low-coppergrown cells and absent in the higher-copper-grown cells. Similar results were obtained with analysis of

Chapter 12 Soluble Electron Transfer Catalysts of Cyanobacteria plastocyanin mRNA. Thus, both the genes for plastocyanin and cytochrome appear to be regulated in their expression at the level of transcription by very low concentrations ofcopper. Bovy et al. (1992) reached the conclusion that copper regulates the expression of both of these genes at the initiation of mRNA synthesis in Anabaena sp. strain PCC 7937. These authors found that the copper-dependent changes in mRNA levels were abolished by adding chloramphenicol to the cells from which they inferred a possible de novo synthesis ofa trans-acting element in the regulatory process. The mRNAs for neither plastocyanin nor cytochrome showed altered stability under varying copper concentrations. Caution is still advisable in generalizations about the regulation of plastocyanin synthesis as evidenced by studies of eucaryotic algae. In Scenedesmus obliquus the regulation is like that of Anabaena sp. (Li and Merchant, 1992) while in Chlamydomonas reinhardtii, regulation is achieved at the posttranslational stage (Merchant and Bogorad, 1986).

B. Protein Chemistry Studies of the protein chemistry of cyanobacterial plastocyanins may have been impeded by difficulties in the purification of this protein from this source. Unfortunately, all ofthe cyanobacteria collected thus far from natural blooms or commercial cultures have failed to yield any plastocyanin. Fortunately, there are new isolation procedures that improve the convenience, yield and purity of isolations from laboratory-cultured cells. Tan and Ho (1989) have devised a procedure involving absorption of plastocyanin onto a Sephacryl S-200 column from a solution with subsequent elution at a lower salt concentration. The high salt concentration protects acidic plastocyanins of unicellular cyanobacteria from loss of copper. Christensen et al. (1990) have published an excellent procedure for the isolation of the basic plastocyanin from Anabaena variabilis. The total cell extract is diluted to a very low ionic strength (a conductivity below 0.25 mS) which allows the plastocyanin to bind to an S-Sepharose Fast-Flow column from which it is eluted with 40 mM NaCl. As the salt concentration is lowered, the plastocyanin is absorbed on a second S-Sepharose column, and it is then eluted by an NaCl gradient. This very simple and rapid procedure gives plastocyanin in high yield and of high purity. Morand and Krogmann (1993) describe a purification of

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plastocyanin from spinach which may be useful for cyanobacteria. For large amounts ofcyanobacterial plastocyanin, it would be very nice to overproduce the protein in E. coli that had been transformed with the petE gene behind appropriate promoters. This has been done for the precursor apoprotein of a higher plant plastocyanin (Hibino et al., 1991). Perhaps the lack of abundant plastocyanin is responsible for the lack of experiments on the structure ofthe cyanobacterial protein. Overproduction of the Synechocystis sp. strain PCC 6803 plastocyanin in E. coli has recently been achieved, but reconstitution with copper has not yet been investigated (J. Zhao and D. A. Bryant, personal communication). Three dimensional structures of plastocyanin (see Fig. 1D) have been determined for the proteins from poplar (Guss et al., 1986) and from the green alga Enteromorpha prolifera (Collyer et al., 1990). Moore et al. (1988) solved the three-dimensional structure of plastocyanin from the green alga Scenedesmus obliquus by NMR spectroscopy. The structures derived from these two methods and for the various organisms are quite similar to one another. The algae seem to have a less concentrated negative charge on the ‘east-face acid patch’ in the residue 58 to 60 sequence ofthe poplar structure. While there are many studies of the interaction of plastocyanin with its reaction partners, most have been done with enzymes from higher plants. Morand et al. (1989) and Bagby et al. (1990) have nicely summarized the many physical and chemical experiments that indicate that the ‘east-face acid patch’ of plastocyanin is the site for receiving electrons from cytochrome f and the ‘north-face hydrophobic patch’ of plastocyanin is the site for donating electrons to PS I. Roberts et al. (1991) have used computer graphic alignments of protein electrostatic fields and a systematic orientational search of intermolecular electrostatic energies using the crystal structures of plastocyanin and tuna cytochrome c. Their data is consistent with the earlier work which indicates the ‘east-face acid patch’ reaches toward the heme edge crevice of the cytochrome. A crystal structure of a higher-plant cytochrome f has recently appeared (Martinez et al., 1992; W. A. Cramer, personal communication) and this should strengthen confidence in the models of cytochrome f - plastocyanin interaction. Zhou et al. (1992) have developed impressive new physical evidence for the plastocyanin-cytochrome docking

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interactions. He et al. (1991) and Modi et al. (1992, 1993) have used site-directed mutagenesis to identify tyrosine 83 of spinach plastocyanin (Fig. 2) as a participant in both binding to and electron transfer from cytochrome f. Christensen et al. (1993) have extended some of these observations to the equivalent tyrosine in Anabaena variabilis plastocyanin. Note that the seven carboxyl residues in the ‘east-face acid patch’ of spinach plastocyanin have, in Anabaena variabilis, been diminished to one carboxyl group and one lysine. Reduced plastocyanin donates electrons to P700 and, in this process with higher plant PS I particles, plastocyanin can be trapped as a covalently cross-linked complex with the 19-kDa protein which is the product ofthe psaF gene (Wynn and Malkin, 1988; see Chapter 10 for further details).

C. Physiological Response to Copper Deprivation Paul Wood (1978) realized that when the alga Chlamydomonas reinhardtiiwas subjectedto copper deprivation, plastocyanin was replaced by cytochrome In cell-free reconstitution of PS I activity, the two proteins were virtually interchangeable. Sandmann and Böger (1980) confirmed this in the cyanobacteria Anabaena variabilis and Plectonema boryanum. Sandmann (1986) surveyed fifteen additional species of cyanobacteria and found that four of these (Phormidium foveolarum, Spirulina platensis, Tolypothrix sp. and Fischerella muscicola) did not make plastocyanin in response to copper. This might be due to an environmentally driven gene deletion. Consider that Spirulina maxima has been growing in Lake Texcoco near Mexico City for a very long time. There is recorded human witness to its presence there in 1520, and it is likely to have been there earlier. The waters of the lake have been alkaline — pH 10 or higher — for much ofthat time and copper is not soluble in that condition. Thus, the gene for plastocyanin would be useless for many generations. Synechococcus sp. strain PCC 7942 is a unicellular species that can be added to the list of cyanobacteria that fail to make plastocyanin and produce the substitute cytochrome in the presence of copper. Someday these organisms should be examined for remnants of the plastocyanin gene. Sandmann noted in his survey that Pseudanabaena foveolarum and Calothrix membranacea contained small but appreciable amounts of cytochrome in

cells with ample copper and generous levels of plastocyanin. Earlier, in the section on gene regulation, substantial evidence for the suppression of cytochrome gene transcription by copper was cited. In a few species, this suppression may be less complete or the residual cytochrome may be extraordinarily persistent. In higherplants, only plastocyanin is found and no trace ofthe cytochrome has been reported. Perhaps the gene encoding cytochrome shouldbe sought in transitional species. It seems plausible that plants which live in contact with the insoluble stratum may have an efficient mechanism for collecting copper from itsinsolubleforms.Thiswouldsolve theproblem of copper availability as the enterochelins have vastly improved iron availability for bacteria and plants. Still, the triumphal persistence ofthe copper protein plastocyanin over cytochrome in higher plants suggests that it mayprovide some positive advantage.

V. Cytochrome Cytochrome (formerly or can replace plastocyanin in some cyanobacteria and algae experiencing copper depletion and it permanently replaces plastocyanin in some species. Like plastocyanin, cytochrome catalyzes the transfer of electrons from cytochrome f to P700. There is substantial evidence that cytochrome serves as an electron donor to the respiratory cytochrome oxidase of cyanobacteria. The redox potential varies from +330 to +380 mV. Cytochrome is a protein of approximately ninety amino acids with a single iron heme covalently held by thioether bonds at a characteristic -Cys-X-X-Cys-His- site. The isoelectric point of this protein varies from about 3.7 in unicellular cyanobacteria to 9.3 in filamentous species.

A. Genes Laudenbach et al. (1990) isolated the petJ gene for cytochrome from Synechococcus sp. strain PCC 7942. The gene was isolated through the use of oligonucleotide probes designed from the amino acid sequence of the protein. The gene encodes a preprotein of 111 amino acids with a probable 24 residue amino-terminal signal sequence which presumably facilitates passage ofthe protein into the

Chapter 12 Soluble Electron Transfer Catalysts of Cyanobacteria thylakoid lumen. The gene is present in a single copy. Copper had no influence on the amount of mRNA from this gene. It should be noted that Synechococcus sp. strain PCC 7942 has neither plastocyanin nor a detectable gene encoding it, so there should be no need for copper regulation of the cytochrome gene. The petJ gene encoding cytochrome was disrupted by insertionally inactivated with a drugresistance cartridge, and this was used to transform Synechococcus sp. strain PCC 7942 cells into a mutant strain which no longer contained the wildtype gene or cytochrome Remarkably, these mutant cells grew only slightly more slowly than wild-type cells at low cell densities under non-saturating light conditions. The mutant cells evolved oxygen at various light intensities at the same rate as wild-type cells. The mutant did show a diminished efficiency in oxidizing cytochrome f on its PS II side. Thus it appears that cytochrome is not required for photosynthesis and perhaps not for respiration in this organism. The long arguments identifying plastocyanin (or its alternate, cytochrome as a mobile electron carrier seem to preclude a direct transfer of electrons from cytochrome f to P700. One supposes then that some other mobile electron carrier can fill the gap. Zhang et al. (1994) have confirmed the Laudenbach et al. observations in another cyanobacterium, Synechocystis sp. strain PCC 6803. The petJ gene was cloned, sequenced and used to construct a deletion mutant lacking the cytochrome. Plastocyanin was eliminated from these cells by copper deprivation, yet the mutant grew photoautotrophically and steadystate electron transport from water in the presence of bicarbonate was not affected, despite the absence of both plastocyanin and cytochrome Copper appears to regulate the transcription ofthe gene for cytochrome Zhang et al. (1992) have made a thorough study of cytochrome and plastocyanin synthesis in response to copper concentration using both immunological and spectrophotometric assays. The mRNAs for these proteins were observed by Northern blot analysis. Synechocystis sp. strain PCC 6803 grown at 20 to 30 nanomolar copper had only cytochrome and the mRNA encoding this protein. Cells grown at one micromolar copper had only plastocyanin and the mRNA encoding this protein. Cells grown at 0.3 micromolar copper had both catalysts and their corresponding mRNAs. Thus, copper stimulatespetE

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gene transcription and diminishes petJ gene transcription. Bovy et al. (1992) have reached similar conclusions about the regulation of these genes in Anabaena sp. strain PCC 7937 and have increased our insight by finding that chloramphenicol blocks the copper-induced changes in mRNA levels. This suggests that copper prompts the de novo synthesis of at least one trans-acting element in regulating the initiation of transcription of these genes. Further, they could find no influence ofcopper on the stability ofeither the plastocyanin orcytochrome mRNAs.

B. Protein Chemistry Cytochrome was first described by Holton and Myers in 1967. The protein is easily recognized by its visible color after removal of the phycobiliproteins byprecipitationwith 40-50% and is easily purified by ion-exchange chromatography. Further purification can be achieved by adsorption at high ammonium sulfate concentration on Sephacryl S500 columns (Tan and Ho, 1993). Cytochrome has a low isoelectricpoint when isolated from unicellular and simple filamentous forms of cyanobacteria, and so it is purified on DEAE-cellulose. The cytochrome has a high isoelectric point in complex filamentous species and must be purified on CM-cellulose (Ho and Krogmann, 1984). There is a parallel variation in isoelectric points of the plastocyanins from various cyanobacteria and this alteration clearly influences the interaction of plastocyanin or cytochrome with P700 (Davis et al., 1980). Wynn et al. (1989b) were able to crosslink cytochrome to a 17-kDa protein in a cyanobacterial PS I reaction center. This ‘docking’ polypeptide was identified as the product of the psaF gene in the PS I complex (see Chapter 10). Perhaps comparisons of amino acid sequences deduced for PsaF proteins of unicellular and filamentous cyanobacteria will provide clues about the coevolution of electron carriers and their docking sites. Widger (1991) has made tentative identification of possible binding sites of plastocyanin and cytochrome on cytochrome f from unicellular and filamentous cyanobacteria using the difference in pIs of the plastocyanins and cytochrome as a clue. A crystal structure of cyanobacterial cytochrome has been determined (Ludwig et al., 1983; this structure has been greatly refined and Dr. Ludwig kindly provided the data for the structure shown in Fig. 2), and it will be quite useful in modeling the

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interaction of this cytochrome with cytochrome f in future studies.

C. Physiology of Cytochrome While this review has dealt with cytochrome as a catalyst of photosynthesis, there is very strong evidence that this cytochrome participates in the respiration of cyanobacteria as well (see Chapter 13). Several isoforms of cytochrome have been observed (Ho and Krogmann, 1984) and one can observe variations in the proportions of these isoforms in response to growth conditions. Since the gene for this cytochromeis foundasasinglecopy,theisoforms may be the result of post-transcriptional modifications that relate to different physiological roles. The important question of the moment is what replaces cytochrome when the petJ gene has been deleted and there is no plastocyanin present. Perhaps another cytochrome fills the role in both respiration and photosynthesis. Malakhou et al. (1993) have described a gene for a new cytochrome in cyanobacteria. Holton and Myers (1967) had described a unique cytochrome of very low abundance in their photoautotrophically grown Synechococcus sp. strain PCC 6301. In considering cytochrome as a shared catalyst of photosynthesis and respiration, recall that Sandmann (1985) haspublished a study ofthe effects of iron deficiency on these processes in cyanobacteria. Photosynthesis can be greatly diminished with very little effect on respiration in Synechocystis sp. strain PCC 6714. If a cytochrome serves both processes, greater care may be needed to detect the effects of a gene interruption on respiratory activity.

VI. Low-Potential Cytochrome c A low-potential (–260 mV) cytochrome isfound in many cyanobacteria and in several eucaryotic algae. This protein has an apparent mass of 15 kDa upon electrophoresis in the presence of SDS, and it consists of approximately 130 amino acid residues with a single heme. The amino acid sequence reveals that two segments of the polypeptide are similar in sequence to two segments of cytochrome The isoelectric point varies from 4.1 to 4.8 in various genera. This cytochrome, or variants ofit, may reside

in three distinct cellular locations: as a soluble protein in the cytoplasm, and perhaps the periplasm, and as a tightly held constituent of the PS II core complex. While its in vivo function is not established, the lowpotential cytochrome is rapidly reduced by ferredoxin II and it can be oxidized by hydrogenase. ‘Low-potential’ is a better identification than the position ofthe alpha band which has been reported at 548, 549, and 550 nm for this cytochrome.

A. Protein Chemistry and Location in the Cell Holton and Myers (1963, 1967a,b) first isolated and characterized cytochrome from Synechococcus sp. strain PCC 6301. The cytochrome was found in high abundance, comparable to ferredoxin, at a ratio of one per 200 chlorophyll molecules. Dithionite but not ascorbate reduces this cytochrome, which is autooxidizable.Thepurificationprocedure of Holton and Myers has not been improved upon. The cytochrome precipitates in ammonium sulfate between 40 and 50% ofsaturation. The low pI (Alam et al., 1984) allows absorption onto DEAE and careful elution is required to remove contaminating phycobiliproteins. This protein is often missed in studies ofthe soluble proteins ofcyanobacteria since it is obscured by the far more abundant blue proteins. D. Sherman found that immunogold staining with an antibody to cytochrome of sections of Synechococcus sp. strain PCC 7942 indicates that the cytochrome is uniformly distributed in the cytoplasm of the cell. Laudenbach et al. (1990) published an electrophoretogram of cytochromes from Synechococcus sp. strain PCC 7942 which indicates that an appreciable fraction of the cytochrome is in the periplasm of these cells. There is a long series of observations placing a lowpotential cytochrome c near PS II in particles released from thylakoids by detergent disruption. Krinner et al. (1982), in the course ofisolating a cytochrome complex from Anabaena variabilis, released particles from thylakoids by detergent treatment and, in a subsequent detergent wash of these particles, released a c-type cytochrome with a mass of 15 kDa into solution. Bowes et al. (1983) found a similar cytochrome in detergent washed PS II particles from the thermophilic Phormidium laminosum. Redlinger and Gantt (1983) reported a similar experience in well-washed photosynthetic membranes from the

Chapter 12 Soluble Electron Transfer Catalysts of Cyanobacteria red alga Porphyridium cruentum. The same alga has also yielded a readily soluble form of low-potential cytochrome (Evans and Krogmann, 1983). In 1984, Omata and Murata found they could release low potential cytochrome c from thylakoid membranes of Synechococcus sp. strain PCC 6301 only after sonication. Recently, Hoganson et al. (1990) reported on new experiences with the bound lowpotential cytochrome c of Synechococcus sp. strain PCC 6301. They isolated thylakoids and washed them thoroughly with 2 mM Mes buffer at pH 6 to remove 95% of the phycocyanin (and probably a similar fraction of any soluble low potential cytochrome c). Treatment of the washed membranes with 1 M NaCl released low-potential cytochrome c which was purified by DEAE chromatography. They note that Holton and Myers (1967) had seen several peaks in the elution of this cytochrome from DEAE. Hoganson et al. (1990) report variations in the redox potential from –280 to –314mV (cf –260 mV of Holton and Myers, 1967) depending on the storage history ofthe sample. The chromatographic behavior and variance in redox potentials are the only hints so far that there may be multiple forms of this cytochrome. When Cohn et al. (1989) prepared and sequenced the low-potential cytochrome c from Microcystis aeruginosa, there was no indication of heterogeneity of the cytochrome which had been extracted from the cells with distilled water. The most recent piece added to this complex picture of the location of the low-potential cytochrome c is an elegant demonstration that this protein is held to a PS II core complex of Synechococcus vulcanus in an amount stoichiometric to the intrinsic CP43 and CP47 proteins and the extrinsic 33-kDa protein (Shen et al., 1992). The cytochrome adheres through treatment with several detergents and is released from the core particles by 2 M NaCl or 1 M which indicates it is an extrinsic constituent of the core. The identification of the cytochrome is unambiguous and its one-to-one stoichiometry with core constituents make it a promising candidate for participation in electron transfer in or near PS II. Genes (petK) encoding cytochrome have recently been identified on the chloroplast genome of the red alga Porphyra purpurea (Reith and Munholland, 1993) and on the cyanelle genome of the taxonomically ambiguous flagellate Cyanophora paradoxa (V. L. Stirewalt and D. A. Bryant, personal

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communication). Additionally, the petK gene of Synechocystis sp. strain PCC 6803 has been cloned (C. Kang, P. Chitnis, and D. W. Krogmann, unpublished results). All of these sequences predict leader sequences which presumably target at least some of the protein to the thylakoid lumen and/or periplasmic space.

B. Physiological Role Shen et al. (1993) have developed evidence that the low-potential cytochrome c bound to the PS II core is a functional component of the oxygen evolving complex. When the cytochrome and a 12-kDa protein are removed from the core complex, oxygen evolution diminishes; reconstitution with these two proteins restores maximal activity, while either one alone has only a slight effect. Matsuura and Fujita (1986) have provided a very interesting insight into the function of low-potential cytochrome c of Synechococcus sp. strain PCC 6301. They grew cells in light ofdifferent colors and found that the low-potential cytochrome c concentration varied in parallel with PS I and not PS II concentration. They suggested that low-potential ofthe cytochrome would allow it to reduce plastoquinone or cytochrome in response to the demand of PS I. Another approach to the problem of low-potential cytochrome function was taken by Pulich in 1977. He found that the cytochrome could stimulate the oxidation of NADPH in crude extracts and concluded that oxidoreductase was the catalyst responsible for this. Krogmann (1991) confirmed this and found the reaction was stimulated by ferredoxin. Ferredoxin II was ten-fold more active than ferredoxin I in delivering electrons to the lowpotential cytochrome c. In these experiments, the reduced cytochrome could deliver electrons to a crude hydrogenase preparation. Asada et al. (1993) have found that a partially purified hydrogenase and a c-type cytochrome from Microcystis aeruginosa could evolve hydrogen gas upon addition of hydrosulfite. Finally, it should be noted that Kinzel and Peschek (1983) have provided evidence by reconstruction of a cell-free system that the low-potential cytochrome c can serve as a cofactor of cyclic phosphorylation. Since cyclic phosphorylation can be elicited by a variety of redox catalysts, the lack of specificity of

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the thylakoids denies certainty to this experiment but a role for this cytochrome in cyclic phosphorylation is plausible. One must acknowledge that since the low potential cytochrome c can reside in several compartments ofthe cell, it may, like cytochrome have several functions. VII. Hydrogenase Hydrogenase is the least known and most difficult to study of the redox catalysts of cyanobacteria. There are at least two forms ofthe enzyme in cyanobacteria —a soluble, ‘reversible’ hydrogenase and a membrane bound, ‘uptake’ hydrogenase. One of the soluble enzymes appears to consist of two nonidentical subunits which form an easily dissociable dimer, one subunit of which binds electron donors for hydrogen formation and the other binds hydrogen. There is evidence that a nickel-iron center is involved in catalysis. The lability of the enzyme has surely impeded biochemical characterization.

A. Genes Genes potentially encoding the small subunit of the soluble hydrogenase ofAnabaena cylindrica (Ewart et al., 1990) and of Synechococcus sp. strain PCC 6716 (Van der Oost et al., 1989) have been isolated and sequenced. The sequences show no similarity to sequences ofthe genes for this subunit in a variety of otherbacteria. TheA. cylindrica gene was recognized by a probe designed from the amino-terminal sequence of the purified protein and the purification of this protein is surely a milestone in hydrogenase research. The gene predicted the probe sequence at its 5'-end and was of the size predicted from the size of the protein. The Synechococcus sp. strain PCC 6716 gene was isolated using the A. cylindrica gene as a probe. The uniqueness ofthese sequences prompts the suggestion that the soluble enzyme from cyanobacteria represents a distinct class of hydrogenases. Smith’s great work in purifying the enzyme to homogeneity needs repeating so that the amino acid sequences can be confirmed and extended. There is an extensive literature on the nickelcontaining hydrogenases and theirgenes from bacteria as diverseasDesulfovibriogigas andRhodospirillum rubrum. The reviews of Przybly et al. (1992) and of Reeve and Beckler (1990) provide good access to this information.

B. Protein Chemistry Two distinct hydrogenases have been recognized in heterocystous cyanobacteria and may be present in other cyanobacteria as well (Houchins and Burris, 1981). A reversible hydrogenase is found in heterocysts and in vegetative cells including those of non-heterocystous andapparentlynon-nitrogen fixing cyanobacteria. The enzyme is increased in many strains by an anaerobic induction period. In Spirulina maxima and Synechocystis sp. strain PCC 6803, the enzyme is present in extraordinarily high levels in cells growing in aerobic, photoautotrophic culture. An uptake hydrogenase is present in both heterocysts and vegetative cells and is membrane-bound; thus, it has been studied very little thus far. The uptake hydrogenase seems incapable of evolving hydrogen and its activity is saturated by a low partial pressure of hydrogen. Purification ofthe reversible hydrogenase has been plagued by the lability of this enzyme to oxygen inactivation. However, the enzymes from some cyanobacteria seem less vulnerable than the hydrogenases ofsome eucaryotic algae. The inclusion of strong reducing agents—hydrosulfite and/or dithiothreitol—in the preparative media does much to mitigate this problem with the cyanobacterial enzyme. An early partial purification (Hallenbeck and Bennemann, 1978) made clear the inability of Anabaena cylindrica ferredoxintoserve as anelectron donor to Anabaena cylindrica hydrogenase. This is in contrast to the ability of hydrogenase from Clostridium pasteurianum to accept electrons from various ferredoxins including those ofcyanobacteria (Fitzgerald et al., 1980). This suggested thatAnabeana cylindrica compartmentalizes its reductant pool by electron-carrier specificity, since nitrogenase in this organism is readily reduced by ferredoxin. Belkin et al. (1981) noted two forms of the reversible hydrogenase, distinct in chromatographic and catalytic properties, during purification ofthe enzyme from Oscillatoria limnetica. The separation ofthese isozymes is also described in Rao and Hall (1988). While a single form of this enzyme is more usual in purification, differential lability or differential induction of different forms are real possibilities. Laczkó (1986) found that low-light-grown cells of Anabaena cylindrica released most of the reversible hydrogenase to the supernatant after sonication while high-light-grown cells released only 30% of this activity and the rest remained membrane-bound.

Chapter 12 Soluble Electron Transfer Catalysts of Cyanobacteria Asada et al. (1987) described a nice preparation of hydrogenase from the unicellular Microcystis aeruginosa, and they saw hydrogen-evolving activity in the membrane pellet. This observation will need careful study in the future. In 1989, two laboratories described procedures that resulted in pure hydrogenase from cyanobacteria. Ewart and Smith (1989) have given an excellently detailed account of the purification of the soluble hydrogenase from Anabaena cylindrica. They show very clearly that the enzyme is a readily dissociable dimer oftwo nonidentical subunits with masses of 42 and 50 kDa. One subunit contains a hydrogen binding site and catalyzes an exchange of tritium gas into water. The other subunit carries the site for utilizing electrons from a reductant to form hydrogen and catalyzes the dithionite plus methyl viologen reaction which produces hydrogen. Kentemich et al. (1989) purified the reversible hydrogenase from Synechococcus sp. strain PCC 6301. Classical methods gave a 250-fold purification and a final polyacrylamide gel electrophoresis step brought the enzyme to homogeneity. It is interesting to note that this enzyme was more stable at room temperature than at 4° or 0 °C. The enzyme has subunits of 56 and 17 kDa, as is typical of the NiFeS-hydrogenases of other bacteria. An antibody to the enzyme was produced and used in immunogold labeling of thin sections. Most of the enzyme appears to be associated with the cytoplasmic membrane. In a subsequent paper, Kentemich et al. (1991) show clearly that the reversible hydrogenase, previously identified as soluble, is bound to the cytoplasmic membrane. This enzyme is probably a peripheral protein held by weak hydrophobic interactions which are easily disrupted by the techniques used to break open the cell. The authors note that the low of this hydrogenase for hydrogen suggests that the enzyme functions in hydrogen utilization. The enzyme appears to be on the periplasmic face ofthe cytoplasmic membrane, where it could strip protons and electrons from hydrogen to be used to form a proton gradient to drive ATP synthesis and for electron transport processes in the cytoplasmic membrane. Lindblad and Sellstedt (1990) used antibodies formed against the uptake hydrogenase of Alcaligenes latus to show the antigen is rather evenly distributed on the internal membranes in the cytoplasm of both heterocysts and vegetative cells of Nostoc sp. strain PCC 73102. There is a higher concentration ofthe enzyme in regions where vegetative cells are in contact with one another.

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One wishes for more data on purified hydrogenase from cyanobacteria. The participation of nickel in the catalytic activity of the enzyme is indicated by the relation ofhydrogenase activity to the availability of nickel in the growth medium (Almon and Böger, 1984; Dayday et al., 1985). A metal analysis of the enzyme would be reassuring. Smith et al. (1992) have determined the reduction potential of hydrogenase from Clostridium pasteurianum –377 mV) and a similar measurement for the cyanobacterial enzyme is needed.

C. Physiology of Hydrogen Metabolism There is a fine review of the physiology and biochemistry of hydrogen metabolism published by Houchins in 1984. The enzyme nitrogenase reduces protons to produce hydrogen and this complicates the study of hydrogenase in nitrogen-fixing organisms. In general, the excretion of hydrogen from the cell is a very expensive process since much valuable energy is available from the oxidation of hydrogen. Some nitrogen fixers mitigate this loss by increasing their ‘uptake’ hydrogenase to recapture the hydrogen and oxidize it to produce ATP. Work in this area has been thoroughly reviewed (Scherer et al., 1988). This special case of hydrogen cycling here is consigned to the process ofnitrogen fixation; attention will be focused more on hydrogen metabolism in organisms that are not fixing nitrogen in the hope that this simplification will aid clarity. Firstly, consider the process ofhydrogen production by cyanobacteria. Many cyanobacteria, when incubated in dark, anaerobic conditions, will begin to produce hydrogen. Induction of hydrogen production varies from one to 48 hours. Hydrogen production is usually low—perhaps a micromole (mg dry weight of Loading the cells with carbohydrate reserves may increase the rate of dark hydrogen production (Almon and Böger, 1988). The paths of carbohydrate transformation in cyanobacteria that allow this hydrogen production are being examined (Heyer et al., 1989; Van der Oost et al., 1989; Heyer and Krumbein, 1991). There is interesting evidence that such fermentation occurs in nature. Zohary (1989) has studied the hyperscums of Microcystis aeruginosa that accumulate to a depth of one meter on the Heartbeespoort Dam reservoir in South Africa. Buoyancy compresses colonies at the surface into a very thick paste at a concentration of 100 mg chlorophyll per milliliter. The cells remain

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viable for up to three months and resume photosynthesis after one day’s acclimatization in dilute suspension in moderate light. Cells in the hyperscum lack light and oxygen. Trapped gas bubbles contain traces of hydrogen, and up to 28% methane which is likely to be produced by methanogenic bacteria using the hydrogen produced by the Microcystis aeruginosa. Acetic acid accumulates in the hyperscum to a concentration of 1 mM and is a likely product of the fermentation. While the hyperscum provides dramatic clues for fermentation by cyanobacteria in nature, it is a rare event and so hardly justifies the long persistence of this survival mechanism. However, many cyanobacteria and microscopic algae in the temperate zone pass the winter buried in sediment without light or oxygen. Thus, fermentation by these organisms insures winter survival. As suggested by Van der Oost et al. (1987), the physiological function of the reversible hydrogenase is the disposal of excess reduction equivalents under anaerobic conditions. The immediate biochemical value of the fermentation is documented by several studies (Mahro et al., 1986; Kumazawa et al., 1987) that correlate hydrogen evolution with maintenance of higher ATP levels during dark, anaerobic incubation. Light-driven hydrogen production by cyanobacteria and algae is also observed. A dark, anaerobic adaptation period is usually required. Illumination accelerates the slow, dark hydrogen production. There is simultaneous oxygen production and this may eventually inactivate the hydrogenase since often the light-stimulated hydrogenproduction is not sustained. A striking exception is found in the report of Mitsui and Kumazawa (1977), who described a marine cyanobacterium (strain Miami BG7) that sustained photohydrogen production for several days. Kuwada et al. (1988) have isolated a nonheterocystous Lyngbya sp. with a high rate of photohydrogen production that persists for five days. In both cases, hydrogen production was achieved by deprivation of nitrogen and was suppressed by addition of fixed nitrogen. This makes it likely that the nitrogenase is one source of hydrogen. In the Lyngbya sp. experiments, cessation of hydrogen production occurred when the cell glycogen level was nearly exhausted. Recently, Kuwada et al. (1991) have found that when the Lyngbya sp. are switched from growing conditions through a stage for nitrogenase induction to hydrogen producing conditions, PS II was

inactivated and some of its proteins degraded. Thus, the cells degrade carbohydrate reserves, pass the electrons through PS I and deliver them to the reversible hydrogenase and/or nitrogenase and excrete them as hydrogen. The rapid excretion of such an energy-rich product seems undesirable in the normal course ofevents so this process may not be normal to the cells. A process that resembles the foregoing is seen in the oxidation of sulfide by Oscillatoria limnetica (Belkin and Padan, 1978; Sybesma et al., 1986). Electrons from sulfide are delivered to PS I which drives them to a low potential where they join protons to form hydrogen. Shahak et al. (1992) have characterized the sulfide-quinone reductase that initiates this sequence. Laczkó (1986) has described another type of photohydrogen evolution in cyanobacteria that involves both photosystems. Anabaena cylindrica was grown with ammonia to eliminate nitrogenase and at high light intensity to build up carbohydrate reserves. After a two-hour anaerobic incubation, the cells evolved both oxygen and hydrogen when illuminated. The hydrogen production was inhibited by DCMU and proceeded at a higher rate in light absorbed by phycocyanin than in light absorbed by chlorophyll. PS II was clearly involved. Laczkó (1986) suggests that the high-light-grown cells excrete part of the photosynthetically produced excess electrons via the reversible hydrogenase activity. Cyanobacteria consume hydrogen in laboratory experiments. Eisbrenner et al. (1981) recognized two distinct pathways of hydrogen consumption. One pathway is the oxyhydrogen reaction — an oxygendependent hydrogen uptake process that may proceed through the respiratory chain but can proceed at a rate that is considerably higher than the rate of respiration. The second pathway is a light-dependent, DCMU-sensitive consumption of hydrogen with concomitant reduction of Hydrogen uptake in the light can be enhanced by nitrate and nitrite (Houchins and Burris, 1981a). These reactions will be better understood when more biochemical details are added to their description. Perhaps they reflect the extraordinary value of hydrogen as a rich source of energy for the cell. The competition for hydrogen is fierce in mixed populations ofanaerobes in nature. Cyanobacteria do well by recycling all the hydrogen they can and releasing as little as possible from the cell. In addition the uptake mechanisms may reflect a need by the cell to purge itself of oxygen.

Chapter 12 Soluble Electron Transfer Catalysts of Cyanobacteria

D. A Speculative Model of Hydrogen Metabolism in Cyanobacteria In seeking a function for the low-potential cytochrome c of cyanobacteria, one realized that the only other low-potential cytochrome c found to date is the cytochrome of Desulfovibrio sp. These are anaerobic heterotrophs that either reduce sulfate or evolve hydrogen to dispose of excess reducing power. The low-potential cytochrome donates electrons to hydrogenase in a reaction more easily measured than the one between the analogous cyanobacterial catalysts. However, there is a growing convergence of similarities between the facts of cyanobacterial hydrogen metabolism and the model developed to describe similar processes in Desulfovibrio sp. Odom and Peck (1984) have developed and extensively documented an elegant model to describe the hydrogen metabolism of the Desulfovibrio sp. In the cytoplasm of the cell, fermentation converts carbohydrate into acetate, and hydrogen. The

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hydrogen diffuses out to the periplasm where a hydrogenase may capture it and strip the electrons away from the protons if an electron sink is available. Low-potential cytochrome is also present in the periplasm to mediate electron transfer. Sulfate is an electron sink and the electrons from the periplasmic hydrogenase may be directed to it and reduce the sulfate to sulfide. Nitrate reduction to ammonia is another mechanism of electron utilization. Protons are left to accumulate in the periplasm. These protons form a gradient which is used to drive ATP synthesis by coupling factors that protrude from the cytoplasmic membrane into the cytoplasm. The diagram shown in Fig. 3 combines the Desulfovibrio sp. model of Odom and Peck (1984) with some of the data from cyanobacteria on hydrogenase and the low-potential cytochrome. In cyanobacteria, it is plausible but not yet established, that the hydrogen-producing fermentation occurs in the cytoplasm. Kentemich et al. (1991) have located a powerful reversible hydrogenase on the periplasmic

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face of the cytoplasmic membrane and low-potential cytochrome c is available in the periplasm, possibly to assistin electron transfertothe sinks. Houkins and Burris (1981b) observed stimulation of light-driven hydrogen uptake in Anabaena sp. strain PCC 7120 bynitrate,nitrite andsulfate. Peschek (1988a; 1988b) has established the presence of coupling factors in the cyanobacterial membrane and a respiratory chain there as well which could participate in the oxyhydrogen reaction. Onemustgobeyond theDesulfovibriosp.analogy to explain light-driven hydrogen consumption and production. The uptake hydrogenase on the thylakoid membrane might couple through the membranebound, low-potential cytochrome c to PS I. Photohydrogen production might proceed by conventional photolysis of water generating reduced ferredoxinwhich couldbecoupled toahydrogenase. Many more experiments are needed to confirm or deny these possibilities. The diagram in Fig. 3 illustrates some of these possibilities for hydrogen metabolism in cyanobacteria. References Alam J, Sprinkle J, Hermodson MA and Krogmann DW (1984) Characterization of cytochrome from cyanobacteria. Biochim Biophys Acta 766: 317–321 Alam J, Whitaker RA, Krogmann DW and Curtis S (1986) Isolation and sequence of the gene for ferredoxin I from the cyanobacterium Anabaena sp. strain PCC7120. J Bacteriol 168: 1265–1271 Almon H and Böger P (1984) Nickel-dependent uptakehydrogenase activity in the blue-green algaAnabaena variabilis. Z Naturforsch 39c: 90–92 Almon H and Böger P (1988) Hydrogen metabolism of the unicellular cyanobacterium Chroococcidiopsis thermalis ATCC 29380. FEMS Microbiol Lett 49: 445–449 Apley EC and Wagner R (1988) Chemical modification of the active site of reductase and conformation of the binary reductase complex in solution. Biochim Biophys Acta 936: 269–279 Asada Y, Miyake M and Tomizuka N (1992) Hydrogenase and mediated hydrogen metabolism in some cyanobacteria. Photosynth Res 34: 128 Asada Y, Kawamura S and Ho KK (1987) Hydrogenase from the unicellular cyanobacterium, Microcystis aeruginosa. Phytochem 26: 637–640 Bagley S, Driscoll PC, Goodall KG, Redfield C and Hill HAO (1990) The complex formed between plastocyanin and cytochrome c: investigation by NMR spectroscopy. Eur J Biochem 188: 413–420 Belkin S and Padan E (1978) Sulfide dependent hydrogen evolution in the cyanobacterium Oscillatoria limnetica. FEBS Lett 94: 291–294 Belkin S, Rao KK and Hall DO (1981) Partial purification and

characterization of two hydrogenase forms from Oscillatoria limnetica. Biochem Int 3: 301–309 Böhme H and Haselkorn R (1988) Molecular cloning and nucleotide sequence analysis of the gene coding for heterocyst ferredoxin from the cyanobacterium Anabaena PCC 7120. Mol Gen Genet 214: 278–285 Böhme H and Haselkorn R (1989) Expression of Anabaena ferredoxin genes in E. coli. Plant Mol Biol 12: 667–672 Böhme H and Schrautemeier B (1987) Comparative characterization of ferredoxins from heterocysts and vegetative cells of Anabaena variabilis. Biochim Biophys Acta 891: 1–7 Bottin H and Lagoutte B (1992) Ferredoxin and flavodoxin from the cyanobacterium Synechocystis sp PCC 6803. Biochim Biophys Acta 1101: 48–56 Bovy A, de Vrieze G, Borrias M and Weisbeek P (1992) Isolation and sequence analysis of a gene encoding a basic cytochrome from the cyanobacterium Anabaena sp. PCC 7937. Plant Mol Biol 19: 491–492 Bovy A, de Vrieze G, Borrias M and Weisbeek P (1992) Transcriptional regulation of the plastocyanin and cytochrome genes from the cyanobacterium Anabaena sp. PCC 7937. Mol Microbiol 6: 1507–1513 Bowes JM, Stewart AC and Bendall DS (1983) Purification of Photosystem II particles from Phormidium laminosum using the detergent Biochim Biophys Acta 725: 210–219 Briggs LM, Pecoraro VL and McIntosh L (1990) Copper induced expression, cloning and regulatory studies of the plastocyanin gene from the cyanobacterium Synechocystis sp. PCC 6803. Plant Mol Biol 15: 633–642 Cammack R, Rao K, Bargeron CP, Hutson KG, Andrew PW and Rogers LJ (1977) Midpoint potentials of plant and algal ferredoxins. Biochem J 168: 205–209 Carillo N, Ceccarelli EA, Krapp AR, Boggio S, Ferreya RG and Viale AM (1992) Assembly of plant oxidoreductase in Escherichia coli requires GroE molecular chaperones. J Biol Chem 267: 15537–15541 Christensen HEM, Conrad LS and Ulstrup J (1990) A new procedure for the fast isolation and purification of plastocyanin from the cyanobacterium Anabaena variabilis. Photosynth Res 25: 72–76 Christensen HEM, Conrad LS, Mikkelsen, KU and Ulstrup (1992) Electron transport routes in native and modified plant and cyanobacterial plastocyanins. Photosynth Res 34: 154 Conn CL, Sprinkle JR, Alam J, Hermodson M, Meyer T and Krogmann DW (1989) The amino acid sequence of the lowpotential cytochrome from the cyanobacterium Microcystis aeruginosa. Arch Biochem Biophys 270: 227–235 Collyer CA, Guss JM, Sugimura Y, Yoshizaki F and Freeman HC (1990) Crystal structure of plastocyanin from a green alga Enteromorpha prolifera. J Mol Biol 211: 617–632 Cozens AL and Walker JE (1987) The organization and sequence of the genes for ATP synthase subunits in the cyanobacterium Synechococcus 6301. Support for an endosymbiotic origin of chloroplasts. J Mol Biol 194: 359–383 Davis DJ, Krogmann DW and San Pietro A (1980) Electron donation to Photosystem I. Plant Physiol 65: 697–702 Davis DJ (1990) Tryptophan fluorescence studies of ferredoxinreductase indicate the presence of tryptophan in or near the ferredoxin binding site. Arch Biochem Biophys 276: 1–5

Chapter 12 Soluble Electron Transfer Catalysts of Cyanobacteria Dayday A, Mackerras AH and Smith GD (1985) The effect of nickel on hydrogen metabolism and nitrogen fixation in the cyanobacterium Anabaena cylindrica. J Gen Microbiol 131: 231–238 de Lorimier R, Bryant, DA and Stevens SE Jr (1990) Genetic analysis of a 9 kDa phycocyanin-associated linker polypeptide. Biochim Biophys Acta 1019: 29–41 Eisbrenner G, Roos P and Bothe H (1981) The number of hydrogenases in cyanobacteria. J Gen Microbiol 125:383–390 Evans P and Krogmann DW (1983) Three c-type cytochromes from the red alga Porphyridium cruentum. Arch Biochem Biophys 227: 494–510 Ewart GD, Reed KC and Smith GD (1990) Soluble hydrogenase of Anabaena cylindrica. Cloning and sequencing of a potential gene encoding the tritium exchange subunit. Eur J Biochem 187:215–223 Ewart GE and Smith GD (1989) Purification and properties of soluble hydrogenase from the cyanobacterium Anabaena cylindrica. Arch Biochem Biophys 268: 327-337 Fillat MF, Borrias WE and Weisbeck PJ (1991) Isolation and overexpression in Escherichia coli of the flavodoxin gene for Anabaena PCC 7119. Biochem J 280: 187–191 Fillat MF, Sandman G and Gomez-Moreno C (1988) Flavodoxin from the cyanobacterium Anabaena PCC 7119. Arch Microbiol 150:160–164 Fillat MF, Edmondson DE and Gomez-Moreno C (1990a) Structural and chemical properties of a flavodoxin from Anabaena PCC 7119. Biochim Biophys Acta 1040: 301–307 Fillat MF, Bakker HAC and Weisbeck PJ (1990b) Sequence of the FNR gene from Anabaena PCC 7119. Nucl Acids Res 18: 7161 Fillat MF, Edmondson DE and Gomez-Moreno C (1991) Light dependent deactivation/reactivation of Anabaena variabilis ferredoxin–NADP reductase. Biochem J 274: 781–786 Fitzgerald MP, Rogers LJ, Rao KK and Hall DO (1980) Efficiencies of ferredoxins and flavodoxins as mediators in systems for hydrogen evolution. Biochem J 192: 665–672 Fitzgerald MP, Hussain A and Rogers LJ (1978) A constitutive flavodoxin from a eukaryotic alga. Biochem Biophys Res Commun 81: 630–635 Fukuyama K, Wakabayashi S, Matsubara H and Rogers LJ (1990) Tertiary structure of oxidized flavodoxin from an eukaryotic red alga Chondrus crispus at 2.35 Angstrom resolution – localization of charged residues and implications for interaction with electron transport. J Biol Chem 265: 15804–15812 Fukuyama K, Matsubara H and Rogers LJ (1992) Crystal structure of the oxidized flavodoxin from a red alga Chondrus crispus refined at 1.8 Ångstrom resolution – description of the flavin mononucleotide binding site. J Mol Biol 225: 775–789 Guss JM, Harrowell PR, Murata M, Norris VA and Freeman HC (1986) Crystal structure analysis of reduced poplar plastocyanin at six pH values. J Mol Biol 192: 361–387 Hall DO, Rao KK and Cammack R (1971) A stable and easily extractable plant-type ferredoxin from the blue-green alga Spirulina maxima. Biochim Biophys Res Commun 47: 798– 802 Hallenbeck PC and Beneman JR (1978) Characterization and partial purification of the reversible hydrogenase of Anabaena cylindrica. FEBS Lett 94: 261-264 Hase T, Mawtani S and Mukohata Y (1991) Expression of maize

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ferredoxin cDNA in Escherichia coli. Plant Physiol 97:1395– 1401 Hase T, Masubara H, Hutber GN and Rogers LJ (1982) Amino acid sequences of Nostoc strain MAC ferredoxins I and II. J Biochem 92: 1347–1355 Hase T, Kimata Y, Yonekura K, Matsumura T and Sakakibara H (1990) Molecular cloning and differential expression of the maize ferredoxin gene family. Plant Physiol 96: 77–83 He S, Modi S, Bendall DS and Gray JC (1991) The surface exposed tyrosine residue Tyr 83 of pea plastocyanin is involved in both binding and electron transfer reactions with cytochrome f. EMBO J 10: 4011–4016 Heyer H, Stal L and Krumbein WE (1989) Simultaneous heterolactic and acetate fermentation in the marine cyanobacterium Oscillatori limnosa incubated anaerobically in the dark. Arch Microbiol 151: 558–564 Heyer H and Krumbein WE (1991) Excretion of fermentation products in dark and anaerobically incubated cyanobacteria. Arch Microbiol 155: 284–287 Hibino T, de Boer AD, Weisbeek PJ and Takabe T (1991) Reconstitution of mature plastocyanin from precursor apoplastocyanin expressed in E. coli. Biochim Biophys Acta 1058: 107–112 Ho KK and Krogmann DW (1982) Photosynthesis. In: Carr NG and Whitton BA (eds) The Biology of Cyanobacteria, pp 191– 214. Blackwell Scientific Publications, Oxford Ho KK and Krogmann DW (1984) Electron donors to P700 in cyanobacteria and algae, an instance of unusual genetic variability. Biochim Biophys Acta 766: 310–316 Hodges M, Miginiac-Maslow M, Le Marechal P and Rémy R (1990) The ATP dependent post translational modification of reductase. Biochim Biophys Acta 1052: 446–152 Hoganson CW, Lagenfelt G and Andreasson LE (1990) EPR and redox potentiometric studies of cytochrome of Anacystis nidulans. Biochim Biophys Acta 1016: 203–206 Holton RW and Myers J (1963) Cytochromes of a blue-green alga: extraction of a c-type with a strongly negative redox potential. Science 142: 234–235 Holton RW and Myers J (1967a) Water soluble cytochromes from a blue-green alga. I. Extraction, purification, and spectral properties of cytochromes c and Anacystis nidulans) Biochim Biophys Acta 131: 362–374 Holton RW and Myers J (1967b) Water soluble cytochromes from a blue-green alga. II. Physicochemical properties and quantitative relationships of cytochrome and Anacystis nidulans). Biochim Biophys Acta 131: 375–384 Houchins JP (1984) The physiology and biochemistry of hydrogen metabolism in cyanobacteria. Biochim Biophys Acta 768: 227–255 Houchins JP and Burris RH (1981) Occurrence and localization of two distinct forms of hydrogenase in the heterocystous cyanobacterium Anabaena sp. 7120. J Bacteriol 146:209–214 Houchins JP and Burris RH (198la) Light and dark reactions of the uptake hydrogenase in Anabaena 7120. Plant Physiol 68: 712–716 Houchins JP and Burris RH (1981b) Physiological reactions of the reversible hydrogenase from Anabaena 7120. Plant Physiol 68:717–721 Hutber GN, Smith AJ and Rogers LJ (1978) Comparative biological activities of two ferredoxins and a flavodoxin from

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the cyanobacterium Nostoc strain MAC. FEMS Microbiol Lett 4: 11–14 Hutson KG, Rogers LJ, Haslett BG, Boulter D and Cammack R (1978) Comparative studies on two ferredoxins from the cyanobacterium Nostoc strain MAC. Biochem J 172: 465–477 Jacobson BL, Chae YK, Böhme H, Markley JL and Holden HM (1992) Crystallization and preliminary analysis of oxidized, recombinant heterocyst [2Fe-2S] ferredoxin from Anabaena 7120. Arch Biochem Biophys 294: 279–281 Karplus PA, Daniels MJ and Herriott JR (1991) Atomic structure of reductase: prototype for a structurally novel flavoenzyme family. Science 251: 60–66 Kentemich T, Bahnweg M, Mayer F and Bothe H (1989) Localization of the reversible hydrogenase in cyanobacteria. Z Naturforsch 44C: 384–391 Kimata Y and Hase T (1988) Localization of ferredoxin isoproteins in mesophyll and bundle sheath cells in maize. Plant Physiol 89: 1193–1197 Knaff DB and Hirasawa M (1991) Ferredoxin-dependent chloroplast enzymes. Biochim Biophys Acta 1056: 93–125 Krinner M, Hauska G, Hurt E and Lockau W (1982) A cytochrome with plastoquinone-cytochrome c oxidoreductase activity from Anabaena variabilis. Biochim Biophys Acta 681: 110–117 Krogmann DW (1991) The low-potential cytochrome c of cyanobacteria and algae. Biochim Biophys Acta 1058: 35–37 Kumazawa S, Skjoldal HR and Mitsui A (1987) Dark hydrogen evolution and adenine nucleotides of subtropical marine unicellular green algae during anaerobic incubation. Plant Cell Physiol 28: 653–661 Kuwada Y, Inoue Y, Koike H and Ohta Y (1991) Functional and structural changes of PS II of Lyngbya sp. under hydrogenproducing conditions. Agric Biol Chem 55: 299–305 Kuwada Y, Nakatsukasas M and Ohta Y (1988) Isolation and characterization of a nonheterocystous cyanobacterium, Lyngbya sp. isolate no. 108, for large quantity hydrogen production. Agric Biol Chem 55: 299–305 Laczkó I (1986) Appearance of a reversible hydrogenase activity in Anabaena cylindrica grown in high light. Physiol Plant 67: 634–637 Laudenbach DE, Herbert SK, McDowell C, Fork DC, Grossman AR and Straus NA (1990) Cytochrome is not required for photosynthetic activity in the cyanobacterium Synechococcus. Plant Cell 2: 913–924 Laudenbach DE, Reith ME and Straus NA (1988) Isolation, sequence analysis, and transcriptional studies of the flavodoxin gene from Anacystis nidulans R2. J Bacteriol 170: 258–265 Lee IS, Hase T, Matsubara H, Ho KK and Krogmann DW (1983) Amino acid sequence of ferredoxin I from Aphanizomenon flos–aquae. Biochim Biophys Acta 744: 53–56 Leonhardt KG and Straus NA (1989) Sequence of the flavodoxin gene from Anabaena variabilis. Nucl Acids Res 17: 4384 Leonhardt KG and Straus NA (1992) The iron stress operon involved in photosynthetic electron transport in the marine cyanobacterium Synechococcus SpPCC-7002. J Gen Microbiol 138: 1613–1621 Li HH and Merchant S (1992) Two metal-dependent steps in the biosynthesis of Scenedesmus obliquus plastocyanin. J Biol Chem 267: 9368–9375 Ludwig ML, Pattridge KA, Powers TB, Dickerson RE and

Takano T (1983) Structure analysis of ferricytochrome c from the cyanobacterium Anacystis nidulans. In: Hu C and Eaton WA (eds) Electron Transport and Oxygen Evolution, pp. 27– 32. Elsevier Scientific Publishers, Amsterdam Mahro B, Küsel, AC and Grimme LH (1986) The significance of hydrogenase activity for the energy metabolism ofgreen algae: anaerobiosis favors ATP synthesis in cells of Chlorella with active hydrogenase. Arch Microbiol 144: 91–95 Malakhov MP, Wada H, Los DA and Murata N (1992) A new cytochrome gene of Synechococcus PCC 6803. Photosynth Res 43: 134 Martinez SE, Szczepaniak A, Huang D, Smith JL and Cramer WA (1992) Crystallographic studies of the lumen-side domain of turnip cytochrome f. Photosynth Res 43: 152 Marques S, Florencio FJ, and Candu P (1992) Purification of the ferredoxin–glutamate synthase from the unicellular cyanobacterium Synechococcus sp PCC 6301. Eur J Biochem 206: 69–77 Matsuura K and Fujita Y (1986) Chromatic regulation of cytochrome composition and electron transfer from the cytochrome to Photosystem I in Anacystis nidulans Tx 20. Plant Cell Physiol 27: 685–691 Medina M, Peleato ML, Mendez E and Gomez-Moreno C (1992) Identification of specific carboxyl groups on Anabaena PCC 7119 flavodoxin which are involved in the interaction with reductase. Eur J Biochem 203: 373–379 Medina M, Mendez E and Gomez-Moreno C (1992) Identification of argenyl residues involved in the binding of ferredoxinNADP from Anabaena sp. PCC 7119 to its substrates. Arch Biochem Biophys 299: 281–286 Medina M, Mendez E and Gomez-Moreno C (1992) Lysine residues of reductase from Anabaena sp 7119 involved in substrate binding. FEBS Lett 298: 25–28 Merchant S and Bogorad L (1986) Regulation by copper of the expression of plastocyanin and cytochrome in Chlamydomonas reinhardtii. Mol Cell Biol 6: 462–469 Mitsui A and Arnon D (1971) Crystalline ferredoxin from a bluegreen alga, Nostoc sp. Physiol Plant 25: 135–140 Mitsui A and Kumazawa S (1977) Hydrogen production by marine photosynthetic organisms as a potential energy source. In: Mitsui A, Miyachi S, San Pietro A and Tamura S (eds) Biosolar Energy Conversion, pp 23–51. Academic Press, New York Modi S, He S, Grey JC and Bendall DS (1992) The role of surface exposed Tyr 83 of plastocyanin in electron transfer from cytochrome c. Biochim Biophys Acta 1101: 64–68 Modi S, Nordling M, Lundberg LG, Hanson O, and Bendall DS (1992) Reactivity of cytochrome c and cytochrome f with mutant forms of spinach plastocyanin. Biochim Biophys Acta 1102: 85–90 Modi S, McLaughlin E, Bendall DS, He S and Gray JC (1993) The binding site of plastocyanin and role of Tyr-83 in electron transfer with cytochrome f. Photosynth Res 34: 153 Moore JM, Case DA, Chazin WJ, Gippert GP, Havel TF, Powels R and Wright PE (1988) The three dimensional solution structure of plastocyanin from the green alga Scenedesmus obliquus. Science 240: 314–317 Morand LZ, Frame MK, Covlert KK, Johnson DA, Krogmann DW and Davis DJ (1989) Plastocyanin cytochromefinteraction. Biochemistry 28: 8039–8047

Chapter 12 Soluble Electron Transfer Catalysts of Cyanobacteria Morand LZ and Krogmann DW (1993) Large scale preparation of pure plastocyanin from spinach. Biochim Biophys Acta 1141: 105–106 Mulligan ME, Buikema WJ and Hasclkorn R (1988) Bacterialtype ferredoxin genes in the nitrogen fixation regions of the cyanobacterium Anabaena sp strain PCC 7120 and Rhizobium meliloti. J Bacteriol 170: 4406–4410 Odom JM and Peck HD (1984) Hydrogenase, electron transfer proteins and energy coupling in the sulfate reducing bacterium Desulfovibrio. Ann Rev Microbiol 38: 551–592 Oh BH and Markley JL (1990) Multinuclear resonance studies of the two iron-two sulfur ferredoxin from Anabaena species strain PCC 7120. (a series of 3 papers). Biochemistry 29: 3993–4017 Omata T and Murata N (1984) Cytochromes and prenylquinones in preparations of cytoplasmic and thylakoid membranes from the cyanobacterium (blue-green alga) Anacystis nidulans. Biochim Biophys Acta 766: 395–402 Pardo MB, Gomez-Moreno C and Peleato ML (1989) Effects of iron deficiency on ferredoxin levels in Anabaena variabilis PCC 6309. Arch Microbiol 153: 528–530 Paulsen KE, Stankovich MT, Stockman BJ and Markley JL (1990) Redox and spectral properties of flavodoxin from Anabaena 7120. Arch Biochem Biophys 280: 68–73 Peschek GA, Nitschmann WH and Czerny T (1988) Respiratory proton exclusion and plasma membrane energization. Meth Enzymol 167: 361–380 Peschek GA, Molitor V, Trnka M, Wastyn M and Eber W (1988) Characterization of cytochrome c oxidase in isolated and purified plasma and thylakoid membranes from cyanobacteria. Meth Enzymol 167: 437–448 Przybyla AE, Robbins J, Menon N and Peck HD (1992) Structurefunction relationships among nickel-containing hydrogenases. FEMS Microbiol Rev 88: 109–136 Pueyo, JJ and Gomez-Moreno, C (1991) Characterization of the cross-linked complex formed between reductase and flavodoxin from Anabaena PCC 7119. Biochim Biophys Acta 1059: 149–156 Pueyo JJ, Revilla C, Mathew SG and Gomez-Moreno C (1992) Complex formation between ferredoxin and ferredoxin-NADP reductase from Anabaena PCC 7119: crosslinking studies. Arch Biochem Biophys 294: 367–372 Pulich W (1977) Cytochrome in Nostoc sp. (Cyanophyceae): an electron acceptor from reduced NADP in the dark. J Phycol 13:40–45 Puyeo JJ and Gomez-Moreno C (1991) Purification of ferredoxinNADP reductase, flavodoxin and ferredoxin from a single batch of the cyanobacterium Anabaena PCC 7119. Prep Biochem 21: 191–204 Rao KK and Hall DO (1988) Hydrogenases: isolation and assay. Meth Enzymol 167: 501–509 Redlinger T and Gantt E (1983) Photosynthetic membranes of Prophyridium cruentum. An analysis of chlorophyll-protein complexes and heme-binding proteins. Plant Physiol 73: 36– 40 Reeve JN and Beckler GS (1990) Conservation of primary structure in prokaryotic hydrogenases. FEMS Microbiol Rev 87:419–124 Reith ME, Laudenbach DE and Straus NA (1986) Isolation and nucleotide sequence analysts of the ferredoxin I gene from the

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bacterium Anabaena variabilis. FEBS Lett 184: 304–308 Schrautemeier B and Böhme H (1992) Coding sequence of a heterocyst ferredoxin gene (fdxH) isolated from the nitrogenfixing cyanobacterium Calothrix sp PCC 7601. Plant Mol Biol 18:1005–1006 Serre L, Medina M, Gomez-Moreno C, Fontecillacamps JC and Frey M (1991) Crystals of Anabaena PCC 7119 ferredoxinNADP reductase. J Mol Biol 218: 271–272 Shahak Y, Hauska G, Arieli B, Herrmann I, Taglicht D and Padan E (1992) Sulfide-quinone reductase (SQR) drives anoxygenic photosynthesis in prokaryotes Photosynth Res 34: 152 Shen J-R, Ikeuchi M and Inoue Y (1992) Stoichiometric association of extrinsic cytochrome and 12 kDa protein with a highly purified oxygen-evolving Photosystem II core complex from Synechococcus vulcanus. FEBS Lett 301: 145– 149 Shen J-R and Inoue Y (1992) Enhancement of evolution by extrinsic cytochrome and a 12 kDa protein in a purified PS II complex from a thermophilic cyanobacterium. Photosynth Res 34: 150 Shin M, Sukenobu M, Oshino R and Kitazume Y (1977) Two plant-type ferredoxins from a blue-green alga, Nostoc verrucosum. Biochim Biophys Acta 460: 85–93 Skjeldal L, Westler WM, and Markley JL (1990) Detection and characterization of hyperfine-shifted resonances in the proton NMR spectra of Anabaena 7120 ferredoxin. Arch Biochem Biophys 278: 482–485 Smillie RM (1965) Isolation of two proteins with chloroplast ferredoxin activity from blue-green algae. Biochem Biophys Res Commun 20:621–629 Smith ET, Ensign SA, Ludden PW and Feinberg BA (1992) Direct electrochemical studies of hydrogenase and carbon monoxide dehydrogenase. Biochem J 285: 181–185 Smith WW, Patridge KA, Ludwig ML, Petsko GA, Tsernoglov D, Tanaka M and Yasunobu KT (1983) Structure of oxidized flavodoxin from Anacystis nidulans. J Mol Biol 165: 737–755 Stockman BJ, Krezel AM, Markley JL, Leonhardt KG and Strauss NA (1990) Hydrogen-1, carbon-13 and nitrogen-15 NMR spectroscopy of Anabaena 7120 flavodoxin: assignment of b-sheet and flavin binding site resonances and analysis of protein-flavin interactions. Biochemistry 29: 9600–9609 Susor WA and Krogmann DW (1966) Triphosphopyridine nucleotide photoreduction with cell-free preparations of Anabaena variabilis. Biochim Biophys Acta 120: 65–72 Sybesma C, Schowanek D, Sloten L and Walravens N (1986) Anoxygenic photosynthetic hydrogen production and electron transport in the cyanobacterium Oscillatoria limnetica. Photosynth Res 9: 149–158 Tan S and Ho KK (1989) Purification of an acidic plastocyanin from Microcystis aeruginosa. Biochim Biophys Acta 973: 111 – 117 Tan S and Ho KK (1993) Use of adsorption chromatography on Sephacryl S-500 for improved purification of cytochrome and ferredoxin from cyanobacteria. J Biochem Biophys Meth 8: 27–33 Tsukihara T, Fukuyama K, Nakamura M, Katsube Y, Tanaka N, Kakudo M, Wada K, Hase T and Matsubara H (1981) X-ray analysis of a [2Fe-2S] ferredoxin from Spirulina platensis. Main chain fold and location of side chains at 2.5 Å resolution. J Biochem 90: 1763–1773

Tsukihara T, Fukuyama K, Mizushima M, Harioka T, Kusonoki M, Katsube Y, Hase T and Matsubara H (1990) Structure of the [2Fe-2S] ferredoxin I from the blue-green alga Aphanothece sacrum at 2.2Å resolution. J Mol Biol 216:399–410 Van der Oost J, van Walraven HS, Bogerd J, Smit AB, Ewart GD and Smith GD (1989) Nucleotide sequence of the gene proposed to encode the small subunit of the soluble hydrogenase of the thermophilic unicellular cyanobacterium Synechococcus PCC 6716. Nucl Acid Res 17: 10098 Van der Oost J, Bulthuis BA, Feitz S, Krab K and Kraayenhof R (1989) Fermentation metabolism of the unicellular cyanobacterium Cyanothece PCC 7822. Arch Microbiol 152: 415– 419 Van der Oost J, Kannerworff WA, Krab K and Kraayenhof R (1987) Hydrogen metabolism of three unicellular nitrogenfixing cyanobacteria. FEMS Microbiol Lett 48: 41–45 Van der Plas J, de Groot RP, Woortman MR, Cremers F, Borrias M, van Arkel G, and Weisbeck P (1988) Genes encoding ferredoxins from Anabaena sp. PCC 7937 and Synechococcus sp. PCC 7942: Structure and regulation. Photosynth Res 18: 179–204 Van der Plas J, Bovy A, Kruyt F, de Vrieze G, Dassene E, Klein B and Weisbeck P (1989) The gene for the precursor of plastocyanin from the cyanobacterium Anabaena sp. PCC 7937. Isolation, sequence and regulation. Mol Microbiol 3: 275–284 Van der Plas J, de Groot RP, Woortman MR, Weisbeck PJ and van Arkel G (1986) Coding sequence of a ferredoxin gene from Anacystis nidulans R2 (Synechococcus PCC 7942). Nucl Acids Res 14: 7804 Van der Plas J, de Groot R, Woortman M, Cremers F., Borrias M, van Arkel G and Weisbeek, P (1988) Genes encoding ferredoxins from Anabaena sp PCC7937 and Synechococcus sp. PCC7942. Structure and regulation. Photosynth Res 18: 179–204 Wada K, Tamura T, Matsubara H and Kodo K (1983) Spirulina reductase. Further characterization with an improved preparation. J Biochem 94: 387–393 Wada K, Matsubara H, Chain RK and Arnon D (1981) A comparative study of the biological activities of two molecular species of chloroplast-type ferredoxins. Plant Cell Physiol 22: 275–281 Walker MC, Pueyo JJ, Gomez-Moreno C and Tollin G (1990) Comparison of kinetics of reduction and intramolecular electron transfer in electrostatic and covalent complexes of ferredoxinreductase and flavodoxin from Anabaena PCC 7119. Arch Biochem Biophys 281: 76–83 Widger W (1991) The cloning and sequencing of Synechococcus sp. PCC 7002 petCA operon: implications for the cytochrome binding domain of cytochrome f. Photosynth Res 30: 71–84 Wood RM (1978) Interchangeable copper and iron proteins in algal photosynthesis. Eur J Biochem 87: 9–19 Wynn RM, Omaha J and Malkin R (1986b) Structural and functional properties of the cyanobacterial Photosystem I complex. Biochemistry 28: 5554–5560 Wynn RM and Malkin R (1988) Interaction of plastocyanin with Photosystem I: a chemical cross-linking study of the polypeptide that binds plastocyanin. Biochemistry 27: 5863–5869 Yakunin AF, Chai K, Laurinavichene TU and Gogotov IN (1990) Purification and properties of two ferredoxins of the

Chapter 12 Soluble Electron Transfer Catalysts of Cyanobacteria nitrogen fixing heterocystic cyanobacterium Anabaena sphaerica. Biokhimiya (English translation) 55: 61–65 Yakunin AF, Chan VN and Gogotov IN (1991) Synthesis of two ferredoxins and nitrogenase by the heterocystic cyanobacterium Anabaena sphaerica. Biokhimiya (English translation) 60: 427–430 Yakunin AF, Hallenbeck PC, Troshina OY and Gogotov IN (1993) Purification and properties of a bacterial-type ferredoxin from the nitrogen-fixing cyanobacterium Anabaena variabilis ATCC 29413. Biochim Biophys Acta 1163: 124–130 Yao Y, Tamura T, Wada K, Matsubara H and Kodo K (1984) Spirulina reductase. The complete amino acid sequence. J Biochem 95: 1513–1516 Zanetti G, Morelli D, Ronchi S, Negri A, Aliverti A and Curti B (1988) Structural studies on the interaction between ferredoxin and oxidoreductase. Biochemistry 27: 3753–3759 Zhang L, McSpaddin B, Pakrasi HB and Whitmarsh J (1992)

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Chapter 13 Cyanobacterial Respiration G. Schmetterer Institute of Physical Chemistry, University of Vienna, Währingerstrasse 42, A-1090 Wien, Austria Summary I. Introduction II. Primary Electron Donors to the Respiratory Electron Transport Chain A. NADPH B. NADH C. D. Other Primary Electron Donors III. Primary Oxidoreductases A. NADPH Dehydrogenase(s) 1. Ferredoxin-NADP+ Oxidoreductase (FNR) 2. Other Membrane-Bound NADPH Dehydrogenase(s) B. NADH Dehydrogenases 1. Mitochondrial-Type NADH Dehydrogenase 2. Bacterial-Type NADH Dehydrogenase 3. Soluble NADH Dehydrogenase C. Hydrogenase(s) IV. Quinone Pool complex V. Cytochrome VI. Peripheral Intermediate Electron Carriers A. Cytochrome B. Plastocyanin C. Other Possible Peripheral Electron Carrier(s) VII. Terminal Oxidases A. Cytochrome c Oxidase, B. Cyanide-Sensitive Alternative Terminal Oxidase(s) C. Cyanide-Insensitive Alternative Terminal Oxidase(s) VIII. Conclusion A. Biological Function of Cyanobacterial Respiration B. Respiration and Photosynthesis C. Final Remarks Acknowledgments References

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Summary Cyanobacterial respiration is unique in several respects. Oxygenic photosynthesis and aerobic respiration are not separated in different organelles as in plants, but are active in the same compartment. Moreover, at least some cyanobacteria contain two distinct and complete respiratory chains, with one being found in each of their bioenergetically active membranes: the cell membrane and the intracellular membranes or thylakoids. In contrast, photosynthetic electron transport is generally present only in the intracellular membranes. Components not involved in respiration in other organisms are parts of the respiratory electron transport chain, such as NADPH, plastoquinone, and a chloroplast-type cytochrome complex. D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 409–435. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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Several structural genes for components of the respiratory chain have been cloned. Even in those cyanobacteria that contain two distinct respiratory chains, almost all genes are present in only one copy per chromosome. How one gene directs its gene product into two different membranes is an intriguing yet unanswered question. The amounts of several components of the respiratory chain are regulated by external conditions: e.g., cytochrome and plastocyanin by the concentration of and cytochrome oxidase by the ionic strength ofthe medium and (in heterocyst forming strains) by the availability of combined nitrogen. The elucidation of these mechanisms of gene regulation, largely by applying the methods of molecular biology to cyanobacteria, will be among the topics of the future studies in cyanobacterial respiration.

I. Introduction Cyanobacteria are by definition procaryotes capable of oxygenic photosynthesis. From this definition the overwhelming importance of bioenergetic processes in cyanobacteria is already evident. However, while cyanobacteria have become extremely useful for the elucidation structural and mechanistic details of oxygenic photosynthesis, much less attention has been devoted to the fact that these cells are unique in their bioenergetics in yet another way: they are the only cells capable of performing aerobic respiration and oxygenic photosynthesis in the same compartment. Thus, cyanobacteria are especially interesting in that they must regulate these two processes so as to avoid futile cycles. These would inevitably occur if both processes were equally active in the light. The stage is not yet set for a complete investigation of the interaction between photosynthesis and respiration; the number of gene products involved for neither process is known so far with certainty, and moreover, it may be said that in this respect respiration lags farbehind photosynthesis. This review will concentrate on all components thought to play a role in cyanobacterial respiratory electron transport, irrespective of whether such components are solely active in respiration or also part of the photosynthetic electron transport chain. In the latter case, only aspects concerning respiration or the interaction between respiration and photosynthesis will be highlighted, since details from the perspective of photosynthesis can be found in other chapters of this book. Several reviews have been published dealing exclusively with different aspects of cyanobacterial respiration. The older literature is covered in reviews Abbreviations: CM – cytoplasmic membrane; DBMIB – 2,5dibromo-3-methyl-6-isopropyl-p-benzoquinone; FNR – ferredoxin oxidoreductase; HQNO – 2-n-heptyl-4hydroxyquinoline-N-oxide; ICM – intracytoplasmic membranes.

by Binder (1982), and especially by Peschek (1984a, 1987). Two reviews by Scherer et al. (1988a) and Scherer (1990) concentrate on the interaction between photosynthetic and respiratory electron transport chains. The review by Matthijs and Lubberding (1988) is largely concerned with the localization of the respiratory electron transport chain. The function of respiration in the protection of nitrogenase from damage by has been reviewed quite recently (Fay 1992) and will therefore not be dealt with here. In cyanobacteria, ‘respiration’ has been used for several quite different processes so that a clear definition of what shall be understood by this term will be attempted. In cyanobacteria, a dedicated respiratory electron transport chain does not seem to exist; there is ample evidence that several gene products are used in both respiratory and photosynthetic electron transport pathways. Also, there is now significant evidence that the respiratory chain in cyanobacteria is branched, and it may turn out that only a single substance (possibly plastoquinone), but no chemical reaction, is common to all respiratory electron transport pathways. Therefore, no specific sequence of reactions can be said to make up the respiratory electron transport chain (as e.g. in mitochondria). Clearly, ‘respiration’ is a bioenergetic process, and therefore, respiratory electron transport must be coupled to the generation of ATP. Furthermore, it is essentially a membrane-bound process, although several components may be peripheral membrane proteins and thus soluble upon isolation ofthe membranes. In this review, therefore, ‘respiration’ is considered to be a membrane-bound electron transport process leading to the formation ofATP in the dark. Note that this definition does not contain any specific electron carrier, since neither the primary reductant, nor the terminal oxidant must always be the same. Although only one terminal oxidant, namely is currently known in cyanobacteria, the existence of other terminal oxidants cannot be excluded at present.

Chapter 13 Cyanobacterial Respiration According to the definition above, oxygen uptake in the dark is not necessarily due to respiration. The amino acid oxidations described by some authors (e.g., Pistorius et al. 1979, Barsky et al. 1985) are processes that reduce in the dark, but most likely are not respiratory, since there is no evidence that these reactions are coupled to ATP formation. The term ‘photorespiration’ for the production of glycolic acid by ribulose bisphosphate carboxylase-oxygenase is not well chosen, since this process is operative only in the light. A third process, namely the socalled ‘cyanide insensitive respiration,’ probably also carries its name without justification, because apparently it is also not coupled to ATP production (see Section VII C). A fourth example of possible misuse of the term respiration has been pointed out by Peschek (1984a): formation from prelabeled cells is not necessarily coupled to respiration, especially when the release is measured in the light (Scherer and Böger 1982, Scherer et al. 1982). Since respiration is a membrane-bound process, much attention has been devoted to the question: Which of the membranes in cyanobacteria contain respiratory electron transport chains? With one single exception (Gloeobacter violaceus, which contains no ICM (Rippka et al. 1974)), cyanobacteria contain three different membranes (Wolk 1973): 1. the outer membrane typical of gram-negative bacteria, which is not bioenergetically active and which will not be dealt with further here (see Chapter 6); 2. the cytoplasmic membrane (also called cell membrane, CM) and 3. the intracytoplasmic membranes (also called thylakoids, ICM). This review will assume that the latter two membranes are distinct, mainly as a result ofthe technique for the separation of the two membranes developed by Omata and Murata (1983) and in conjunction with their results on the different composition of the two membrane preparations (Omata and Murata 1984a, 1985). However, it should be emphasized that the relationship between the two membranes in vivo is far from clear. Electron microscopic studies by Nierzwicki-Bauer et al. (1983) on Synechococcus sp. strain PCC 7002 (Agmenellum quadruplicatum strain PR-6) showed that in this strain the ICM converge at several points close to the cell periphery and thus the CM. Unfortunately, their otherwise very detailed pictures do not allow one to decide whether actual membrane fusions exist between ICM and CM. Very interesting electron microscopic pictures of several different cyano-

411 bacteria (Anabaena cylindrica, Dermocarpa violaceae, Gloeocapsa alpicola, Pleurocapsa minor) have been published by Kunkel (1982). The ICM are shown to converge at distinct structures called thylakoid centers that lie near the cell periphery but are apparently not contiguous with the CM. Unfortunately, biochemical data on the nature of these thylakoid centers are not yet available. In contrast, Hinterstoisser et al. (1988) and Peschek et al. (1989a) produced evidence that the CM of Synechococcus sp. strain PCC 6301 contains precursors of chlorophyll synthesis but no chlorophyll. From these results one could conclude that an intimate connection between CM and ICM may exist. Inthin-section electron micrographspublished by Wildman and Bowen (1974), the thylakoids for some species, such as Arthrospira jenneri, can clearly be seen to arise as infoldings from the cytoplasmic membrane, but such evidence has rarely been obtained for other cyanobacterial species. Therefore, the final settlement of the question of the localization of the respiratory chain in cyanobacteria will have to be preceded by a clarification of the membrane topology in vivo. A note of caution seems necessary before going into further detail. Many different aspects of cyanobacterial respiration have unfortunately been investigated in a variety of different genera and strains and under different growth conditions. No cyanobacterium exists in which a complete picture is possible. It is not necessarily justified to extrapolate the results obtained in one strain to another, or to all cyanobacteria in general, and not very many comparative data exist. Only for lack of data, this review will construct a picture of cyanobacterial respiration from results assembled on a number of strains, and the reader is alerted to this currently necessary shortcoming. In view of the rather limited resources dedicated to cyanobacterial respiration (when compared to photosynthesis, for instance), it seems imperative to concentrate in the future on a few well-characterized strains that should be amenable to genetic manipulation. A major obstacle to presenting a complete and correct picture of cyanobacterial respiration is that there are too few data on respiration in intact cells. Since several components of the respiratory chain in cyanobacteria are peripheral membrane proteins (ferredoxin oxidoreductase, cytochrome c, plastocyanin) and easily lost during the preparation

412 of membranes, the reactions performed by such membrane preparations do not necessarily represent the in vivo situation. It seems that different membrane preparations contain variable amounts of these peripheral membrane proteins; although it is rarely done, it would be useful to measure the content of these proteins in the membrane preparations. Fig. 1 attempts to summarize all data on the components of cyanobacterial respiration obtained in the different strains investigated. Not all components exist with certainty and those of dubious status have been marked with a question mark. Arrows denote redox reactions. Arrowheads point to the oxidants being reduced by the reductant. Figure 1 gives only qualitative but no quantitative relations between the components and the reactions between them. It is certain that not all components of the respiratory chain are present in equimolar amounts (e. g., see Nanba and Katoh 1985). Furthermore, it should be kept in mind that not all reactions or components are present in all strains or cell types and there must be regulatory mechanisms that activate or deactivate certain reactions under different external conditions. In this respect Fig. 1 is certainly not complete, since there are some well-founded observations that cannot be explained within its framework. A striking example of this is the work by Smart et al. (1991) on mutants of Synechocystis sp. strain PCC 6803 lacking Photosystem I. Synechocystis sp. strain PCC 6803 is capable of chemoheterotrophic growth under so-called light-activated conditions (5 min of light per 24 h; Anderson and McIntosh 1991a). Using this capacity Smart et al. (1991) constructed a mutant lacking the psaA gene and thus a functional Photosystem I complex. This mutant was viable under light-activated heterotrophic growth conditions but was unable to grow in continuous light of The authors suggest that photoinhibition is responsible for this. Fig. 1, however, would qualitatively predict that growth of such a mutant in the light should be possible using Photosystem II, the quinone pool, the cytochrome complex (thus allowing ATP synthesis), and terminal oxidase(s). Indeed, Shen et al. (1993) were able to construct a Photosystem I minus mutant that can grow under continuous light even at but only after a lengthy period of adaptation, the olecular nature of which remains unknown.

G. Schmetterer II. Primary Electron Donors to the Respiratory Electron Transport Chain NADH is the primary electron donor for the respiratory chain in mitochondria. It was quite unusual, therefore, that early work on the respiratory chain in cyanobacteria indicated NADPH as the major donor (Biggins, 1969). In the meantime it has become clear that both NADH and NADPH are able to function as electron donors in cyanobacterial respiration. Additionally, can feed electrons into the respiratory chain via one or more hydrogenases.

A. NADPH It has long been known that the main source for NADPH in the dark is the oxidative pentose phosphate pathway (Cheung and Gibbs, 1966; Pelroy and Bassham, 1972, Pelroy et al., 1972, Winkenbach and Wolk, 1972; Wolk, 1973). An exogenous sugar, such as glucose or fructose, or more importantly glycogen, the major storage polysaccharide accumulated in the light in all cyanobacteria can serve as the source of the electrons (In heterocysts the primary source of reducing equivalents is still not known with certainty.). Early work indicating that NADPH might be the major donor for respiratory electron transport seemed to fit well with this fact. Especially the experiment performed by Biggins (1969) on intact cells of Synechococcus sp. strain PCC 6301 showed that NADPH and not NADH is important for respiratory activity. Biggins (1969) observed that on transition from aerobiosis to anaerobiosis the amount of NADPH in the cells increased (and and ATP decreased) while the amount of NADH remained unaltered. Concomitantly the ATP level of the cells decreased. There are two observations with mutants of Synechococcus sp. strains that corroborate Biggins’ hypothesis that NADPH is the primary reductant for respiration. One was described by Doolittle and Singer (1974). These workers isolated two mutants of Synechococcus sp. strain PCC 6301: One was lacking 6-phosphogluconate dehydrogenase, the second enzyme in the degradation of glucose by the oxidative pentose phosphate pathway; the second one was lacking this enzyme and additionally glucose 6-phosphate dehydrogenase, the first enzyme in the oxidative pentose phosphate pathway. The mutant lacking both enzymes is expected to have very little capacity to produce NADPH in the dark. Both mutants

Chapter 13 Cyanobacterial Respiration 413

G. Schmetterer

414 showed abnormally low respiratory uptake, and the single mutant (but not the double mutant) displayed a low survival rate in the dark. More recently, a similar experiment was performed by Broedel and Wolf (1990) with Synechococcus sp. strain PCC 7942. They cloned the gnd gene encoding 6-phosphogluconate dehydrogenase and insertionally inactivated the gene with the nptII gene of transposon Tn903. Their mutant had exactly the same phenotype as described by Doolittle and Singer (1974). Unfortunately, a detailed analysis of the respiratory capacities of this mutant was not performed. Although only performed on strains of one genus, the results mentioned above seem to indicate a special importance for NADPH in cyanobacterial respiration. Schluchter et al. (1993) showed that a mutant of Synechococcus sp. strain PCC 7002 lacking theNdhF subunit ofthe mitochrondrial-type NADH dehydrogenase displays low respiratory uptake. This does not necessarily contradict the postulated role for NADPH if the proposition by Howitt et al. (1993) that this enzyme oxidizes both NADH and NADPH in Anabaena sp. strain PCC 7120 is corroborated for other cyanobacteria. Still, the experiment by Biggins (1969) cannot be satisfactorily explained with the information currently available.

B. NADH Although it is now virtually certain that cyanobacteria oxidize not only NADPH but also NADH in the dark (see below), practically nothing is known about the dark reduction of The major source for NADH in mitochondria is undoubtedly the citric acid cycle, which has long been known to be incomplete in cyanobacteria, since dehydrogenase has never been found in any cyanobacterium (Smith et al., 1967, Pearce et al., 1969). Two apparent possibilities for NADH production thus are: 1. cyanobacteria may contain all enzymes necessary for the glycolytic pathway, but the amount of 6-phosphofructokinase seems to be low (Neuer and Bothe, 1982, Fewson et al., 1962, Pearce and Carr, 1969, Pelroy et al., 1972) so that the quantitative contribution ofthis pathway is not clear currently (In this connection it may be added that the results described by Peschek (1987), showing with the aid of positionally labeled exogenous and that the degradation of these sugars in the dark in several cyanobacteria follows different pathways, also has not found its final interpretation.);

2. transhydrogenase activity with NADPH and as substrates. Ferredoxin oxidoreductase possesses such an activity (Forti, 1971), but although the participation of this enzyme in cyanobacterial respiration has been investigated (Scherer et al., 1988b), the relevance of the transhydrogenase reaction for NADH driven electron transport remains obscure. However, Leach and Carr (1970) found negligible transhydrogenase activity in Anabaena variabilis, and no transhydrogenase activity could be detected in Synechococcus sp. strain PCC 7002 by the acetylpyridine adenine nucleotide assay (Schluchter et al., 1993).

C. Although exogenous molecular hydrogen is able to donate electrons to the respiratory chain in many different cyanobacteria, this reaction is probably of special importance in heterocysts. Since the reduction of by nitrogenase is inevitably accompanied by reduction of by the same enzyme, reoxidation of through the respiratory chain has two advantages: 1. the reducing power is not wasted but instead is used to generate ATP; 2. at the same time which irreversibly deactivates nitrogenase, is removed. This topic has quite recently been reviewed in detail (Fay, 1992).

D. Other Primary Electron Donors Few data exist on other primary electron donors for the respiratory chain of cyanobacteria. Biggins (1969) found a succinate-O2 oxidoreductase activity inhibitable by cyanide in membranes from Phormidium luridum. Succinate also supported low levels of oxidative phosphorylation in membranes from Anabaena variabilis (Leach and Carr, 1970). Peschek (1980a) showed that succinate is oxidized by membranes from Synechococcus sp. strain PCC 6301 through an HQNO- and cyanide-sensitive pathway. Succinate (and similarly NADH and NADPH) competed with H2 for a common acceptor in the respiratory chain. This common acceptor was probably the quinone pool, since Synechococcus sp. strain PCC 6301 membranes also catalyzed the reduction ofmenadione by succinate, H2, NADH, or NADPH (Peschek, 1980b). Omata and Murata (1985), however, detected no succinate-cytochrome c oxidoreductase in either CM or ICM of Synecho-

Chapter 13 Cyanobacterial Respiration coccus sp. strain PCC 6301. In Synechocystis sp. strain PCC 6714, low levels of succinate-cytochrome c oxidoreductase were detectable in the ICM but not in the CM (Omata and Murata, 1985).

III. Primary Oxidoreductases Enzymes oxidizing both NADH and NADPH are not common, and in cyanobacteria there is evidence for distinctive respiratory oxidoreductases for NADH and NADPH (see below). A membrane-bound NAD(P)H-quinone oxidoreductase from Synechocystis sp. strain PCC 6714, however, had a dehydrogenase activity ratio of 1.4 for NADPH/ NADH oxidation (Sandmann and Malkin 1983a). Since both the and were different for the two reduced pyridine nucleotides and there is evidence for distinctive membrane-bound NADPH and NADH dehydrogenases (see below), this enzyme preparation may have contained a mixture of both. However, Howitt et al. (1993) found in the CM of Anabaena sp. strain PCC 7120 an enzyme that cross-reacted with antibodies against mitochondrial NADH dehydrogenase, but could oxidize both NADH and NADPH. One purified soluble enzyme that was claimed to be an NAD(P)H oxidase from Microcystis aeruginosa (Viljoen et al., 1985) may actually have been FNR, even though these authors could not detect reduction of with reduced ferredoxin. No soluble NADPH dehydrogenase other than FNR was found by Scherer et al. (1988b) in Anabaena variabilis, and the enzyme characterized by Viljoen et al. (1985) has a high preference for NADPH over NADH (activity ratio of about 20 with 1, 4naphthoquinone as acceptor) and showed the isoforms characteristic of FNR (see below).

A. NADPH Dehydrogenase(s) The reactions participating in respiratory oxidation of NADPH have not yet been completely elucidated. There is evidence that one of the NADPH dehydrogenases active in respiratory electron transport is FNR (Scherer et al. 1988b). FNR is a peripheral membrane protein and easily lost from membrane preparations. This may at least partially account for some earlier results on respiratory NADPH oxidation. NADPH oxidation with by isolated membranes from different species was not inhibited, or was only partially inhibited, by cyanide [Phormidium luridum

415 (Biggins, 1969); Anabaena variabilis (Leach and Carr, 1970, Stürzl et al. 1984); heterocysts of Anabaena variabilis (Houchins and Hind, 1982); Synechococcus sp. strain PCC 6301 (Peschek, 1980a); Mastigocladus laminosus (Binder et al., 1984)]. Matthijs et al. (1984a, 1984b), however, demonstrated that the cyanide sensitivity of NADPH oxidation by in Plectonema boryanum depended on the preparation method: e.g., membranes isolated by sonication were inhibited about 80%, whereas membranes isolated by osmotic shock were only inhibited about 30%. This could reflect a different participation of CM and ICM in the reactions but could additionally be caused by different electron transport pathways made possible by disintegrating the cells to a different degree. Cyanide-insensitive NADPH oxidation by membrane preparations cannot yet be satisfactorily explained. Cyanide-sensitive NADPH oxidation, however, is a genuine respiratory reaction. This was demonstrated by Matthijs et al. (1984b) by showing that oxidation of NADPH coupled to reduction of was accompanied by oxidative phosphorylation in ‘leaky spheroplasts’ of Plectonema boryanum. A similar experiment was described Scherer et al. (1987) with Anabaena variabilis.

1.

Oxidoreductase (FNR)

FNR is present in chloroplasts and cyanobacteria. Its main function is undoubtedly the reduction of with electrons obtained from ferredoxin in noncyclic photosynthetic electron transport. There is now some evidence that FNR also acts as an NADPH dehydrogenase active in respiratory electron transport. If this is correct, a respiratory function of FNR in the ICM appears likely in view of its association with photosynthesis, but no information concerning a possible electron transfer from FNR to the quinone pool of the CM is available. FNR is a peripheral membrane protein and thus can be removed from membranes by washing. Schrautemeier et al. (1985) found that membranes from Anabaena variabilis heterocysts washed free of FNR had lost their ability to use NADPH as the electron donor to Photosystem I. Washing membranes from Anabaena variabilis also inhibited NADPH-dependent oxygen uptake (Alpes et al., 1985) and NADPH-dependent oxidative phosphorylation (Schrautemeier et al., 1988). FNR constituted the onlyNADPH-menadione oxidoreductase found in Anabaena variabilis. Thus,

G. Schmetterer

416 Schrautemeier et al. (1985) concluded that FNR is the respiratory NADPH dehydrogenase in respiratory electron transport. Antibodies generated against the purified FNR, however, were not able to inhibit NADPH-dependent uptake by isolated membranes (Scherer et al., 1988b), so that the possibility of another membrane-bound NADPH dehydrogenase remains. In addition to the reduction of by reduced ferredoxin FNR catalyses the oxidation of NADPH with concomitant reduction of several different substrates (Forti, 1971; Shin, 1971; Schrautemeier et al., 1985; Matthijs and Lubberding, 1988): 1. NADPH to ferricyanide or indophenol dyes (diaphorase activity). 2. Transhydrogenase activity (e.g., NADPH to or NADPH to 3. NADPH to cytochrome f. 4. NADPH to b-type cytochromes. 5. NADPH to cytochrome c (ferredoxindependent). 6. NADPH to plastocyanin. 7. NADPH to quinols. At present it is hard to assess which of these reactions have in vivo significance in cyanobacteria, since recent work has showed that although there is only one gene for FNR ( at least in Synechococcus sp. strain PCC 7002; Schluchter and Bryant, 1992), there are several forms of FNR that have not been fully characterized so far. Purified FNR from Anabaena variabilis displays several isoforms (Sancho et al., 1988; Scherer et al., 1988b). Schluchter and Bryant (1992) cloned the unique petH gene, encoding FNR, from Synechococcus sp. strain PCC 7002 and found that there is only one transcript from this gene. Thus, they concluded that the isoforms must have arisen post-translationally. Indeed their derived amino acid sequence is about 110 amino acids longer than would have been expected from previous work on the isolated protein from Spirulina sp. (Yao et al., 1984), Anabaena variabilis (Sancho et al., 1988) and the incomplete DNA sequence from Anabaena sp. strain PCC 7119 (Fillat et al., 1990; M. Fillat, personal communication). Since Scherer et al., (1988b) found two isoforms of 52 kDa and 34 kDa (and three separate isoforms of the 34-kDa species in non-denaturing polyacrylamide gel

electrophoresis), Schluchter and Bryant (1992) concluded that FNR may be easily proteolytically degraded. Besides this proteolysis (which may or may not have biological significance), there exist two other isoforms of FNR – one membrane-bound and one soluble as first demonstrated in chloroplasts (Carrillo and Vallejos, 1983, Matthijs et al., 1986). It was suggested that the soluble form might be active as a oxidoreductase and the membrane bound form as an NADPH-quinone oxidoreductase with a switch-like mechanism that favors the bound state when the membranes are highly energized (Carrillo and Vallejos, 1983). Matthijs and Lubberding (1988) discussed the possible existence of similar forms in cyanobacteria. In the cyanobacteria Synechococcus sp. strains PCC 6301 and PCC 7002 the water-soluble form of FNR is actually associated with the peripheral rods of the phycobilisomes by way of the amino-terminal domain which structurally similar to the phycocyaninassociated linker polypeptide CpcD (Schluchter and Bryant, 1992; see Chapter 7). Schluchter and Bryant (1992) also found a hydrophobic, membraneassociated form of FNR that was suggested to be acylated. This latter form could be the NADPH dehydrogenase active in respiratory electron transport. Schluchter and Bryant (1992) also suggested that acylation might be important for targeting the enzyme into the ICM and/or CM, but no data are yet available to prove or refute this interesting possibility. Several of the FNR reactions listed above could be of significance for cyanobacterial respiration. The possible importance of the transhydrogenase activity has been mentioned above (Section II B). In view of the observation by Peschek and Schmetterer (1982) that oxidation of NADPH by membranes from Nostoc sp. strain MAC depended on the presence of quinones, Figure 1 shows that the quinone pool is the electron acceptor for FNR-catalyzed NADPH oxidation. Figure 1 also demonstrates why the direct reduction of cytochrome or plastocyanin is unlikely to occur in vivo, although FNR has a high affinity for these substances and the reaction has been well studied (e.g., see Pueyo et al., 1992). While FNR must be localized on the cytoplasmic side of the ICM, cytochrome c and plastocyanin occur in the intrathylakoid space. After breaking the cells, or more exactly the ICM, these reactions become possible, however. Attempts by Schluchter and Bryant (1992) to

Chapter 13 Cyanobacterial Respiration inactivate the petH gene by insertion of a kanamycin resistance cassette only led to merodiploids containing both intact kanamycin-resistance and petH genes, so it seems that this gene is essential in Synechococcus sp. strain PCC 7002. It would be very interesting to repeat this experiment in a strain that is able to grow chemoheterotrophically in the dark, unlike Synechococcus sp. strain PCC 7002, which only grows photoautotrophically or photoheterotrophically (Rippka et al., 1979).

2. Other Membrane-Bound NADPH Dehydrogenase(s) FNR is probably not the only respiratory NADPH dehydrogenase in cyanobacteria. Schrautemeier et al. (1985) had observed that washing membranes from heterocysts of Anabaena variabilis led to loss of NADPH-, but not NADH-oxidase activity. However, Omata and Murata (1985) noted that removal of NADPH-DBMIB oxidoreductase by washing with NaCl was strain-dependent and also depended upon whether CM or ICM were investigated. In both Synechococcus sp. strain PCC 6301 and Synechocystis sp. strain PCC 6714, the activity in the ICM was largely lost by washing, consistent with the notion that the reaction is due to FNR. The activity of the CM, however, was much more sensitive to washing in Synechococcus sp. strain PCC 6301 than in Synechocystis sp. strain PCC 6714, and in both strains residual NADPH oxidase reactivity remained after washing (especially in the CM). The CM and ICM displayed similar activities for both strains. In Synechococcus sp. strain PCC 6301 Molitor and Peschek (1988) found considerable NADPHdriven cytochrome c reduction in both CM and ICM; the specific reactivity was similar for both membranes. A similar experiment performed by Wastyn et al. (1988) on Anabaena sp. strain ATCC 29413 gave a different result. They separated vegetative cells and heterocysts and isolated CM and ICM from both cell types. Only the CM from either vegetative cells or heterocysts, but not the ICM, displayed NADPH-driven cytochrome c reduction. Most of these data can be reconciled with the proposal of Howitt et al. (1993) that the mitochondrial-type NADH dehydrogenase in the CM of Anabaena sp. strain PCC 7120 also oxidizes NADPH. Therefore, Fig. 1 displays NADH- and NADPH dehydrogenases in the CM in close proximity. The varying results on NADPH dehydrogenases in the ICM may simply be due to different attachment stabilities of FNR to the

417

membranes so that currently there is no need to postulate an additional membrane-bound NADPH dehydrogenase in the ICM.

B. NADH Dehydrogenases Although the experiment by Biggins (1969), described in Section II A, implies that NADPH and not NADH is the major primary reductant for cyanobacterial respiration, the respiratory chain starting with NADH is better understood than that starting with NADPH. A clear distinction that the respiratory electron transport starting with either NADH or NADPH as donors may proceed through different primary oxidoreductases was first presented by Matthijs et al. (1984 a). They showed that oxidation of NADH, but not NADPH, was specifically inhibited by antimycin A in Plectonema boryanum. NADH oxidation has been shown to be coupled to oxidative phosphorylation in Mastigocladus laminosus (Frei et al., 1984) and Anabaena variabilis (Scherer et al., 1987). NADH oxidation (with DBMIB as the electron acceptor) was shown to be catalyzed by both CM and ICM (Omata and Murata, 1985). The relation of the reactivities between CM and ICM depended on the strain, however. In Synechococcus sp. strain PCC 6301, the reactivities were similar for both membranes, but in Synechocystis sp. strain PCC 6714 the CM contained 15-times the activity of the ICM (Omata and Murata, 1985). In Synechococcus sp. strain PCC 6301 NADH oxidation with cytochrome c as the electron acceptor was found in both membranes and was shown to be about 3-times higher in CM than in ICM (Molitor and Peschek, 1986). From Anabaena sp. strain ATCC 29413, Wastyn et al. (1988) isolated CM and ICM from both vegetative cells and heterocysts. All four membranes oxidized NADH with cytochrome c as acceptor, but to a very different degree. The CM ofboth cell types have much higher rates that the ICM 430, 170, 5, 20 nmol cytochrome c per min per mg protein). The major function of FNR in cyanobacteria is photosynthetic reduction of a process necessarily taking place at the ICM. Hence, one could speculate that in Anabaena variabilis the in vivo respiratory electron transport chain of the CM works largely with NADH and the respiratory chain of the ICM largely with NADPH as the electron donor. Two different membrane-bound NADH oxidases have been identified in cyanobacteria, one of the

418 ‘bacterial type’ and one of the ‘mitochondrial type’. There is also evidence for a soluble NADH dehydrogenase.

1. Mitochondrial-Type NADH Dehydrogenase The primary oxidoreductase of mitochondria is an NADH dehydrogenase that catalyzes the electron transfer from NADH to the quinone pool. It consists of more than 40 subunits and is sensitive to rotenone (Weiss et al., 1991; Walker, 1992). Several procaryotes have been shown to have related NADH dehydrogenases (Type-1 dehydrogenases; see Yagi, 1991) which contain non-covalently bound flavin mononucleotide and multiple iron-sulfur centers as prosthetic groups and whose activity is coupled to proton translocation. A similar enzyme is thought to reside in the thylakoids of chloroplasts (Nixon et al., 1989; Peltier and Schmidt, 1991). Chloroplast genomes contain eleven genes with sequence similarity to components of the mitochondrial NADH dehydrogenase complex (Ohyama et al., 1988; Shinozaki et al., 1986). Genes encoding the corresponding subunits in cyanobacteria have been isolated using hybridization probes derived from these genes. The ndhC-ndhK1-ndhJ operon was cloned from Synechocystis sp. strain PCC 6803 using an ndhJ probe from maize (Steinmüller et al., 1989). Three transcripts were identified that respectively contained the sequences coding for all three subunits, for ndhK and ndhJ only, and for ndhJ only. The ndhK gene was originally found in chloroplasts and called psbG (Steinmetz et al., 1986) but was later shown to be a subunit of the Type-1 NADH dehydrogenase in chloroplasts (Nixon et al., 1989) and cyanobacteria (Berger et al., 1991). In Synechocystis sp. strain PCC 6803 Mayes et al. (1990) found a second copy of the psbG gene, that should properly be called ndhK2, that was not linked to the operon described by Steinmüller et al. (1989) and indeed was located on a plasmid (Steinmüller and Bogorad, 1990). In two independent mutants lacking the ndhK1 gene, the ndhK2 gene that is not normally expressed in wild type cells had been activated (Steinmüller et al., 1991). The mutants displayed the same phenotype as the wild type when grown under photoautotrophic conditions, but grew slower under mixotrophic conditions; photoheterotrophic conditions were not tested (Steinmüller et al., 1991). In contrast, Synechococcus sp. strain UTEX 625 (Mayes et al.,

G. Schmetterer 1990) and Anabaena sp. strain PCC 7120 (Howitt et al., 1993) were reported to have only one ndhK gene. Other subunits of the Type-1 NADH dehydrogenase described so far from cyanobacteria are the ndhH gene from Synechocystis sp. strain PCC 6803, a single-copy gene yielding only one transcript, (Steinmüller, 1992a), and the ndhF gene from Synechococcus sp. strain PCC 7002 (Schluchter and Bryant, 1992; Schluchter et al., 1993). Takahashi et al. (1991) have isolated the ndhAIGE operon of Plectonema boryanum, and Ellersiek and Steinmüller (1992) have isolated the same operon, as well as the ndhD gene, from Synechocystis sp. strain PCC 6803. Ogawa (1991,1992) has isolated the ndhB and ndhL genes from Synechocystis sp. strain PCC 6803. Anderson and McIntosh (1991b) cloned and sequenced a DNA fragment from Synechocystis sp. PCC 6803 that contained the sequences similar to ndhE-psaC-ndhD genes. Although originally reported to be present in single copies, psaC, ndhE, and at least a portion of ndhD are present in two copies per chromosome in this cyanobacterium (Anderson and McIntosh, 1991b; Ellersiek and Steinmüller, 1992; Steinmüller, 1992b). Type-1 NADH dehydrogenase has been found to be located both in the CM and in the ICM of Synechocystis sp. strain PCC 6803 using antibodies prepared against the ndhK- and ndhJ- coded proteins in immunoblots (Berger et al., 1991). Measuring the oxidation of NADH with DBMIB as the electron acceptor, it was shown that the specific activity was about 12 times higher in the CM than in the ICM. 40 rotenone inhibited 50% of the activity in the ICM and 32% in the CM. Ogawa (1992) found the NdhL subunit of Synechocystis sp. strain PCC 6803 both in CM- and in ICM-preparations, but argued that NdhL may actually be located only in the ICM, its occurrence in the CM-preparation being due to contamination with ICM. In contrast, Howitt et al. (1993) report that Type-1 NADH dehydrogenase is confined to the CM in Anabaena sp. strain PCC 7120. This would fit well with experiments by Kraushaar et al. (1990) who found that NADH oxidation with cytochrome c as the electron acceptor was highly inhibited by rotenone only in the CM but not in the ICM in the closely related Synechocystis sp. strain PCC 6714. In three other strains (Synechococcus sp. strain PCC 6301, Anabaena sp. strain PCC 7937 (ATCC 29413), and Nostoc sp. strain PCC 8009) NADH oxidation, sensitive to low concentrations of rotenone,

Chapter 13 Cyanobacterial Respiration was mostly identified in the CM (Kraushaar et al., 1990). Clearly, the intracellular location of Type-1 NADH dehydrogenase needs further clarification. An important step will be the purification of the protein complex, which will also allow researchers to decide whether this enzyme really oxidizes both NADH and NADPH (Howitt et al., 1993). Recently, Berger et al. (1993) reported for the first time the purification of a subcomplex consisting of subunits NdhH, -I, -J, and -K from Synechocystis sp. strain PCC 6803. Mi et al. (1992a, b) have suggested that, in addition to the role the NADH dehydrogenase plays in respiratory electron transport, this enzyme is the point ofreentry of electrons from the acceptor side of Photosystem I for cyclic electron transport. However, Yu et al. (1992, 1993) have recently provided evidence for a novel cyclic electron transportpathway that does not require a functional NADH dehydrogenase in Synechococcus sp. strain PCC 7002. Moreover, if the Type-1 NADH dehydrogenase has substantial NADPH plastoquinone oxidoreductase activity and is present in the ICM, the NADH dehydrogenase could contribute to a type of ‘cyclic electron transport.’ In addition to characterization of the substrate specificity of this enzyme, the physiological and biochemical properties ofmutants devoid of this enzyme, should clarify this issue.

2. Bacterial-Type NADH Dehydrogenase Several bacteria have been found to contain socalled Type-2 respiratory NADH dehydrogenases that are much simpler than those found in mitochondria [e.g., Rhodopseudomonas capsulatus (Ohshima and Drews, 1981); Escherichia coli (Jaworowski et al., 1981); Paracoccus denitrificans (George and Ferguson, 1984)]. These enzymes only have one or two subunits, employ flavin adenine nucleotide as a cofactor, and act as simple oxidoreductases (Yagi, 1991). The cyanobacterium Anabaena variabilis strain ATCC 29413 contains such a Type-2 NADH dehydrogenase, and the enzyme was purified by Alpes et al. (1989). Two subunits, a major one of 17 kDa and a minor one of 52 kDa were identified. As in other Type-2 NADH dehydrogenases, flavin adenine nucleotide is a cofactor of the enzyme necessary for activity. The enzyme catalyzes oxidation of NADH with concomitant reduction of quinones – ubiquinone-1, which does not naturally occur in cyanobacteria, being the best

419 substrate found. Even high concentrations of rotenone inhibit the enzyme only by 80%, while at of the same inhibitor almost no inhibition is measured (Alpes et al., 1989). No other NADH dehydrogenase was observed in this strain by Alpes et al. (1989), in contrast to Kraushaar et al. (1990) who measured NADH oxidation sensitive to low concentrations of rotenone characteristic of mitochondrial-type NADH dehydrogenase in the same strain. One difference between the two studies is that Alpes et al. (1989) grew their cell under 2fixing conditions, while Kraushaar et al. (1990) used a medium containing nitrate. Howitt et al. (1993) also reported a Type-2 NADH dehydrogenase in Anabaena sp. strain PCC 7120. In addition, they found this enzyme to be localized only in the ICM.

3. Soluble NADH Dehydrogenase There are reports in the literature showing that cyanobacteria may contain considerable amounts of soluble NADH dehydrogenase activity. Since both mitochondrial (Type-1) and bacterial (Type-2) NADH dehydrogenases are membrane-bound enzymes, the soluble NADH dehydrogenase activities observed in Synechococcus sp. strain PCC 6301 (Omata and Murata, 1985) and Synechocystis sp. strain PCC 6714 (Sandmann and Malkin, 1983a; Omata and Murata, 1985) probably are due to one or more different enzyme(s). Sandmann and Malkin (1983) partially purified such an enzyme and found it to be inhibited 88% by 100 mM rotenone and 95% by o-phenanthroline. The NADH dehydrogenase activity ofthe soluble fraction is considerably higher than that of the membranes prepared from the same cells (Omata and Murata, 1985; Sandmann and Malkin, 1983a). The soluble dehydrogenase may be important but its biological function, and thus its possible involvement in respiratory electron transport, are unknown.

C. Hydrogenase(s) Cyanobacteria contain at least two different hydrogenases able to oxidize ; these are the ‘uptake’ hydrogenase and the ‘reversible’ hydrogenase (reviewed by Houchins, 1984). Reversible hydrogenase catalyzes both the uptake and evolution of and is also called soluble hydrogenase because it is found in the soluble fraction on breaking the cells (Houchins, 1984). Recently, however, immunological

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420 methods (Kentemich et al., 1989) and gentle lysis of the cells (Kentemich et al., 1991) produced evidence that reversible hydrogenase of Synechococcus sp. strain PCC 6301 may be associated with the cell periphery and specifically the CM. Although two genes possibly coding for subunits of the reversible hydrogenase have been cloned and sequenced (Anabaena cylindrica, Ewart et al., 1990; Synechocystis sp. strain PCC 6714, Van der Oost et al., 1989), little is known about its biological function. Since reversible hydrogenase apparently is active mainly under anaerobic conditions, and indeed is sensitive to (Houchins 1984), its involvement in respiration is unlikely. Uptake hydrogenase is a membrane-bound enzyme that is able to feed electrons from into the respiratory chain. This was first shown by Peterson and Burris (1978) in heterocysts of Anabaena sp. strain PCC 7120. They demonstrated that can donate electrons to in the dark in a reaction sensitive to CO and cyanide and coupled to oxidative phosphorylation. Oxyhydrogen reactions were also discovered in Synechococcus sp. strain PCC 6301 (Peschek, 1979a, 1979b, 1980a), in heterocysts of Anabaena cylindrica (Eisbrenner and Bothe, 1979), and in vegetative cells of Anabaena sp. strain PCC 7120 (Houchins and Burris, 1981a, 1981b); however, significant activity was only observed when the uptake hydrogenase had been induced by or other anaerobic conditions. The reaction in Anabaena cylindrica was inhibited by DBMIB, making the quinone pool a likely intermediate (Eisbrenner and Bothe, 1979). Furthermore, electron transport from to Photosystem I, also in a DBMIB-dependent fashion, was also found to occur in A. cylindrica (Eisbrenner and Bothe, 1979). In membranes from heterocysts of Anabaena variabilis, reduced in a light-dependent reaction, when either cytochrome c or plastocyanin were added as electron carriers between the cytochrome complex and Photosystem I (Schrautemeier et al., 1985). In Synechococcus sp. strain PCC 6301 the oxyhydrogen reaction was found to be inhibited by HQNO (2-nheptyl-4-hydroxyquinoline-N-oxide) and both CO and cyanide, suggesting the participation of the cytochrome complex and a cyanide-sensitive terminal oxidase (Peschek, 1979b, 1980a). In membrane particles from Synechococcus sp. strain PCC 6301, from which the quinones had been extracted with n-pentane, addition of plastoquinone or phylloquinone restored the oxyhydrogen reaction

(Peschek, 1980b). Peschek (1980b) also showed that could donate electrons to menadione. Taken together, the above-mentioned results imply that respiratory electron transport with as the primary electron donor proceeds through an uptake hydrogenase acting as an oxidoreductase that is located in the ICM. However, the possibility that uptake hydrogenase is also present in the CM cannot be rigorously excluded.

IV. Quinone Pool The quinone pool of cyanobacteria is thought to be the central component common to all respiratory and photosynthetic electron transport path ways. Keeping this is mind, the amount of data available on its chemical composition and function are rather limited. Quantitatively, plastoquinone-9 is present in higher amounts than any other component of the cyanobacterial electron transport chains (Nanba and Katoh, 1985) and it probably forms a redox buffer. The involvement of the quinone pool in the cyanobacterial respiratory electron transport chain was first shown by Leach and Carr (1970) by the demonstration of an NADPH-menadione oxidoreductase in membranes from Anabaena variabilis. Eisbrenner and Bothe (1979) used isolated heterocysts of Anabaena variabilis to measure light-dependent ethine reduction and dark consumption of both with as the electron donor. Both processes were inhibited by DBMIB, an inhibitor of quinol oxidation (Trebst and Harth, 1970), thus providing for the first time evidence that the quinone pool was common to respiratory and photosynthetic electron transports. DBMIB also inhibited the reduction of cytochrome in membranes from Synechococcus sp. strain PCC 6301 with NADH, or NADPH as the electron donors (Peschek, 1982). In membranes from Nostoc sp. strain MAC, Peschek and Schmetterer (1982) obtained further evidence for this. They measured spectroscopically the oxidation of cytochrome f by diphenylcarbazide with Photosystem I-light (707 nm), followed by its re-reduction by Photosystem IIlight (674 nm). The re-reduction, but not the oxidation of cytochrome f, was inhibited when the membranes were depleted of quinones by w-pentane extraction, and this inhibition could be reversed by the addition of plastoquinone. Similarly, in the presence of either NADH or NADPH, cytochrome f could be oxidized by and re-reduced when cyanide was added, but

Chapter 13 Cyanobacterial Respiration the re-reduction was inhibited in the absence of plastoquinone. In membranes from Synechococcus sp. strain PCC 6301 photosynthetic and respiratory electron transport inhibited each other by competing for electrons from a common donor (Peschek and Schmetterer, 1982). reduction dependent upon Photosystem I-light was inhibited by and consumption of with NADPH as the electron donor decreased when the Photosystem I-light was turned on. Final proof that the quinone pool is common to photosynthetic and respiratory electron transport chains came from a series of experiments performed by Katoh’s group on a thermophilic Synechococcus sp. These experiments are among the few performed with intact cells and therefore characterize the in vivo situation. Hirano et al. (1980) flash-photooxidized cytochrome and measured the ensuing re-reduction in the dark with either glycogen (previously accumulated in the light) or exogenously added fructose (used with starved cells) as the electron donor. This re-reduction was inhibited by DBMIB, but was not influenced by DCMU, showing directly that the quinone pool is part of the electron transport chain from primary reductants (either NADPH or NADH, see Sections II A and II B) to cytochrome Using the I-D dip, an early transient of fluorescence induction by strong red light that is a direct probe for the redox state of the quinone pool, Aoki and Katoh (1982) demonstrated that plastoquinone could be reduced by endogenous glycogen or exogenous fructose, and was oxidized by a cyanide sensitive oxidase. By the same technique, the steadystate redox level of the quinone pool under aerobic conditions in the dark was found to be mostly reduced due to the steady delivery of electrons from the dehydrogenase(s), as long as either endogenous or exogenous carbohydrates were present as electron donors (Aoki and Katoh, 1982). Nanba and Katoh (1983, 1984) studied the kinetics of re-reduction of flash-oxidized cytochrome and obtained evidence that the plastoquinone pool transfers its electrons to the cytochrome complex and that this transfer is inhibited by DBMIB. Omata and Murata (1984a) showed that prenylquinones are present in both CM and ICM from Synechococcus sp. strain PCC 6301, but in different relative amounts. Plastoquinone-9 made up 71% of quinones in ICM, but 95% in CM; in ICM 26% was phylloquinone, in CM only 5%; and 5'-monohydroxyphylloquinone, which constituted 3% in the

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ICM, was present in very low levels in CM. Interestingly, Mühl and Löffelhardt (1982) detected 4-hydroxyphenylpyruvate dioxygenase, an intermediate enzyme in the biosynthesis of plastoquinone, in both of two membrane fractions from Synechococcus sp. strain PCC 6301 that probably corresponded to CM and ICM, the ICM having about four times the activity of the CM. In reconstitution experiments using quinone-depleted membranes from Synechococcus sp. strain PCC 6301, Peschek (1980b) found phylloquinone to be best in restoring NADPH- or c reductase and also NADPH- or consumption. Peschek and Kuntner (1987) extended these observations to Anabaena sp. strain ATCC 29413 and Nostoc sp. strain PCC 8009 with essentially the same results. However, when photosynthetic electron transport (from to or from diphenylcarbazide to 2,6-dichlorophenolindophenol) was investigated in quinone-depleted membranes from Synechococcus sp. strain PCC 6301, Anabaena sp. strain ATCC 29413, and Nostoc sp. strain PCC 8009, reconstitution gave higher activities with plastoquinone-9 than with phylloquinone (Peschek and Kuntner, 1987). Sandmann and Malkin (1983a), showed that naphthoquinone was a better electron acceptor than different benzoquinone derivatives for a membranebound NAD(P)H dehydrogenase from Synechocystis sp. strain PCC 6714. Furthermore, 1,4-naphthoquinone was the best activator found for NADHcytochrome c oxidoreductase in CM from Synechocystis sp. strain PCC 6714 (Omata and Murata, 1985) and 1,4-naphthoquinone was the best electron acceptor for the NADPH dehydrogenase isolated from Microcystis aeruginosa (Viljoen et al., 1985). Krinner et al. (1982) isolated the cytochrome complex from Anabaena variabilis and tested different quinones for their ability to act as electron donors. Plastoquinone-9 was the most effective one found, but phylloquinone or other naphthoquinones were not tested. Aoki and Satoh (1983) estimated that the sizes of the quinone pool for respiratory and photosynthetic electron transport in a thermophilic Synechococcus sp. were equal. If indeed plastoquinone-9 is the major photosynthetic quinone, then respiratory electron transport would make use of both phylloquinone and plastoquinone-9. Peschek and Kuntner (1987) speculated therefore, that the electron donors to the quinone pool are responsible for the differential effects of the two quinones, possibly by having different affinities for them.

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V. Cytochrome

complex

In chloroplasts, the cytochrome complex acts as a plastoquinol-plastocyanin oxidoreductase. In mitochondria, the structurally and functionally related cytochrome complex acts as a ubiquinolcytochrome c oxidoreductase. In cyanobacteria only a cytochrome complex exists that functions both in respiratory and photosynthetic electron transport chains (for a detailed discussion of this complex, see Chapter 9). As in chloroplasts, isolated cyanobacterial cytochrome complexes contain four subunits, cytochrome (or subunit IV, Rieske FeSprotein, and cytochrome f, a c-type cytochrome (Anabaena variabilis, Krinner et al., 1982; Spirulina sp., Minami et al., 1989). All subunits are integral membrane proteins. The corresponding genes have been cloned and sequenced from Nostoc sp. strain PCC 7906. The petC (Rieske FeS protein) and petA (cytochrome f) form one operon (Kallas et at., 1988a), while petB (apocytochrome and petD (subunit IV) form a second operon (Kallas et al. 1988b). All genes are present in only one copy per chromosome. A similar gene arrangement was found in Synechococcus sp. strain PCC 7002 in which only single copies ofeach gene occur per chromosome (Brand et al., 1992). The single copies of the petD and petB genes from Synechocystis sp. strain PCC 6803 have been isolated, and attempts to delete the petD gene in this strain were not successful, indicating that this gene and thus the cytochrome complex are essential for cell viability (Osiewacz, 1992). The petC-petA genes from Synechocystis sp. strain PCC 6803 have also been described (Mayes and Barber, 1991). The electron donor to the cytochrome complex most likely is plastoquinone-9 (see Section IV). Thus electrons can be donated both from Photosystem II and from one of the dehydrogenases (cf. Section IV). This was shown in Synechococcus sp. strain PCC 7002 (Agmenellum quadruplicatum strain PR6) by Myers (1986), in Nostoc sp. strain MAC by Peschek and Schmetterer (1982), in Synechococcus sp. strain PCC 6301 by Peschek (1983), and in Synechocystis (Aphanocapsa) sp. strain PCC 6714 by Sandmann and Malkin (1984). Re-reduction in the dark of flash oxidized cytochrome f has been demonstrated in a thermophilic Synechococcus sp. by Nanba and Katoh (1983, 1984), who also obtained kinetic evidence for the participation of the Rieske FeS protein. Re-reduction in the dark of flash oxidized

cytochrome was found in Anabaena sp. strain PCC 7120 by Houchins and Hind (1983) and in Synechocystis sp. strain PCC 6714 by Matsuura et al. (1988). The cytochrome complex donates electrons either to cytochrome or plastocyanin, depending on the strain and the growth conditions (see Section VI and Chapter 12). Oxidation ofthe cytochrome complex by both in the dark and by Photosystem I in the light has been demonstrated in heterocysts of Anabaena variabilis (Böhme and Almon, 1983), in Synechococcus sp. strain PCC 6301 by Peschek (1983a), and in Nostoc sp. strain MAC by Peschek and Schmetterer (1982). Evidence that electron transport through the cytochrome complex of Synechocystis sp. strain PCC 6714 occurs by the Qcycle mechanism (Mitchell 1976) was presented by Matsuura et al. (1988). Inhibitors have frequently been used to show the common function ofthe cytochrome complex in photosynthetic and respiratory electron transport (Peschek, 1979b; Houchins and Burris, 1981c; Houchins and Hind, 1982; Sandmann and Malkin, 1983b; Matthijs et al., 1984a; Stürzl et al., 1984; Schrautemeier et al., 1984; Binder et al., 1984; Abdourashitova et al., 1985; Scherer et al., 1987; Matsuura et al., 1988). Among the inhibitors employed, the site of action has been determined for DBMIB (competeswithplastoquinoneforthebinding site on cytochrome Nanba and Katoh, 1984) and HQNO (binds at the site of cytochrome Matsuura et al., 1988). However, the action of antimycin A, an inhibitor of the mitochondrial cytochrome complex, is not completely clear. Membranes from Anabaena variabilis (Leach and Carr, 1970) and Plectonema boryanum (Matthijs et al., 1984a) oxidized NADH (with in an antimycin A sensitive way, but NADPH oxidation (also with was barely affected. The periplasmic reduction of ferricyanide by intact cells of Synechococcus sp. strain PCC 6301, a reaction presumably involving the cytochrome complex, was also inhibited about 50% by antimycin A (Peschek et al., 1988). In contrast, electron transport by isolated membranes from NADH or NADPH to Photosystem I (or from duroquinol to oxidized cytochrome c ) in Synechocystis (Aphanocapsa) sp. strain PCC 6714 (Sandmann and Malkin, 1983b, 1984); from NADH to Photosystem I or from NADH to in Anabaena sp. strain ATCC 29413 (Scherer et al., 1987); and from NADH, NADPH or succinate to in

Chapter 13 Cyanobacterial Respiration Synechococcus sp. strain PCC 6301 (Peschek, 1980a) was completely insensitive to antimycin A. A possible explanation for these discrepancies is offered by Kraushaar et al. (1990; see below). The cytochrome complex, being a component of the photosynthetic electron transport chain, is obviously localized in the ICM, but recent evidence also shows its presence in the CM. Its occurrence in the CM was suggested by the demonstration that exogenous ferricyanide, which cannot enter cells, is reduced by intact cells of Synechococcus sp. strain PCC 6301 (Craig et al., 1984; Peschek et al., 1988). Clear evidence for the presence of the cytochrome complex in both CM and ICM came from immunological data (Kraushaar et al., 1990). Antibodies against chloroplast cytochrome f, cytochrome and subunit IV cross-reacted with both CM and ICM from Synechocystis sp. strain PCC 6714, and antibodies against chloroplast cytochrome f, cytochrome and Rieske FeS protein cross-reacted with both CM and ICM from Synechococcus sp. strain PCC 6301 (Kraushaar et al., 1990). In addition, Kraushaar et al. (1990) noted that the reduction of cytochrome c with plastoquinol9 was inhibitable by antimycin A only in CM but not in ICM of Synechococcus sp. strain PCC 6301. An explanation for how the single-copy genes for each of the four subunits give rise to the complex in both the ICM and CM, and especially for how the complexes in CM and ICM come to have different properties (namely sensitivity towards antimycin A) must await further investigations.

VI. Peripheral Intermediate Electron Carriers Krinner et al. (1982) showed that their purified cytochrome complex catalyzed plastoquinol-9 oxidation with concomitant reduction of either cytochrome c or plastocyanin. Therefore, subsequently it has been generally assumed that either cytochrome c or plastocyanin, both peripheral membrane proteins that are found in the soluble fraction on breaking the cells, act as the intermediate electron carriers between the cytochrome complex and either Photosystem I (photosynthetic electron transport) or a terminal oxidase (e.g., of the respiratory electron transport). In reality, very little is known about the function of the intermediate electron carriers, and for two strains the above assumption has been shown to be incorrect (see

423 below). Sandmann and Böger (1980) discovered that in some cyanobacteria, the contents of cytochrome and plastocyanin are regulated by the concentration in the growth medium. Indeed, three groups of cyanobacteria can be distinguished in this respect (Sandmann, 1986): 1. those forming cytochrome only, independent of the concentration of the growth medium; 2. those forming cytochrome in the absence of and plastocyanin in the presence of and 3. those forming cytochrome both in the absence and in the presence of and additionally plastocyanin in the presence of Thus the ability to synthesize cytochrome seems to be universal among cyanobacteria, while gene(s) for plastocyanin can be absent (see below).

A. Cytochrome Synechococcus sp. strain PCC 6301 contains three different soluble cytochromes c (Holton and Myers, 1967). Cytochromes (or and are apparently present in all cyanobacteria. Cytochrome is present in so extremely small amounts, that its real existence must remain doubtful. Cytochrome has a very negative redox potential, is absent from heterocysts of Nostoc muscorum (Anabaena sp. strain PCC 7119; Almon and Böhme, 1980), and may thus be a component of Photosystem II. It is most likely not involved in respiration and will not be considered further. Participation of cytochrome c in the respiratory electron transport chain was first shown when Biggins (1969) measured oxidation of reduced cytochrome c (apparently from a eucaryotic source) in membranes from Phormidium luridum. Dark reduction of cytochrome c (probably also of eucaryotic origin) with either NADPH or NADH as electron donors was reported by Leach and Carr (1970) by membranes from Anabaena variabilis. Even today, most measurements of cytochrome c reductase and cytochrome c oxidase activities in cyanobacteria are performed with eucaryotic cytochrome c (commonly from horse heart), so that no information concerning the in vivo cytochromes c can be derived from these experiments. Lockau (1981) demonstrated that reduced cytochrome from Anabaena variabilis can be oxidized by in the dark and by methyl viologen in the light in membranes from the same organism. In membrane preparations from Nostoc muscorum,

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cytochrome from the same strain, but not from other sources including other cyanobacteria, was able to reconstitute photosynthetic (from to ) electron transport (Stürzl et al., 1982). Respiratory electron transport (from NADPH to in the same membranes, however, could be reconstituted with any of the cytochromes c tested, including the one from N. muscorum (Stürzl et al., 1982). Stürzl et al. (1982) also measured the residual amount of cytochrome present in their membrane preparations and showed it to depend on the isolation procedure. Light-driven electron transport from either or NADH to NADPH (via ferredoxin and FNR) in membranes from Anabaena variabilis heterocysts could be restored by the addition of cytochrome or plastocyanin (Schrautemeier et al., 1985). Antibodies against cytochrome from Nostoc muscorum strain ATCC 29151 inhibited both respiratory (from NADPH to and photosynthetic (from to electron transport in membranes from the same organism (Alpes et al., 1984). By flash-oxidizing Photosystem I (P700) in intact cells of a thermophilic Synechococcus sp., Nanba and Katoh (1983, 1984, 1985) obtained direct evidence for the function of cytochrome in mediating electron transfer from cytochrome f to Photosystem I. In membranes from Synechocystis (Aphanocapsa) sp. strain PCC 6714, flash-oxidation of cytochrome in the presence of either duroquinol or NADH is enhanced when cyanide is also added (Sandmann and Malkin, 1984), showing that cytochrome is an electron donor to Photosystem I that competes for electrons moving to the terminal respiratory oxidase, but not necessarily that cytochrome is a common component of the respiratory and photosynthetic electron transport chains under these conditions (although this may well be the case). Clearly, cytochrome is a possible donor to Photosystem I and cytochrome c oxidase. Unfortunately, no in vivo data are available showing definitively that cytochrome also is the in vivo donor to cytochrome c oxidase. Therefore, it is difficult to interpret the results by Peschek et al. (1981 a), Kienzl and Peschek (1982) and Moser et al. (1990). These authors studied the effect of c-type cytochromes from different sources in the cytochrome c oxidase reactions of membranes from different cyanobacteria. Especially striking was the fact that cytochrome from Synechococcus sp. strain PCC 6301 had no activity with the cytochrome c oxidase from the same organism (Kienzl and Peschek, 1982). Moser

G. Schmetterer et al. (1990) resolved this puzzling observation by showing that the reactivity of cyanobacterial cytochromes towards cyanobacterial cytochrome c oxidases depended on the ionic strength of the assay medium. Cytochromes from Synechococcus sp. strain PCC 6301, Synechocystis sp. strain PCC 6803, and Microcystis sp. reacted only at high ionic strength(100 mM phosphate) with cytochrome c oxidases from Synechococcus sp. strain PCC 6301 or Synechocystis sp. strain PCC 6803, while cytochromes c from horse heart or Nostoc sp. strain PCC 8009 were active only under low ionic strength (10 mM phosphate). Moser et al. (1990) attributed this to the different isoelectric points ofthe cytochromes involved – the former three being acidic, the latter two basic. Indeed, the isoelectric points ofcyanobacterial cytochromes varywidely (between 3.8 and 9.3) (Ho and Krogmann, 1984). The significance ofthis is unknown, but the isoelectric points of both cytochrome and plastocyanin from the same cyanobacterium seem to be similar (Ho and Krogmann 1984). Since at least some cyanobacteria appear to have a complete respiratory chain both in CM and ICM, cytochrome should be located both in the periplasmic space and in the intrathylakoid space, as pointed out by Peschek (1984a). The genes for apocytochrome have been cloned from Synechococcus sp. strain PCC 7942 (Laudenbach et al., 1990) from Anabaena sp. strain PCC 7937 (Bovy et al., 1992a,b) and from Synechocystis sp. strain PCC 6803 (Zhang et al., 1994). All are single-copy genes so that the question of targeting cytochrome into two different compartments remains open. Gentle extraction with EDTA showed that the periplasm contains cytochrome in Anabaena variabilis strain ATCC 29413 grown in the absence of (Serrano et al., 1990) and in Nostoc sp. strain PCC 8009 grown in the presence of (Obinger et al., 1990). Immunogold-labeling in Anabaena variabilis demonstrated that 75% of the cytochrome in this strain is associated with the cell periphery (presumably the periplasmic space) and 25% with the ICM (presumably the intrathylakoid space). This was unexpected, since ICM contain the photosynthetic electron transport chain, whose activity is much greater than that ofthe respiratory chain. Moser et al. (1990) demonstrated that both CM and ICM of Synechococcus sp. strain PCC 6301 and Synechocystis sp. strain PCC 6803 oxidized cytochrome from the same organism, but to a very different

Chapter 13 Cyanobacterial Respiration extent. Most cytochrome c oxidase activity in Synechococcus sp. strain PCC 6301 was in the CM, while in Synechocystis sp. strain PCC 6803 the activity was in the ICM. The cytochrome isolated from the periplasm of Nostoc sp. strain PCC 8009 was oxidized by both CM and ICM from the same organism (Obinger et al., 1990). Recent results have shown that in Synechococcus sp. strain PCC 7942, a strain containing no plastocyanin (Aitken 1976, Van der Plas et al., 1989, Laudenbach et al. 1990), cytochrome is also dispensable for photosynthetic (Laudenbach et al., 1990) or respiratory (G. Schmetterer and E.M. Wanzenböck, unpublished results) electron transport (see Section VI C). Similarly, a Synechocystis sp. strain PCC 6803 derivative containing neither plastocyanin nor cytochrome had almost normal photosynthetic and respiratory activities (Zhang et al., 1994). It will be interesting to determine whether a recently discovered, membrane-intrinsic cytochrome (N. Murata, personal communication) can substitute for the soluble cytochrome in light of a similar situation in purple photosynthetic bacteria.

B. Plastocyanin It has frequently been stated that cytochrome c and plastocyanin can substitute for each other in photosynthetic and respiratory electron transport chains. While this is undoubtedly true for photosynthesis, no data are available to corroborate this claim for respiration. In fact, as far as I was able to ascertain, the only experimental data possibly showing an involvement of plastocyanin in cyanobacterial respiration are presented in Figure 4 of Lockau (1981). In these studies total membranes (separation of CM and ICM was not yet possible at the time) from Anabaena sp. strain ATCC 29413 were shown to consume in the presence of plastocyanin at both pH 7.5 and pH 6.2, and this uptake was inhibited by 125 mM NaCl. It would be highly interesting to know which terminal oxidase catalyzed this reaction. It appears that the most urgent, open question with respect to cyanobacterial respiration is clarification of respiratory electron transport in cells containing plastocyanin but no cytochrome since this is the case for some frequently used heterotrophically grown strains (e.g., Synechocystis sp. strain PCC 6803(Briggs et al., 1990, Zhang et al., 1992) and Anabaena sp. strain PCC 7937 (Bovy et al., 1992b)).

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C. Other Possible Peripheral Electron Carrier(s) Laudenbach et al. (1990) constructed a mutant of Symchococcus sp. strain PCC 7942 that lacks the cytA gene coding for apocytochrome and thus contains no detectable soluble cytochrome c protein other than cytochrome (see above). This strain displayed normal photosynthetic activity (except for a decreased efficiency of cytochrome f oxidation). Since Synechococcus sp. strain PCC 7942 contains no plastocyanin (see Section VI A), there appears to be no electron carrier between the cytochrome complex and Photosystem I in the mutant. This can only be explained in two ways: Either direct electron transfer between the cytochrome complex and Photosstem I is possible, or there exists an alternative peripheral electron carrier, tentatively called ‘X’ in Fig. 1. The same mutant was analyzed by Schmetterer and Wanzenböck (unpublished) for respiratory activity. No phenotypic difference between the wild type and the cytA mutant could be observed in any of a number of respiratory reactions. Especially striking was the observation that below about 5 mM when the uptake rate of was not zero-order in concentration, the apparent for was exactly the same in wild type and mutant. Apparently cytochrome is not a component of the (major) respiratory chain in Synechococcus sp. strain PCC 7942. Quite recently Zhang et al. (1994) constructed a mutant of Synechocystis sp. strain PCC 6803 lacking both cytochrome (by inactivation of the single gene) and plastocyanin (by growth under very low concentration). These cells also had almost normal respiratory activity. If cytochrome c oxidase (of the see Section VII A) is the major terminal oxidase, it might also be reducible by another electron carrier (‘X’?). Otherwise a different terminal oxidase may be the major respiratory terminal oxidase (cf. Section VII B). Again, it will be interesting to determine whether the recently discovered membrane-intrinsic cytochrome (N. Murata, personal communication) might be the component ‘X’ mentioned above. A result from the earlier literature may be reinterpreted in the sense that there is an alternative electron donor between cytochrome f and Photosystem I. Nanba and Katoh (1985) observed that at low light intensities, almost 80% of the cytochrome remained reduced after flash-oxidizing P700. Instead of the expected very fast re-reduction of P700, P700 was re-reduced more slowly than when

426 90% of the cytochrome had been flash-oxidized with high light intensity. It is tempting to assume that, depending on the redox state of cytochrome an alternative electron carrier (‘X’) can catalyze the cytochrome f-P700 oxidoreductase reaction.

VII. Terminal Oxidases Only one terminal electron cyanobacterial respiration is known. Thus, the respiratory chain in cyanobacteria must end in one or more terminal oxidases. Of these, only cytochrome c oxidase has been well characterized. There is evidence, however, for the occurrence of at least two more terminal oxidases.

A. Cytochrome c Oxidase, The earliest investigations on cyanobacterial respiration noted that uptake in the dark was sensitive to cyanide, azide and CO (Webster and Frenkel, 1953). Therefore, it was (correctly) assumed that the terminal oxidase might be a hemoprotein. Evidence for cyanide-sensitive cytochrome c oxidase activity in cyanobacteria was first obtained by Biggins (1969) in Synechococcus sp. strain PCC 6301 and Phormidium luridum and by Leach and Carr (1970) in Anabaena variabilis. Cytochrome c oxidase activity was higher in heterocysts than in vegetative cells of Anabaena sp. strain PCC 7120 (Houchins and Hind, 1984). Cytochrome was detected spectroscopically first in Synechococcus sp. strain PCC 6301 (Peschek 198la), and then in many other cyanobacteria (Peschek, 1981b; Peschek et al., 1982a). An EPR signal characteristic of cytochromes was detected in Synechococcus sp. strain PCC 6311 (Fry et al., 1985), Synechococcus sp. strain PCC 6301 (Fry and Peschek, 1988), and Plectonema boryanum (Matthijs et al., 1984c). In at least some cyanobacteria, cytochrome c oxidase is present in both CM and ICM. The first indication for this came from the temperaturedependence of respiratory andphotosynthetic electron transport reactions, the former typically showing two and the latter only one discontinuity in Arrhenius plots in Synechococcus sp. strain PCC 6301 (Peschek et al., 1981b; Peschek et al. 1982b) and in Anabaena sp. strain ATCC 29413 (Scherer et al., 1981). Intact spheroplasts from Synechococcus sp. strain PCC 6301 oxidized exogenous cytochrome c in the dark

G. Schmetterer (Peschek et al., 1982c) coupled to proton extrusion (Peschek, 1983b, 1984b). Intact cells of Synechococcus sp. strain PCC 6301 oxidized exogenous ferrocyanide with concomitant extrusion of protons (Peschek et al., 1988). In intact cells, respiratory electron transport was also shown to be coupled to proton extrusion, which at least partially (dependent on the strain) was due to direct proton translocation by the respiratory chain (Nitschmann and Peschek, 1984, 1985; Peschek et al., 1985a, 1985b; Erber et al., 1986; Peschek, 1987). Physical separation of CM and ICM did not show consistent results. In Synechococcus sp. strain PCC 6301 Omata and Murata (1985) found cytochrome c oxidase activity only in ICM, while Molitor and Peschek (1986) found activity both in CM and in ICM fractions. The difference was found to be due to the growth conditions. High ionic strength (0.4-0.5 M NaCl) of the growth medium leads to a greatly enhanced synthesis of cytochrome c oxidase (Fry et al., 1986; Molitor et al., 1986; Molitor et al., 1990). In this connection it is interesting that in Synechococcus sp. strain PCC 6301 and Synechocystis sp. strain PCC 6714, both CM and ICM reacted with cytochrome from the same strain only under elevated ionic strength 100 mM phosphate; Moser et al., 1991). Several different strains have been tested for the presence of cytochrome c oxidase activity in CM and ICM (Peschek et al., 1989 b). Except for Gloeobacter violaceus strain PCC 7421, a strain that has no ICM, all strains showed activity in both membranes, albeit in very different relative amounts. Synechococcus sp. strain PCC 6301 had 95% of the total activity in CM, Synechocystis sp. strain PCC 6714 had only 3.5% of the total activity in CM, and other strains had intermediate values. In Anabaena sp. strain ATCC 29413, heterocysts had much higher cytochrome c oxidase activities than vegetative cells: 10 times higher in CM, and 50 times higher in ICM (Wastyn et al., 1988), clearly demonstrating the importance of respiration in heterocysts (see Fay, 1992). It is remarkable that all strains investigated by Peschek’s group for cytochrome c oxidase activity were grown in media containing Some of the strains (e.g., Synechocystis sp. strains PCC 6714 and PCC 6803) investigated do not produce cytochrome under these conditions (see Sandmann, 1986). Immunological methods have yielded independent evidence for the occurrence of cytochrome c oxidase in both CM and ICM. Antibodies prepared against the two-subunit form of cytochrome c oxidase

Chapter 13 Cyanobacterial Respiration from Paracoccus denitrificans (Ludwig and Schatz, 1980), cross-reacted with the corresponding subunits in Synechococcus sp. strain PCC 6301 both in CM and ICM (Trnka and Peschek, 1986; Molitor et al., 1987; Peschek et al., 1989c). Two groups reported partial purifications of cyanobacterial cytochrome c oxidase (Wastyn et al., 1987; Häfele et al., 1988). Quite recently, Niederhauser (1992) obtained evidence that the cytochrome c oxidase from Synechococcus sp. strain PCC 6301 contains four subunits. He purified the enzyme extensively and found only four polypeptides by polyacrylamide gel electrophoresis in the presence of sodium dodecylsulfate. Antibodies separately prepared against isolated cytochrome c oxidase subunits I, II, and III from Paracoccus denitrificans and mitochondrial subunit IV cross-reacted with only one polypeptide in immunoblots prepared with the enzyme. Tano et al. (1991) cloned the gene for subunit II and part of subunit I of cytochrome c oxidase from Synechococcus vulcanus. More recently, these authors have reported the complete nucleotide sequences for the coxBAC operon (Sone et al., 1992). The complete cox locus from Synechocystis sp. strain PCC 6803, containing the coxB, coxA, and coxC genes in that order, coding for subunits II, I, and III, respectively, has also been isolated and sequenced (Alge and Peschek, 1993a,b; Alge et al., 1994; Schmetterer et al., 1994). All sequenced subunits proved to show high sequence similarity with all other known cytochrome c oxidases. These sequences finally prove the existence of an cytochrome c oxidase in cyanobacteria. In Synechocystis sp. strain PCC 6803, only one copy for the cox locus was found (Alge et al., 1994) Therefore, cytochrome oxidase and a few other enzymes related to respiratory electron transport (see Sections II B and V) are apparently targeted from single-copy genes into both CM and ICM, which are otherwise completely different (Omata and Murata, 1983; 1984a,b; 1985). Targeting into only one membrane and subsequent transport or partitioning into the other one is also possible. This point will need clarification in the future. Very little is known about the regulation of cytochrome c oxidase expression. Two interesting mutants of Synechocystis sp. strain PCC 6803 were described by Jeanjean et al. (1990). Originally selected for sensitivity to growth in high (0.5 M) NaCl, the mutants had unaltered cytochrome c

427

oxidase activity in the CM, but only about half the cytochrome c oxidase activity in ICM, when compared with the wild-type strain grown under low-salt conditions.

B. Cyanide-Sensitive Alternative Terminal Oxidase(s) Recently, Schmetterer et al. (1994) have obtained evidence for a cyanide-sensitive alternative terminal oxidase distinct from cytochrome c oxidase. Starting from the cox locus cloned from Synechocystis sp. strain PCC 6803, a homozygous mutant was constructed that lacks most of the coxA gene and about half of the coxC gene (coding for subunits I and III of the cytochrome c oxidase) and that carries a kanamycin resistance cassette instead. As expected, this mutant showed no cytochrome c oxidase activity either in CM or ICM. However, intact cells of the mutant consumed almost control amounts of in the dark, and this consumption was sensitive to HQNO and cyanide. This implies the existence of another cyanide-sensitive terminal oxidase in ynechocystis sp. strain PCC 6803. One possibility would be a b-type cytochrome similar to the cytochrome o oxidase in Escherichia coli (Kita et al., 1984). This enzyme is a oxidoreductase and is sensitive to both cyanide and HQNO. Another possibility would be a cyanide-sensitive terminal oxidase that accepts electrons through the cytochrome complex. Both possibilities are considered in Fig. 1, as is the location in CM and ICM, about which no information is available.

C. Cyanide-Insensitive Alternative Terminal Oxidase(s) Although ‘cyanide-insensitive respiration’, i.e. uptake in the dark in the presence of significant amounts of cyanide (e.g. 1–5 mM) occurs in cyanobacteria, its mechanism is little understood. A well defined cyanide-insensitive ‘respiratory’ pathway, that ends in the so-called ‘alternative oxidase,’ exists in higher plants. The gene encoding this alternative oxidase has recently been cloned from Sauromatum guttatum (Rhoads and McIntosh, 1991), but there is no evidence that a similar alternative oxidase exists in cyanobacteria. Peschek (1980a) identified several inhibitors for cyanide-insensitive uptake in Synechococcus sp. strain PCC 6301, bathophenanthroline being most

428 effective. Scherer et al. (1988d) found cyanideinsensitive respiration in all of nine strains tested except in Phormidium foveolarum, as had been noted before for Phormidium luridum by Biggins (1969). However, no ATP synthesis was detectable in the presence of 1 mM cyanide so that ‘cyanide-insensitive respiration’ is not a respiratory process according to our definition (Section I.).

VIII. Conclusion

A. Biological Function of Cyanobacterial Respiration Respiration has been found in all cyanobacteria tested for this activity, even in those that are obligate photo(auto)trophs. Most authors state that the probable function of respiration is the generation of a minimum amount of energy necessary for survival in the dark. Actually, there exist very little data to corroborate this. In fact, one report specifically claims that dark-survival does not depend on the presence of respiratory activity. Doolittle and Singer (1974) isolated a mutant of Synechococcus sp. strain PCC 6301, a strain that is an obligate photoautotroph (!), which lacks both glucose 6-phosphate dehydrogenase and 6-phosphogluconate dehydrogenase. The reactions catalyzed by these two enzymes being the major source of reductant (in the form of NADPH) in the dark, it was not surprising to find that these mutants had very low uptake rates in the dark and thus probably very low respiratory electron transport rates. However, this mutant survived in the dark nearly as well as the wild-type strain, in contrast to mutants lacking 6-phosphogluconate dehydrogenase only (Doolittle and Singer, 1974, Broedel and Wolf, 1990). Furthermore, at least some cyanobacteria can survive well under anaerobic conditions (Paerl and Bebout, 1988; Heyer, 1989). It is tempting to speculate that besides the bioenergetic aspect, respiration in cyanobacteria might have a more specific function under certain ecological conditions. This is certainly true in heterocysts, where cytochrome c oxidase apparently acts as a scavenger for to protect the nitrogenase (see Fay, 1992). In the unicellular cyanobacterium Gloeothece sp. strain PCC 6909, that is able to fix nitrogen and perform oxygenic photosynthesis in the same cell even during continuous illumination

G. Schmetterer (Gallon et al., 1974), respiration has been suggested to provide the major source of ATP for nitrogen fixation both in the light and in the dark (Maryan et al., 1986). Furthermore, there exists the theory that respiration (at least with as the terminal electron acceptor) originated in cyanobacteria (the conversion hypothesis of Broda, 1974), since cyanobacteria are thought to have been the first organisms to produce massive amounts of free and their respiratory electron transport can be thought of as being made from photosynthetic electron transport with the addition of only a single additional electron carrier, the terminal oxidase (see Fig. 1). Since the affinity of modern terminal oxidases (such as cytochrome c oxidases) for is very high in Anabaena variabilis about Jensen and Cox, 1983), the original function of respiration could also have been to regulate the intracellular concentration of evolved by Photosystem II. It appears that a definite clarification of the function(s) of cyanobacterial respiration in natural ecosystems requires a viable respiration-deficient mutant of a cyanobacterium that will either have to be constructed in the laboratory or (less likely) be found in nature.

B. Respiration and Photosynthesis A problem that has attracted quite some attention is the existence ofinteractions between photosynthetic and respiratory electron transport chains. Early observations (Hoch et al., 1963; Jones and Myers, 196) showed that light, in the presence of the Photosystem II inhibitor DCMU, inhibits uptake. This was interpreted to mean that photosynthesis and respiration share common components, and many similar observations followed. The existence of complete respiratory chains in both CM and ICM at least in some cyanobacteria, however, sheds new light on the question of the interaction between respiratory and photosynthetic electron transport chains. Clearly, these interactions must be quite different for the CM and the ICM. There is now convincing evidence that the CM contains no chlorophyll. This was originally postulated by Peschek and Schmetterer (1978), when they showed that only the ICM but not the CM could be degraded by photooxidation. The technique for the separation of CM and ICM developed by Omata and Murata (1983) showed this to be correct. Completely purified CM contains no trace of chlorophyll although the resence of precursors of chlorophyll has been reported

Chapter 13 Cyanobacterial Respiration (Hinterstoisser et al., 1988; Peschek et al., 1989a). Therefore, photosynthetic electron transport is confined to the ICM and only in the ICM is the question of whether respiratory and photosynthetic electron transport chains share common components meaningful. The evidence presented above clearly favors an affirmative answer. In fact, it can be said that in the ICM of cyanobacteria both respiration and photosynthesis are part of the branched bioenergetic membrane-bound electron transport chain, the only difference being that photosynthetic branches (comprising essentially Photosystems I and II) are only active in the light. Evidence for a possible interaction between photosynthetic electron transport and the respiratory chain in the CM comes from light-induced proton efflux. This was first reported by Scholes et al. (1969). Its mechanism(s) is not yet fully clear. Both a proton-translocating respiratory terminal oxidase (see Section VII A) and an ATPase [in Synechococcus sp. strain PCC 6301 (Peschek et al. 1986) and in Anabaena variabilis (Scherer and Böger, 1984; Scherer et al., 1984; Nitschmann and Peschek, 1985)] have been found in the CM so that at least two mechanisms for the interaction between photosynthetic and CM-localized electron transport are possible, which are not mutually exclusive (see also Nitschmann and Peschek, 1985). ATP formed by the ATPase in the ICM could be hydrolyzed with the ATPase in the CM (Scherer et al., 1988c), and reduced pyridine nucleotides formed by non-cyclic photosynthesis could feed electrons into the respiratory chain in the CM (Hawkesford et al., 1983).

C. Final Remarks Knowledge of cyanobacterial respiration is not yet satisfactory, desite the fact that cyanobacteria play an important role in the biosphere and that the interaction between the two most important bioenergetic processes, oxygenic photosynthesis and aerobic respiration, should attract considerable general interest. The development of cyanobacterial molecular biology and genetics has presented the possibility of defining exactly which proteins are involved in cyanobacterial respiration. However, some of the few reports on cyanobacteria specifically dealing with genes important in respiration have produced quite urprising results that cannot yet be reconciled with the biochemical data mostly obtained on cell fragments (‘membrane preparations’,

429 ‘membrane particles’, ‘spheroplasts’). Clearly, much work lies ahead.

Acknowledgments I would like to thank Dr W. Hessler for his permission to use his personal computer for the preparation of this manuscript. Dr D. Laudenbach kindly sent us Synechococcus sp. strain 158-7, the cytA-derivative of Synechococcus sp. strain PCC 7942 described in Laudenbach et al. (1990). The excellent technical assistance of Mr O. Kuntner is gratefully acknowledged. The work on cyanobacterial respiration in my laboratory has been supported by grants P7015 and S6007 from the Fonds zur Förderung der wissenschaflichen Forschung in Österreich.

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434 in the phototrophic cyanobacteria (blue-green algae)? In: Kotyk A (ed) Highlights in Modern Biochemistry, Vol. 1, pp 893– 902. International Science Publ, Zeist, Netherlands Peschek GA, Wastyn M, Trnka M, Molitor V, Fry IV and Packer L (1989c) Characterization of the cytochrome c oxidase in isolated and purified plasma membranes from the cyanobacterium Anacystis nidulans. Biochemistry 28: 3057–3063 Peltier G and Schmidt GW (1991) Chlororespiration: An adaptation to nitrogen deficiency in Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 88: 4791–4795 Peterson RB and Burris RH (1978) Hydrogen metabolism in isolated heterocysts of Anabaena 7120. Arch Microbiol 116: 125–132 Pistorius EK, Jetschmann K, Voss H and Vennesland B (1979) The dark respiration of Anacystis nidulans. Production of HCN from histidine and oxidation of basic amino acids. Biochim Biophys Acta 585: 630–642 Pueyo JJ, Revilla C, Mayhew SG and Gómez-Moreno C (1992) Complex formation between ferredoxin and reductase from Anabaena PCC 7119: Cross-linking studies. Arch Biochem Biophys 294: 367–372 Rhoads DM and McIntosh L (1991) Isolation and characterization of a cDNA clone encoding an alternative oxidase protein of Sauromatum guttatum (Schott). Proc Natl Acad Sci USA 88: 2122–2126 Rippka R, Waterbury J and Cohen-Bazire G (1974) A cyanobacterium which lacks thylakoids. Arch Mikrobiol 100: 419–436 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 31: 225– 274 Sancho J, Peleato ML, Gomez-Moreno C and Edmondson DE (1988) Purification and properties of oxidoreductase from the nitrogen-fixing cyanobacteria Anabaena variabilis. Arch Biochem Biophys 260: 200–207 Sandmann G (1986) Formation of plastocyanin and cytochrome c-553 in different species of blue-green algae. Arch Microbiol 145: 76–79 Sandmann G and Böger P (1980) Copper induced exchange of plastocyanin and cytochrome c-553 in cultures of Anabaena variabilis and Plectonema boryanum. Plant Sci Lett 17: 417– 424 Sandmann G and Malkin R (1983a) NADH and NADPH dehydrogenases from the blue-green alga, Aphanocapsa. Arch Microbiol 136: 49–53 Sandmann G and Malkin R (1983b) NADH and NADPH as electron donors to respiratory and photosynthetic electron transport in the blue-green alga, Aphanocapsa. Biochim Biophys Acta 725, 221–224 Sandmann G and Malkin R (1984) Light inhibition of respiration is due to a dual function of the cytochrome complex and the plastocyanin/cytochrome c-553 pool in Aphanocapsa. Arch Biochem Biophys 234: 105–111 Scherer S (1990) Do photosynthetic and respiratory electron transport chains share redox proteins? Trends Biochem Sci 15: 458–462 Scherer S and Böger P (1982) Respiration of blue-green algae in the light. Arch Microbiol 132: 329–332 Scherer S and Böger P (1984) Vanadate-sensitive proton efflux by filamentous cyanobacteria. FEMS Microbiol Lett 22: 215–

G. Schmetterer 218 Scherer S, Stürzl E and Böger P (1981) Arrhenius plots indicate localization of photosynthetic and respiratory electron transport in different membrane regions of Anabaena. Z Naturforsch 36c, 1036–1040 Scherer S, Stürzl E and Böger P (1982) Interaction of respiratory and photosynthetic electron transport in Anabaena variabilis Kütz. Arch Microbiol 132: 333–337 Scherer S, Stürzl E and Böger P (1984) Oxygen-dependent proton efflux in cyanobacteria (blue-green algae). J Bacteriol 158: 609–614 Scherer S, Sadowski H and Böger P (1987) Bioenergetic studies of the cyanobacterium Anabaena variabilis. Z Naturforsch 42c: 1280–1284 Scherer S, Almon H and Böger P (1988a) Interaction of photosynthesis, respiration and nitrogen fixation in cyanobacteria. Photosynth Res 15: 95–114 Scherer S, Alpes I, Sadowski H and Böger P (1988b) Ferredoxinoxidoreductase is the respiratory NADPH dehydrogenase of the cyanobacterium Anabaena variabilis. Arch Biochem Biophys 267: 228–235 Scherer S, Riege H and Böger P (1988c) Light-induced proton efflux of the cyanobacterium Anabaena variabilis. In: Rogers LJ and Gallon JR (eds) Biochemistry of the Algae and Cyanobacteria, pp 121–129. Clarendon Press, Oxford, UK Scherer S, Häfele U, Krüger GHJ and Böger P (1988d) Respiration, cyanide-insensitive oxygen uptake and oxidative phosphorylation in cyanobacteria. Physiol Plant 72: 379–384 Shen G, Boussiba S and Vermaas WFJ (1993) Synechocystis sp. PCC 6803 strains lacking Photosystem I and phycobilisome function. Plant Cell 5: 1853–1863 Shinozaki K, Ohme M, Tanaka M, Wakasugi T, Hayashida N, Matsubayashi T, Zaita N, Chungwongse J, Obokata J, Yamaguchi-Shinozaki K, Ohto C, Torozawa K, Meng B-Y, Kusuda J, Takaiwa F, Kato A, Tohdoh N, Shimada H, and Sugiura M (1986) The complete nucleotide sequence of the tobacco chloroplast genome: Its gene organization and expression. EMBO J 5: 2043–2049 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. PCC 7002 and initial characterization of an interposon mutant. J Bacteriol 175: 3343–3352 Schmetterer G, Alge D and Gregor W (1994) Deletion of cytochrome c oxidase genes from the cyanobacterium Synechocystis sp. PCC 6803: Evidence for alternative respiratory pathways. Photosynth Res, in press Scholes PP, Mitchell P and Moyle J (1969) The polarity of proton translocation in some photosynthetic micro-organisms. Eur J Biochem 8: 450–454 Schrautemeier B, Böhme H and Böger P (1984) In vitro studies on pathways and regulation of electron transport to nitrogenase with a cell free extract from heterocysts of Anabaena variabilis. Arch Microbiol 137: 14–20 Schrautemeier B, Böhme H and Böger P (1985) Reconstitution of a light-dependent nitrogen-fixing and transhydrogenase system with heterocyst thylakoids. Biochim Biophys Acta 807: 147–154

Chapter 13 Cyanobacterial Respiration Serrano A, Giménez P, Scherer S and Böger P (1990) Cellular localization of cytochrome in the fixing cyanobacterium Anabaena variabilis. Arch Microbiol 154: 614–618 Shin M (1971) Ferredoxin-NADP reductase from spinach. Meth Enzymol 23: 440–447 Smart LB, Anderson SL and McIntosh L (1991) Targeted genetic inactivation of the Photosystem I reaction center in the cyanobacterium Synechocystis sp. PCC6803. EMBO J 10: 3289–3296 Smith AJ, London J and Stanier RY (1967) Biochemical basis of obligate autotrophy in blue-green algae and thiobacilli. J Bacteriol 94: 972–983 Sone N, Tano H and Ishizuka M (1992) The cytochrome c oxidase genes in thermophilic cyanobacterium Synechococcus vulcanus. In: Murata N (ed) Research in Photosynthesis, Vol II, pp 571–574. Kluwer, Dordrecht Steinmetz AA, Castroviejo M, Sayre RT and Bogorad L (1986) Protein PS II-G. J Biol Chem 261: 2485–2488 Steinmüller K (1992a) Nucleotide sequence and expression of the ndhH gene of the cyanobacterium Synechocystis sp. PCC6803. Plant Mol Biol 18: 135–137 Steinmüller K. (1992b) Identification of a second psaC gene in the cyanobacterium Synechocystis sp. PCC6803. Plant Mol Biol 20: 997–1001 Steinmüller K and Bogorad L (1990) Identification of a psbGhomologous gene in Synechocystis sp. PCC6803. In: Baltscheffsky M (ed) Current Research in Photosynthesis, Vol III, pp 12557–12560. Kluwer, Dordrecht, Netherlands Steinmüller K, Ley AC, Steinmetz AA, Sayre RT and Bogorad L (1989) Characterization of the ndhC-psbG-ORF 157/159 operon of maize plastid DN A and of the cyanobacterium Synechocystis sp. PCC6803. Mol Gen Genet 216: 60–69 Steinmüller K, Ellersiek U and Bogorad L (1991) Deletion of the psbG1 gene of the cyanobacterium Synechocystis sp. PCC6803 leads to the activation of the cryptic psbG2 gene. Mol Gen Genet 226: 107–112 Stürzl E, Scherer S and Böger P (1982) Reconstitution of electron transport by cytochrome c-553 in a cell-free system of Nostoc muscorum. Photosynth Res. 3: 191–201 Stürzl E, Scherer S and Böger P (1984) Interaction of respiratory and photosynthetic electron transport, and evidence for membrane-bound pyridine-nucleotide dehydrogenases in Anabaena variabilis. Physiol Plant 60: 479–483 Takahashi Y, Shonai F, Fujita Y, Kohchi T, Ohyama K and Matsubara H (1991) Structure of a co-transcribed gene cluster, ndh1-frxB-ndh6-ndh4L, cloned from the filamentous cyanobacterium Plectonema boryanum. Plant Cell Physiol 32: 969– 981 Tano H, Ishizuka M and Sone N (1991) The cytochrome c oxidase genes in blue-green algae and characteristics of the deduced protein sequence for subunit II of the thermophilic cyanobacterium Synechococcus vulcanus. Biochem Biophys Res Commun 181: 437–442 Trebst A and Harth E (1970) On a new inhibitor of photosynthetic electron-transport in isolated chloroplasts. Z Naturforsch 25b: 1157–1159 Trnka M and Peschek GA (1986) Immunological identification of cytochrome oxidase in membrane preparations of the cyanobacterium Anacystis nidulans. Biochem Biophys Res Commun 136: 235–241

435 Van der Oost J, Van Walraven HS, Bogerd J, Smit AB, Ewart GD and Smith GD (1989) Nucleotide sequence of the gene proposed to encode the small subunit of the soluble hydrogenase of the thermophilic unicellular cyanobacterium Synechococcus PCC 6716. Nucl Acids Res 17: 10098 Van der Plas J, Bovy A, Kruyt F, De Vrieze G, Dassen E, Klein B and Weisbeek P (1989) The gene for the precursor of plastocyanin from the cyanobacterium Anabaena sp. PCC 7937: Isolation, sequence and regulation. Mol Microbiol 3: 275–284 Viljoen CC, Cloete F and Scott WE (1985) Isolation and characterization of an NAD(P)H dehydrogenase from the cyanobacterium, Microcystis aeruginosa. Biochim Biophys Acta 827: 247–259 Walker JE (1992) The NADH:ubiquinone oxidoreductase from bovine heart mitochondria. Q Rev Biophys 25: 253–324 Wastyn M, Achatz A, Trnka M and Peschek GA (1987) Immunological and spectral characterization of partly purified cytochrome oxidase from the cyanobacterium Synechocystis 6714. Biochem Biophys Res Commun 149: 102–111 Wastyn M, Achatz A, Molitor V and Peschek GA (1988) Respiratory activities and cytochrome oxidase in plasma and thylakoid membranes from vegetative cells and heterocysts of the cyanobacterium Anabaena ATCC29413. Biochim Biophys Acta 953: 217–224 Webster GC and Frenkel AW (1953) Some respiratory characteristics of the blue-green alga, Anabaena. Plant Physiol 28: 63–69 Weiss H, Friedrich T, Hofhaus G and Preis D (1991) The respiratory-chain NADH dehydrogenase (complex I) of mitochondria. Eur J Biochem 197: 563–576 Wildman RB and Bowen CC (1974) Phycobilisomes in bluegreen algae. J Bacteriol 117: 866–881 Winkenbach F and Wolk CP (1973) Activities of enzymes of the oxidative and the reductive pentose phosphate pathways in heterocysts of a blue-green alga. Plant Physiol 52: 480–483 Wolk CP (1973) Physiology and cytological chemistry of bluegreen algae. Bacteriol Rev 37: 32–101 Yagi T (1991) Bacterial NADH-quinone oxidoreductases. J Bioenerg Biomembr 23: 211–225 Yao Y, Tamura T, Wada K, Matsubara H and Kodo K (1984) Spirulina reductase. The complete amino acid sequence. J Biochem 95: 1513–1516. Yu L, Golbeck JH, Zhao J, Schluchter WM, Mühlenhoff U, and Bryant DA (1992) The PsaE protein is required for cyclic electron flow around Photosystem I in the cyanobacterium Synechococcus sp. PCC 7002. In: Murata N (ed) Research in Photosynthesis, Vol I, pp 565–568. Kluwer, Dordrecht Yu, L, Zhao J, Mühlenhoff U, Bryant DA, and Golbeck JH (1993) PsaE is required for cyclic electron flow around Photosystem I in the cyanobacterium Synechococcus sp. PCC 7002. Plant Physiol 103: 171–180 Zhang L, McSpadden B, Pakrasi HB and Whitmarsh J (1992) Copper-mediated regulation of cytochrome and plastocyanin in the cyanobacterium Synechocystis 6803. J Biol Chem 267: 19054–19059 Zhang L, Pakrasi HB and Whitmarsh J (1994) Photoautotrophic growth of the cyanobacterium Synechocystis sp. PCC 6803 in the absence of cytochrome and plastocyanin. J Biol Chem 269: 5036–5042

Chapter 14 The Biochemistry and Molecular Regulation of Carbon Dioxide Metabolism in Cyanobacteria F. Robert Tabita Department of Microbiology and The Biotechnology Center, The Ohio State University, 484 West 12th Avenue, Columbus, Ohio 43210-1192, USA Summary I. Introduction II. Pathways of Carbon Dioxide Metabolism Fixation: Structure, Function, and Regulation of Activity III. Enzymes of A. Ribulose Bisphosphate Carboxylase/Oxygenase (RubisCO) 1. Assembly of Cyanobacterial RubisCO 2. Properties of L Subunits 3. Properties of the S Subunits 4. Attempts to Modify RubisCO Specificity and Activity 5. An Approach to Biological Selection of Mutant Cyanobacterial RubisCO B. Phosphoribulokinase, Phosphoenolpyruvate Carboxylase, and Other Enzymes 1. Phosphoribulokinase 2. Phosphoenolpyruvate carboxylase 3. Other Enzymes C. Regulation of Key Enzymes in Vivo IV. Organization of Reductive Pentose Phosphate Cycle Genes V. Regulation of Expression of Reductive Pentose Phosphate Cycle Genes A. RubisCO Genes Metabolism B. prk and Other Genes Important for Gene Expression in the Environment C. VI. Conclusion Acknowledgments References

437 438 438 439 439 441 443 447 450 451 452 452 453 455 455 457 460 460 461 461 462 462 462

Summary Carbon dioxide is a greenhouse gas whose accumulation in the biosphere has been the cause for increasing concern. is also the source for virtually all organic carbon on Earth and its efficient assimilation is directly related to agricultural productivity. As organisms which often depend on the reduction and assimilation of as their prime source of carbon, cyanobacteria have become important tools for gaining an understanding ofthe biochemical and molecular mechanisms involved. These organisms take on added significance because the entire process, the catalysts employed and their structural genes, are to some extent quite similar to those of higher plants. Because of the relative ease in using molecular techniques and transferring genetic information in cyanobacteria, there are many advantages to these organisms as models for green plant metabolism. There are also differences that make cyanobacteria fascinating in their own right. Over the last few years, there has been a tremendous upsurge in interest in Cyanobacterial fixation research. Important insights relative to the biochemistry of the process have emerged, fueled by the revolution in molecular biology. This chapter thus considers the current state of the field and reviews the many important contributions that have been made on this interesting and important area of cyanobacterial research. D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 437–467. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

438 I. Introduction Carbon dioxide is the preferred source of carbon for cyanobacteria, whether they are capable of photoheterotrophic metabolism or have the ability to grow chemoheterotrophically in the dark. Indeed, those organisms that are capable of significant dark heterotrophic metabolism, such as Nostoc sp. strain MAC, immediatelyinitiatephototrophicmetabolism when light is supplied (Bottomley and Van Baalen, 1978a,b). It will thus be the function of this review to consider the biochemical and molecular mechanisms bywhichcyanobacteriaeffectivelymetabolize carbon dioxide. Much has been learned since the last review on this subject (Tabita, 1987), particularly relative to the key enzymes and the genes which encode these proteins. New genes have been discovered and knowledge relative to the mechanisms cyanobacteria employ to regulate the activity of the various fixation enzymes has increased almost exponentially. Advances concerning the transport of and the role of carboxysomes have also been impressive and are reviewed elsewhere in this volume (Chapter 15). Thus, this chapter will be confined to an in depth review of metabolism, since all cyanobacteria must deal with this ubiquitous gas. II. Pathways of Carbon Dioxide Metabolism Without exception, the Calvin reductive pentose phosphate pathway is the primary route by which cyanobacteria metabolize and reduce The history of the discovery of this metabolic pathway, particularly in cyanobacteria, may be gleaned from papers of the early to mid-1950s (see Norris et al., 1955, and Linko et al., 1957). A perusal of any biochemistry textbook indicates that the Calvin cycle is very much related to the oxidative pentose phosphate pathway or hexose monophosphate shunt, except many of the reactions proceed in the reverse direction. Indeed, the addition of two unique Abbreviations: CA1P – 2-carboxyarabinitol monophosphate; CD – circular dichroism; cpn – chaperonin; CPS – carbamylphosphate synthase; DTT – dithiothreitol; L – large subunit; OTC – ornithine transcarbamylase; PAGE – polyacrylamide gel electrophoresis; PCR – polymerase chain reaction; PEP – phosphoenolpyruvate; PGPase–phosphoglycolate phosphatase; PRK – phosphoribulokinase; RubisCO – ribulose 1, 5bisphosphate carboxylase/oxygenase; RuBP – ribulose 1,5bisphosphate; S – small subunit; SDS – sodium dodecylsulfate.

F. Robert Tabita enzymatic steps, along with some other idiosyncrasies, allows the reductive pentose phosphate pathway to proceed in apurely biosynthetic direction. Obviously reducing the most oxidized source of carbon on Earth to the level of organic carbon does not come cheaply to a photosynthesizing cell. Thus, copious amounts ofreducing power (reduced pyridine nucleotide) and ATP are required and it is readily apparent that the reactions of reduction must be linked to effective energy production through photosynthesis. Indeed, a recurring theme throughout this chapter will be attempts to relate in vivo control of enzymatic activity and gene regulation to net photosynthetic activity. The key and unique reactions of the Calvin cycle that allow biosynthesis to occur are catalyzed by the enzymes phosphoribulokinase (PRK = ATP:Dribulose 5-phosphate 1-phosphotransferase; EC 2.7.1.19) and ribulose 1,5-bisphosphate (RuBP) carboxylase/oxygenase (RubisCO = 3-phospho-Dglycerate carboxylase [dimerizing]; EC 4.1.1.39). RubisCO catalyzes the primary reaction of fixation, that is the actual fixation of onto the enediol form ofRuBP. PRK activity is important for the regeneration ofthe acceptor, RuBP (Fig. 1). Subsequent metabolism of the products of the carboxylase reaction (3-phosphoglyceric acid) via a series ofreductive and transformation reactions leads to the formation of Ru-5-P, the substrate of PRK. The net effect of this cycle is to synthesize one molecule of triose-phosphate from three molecules of When is the gaseous substrate, RubisCO catalyzes an internal monooxygenase reaction to form 2phosphoglycolate as well as 3-phosphoglyceric acid. Glycolate is produced through the action ofa specific phosphoglycolate phosphatase (Section III, B). This compound is further metabolized and/or excreted by cyanobacteria (Fig. 1). Details of the metabolism of glycolate and its significance in cyanobacteria have been reviewed recently (Colman, 1989), and the reader is urged to consider this work for further details of glycolate and photorespiratory metabolism in cyanobacteria. Moreover, it is apparent that cyanobacteria may catalyze anaerobic or anoxygenic photosynthetic metabolism through the reactions of the Calvin pathway. The reader is urged to consult the review of Padan (1979) for further details of this interesting development. Unfortunately, the reasons for poor growth under anoxygenic conditions of photosynthesis have yet to be explained. Finally, the previous review (Tabita, 1987)

Chapter 14

Metabolism

discussed evidence for substantial fixation through an alternative pathway, using the enzymes carbamyl phosphate synthase and ornithine carbamyl phosphate transferase to convert to citrulline, a major product of fixation by some cyanobacteria. Since 1986, considerable additional information has emerged relative to the occurrence of this pathway (Section III B, 3). In addition, much may be assimilated via to form acids, usually aspartate and malate, through the action of phosphoenolpyruvate (PEP) carboxylase. This reaction and its regulation are also discussed (Section III B, 2 and V B). However, since the Calvin pathway is by far the predominant mechanism for assimilating and knowledge has advanced appreciably, the bulk of the following discussion will consider genetic

439

and biochemical aspects of route.

fixation through this

III. Enzymes of Fixation: Structure, Function, and Regulation of Activity

A. Ribulose Bisphosphate Carboxylase/ Oxygenase (RubisCO) In order to learn more about the mechanism of fixation, there has been a primary focus on the structure, function, and general enzymology of RubisCO since it catalyzes the actual fixation of Certainly, the relative ease with which this enzyme may be isolated in homogenous form from

440 eucaryotes (in which it is the major protein found in cell extracts (Miziorko and Lorimer, 1983)) and most procaryotes (Tabita, 1988) has contributed to rather extensive investigations on this enzyme. Recent studies have verified the original contention of Calvin of the importance of the enediol of RuBP in the carboxylasereaction;furtherdetailsofthemechanism have been summarized (Andrews and Lorimer, 1987). In addition to catalyzing the carboxylation of RuBP to yield two molecules of phosphoglyceric acid, it has been found that the same enzyme will catalyze an RuBP-dependent fixation reaction such that the enediol of RuBP is cleaved as a consequence of addition to yield one molecule each of 3-phosphoglycerate and 2-phosphoglycolic acid (Fig. 1). This alternative reaction was discovered after it was found that molecular is a competitive inhibitor with respect to (Bowes and Ogren, 1972). Thus, RubisCO is an enzyme with a dual function in catalyzing two importantsteps inmetabolism,namely fixation and the oxygenolytic cleavage of RuBP to yield phosphoglycolate. In addition, recent studies indicate that pyruvate is a minor product of the reaction catalyzed by several sources of RubisCO, including the enzyme of Synechococcus sp. strain PCC 6301 (Andrews and Kane, 1991). The ability of the enzyme to discriminate between and at any particular concentration of and is referred to as the specificity factor or partition coefficient where and are the for carboxylation and oxygenation, respectively, and and are the Michaelis constants for the oxygenase and carboxylase, respectively. Before carboxylation or oxygenolysis of RuBP can take place, there is an obligatory activation step that must occur in which the enzyme is carbamylated at a specific lysine residue by ‘activator’ this carbamate is stabilized by divalent cations. Thus, the activated enzyme is in effect a ternary complex or It should be stressed that the same polypeptide chain catalyzes both reactions. Aside from its obvious importance in fixation, the enzyme acquired added significance because of the oxygenase function. This is due to the fact that phosphoglycolate, one of the products of the oxygenase reaction, is further oxidatively metabolized to serine in plants, and in the process both and are released (up to 50% ofthe carbon fixed). As a result, growth is drastically reduced because much carbon and nitrogen is wasted. This is the biochemical basis for the poorproductivity

F. Robert Tabita ofcertain important crops such as soybean—i.e., the oxygenase reaction is favored over the carboxylase simply because there is more than in air. Thus, the same enzyme catalyzes the first reaction in the two competing pathways of carbon fixation and respiratory carbon dissipation. Every RubisCO that has been isolated is capable of catalyzing the oxygenase reaction, including the enzyme from anaerobic bacteria, leading to the general belief that the oxygenase function is an inherent property ofthe protein and proceeds as a consequence of the enzymatic formation of the enediol of RuBP (Lorimer and Andrews, 1973). For this reason alone, it is not surprising that this bifunctional enzyme is now one of the most heavily studied enzymes. RubisCO is also undoubtedly the most abundant protein found on Earth (Ellis, 1979). If one could selectively shut off the oxygenase function, or somehow favor the carboxylase reaction by either mutation or chemical means, the practical applications would be enormous. It is well established that RubisCO from all eucaryotes and virtually all procaryotes is a large protein with a of 500–600,000 (Type I RubisCO) (Miziorko and Lorimer, 1983; Tabita, 1988). However, this protein is an exceedingly poor catalyst, having a turnover number of 1000 to 2000 mol fixed of enzyme per min. Given the importance of this enzyme, and its low turnover or it is not surprising that organisms that use as sole carbon source synthesize large quantities of RubisCO. Type I RubisCO is composed oftwo distinct peptides (eight of each): a large catalytic subunit and a small subunit Convincing proof that L is the catalytic subunit came from several active-site directed studies (Miziorko and Lorimer, 1983), as well as the isolation of RubisCO from the bacteria Rhodospirillum rubrum (Tabita and McFadden, 1974a,b) and Rhodobacter sphaeroides (Gibson and Tabita, 1977), which were found to be aggregates of L subunits only (Type II RubisCO). Indeed, the R. rubrum enzyme has become the paradigm for structure-function studies ofRubisCO (Andersson et al., 1989). The solving ofthe structure of the R. rubrum RubisCO, a dimer of L subunits (Tabita and McFadden, 1974b), provided the impetus and baseline information required to determine the structure of the more complex spinach enzyme (Knight et al., 1990). From these studies it became apparent that the core of the enzyme might be considered a tetramer of dimers such that the basic structure of the R. rubrum enzyme is repeated four

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times. There is an extensive interface area between the subunits which bury almost half of the accessible surface areas ofboth L and S. Each S makes contact with three different L subunits from two different dimers and also contacts two neighboring S subunits. From these studies it became apparent that the molecule is a cube-shaped molecule with local 422 symmetry where the 4-fold axis arranges four dimers into a core of eight L subunits The S subunits are clustered into groups of four subunits each Thus, the enzyme is in reality a molecule. A number of important residues of functional significance have been identified, including residues that represent important contact points between subunits. The X-ray structure thus confirmed the initial hypotheses that residues of S that are conserved in evolutionarily diverse S molecules may be important for assembly and/or catalysis by L (Tabita, 1987). More recently, the structure of the Synechococcus sp. strain PCC 6301 RubisCO has been solved (see Fig. 2; Newman and Gutteridge, 1993). The basic ‘plant-like’ enzyme arrangement is notably altered by a nearly two-fold greater solventfilled cavity. The structural basis for the known catalytic ‘idiosyncrasies’of cyanobacterial RubisCO (see Sections III A, 2 to III A, 4) may now be approached with vigor. In cyanobacteria, there is a single Type I enzyme found in all organisms examined; previous findings of only large subunits in some cyanobacteria have not proven to be correct (seeTabita, 1987). Since the large (rbcL) and small (rbcS) subunit genes appear to be part of a single transcriptional unit in all cases (Nierzwicki-Bauer et al., 1984; Shinozaki and Sugiura, 1985), several groups have taken advantage of this gene arrangement by placing the rbcLrbcS genes behind strong Escherichia coli promoters to facilitate the expression and isolation ofrecombinant cyanobacterial RubisCO. From these results, it became apparent that large quantities of fully active and properly assembled enzyme could be obtained, particularly when the Synechococcus sp. strain PCC 6301 genes are placedbehind the lac promoter (Tabita and Small, 1985; Kettleborough et al., 1987; McFadden and Small, 1989; Lee and Tabita, 1990; Gutteridge, 1991) or, to a lesser extent, the pL promoter (Gatenby et al., 1985) or other promoters (Smrcka et al., 1991a). Low yields of correctly assembled recombinant Anabaena sp. strain PCC 7120 RubisCO were also obtained (Gurevitz et al., 1985). In high-density fed-batch fermentations,

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it has been found that up to 2 g of recombinant Synechococcus sp. strain PCC 6301 RubisCO per liter of media may be produced under optimal conditions (B.A. Read, G. Kleman, W.R. Strohl, and F.R. Tabita, unpublished results). Thus, from 9 to 10 liters of a high-density fermentation, with a conservative yield of purified enzyme of 25%, one can in principle isolate about 4 g of pure enzyme. This markedly contrasts to the 1.5 to 2.0 g ofpurified enzyme that were obtained from a 200-liter fermentor (Newman and Gutteridge, 1990).

1. Assembly of Cyanobacterial RubisCO From the above discussion, it is apparent that large quantities of active recombinant RubisCO, containing the proper ratio of large and small subunits, may be obtained. It is actually astounding that such a large and complex protein is properly assembled in E. coli. Recently, however, it has become apparent that a distinct family of accessory proteins, so-called ‘molecular chaperones’ are required to guide or assist oligomeric proteins to assemble into productive or ‘catalytically correct structures’ (Ellis, 1990). These molecular chaperones, or chaperonin proteins, bind to newly synthesized proteins to form stable complexes and prevent ‘incorrect’ or unproductive structures from forming. Molecular chaperones may also facilitate the formation of native protein structures from unfolded biologically inert polypeptides. The complexes are then dissociated by other proteins, usually using the energy obtained from ATP hydrolysis, and thus the chaperones are not found in the final, functional oligomeric protein structure. ForRubisCO, aprotein calledthe RubisCObinding protein was found to be associated with newly synthesized chloroplast-encoded RubisCO L subunits (Barraclough and Ellis, 1980; Roy et al., 1982). Sequence analysis of the RubisCO-binding protein showed remarkable homology to the product of the groEL gene of E. coli, known to be essential for cell viability and the assembly of bacteriophage (Hemingsen et al., 1988). The groEL gene is closely associated with the groES gene, both of which comprise the groE operon ofE. coli (Zeilstra-Ryalls et al., 1991). For plant RubisCO, several studies indicated that the RubisCO-binding protein may be involved in the assembly of the RubisCO structure within the chloroplast organelle (Roy and Cannon, 1988).The R. rubrum RubisCO gene (cbbM) and the cyanobacterial RubisCO genes (rbcLrbcS)

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F. Robert Tabita

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may be expressed in E. coli, with the formation of highly active or RubisCO (closely homologous to plant recombinant enzymes, respectively. Thus, in an important study Goloubinoff et al. (1989) showed that the GroEL (cpn60) and GroES (cpn10) proteins of E. coli were required for the expression of both and RubisCO.An assembly pathway (Fig. 3) to convert nascent unfolded large and small subunits to the structure was proposed, recognizing that the dimer is the basic structural motif (Andersson et al., 1989). However, it is not certain, at this time, whether the chaperonin proteins are required for the assembly of octamersfrom dimers, or if small subunit assembly is chaperonin mediated; free S has been shown to assemble with octamers spontaneously in vitro (Paul et al., 1991). Interestingly, the assembly of cpn60 to itself required and ATP hydrolysis; the reaction is stimulated by cpn 10 (Lissin et al., 1990). In plants and green algae that have been studied extensively, the L and S subunits of RubisCO are encoded by the chloroplast and nuclear genomes,

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respectively. Because of extensive processing of L and S in eucaryotes, problems with assembly, and other complex reasons (Hemmingsen et al., 1988), eucaryotic rbcL and rbcS genes have not been expressed in E. coli to form productive recombinant proteins. However, recent experiments show that plant (Brassica napus) and support the assembly of cyanobacterial (presumably Synechococcus sp.) RubisCO in E. coli strains harboring the proper Brassica napus chaperonins and cyanobacterial RubisCO genes on inducible E. coli expression vectors (Cloney et al., 1992a,b).

2. Properties of L Subunits Various L preparations, which retain the capacity to reconstitute with S, have been obtained after stripping S from L octamers under rather gentle procedures (Andrews and Ballment, 1983; Asami et al., 1983; Incharoensakdi et al., 1985; Jordan and Chollet,

444 1985). However, all of these preparations appeared to retain residual amounts ofS. Andrews (1988) was able to show that recombinant Synechococcus sp. L, prepared from extracts of E. coli, catalyzed weak RubisCO activity, conclusively demonstrating that S is not absolutely required for activity. In this earlier study, however, the L subunits were unstable, perhaps because of poor expression, and were not successfully purified. Lee and Tabita (1990) and Smrcka et al. (1991a) were able to prepare purified and stable recombinant cyanobacterial L using different strategies to express the Synechococcus sp. strain PCC 6301 rbcL gene by itself. Smrcka et al. (1991a) constructed an expression vector where the rbcL gene was under control ofa heat inducible promoter, producing Synechococcus sp. L with a 10-amino acid amino-terminal extension. A similar strategy led to the construction of a vector, under control of the same promoter, that yielded recombinant S with a 6-amino acid extension at the amino-terminus. Lee and Tabita (1990) prepared rbcL and rbcS expression vectors from plasmid pCS75 (Tabita and Small,

F. Robert Tabita 1985), which itself had previously been constructed so that the rbcL and rbcS genes were behind the lac promoter (Fig. 4). The 2.2 kb fragment, that codes for the L and the S subunits of RubisCO of Synechococcus sp. strain PCC 6301, was liberated from pCS75 by digestion with PstI and subsequently cloned into pTZ18R. The 1.5 kbp fragment containing the rbcL gene was liberated by partial digestion with HindIII and cloned into pUC 18. The 0.7 kb HindIIIPstI fragment containing rbcSwas cloned into pUC9. In all plasmid constructs, neither the L nor the S subunit gene was found in the same reading frame as the gene so that the expressed gene products are non-fusion, wild-type proteins. The L subunits from crude extracts of E. coli MV1190 (pBGL520) assembled and exhibited an electrophoretic mobility that approximated that of the holoenzyme in nondenaturing polyacrylamide gels. Trace amounts ofcarboxylase activity were found in crude extracts of E. coli MV1190 (pBGL520), but no activity was found in extracts ofE. coli MV1190 (pBGL535). When L- and S-containing extracts from

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the two strains were mixed, considerable amounts of RubisCO activity were obtained and extracts from cells not containing S did not stimulate activity. These results indicated that the active holoenzyme was reconstituted from the separately expressed recombinant proteins and further indicated that this system could be used to monitor the purification of recombinant L or S subunits. Eventually, L was purified from E. coli MV1190 (pBGL520) crude extracts. However, upon electrophoresis under nondenaturing conditions, the purified L subunits did not exhibit a distinct band like that observed in E. coli MV1190(pBGL520) crude extracts without the addition of either bovine serum albumin or E. coli crude extract (Fig. 5). In the absence of exogenous protein, dissociation of the protein was observed from purified L subunit preparations, which appeared as a diffuse area at the lower part of a non-denaturing gel (Fig. 5, lane 10). At this time it is not clear what causes the apparent dissociation of the L subunits during electrophoresis in the absence of added protein. However, interactions with heterologous proteins might be important to maintain the L in the structure. Perhaps, S subunit binding sites on L,

445 when exposed to solvent, contribute to the loose association of L under the conditions of electrophoresis. Under normal conditions, both subunit genes are cotranscribed in procaryotes, and the enzyme is assembled immediately into a stable holoenzyme (Tabita, 1988). Thus, the effects we have observed with the purified L subunits undoubtedly reflect the ability of S subunits to maintain the enzyme in its most stable conformation. When increasing amounts of crude S were added to purified L, a saturation effect on RubisCO activity was noted (Lee and Tabita, 1990; Gutteridge, 1991; Smrcka et al, 1991a). The purified holoenzyme of Lee and Tabita (1990) had a specific activity of 2470 nmol of fixed By contrast, the specific activity of purified L was only but increased to 310 in the presence of saturating amounts of crude recombinant S and ranged from 200 to (mg of large with different preparations. Thus, the reconstituted RubisCO activity was not as great as that obtained with the native recombinant enzyme. Either the purified L was not as functional as the L subunits of

446 the octameric core of the molecule or the crude S preparation introduced an additional complicating factor. Smrckaetal.(1991a)found withtheirconstruct that S subunits were highly enriched in the 100,000 × g pellet of the E. coli extract. This recombinant S could be extracted with urea and renatured to a form highly competent to combine with L; indeed the specific activity of the reconstituted enzyme was extremely high. It has also recently been found with this expression system that S is enriched in the 100,000 ×g pellet fraction of E. coli MV1190 (pBGL535) and that S could be extracted with urea for subsequent reconstitution with purified L (K. Horken and F.R. Tabita, unpublished results). The Xray structure of in the complete absence of S, as well as site-directed mutagenesis studies, should provide information relative to the conformation required for assembly and give insights to molecular changes that occur when S interacts with the core. The isolation of homogenous recombinant RubisCO has been a stimulus to determine catalytic properties in the absence ofS. Ofcourse, experiments with the and Type II enzymes of R. rubrum and R. sphaeroides have indicated there are considerable differences in catalytic properties compared to Type I RubisCO (reviewed by Tabita, 1988). Since Type II L shows little homology to L from Type I enzymes, the preferred approach would be to undertake comparative experiments with L from enzymes. Obviously, the structure is the most stable aggregate of L that may be isolated since this is the predominant form which assembles and has enzymatic activity. Recent experiments with both and Synechococcus sp. strain PCC 6301 RubisCO, as well as heterologous enzymes comprised of Synechococcus sp. L and Alcaligenes eutrophus S (Lee et al., 1991b) and Synechococcus sp. L and diatom (Cylindrotheca sp. strain N1) S (Read and Tabita, 1992b) indicate that RuBP does not inhibit the activation (carbamylation) of the enzyme by In most cases, the tight binding of RuBP to Type I RubisCO prevents subsequent activation by and The experiments with the hybrid enzymes, using S from enzymes inhibited by RuBP, and the isolated cyanobacterial protein, indicate that neither the absence of S nor the presence of S from different organisms changes the response to RuBP. The response of unactivated enzyme to RuBP must be a distinct property of the particular source of L employed. Thus, there mustbe some precise structural

F. Robert Tabita difference in L subunits from different Type I enzymes; a challenging direction for future experimentation will be to determine the molecular basis for RuBP sensitivity/insensitivity. Inhibition of carbamylated RubisCO with the metabolite 6-phosphogluconate is thought to result from the competition of this compound with RuBP at the active site (Tabita and McFadden, 1972; Chu and Bassham, 1973; Jordan et al., 1983). However, single subunit Type II enzymes from Rhodospirillum rubrum and Rhodobacter sphaeroides are insensitive to this ligand, suggesting that S might influence the binding ofthis and other effectors (reviewed in Tabita, 1988). Thus, experiments were initiated with the fully activated Synechococcus sp. strain PCC 6301 holoenzyme and a heterologous enzyme comprised of Synechococcus sp. L and A. eutrophus S (Lee et al., 1991b). When the activity of each of these enzymes was titrated with increasing quantities of 6-phosphogluconate, it was found that the Synechococcus sp. enzyme was about 12-fold more sensitive than the holoenzyme or the hybrid enzyme. Thus, it is apparent that the previously observed differential effect of 6-phosphogluconate on Type I RubisCO is not due to the presence of S, since Type II RubisCO enzymes containing only L are insensitive to 6-phosphogluconate. Thus the conformation and/or source of L alone is the determining factor. Further experiments indicated that the for of and Synechococcus sp. strain PCC 6301 RubisCO was about the same and that E. coli proteins influenced the elution of complexes from gel filtration columns, emphasizing the requirement for purified preparations of for catalytic studies (Smrcka et al., 1991a). Moreover, RubisCO bound CABP more effectively than (Andrews, 1988; Smrcka et al., 1991b). Experiments with partially purified yielded and values that ranged from 200 to and 300 to (Andrews, 1988; Gutteridge, 1991), respectively, which are, in the case of RuBP, considerably higher than the purified holoenzyme the for the enzyme ranges from (Read and Tabita, 1992a,b). The specificity factor for the and enzymes was about the same, indicating to Gutteridge (1991) that the L subunits might exclusively be involved in the ability of the enzyme to discriminate between the two gaseous substrates, and

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Metabolism

3. Properties of the S Subunits Smrcka et al. (1991a) obtained nearly homogeneous preparations of recombinant S from a urea-extracted particulate fraction of an E. coli lysate. Interestingly, there was a requirement for low levels of urea for maintaining S in solution and urea was required in the reconstitution reaction of S with L; urea concentrations of 5–10 mM did not interfere with assembly or enzyme activity. It was further found by these authors that S affects the apparent for carbamylation, particularly between pH 7.7 to 8.3, and it was hypothesized that S might exert its effect by influencing the specific lysine residue involved in carbamylation (Lys-198). In a subsequent paper (Smrcka et al. 1991b), these authors implicate S in inducing a conformation of L that more effectively binds CABP, and by extension the enediol of RuBP. Presumably, these conformational changes are the result of alterations in the interactions between L to L or L to S. Since S prevents the release of tightly bound inhibitors such as CABP, and perhaps related compounds of physiological significance such as CA1P and XuBP, Smrcka et al. (1991b) suggest that the enzyme RubisCO activase might reverse the influence of S such that these inhibitor compounds may be released from the enzyme. Cyanobacterial RubisCO activase has recently been prepared (Li et al., 1993; Section IV) so that future experiments with purified S, and RubisCO activase, and the compounds noted above, should prove interesting. Comparisons of the deduced amino acid sequence of small subunits from diverse species, from bacteria to plants, indicate that there are several residues that are invariant. Indeed, there are three major regions of sequence homology, an area near the N-terminus, one in the center of the primary structure, and a third region near the C-terminus. There is also a hairpin loop (HPIN) (Knight et al., 1989) found only in S of higher plants and green algae, e.g. in organisms where the rbcL and rbcS genes do not form an operon. Studies with chimeric constructs of pea S lacking HPIN, and a chimera of Synechococcus sp. S that contains the HPIN region, indicate that HPIN is required for assembly with pea L after the chimeric fusion proteins were imported into pea chloroplasts (Wasmann et al., 1989). In addition, Smrcka et al. (1991b) quote unpublished results that indicate that the HPIN sequence might be responsible for the considerably tighter binding of ligands to plant RubisCO compared to cyanobacterial RubisCO.

447

Further studies on this region are awaited with great interest; indeed the and values for diatom (Cylindrotheca sp. strain N1) RubisCO (Read and Tabita, 1992b), which lacks the HPIN region in S, are quite similar to the values obtained for green algal RubisCO, and to some extent plant RubisCO (Jordan and Ogren, 1981). Because of the overall similarity in the primary structure of the cyanobacterial and plant RubisCOs, (Curtis and Haselkorn, 1983; Shinozaki et al., 1983; Reichelt and Delaney, 1983) and due to the fact that recombinant cyanobacterial RubisCO may be produced in large quantities in E. coli, the cyanobacterial enzyme has received considerable attention. Thus, this system has been used in site-directed mutagenesis studies to facilitate investigations of the role that S might have on catalysis and there have been several efforts to determine the role of specific residues in the conserved regions. The initial experiments were performed by Voordouw et al. (1987) who replaced Trp-55 and Trp-58 of the Synechococcus sp. strain PCC 6301 S with phenylalanine residues. The W55F and W58F enzymes, when partially purified, contained the correct ratio of L and S subunits and were found to have about 40% the activity of the wild-type enzyme. There was no change in or the specificity factor McFadden and Small (1989) constructed double and triple amino acid substitutions in consecutive residues of the first conserved region of the Synechococcus sp. strain PCC 6301 S. They found that the activity of the recombinant proteins containing each of the paired substitutions was affected. Values from 6–71% of the activity of the wild-type enzyme were obtained, and the T14A/ S16A and S16A/Y17D double substitutions appeared to be particularly deleterious to activity. Fitchen et al. (1990) prepared single amino acid residue substitutions in key residues of the three conserved regions of the Anabaena sp. strain PCC 7120 S. Mutated rbcS and wild-type rbcL were coexpressed on plasmids in the same E. coli host, resulting in the assembly of RubisCO containing wild-type L and mutant S. None of the resulting recombinant mutant enzymes (E11V, W55R, P59H, and Y84N) contained detectable enzymatic activity in crude extracts. After sucrose density gradient centrifugation, a presumptive species was isolated and found to be devoid of S. The mutant S proteins appeared in a 10,000 ×g particulate fraction of a crude extract, as did significant amounts of the wild-

448 type S, suggesting that the mutants had a great effect on the interaction ofL and S. The substantial amount of wild-type S in the particulate fraction in this system is consistent with the large amounts of particulate recombinant Synechococcus sp. S found in crude extracts from E. coli strains separately expressing rbcS (Smrcka et al., 1991a). When the Anabaena sp. S substitutions were extrapolated to changes in comparable residues in the threedimensional structural model of spinach RubisCO, changes of Glu-11 and Trp-53 to an uncharged or a charged residue, respectively, were found to result in potential steric mispairing ofL and S. The changes in Pro-59 and Tyr-84 are thought to disrupt intersubunit or intrasubunit hydrophobic interactions (Fitchen et al., 1990). However, if Glu-11 of the Anabaena sp. and Synechococcus sp. enzymes (Glu-13 of the spinach enzyme) is important to the integrity of the structure, the Type I enzymes of the chemolithotrophic bacteria Alcaligenes eutrophus (Andersen and Caton, 1987) and Xanthobacter flavus (Meijer et al., 1991), the purple nonsulfur photosynthetic bacterium Rhodobacter sphaeroides (Gibson et al., 1991), and the enzymes from the non-green chromophytic and rhodophytic algae (see Hwang and Tabita, 1991) must somehow compensate, since the primary structure ofS in this region is completely different in the enzyme from these organisms. Lee et al. (1991) also prepared single amino acid substitutions of conserved residues of Synechococcus sp. strain PCC 6301 S. In this study, a 1.5 kb EcoRI fragmentwas removed fromthe mutagenizedplasmid containing the rbcLrbcS genes, leaving a 0.7 kb EcoRI-PstI fragment, containing all of rbcS, and a small part ofthe 3' region ofrbcL. This small fragment was then sequenced and recombined with a 1.5 kb EcoRI fragment from a nonmutagenized rbcLcontaining plasmid. This procedure ensured that there was absolutely no possibility for secondary mutations in the large region of the rbcL gene that was exchanged, and obviated the need to sequence the entire rbcL gene. The 0.7 kb EcoRI-PstI fragment that was mutagenized was completely sequenced to verify that only the target site was changed to yield the expected single amino acid substitution. In all cases, varying effects on enzyme activity were noted, ranging from 30 to 74% ofthe specific activity ofthe wild-type enzyme. In this and subsequent studies (Read and Tabita, 1992a), mutations in each of the three conserved regions yielded two different types of altered RubisCO depending on the residue changed

F. Robert Tabita and the nature of the substitution. In one case, ‘interaction’ mutations were obtained in which the single amino acid substitutions had an effect on the interaction of S with L. These mutant enzymes were found to dissociate on sucrose gradients or upon nondenaturing polyacrylamide gel elecrrophoresis. However, mutant S and wild-type L, separated by sucrose density gradient centrifugation, could reconstitute yielding RubisCO of the same relative activity found in crude extracts. Mutations of this type are S16D and L21E of the first conserved region, Y54S ofthe second conserved region (Lee et al., 1991), and R88Q of the third conserved region (Read and Tabita, 1992a). These studies, and recent X-ray structural investigations ofthe highly conserved spinach enzyme (Knight et al., 1990), indicate that Ser-16, Leu-21, and Tyr-54 play important roles in the interaction of L and S. A second type of modified RubisCO has been obtained. In this instance, although RubisCO activity was substantially affected, the single amino acid substitutions in S do not appear to perturb intersubunit interactions or induce any gross conformational changes. These ‘activity’ mutants include the W55P enzyme (Voordouw et al., 1987; Lee et al., 1991); and the I87V, R88K, F92L, and G91V mutant enzymes, containing changes in the third conserved region (Read and Tabita, 1992a). Because the enzymes maintained their integrity, they were easily purified to homogeneity for subsequent physical and kinetic analyses and each altered enzyme appeared to have the normal complement ofL and S. The four mutants of the third conserved region were analyzed by circular dichroism (CD). The CD spectra from 190 to 250 nm of the G91V, R88K, and I87V mutants differed very little from that of the wild-type enzyme. The only exception was the F92L mutant protein whose CD spectrum differed significantly. In addition, the G91V enzyme showed interesting properties in that it precipitated in 0.1 M phosphate buffer, pH 7.5; subsequent studies showed that the G91V enzyme, unlike the wild-type protein, did not adsorb to phenylSuperose columns in the presence of 1.7 M ammonium sulfate. All four mutants showed a reduction in activity and three ofthe mutant enzymes exhibited values of the wild-type enzyme, with the G91V and F92L enzymes exhibiting 16 and 9% of the wild-type activity, respectively. Although the specificityfactor was not significantly affected for any of these mutants, significant and interesting changes in the Michaelis constants for and

Chapter 14

Metabolism

RuBP were obtained. In particular, the G91V and F92L enzymes showed substantial decreases in and Extrapolation to the three-dimensional structural model of the spinach enzyme provided interesting insights as to the structural basis for the changes in enzymatic properties; this has been fully discussed (Read and Tabita, 1992a). The bottom-line conclusion from these studies is that small subunits substantially influence key catalytic parameters. The most profound effects were on the for both the carboxylase and oxygenase reactions and alterations ofthe Michaelis constants for each substrate. Recent structural studies (Schneider et al., 1990a,b) indicate that S may have the ability to modulate substrate binding by inducing conformational changes at the active site; the mutant studies support this idea. Examples of an interaction mutant and an activity mutant are indicated by the R88K and R88Q enzymes, respectively, which clearly behave differently on sucrose density gradients (Fig. 6). The R88K enzyme remains intact and activity is eluted at the expected position of an enzyme. However, the R88Q enzyme dissociates to populations of L and S, and separate fractions do not have activity unless they are combined to reconstitute the complex. These studies obviously point out that different amino acid substitutions at the same residue can yield both types of mutant enzyme. Paul et al. (1991) separately expressed Synechococcus sp. strain PCC 6301 rbcL and mutant rbcS genes in E. coli and reconstituted either the recombinant mutant or wild-type enzymes as previously described (Lee et al., 1991; Lee andTabita 1990). Residues of the first conserved region were examined using the separate expression/reconstitution system such that effects on enzyme activity could be distinguished from any effect on the interaction between subunits. Although Paul et al. (1991) maintain that the separate expression system is required to quantitate binding, Lee et al. (199la) clearly show that L and S, co-expressed from rbcLrbcS-containing plasmids and purified by sucrose-gradient centrifugation, may be reconstituted to a level ofactivity that is comparable to that obtained in crude extracts. Obviously, if one co-expresses rbcL and mutant rbcS on the same plasmid behind a single promoter and the subsequently assembled recombinant enzyme dissociates into L and S upon purification, it would be a simple manner to quantitate the capacity for mutant S to saturate wild-type L. Thus, the separate expression system is not absolutely

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required. Indeed, working only with crude extracts, one would not detect mutations that affect activity yet do not perturb gross intersubunit interactions (Read and Tabita, 1992a). The separate expression system is most effective in determining the kinetic properties of reconstituted enzyme when purified components are employed (Smrcka et al. 1991b) and might alleviate the large errors inherent in measurements of the binding properties of L and S in crude extracts (Paul et al., 1991). The data of Paul et al. (1991) point out the importance ofThr-14 andTyr-17 of S, which the spinach RubisCO structure model predicts will interact with key residues of L that are at or near residues of the active site. Finally, all the recombinant enzymes containing single mutations of S which could be purified and which were found to influence activity (Voordouw et al., 1987; Lee et al., 1991a; Read and Tabita, 1992a) had no effect on the specificity factor ofthe enzyme. These results seemed at first glance to support the contention that the ability of the enzyme to discriminate between the gaseous substrates is not influenced by S and is a property ofonly L (Andrews et al., 1984; Andrews and Lorimer, 1985). However, only a limited number of mutant enzymes and/or hybrid enzymes have been studied so that a definitive judgment is difficult. Indeed, Chen et al. (1990) provide convincing evidence that a nuclear-encoded

450 protein may influence the specificity factor of Chlamydomonas reinhardtii RubisCO. More recently, experiments with a recombinant hybrid Synechococcus sp. L/Cylindrotheca sp. (diatom) S enzyme [originally prepared via the coexpression of the respective rbcL and rbcS genes on a single vector (Hwang and Tabita, 1989)], indicate that S may play an important role in the ability of the enzyme to discriminate between and (Read and Tabita, 1992b). The of the hybrid enzyme (pVTAC223 enzyme) was about 5% that of the wild-type Synechococcus sp. strain PCC 6301 enzyme and there were significant alterations of the Michaelis constants forRuBP, and Most interesting, however, was the finding that the specificity factor of the hybrid enzyme was increased about 60% over that of the Synechococcus sp. holoenzyme. Another hybrid enzyme, preparedby fusing the Synechococcus sp. strain PCC 6301 rbcL to rbcS from Olisthodiscus luteus, did not result in any meaningful change in specificity factor. These studies also led to the finding that the substrate specificity factor of the Cylindrotheca sp. RubisCO and other chromophytic and rhodophytic enzymes is unusually high (Read and Tabita, 1992c, 1994). The original hypothesis of Schneider et al. (1990a,b) that interactions of S with L may influence substrate specificity (Knight et al., 1990) are thus compatible with the kinetic results obtained with the pVTAC223 hybrid enzyme.

4. Attempts to Modify RubisCO Specificity and Activity As previously discussed, cyanobacterial RubisCO bears close homology to the plant enzyme, yet the cyanobacterial enzyme exhibits a characteristic high and a relatively low specificity factor compared to the enzyme from plants. However, the ease by which the cyanobacterial RubisCO operon may be manipulated to produce recombinant protein in E. coli has become increasingly attractive to RubisCO biochemists since the original expression studies were reported (Christeller et al., 1985; Gatenby et al., 1985; Gurevitz et al., 1985; Tabita and Small, 1985). The importance of the amino-terminus of L was indicated by the studies of Kettleborough et al. (1987), who showed that an alteration at this region through replacement by an unrelated sequence, resulted in a recombinant RubisCO that exhibited a 10-fold increase in Subsequent studies indicated that

F. Robert Tabita thisenzyme,whenactivatedwith showed altered absorption properties that were reflective ofthe altered kinetic properties (Brändén et al. 1990). Haining and McFadden (1990) showed inactivation of the carbamylated Synechococcus sp. strain PCC 6301 recombinant protein by the arginine-specific reagent phenylglyoxal. Inactivation was substantially reduced in the presence of RuBP and the kinetics of inactivation indicated that one or more Arg residues might be essential for activity. Analysis of sequence homologies of a variety of RubisCO enzymes indicated that Arg-292 is completely conserved and recent structural studies implicate this residue in the 5-phosphate CABP binding site (Knight et al., 1990). A number of single amino acid substitutions at residue 292 subsequently confirmed the importance of arginine for activity and/or the correct conformation. A similar approach, based on prior photomodification studies (Mogel and McFadden, 1989) implicated Ser-376, which is positioned to bind CABP or RuBP in the spinach enzyme (Andersson et al. 1989; Knight et al., 1990). Again, a series of site directed Synechococcus sp. strain PCC 6301 mutant proteins were generated, S376C, S376T, and S376A, each of which had their carboxylase activity reduced by 99% or greater (Lee and McFadden, 1992). In all cases the was substantially increased and the authors hypothesized that these alterations may have been due to an effect on the specificity factor. The isolation of mutant Chlamydomonas reinhardtii RubisCO enzymes containing alterations in the specificity factor, all of which mapped in loop 6 ofthe barrel of L, stimulated Parry et al. (1992) to construct enzymes containing changes in residues of this region. The alterations were to the corresponding amino acids in pea and maize RubisCO. In the DKAS338-341EREI mutant, the specificity factor was increased 6% and the A340E mutant exhibited a 17% reduction in specificity factor. The specificity factor ofthe V331A mutant was not changed, although the activity was substantially decreased. The V331A enzyme is particularly interesting because this substitution in the Chlamydomonas reinhardtii enzyme resulted in a 37% decrease in the specificity factor (Chen and Spreitzer, 1989). Since loop 6 is thought to form a flexible flap which folds over the active site during catalysis (Knight et al., 1990), the current results with the Synechococcus sp. enzyme, although stimulating, are somewhat difficult to interpret since all the kinetic constants were not presented, no evidence of the purity of the

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recombinant proteins is given, and the values are about 3- to 5-fold higher than what has been recently reported in both crude and purified preparations of wild-type enzymes (Morell et al., 1992; Paul et al., 1991; Read and Tabita, 1992a). Finally, a recent study appeared which concentrated on residue 306 (residue 309 of the plant enzyme) since this amino acid is the one residue that differs in the RubisCO of closely related genera of and plants. In RubisCO, there is a Met and in the enzyme there is an Ile residue at residue 309. Since the cyanobacterial RubisCO resembles the plant enzyme in its relatively high and since it also contains Ile at this position, Morell et al. (1992) made substitutions in this region of the Synechococcus sp. strain PCC 6301 enzyme to determine the effect on the kinetic properties ofthe resultant recombinant enzyme. The I306M and I306L enzymes showed no difference in assembly properties or in the or although the level of expression for thesemutantenzymes wasextremely low. Substitution of other residues yielded enzymes that were defective in the ability to assembly or fold and did not produce a distinguishable protein band after nondenaturing gel electrophoresis. From the spinach structural model and the data of Morel et al. (1992), it appears that residue 306 (309) must be highly hydrophobic and have a long side-chain in order for the enzyme to be maintained in a proper conformation. Nevertheless, the issue of whether Ile or Met at position 306 contributestodifferencesin or RubisCO cannot be judged by making single substitutions in the Synechococcus sp. enzyme since there are several other differences in the primary structure of cyanobacterial and plant RubisCO.

5. An Approach to Biological Selection of Mutant Cyanobacterial RubisCO A long-term goal has been to develop procedures for isolating mutations that alter the biological properties of cyanobacterial RubisCO. Although ofgreat utility, one of the problems with site-directed mutagenesis is that one needs to know what to mutate and the results thus tend to be somewhat predictable. Random mutagenesis procedures have historically yielded important insights relative to structure and function; however, there is an obvious need for having some means to select for mutations of interest. In considering a feasible approach to this goal, it was reasoned that a RubisCO-deficient strain of an

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organism that normally uses this enzyme to fix might be employed as a host for complementation of randomly mutagenized RubisCO DNA. The use of cyanobacterial strains capable of heterotrophic growth in the dark was considered, since deletion of the key fixation enzyme might still allow for growth under conditions where is not the sole carbon source. However, as pointed out in a previous review (Tabita, 1987), there are several studies which indicate that the level of RubisCO is invariant, even when cyanobacteria are cultured photoheterotrophically or heterotrophically in the dark (Carr, 1973). Thus, deletion of rbcLrbcS genes, even in heterotrophic cyanobacteria, may be lethal. Indeed, attempts to construct rbcL deletion strains in Synechocystis sp. strain PCC 6803 and Synechococcus sp. strain PCC 7002 consistently failed (D. A. Bryant, personal communication). Several other bacteria, however, offer interesting possibilities, particularly representatives of the purple nonsulfur photosynthetic bacteria, organisms which are extremely versatile in their metabolism and which are known to grow under conditions under which RubisCO is both dispensable and indispensable (Tabita, 1988). In addition, the unique arrangement of the RubisCO genes, found downstream from other key Calvin cycle structural genes in Rhodobacter sphaeroides (Tabita et al., 1992) sets this organism apart from other facultative autotrophs for which the Calvin genes have been mapped. Thus, a RubisCO mutant would not exhibit polarity effects on other fixation genes and it might be expected that R. sphaeroides could be complemented by exogenous RubisCO genes from several organisms (Falcone and Tabita, 1991; 1993). To use the R. sphaeroides RubisCO deletion strain (strain 16) as a host for cyanobacterial rbcLrbcS genes, a vector-promoter system had to be developed that would allow one to conveniently express the foreign genes in this background. This was accomplished by D.L. Falcone, who successfully demonstrated complementation of the R. sphaeroides deletion strain to growth using the vectorcontainingtheSynechococcus sp. strain PCC 6301 rbcLrbcS genes (Falcone and Tabita, 1991). Furthermore, the promoter-rbcLrbcS expression vector facilitated determination of the physiological consequences of photosynthetic growth of R. sphaeroides synthesizing foreign (cyanobacterial) RubisCO. This work also suggested that strain 16 could be used to assess the physiological significance of directed or engineered RubisCO

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enzymes. Growth of the strain 16 - Synechococcus sp. strain PCC 6301 rbcLrbcS construct [strain 16 (pRPS-75)] differed significantly from either the wild-type strain HR or strain 16 complemented with bacterial RubisCO genes, in that a pronounced lag occurred when cells were bubbled with argon under photoheterotrophic conditions (Fig. 7). Adding exogenous or eliminating the bubbling with argon prevented the lag and allowed the organism to grow well, suggesting that argon bubbling disperses the endogenous generated by the metabolism of malate. Strain 16 (pRPS-75) grows well and without a lag under phototrophic conditions under an atmosphere of and yet the wild-type strain and other complemented strains (containing noncyanobacterial RubisCO genes) are considerably less sensitive to the endogenous concentration under photoheterotrophic growth conditions. Thus, the requirement for added in strain 16 (pRSP-75) appears to be due to the expression and utilization of the cyanobacterial rbcLrbcS genes. Cyanobacterial RubisCO normally is found in a special environment, the carboxysome. Moreover, cyanobacteria contain active mechanisms that function under low conditions (see Chapter 15), and carbonic anhydrase seems to function to produce high levels of at the site of carboxylation, the carboxysome. For these reasons, the poor of the cyanobacterial enzyme normally does not limit dependent growth. However, in the R. sphaeroides strain 16 environment, there does not appear to be an active concentrating mechanism and there are no carboxysomes. Thus, the cyanobacterial RubisCO in this environment is sensitive to the external levels of supplied to the cells and the organism’s growth is directly related to the ability ofthe enzyme to acquire enough making this an excellent system to screen for mutations in RubisCO function. Indeed, the strain 16(pRPS-75) system is being used to screen for second-site internal suppressor mutations in RubisCO function that overcome a primary mutation, and strains that express cyanobacterial RubisCO with altered affinity are also being selected (F. R. Tabita, unpublished results). These recent studies suggest that any breakthroughs made by using site-directed mutagenesis to affect discrimination by RubisCO could be tested in vivo to determine the physiological consequences of such changes, by noting the response of R. sphaeroides strain 16 complemented with the altered genes. The bacterial system could thus serve as a screen prior to

undertaking similar, more laborious efforts in higher plants. The interesting ‘cyanorubum’ system described by Pierce et al. (1989) also provides a potential way to select for mutants in the R. rubrum RubisCO since this enzyme is expressed in Synechococcus sp. strain PCC 6803 and is subject to oxidative modification (Cook and Tabita, 1988; Cook et al. 1988). The advantage of the R. sphaeroides system is that growth is possible under photosynthetic conditions in the absence of oxygen or under chemoautotrophic conditions in the presence of oxygen (Tabita, 1988); both conditions require RubisCO activity. To reiterate, preliminary results indicate that the strain 16 system is sensitive to RubisCO function in vivo and could provide an exceedingly facile screening system for mutations that might later be introduced into plants.

B. Phosphoribulokinase, Phosphoenolpyruvate Carboxylase, and Other Enzymes 1. Phosphoribulokinase There is a paucity ofinformation on the properties of enzymes, other than RubisCO, important for metabolism (see Tabita, 1987). Although it is one of the two unique enzymes of the Calvin cycle, there have been few detailed investigations ofthe function, structure, and regulation of cyanobacterial phosphoribulokinase (PRK). The light-dependence of PRK

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activity in vitro was first noted in Synechococcus sp. strain PCC 6301 (Duggan and Anderson, 1975), which was consistent with previous steady-state labelingexperimentsperformed inSynechocystis sp. strain PCC 6308 (Pelroy and Bassham, 1972). Subsequent experiments showed that a strong reducing agent such as dithiothreitol (DTT) replaced the requirement for light in vitro for various cyanobacterial PRK enzymes (Duggan andAnderson, 1975; Pelroy and Bassham, 1976; Crawford et al., 1984; Marsden and Codd, 1984; Serra et al., 1989). Unlike the situation in anoxygenic phototrophic and chemolithoautotrophic bacteria (Tabita, 1988), the cyanobacterial enzyme was not regulated by reduced pyridine nucleotide. It is well established that PRK and other enzymes of intermediary metabolism are under the control of light and the ferredoxin/ thioredoxin system in plants (Buchanan, 1980). Key sulfhydryl groups of all of these enzymes are reversibly oxidized and reduced through the lightdependent ferredoxin/thioredoxin system and this is probably the basis for the light and/or DTT-mediated control ofPRK activity in cyanobacteria. The deduced primary structure of PRK from Synechocystis sp. strain PCC 6803 (Su and Bogorad, 1991) indicates that Cys-19 and Cys-41 correspond to the regulatory Cys-16 and Cys-55 residues ofspinach PRK (Krieger and Miziorko, 1986; Milanez et al., 1991). The phenotype of the bright light-sensitive (BRLS) mutant of Synechocystis sp strain PCC 6803 was found to be due to a single-base alteration in the prk gene, resulting in substitution ofphenylalanine for serine at residue 222 of the protein (Su and Bogorad, 1991). In crude preparations, this mutation resulted in a 10-fold reduction in enzymatic activity while the was about 7-fold higher than the wild-type enzyme; the was not altered. The authors postulated that the mutation may affect the hydrophilicity and flexibility of the enzyme since the mutant enzyme migrated slightly differently than the wild-type enzyme upon SDS-PAGE. Further studies with purified enzymes should be most interesting. It is also intriguing that there is not a comparable serine residue in the deduced sequences of phototrophic and chemoautotrophic bacterial PRK enzymes (Kossman et al., 1989; Gibson et al., 1990; Meijer et al., 1990), further substantiating the rather distinct enzymatic properties of these two types of PRK (Tabita, 1980, 1981). The deduced amino acid sequence of the Synechocystis sp. strain PCC 6803 PRK is closely related to that of plants (about 60% identity) but is distantly related to the

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bacterial enzymes (see Fig. 8). There have been two reports of the isolation of purified PRK from cyanobacteria. Marsden and Codd (1984) purified the enzyme from Chlorogloeopsis fritschii. The enzyme was found to have a native molecular weight of 230,000 with a single 40,000 subunit, as determined by SDS-PAGE. ADP, AMP, L-aspartate, PEP, and L-malate all substantially inhibited the enzyme. Curiously, the enzyme from Anabaena cylindrica had a native molecular weight of 72,000 and was found to contain two 43,000 and 26,000 polypeptide chains although the authors were not certain if the smaller protein might have been a copurifying contaminant (Serra et al., 1989). The Anabaena cylindrica enzyme was not inhibited by PEP, aspartate, or malate but was substantially activated by DTT and Anabaena sp. thioredoxin. Finally, immunoelectron microscopy showed that PRK is not localized in the carboxysomes of the cyanelles of Cyanophora paradoxa or Glaucocystis nostochinearum, but is found throughout the thylakoid region of the cyanelles (Mangeney et al., 1987). PRK was also not found in the carboxysomes of Chlorogloeopsis fritschii (Lanaras and Codd, 1982).

2. Phosphoenolpyruvate carboxylase Phosphoenolpyruvate (PEP) carboxylase is an extremely important enzyme for the metabolism of in cyanobacteria since its activity leads to products of fixation which may represent up to 20% of the carbon assimilated in some organisms (Coleman and Colman, 1980). In Coccochloris peniocystis, PEP carboxylase is the only enzyme (Owttrim and Colman, 1986). As discussed in Section V, B., the gene (ppc) encoding PEP carboxylase has been cloned and sequenced from a number of cyanobacteria and studies have been initiated to prepare ppc mutants to assess the contribution and importance of the enzyme to net metabolism (Luinenburg and Coleman, 1990). The enzymology of the cyanobacterial enzyme has not been studied extensively. However, comparison of the deduced amino acid sequence of the cyanobacterial enzyme to the plant and E. coli proteins indicates that there are several homologous regions which may be of functional significance (Katagiri et al., 1985; Luinenburg and Coleman, 1992). Consistent with hypotheses relative to the importance of the Cterminal region in catalysis, the amino acid sequence

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of this part of the molecule is the most conserved (Jiao et al., 1990). The Coccochloris peniocystis enzyme showed Michaelis-Menten kinetics with respect to PEP, and The was which like the for RubisCO is rather high and probably reflects the large light-dependent accumulation of inorganic carbon species, which is at or above 4 mM in this organism (Coleman and Colman, 1981). A residue that has been implicated in the allosteric response of the E. coli enzyme to fructose 1,6-bisphosphate (Thr-232) is conserved in both theAnabaena sp. strain PCC 7120 (Luinenburg and Coleman, 1992) and Synechococcus sp. strain PCC 6301 enzymes, however the cyanobacterial (Coccochloris peniocystis) enzyme is not activated by this ligand (Owttrim and Colman, 1986). Thus, this residue by itself does not appear to be sufficient for allosterism ofthe cyanobacterial enzyme. Finally PEP carboxylase has been found in several strains (Colman, 1989), making itvery likely thatthis enzyme contributes to overall metabolism of all cyanobacteria.

3. Other Enzymes A major amount of may be incorporated into the carbamyl portion of citrulline (reviewed in Tabita, 1987). Subsequent experiments performed by Chen et al. (1987a,b) showed that carbamyl phosphate synthase (CPS) is an important enzyme that links ammonia assimilation to fixation in cyanobacteria. Furthermore, the compound DL-7azatryptophan triggered heterocyst differentiation in Anabaena sp. strain CA and caused a transient accumulation of intracellular citrulline and alanine from bicarbonate or ornithine in Anabaena sp. strain 1F (Chen and Tabita, 1987a,b). These results, in combination with in vitro measurements, implicated CPS and ornithine transcarbamylase activity (OTC) in the intracellular accumulation of citrulline. This glutamine-dependent CPS activity provides an explanation for the lack of detectable glutamate synthase in Anabaena 1F, and along with glutaminase (Chen and Tabita, 1987b), provides for an alternative to the usual ammonia assimilatory pathway of cyanobacteria (Meeks et al., 1977). A recent report describing the properties of a glnA mutant of Synechococcus sp. strain PCC 7002 further indicates the importance of CPS in cyanobacterial nitrogen and carbon metabolism (Wagner et al., 1993). CPS is present in both heterocysts and vegetative

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cells of Nostoc sp. strain PCC 73102 (Lindblad, 1989) as is OTC (Lindblad, 1992; Jansson et al., 1993). Since citrulline has been identified as one of the majorN-containingcompoundstransported from the root to the cycad in the Macrozamia sp.-Nostoc sp. symbiosis (Pate et al., 1988), it is likely that future studies of the regulation of CPS will be extremely interesting. An enzyme that deserves serious consideration for further study in cyanobacteria is phosphoglycolate phosphatase (PGPase). Obviously, this enzyme is required for the further metabolism of phosphoglycolate, the product of the RuBP oxygenase reaction of RubisCO. Norman and Colman (1991) cite evidence that the absence of PGPase is lethal to oxygen-evolving photosynthetic organisms. Under high oxygen concentrations, cyanobacteria excrete substantial amounts of glycolate—up to 60% of the carbon assimilated (Renström and Bergman, 1989). PGPase was initially detected inAnabaena variabilis and Anacystis nidulans (Husic and Tolbert, 1985). More recently, the Coccochloris peniocystis enzyme was shown to have properties similar to PGPase from higher plants and green algae, however the for phosphoglycolate for the cyanobacterial enzyme was relatively high (Norman and Colman, 1991). Other enzymes of the Calvin cycle and other pathways related to metabolism have been studied to some extent; however, since the last review on cyanobacterial fixation (Tabita, 1987), there has been little published that relates to the major theme of the current review. An excellent earlier review ofgeneral cyanobacterial carbon metabolism, including numerous previous studies of additional enzymes, is available (Smith, 1982).

C. Regulation of Key Enzymes in Vivo There have been several studies that relate to the physiological control of enzymes important to assimilation. With regard to RubisCO, any consideration of its regulation in vivo must consider the fact that before RubisCO is functional, for either fixation or fixation, the enzyme must be carbamylated or activated to form the ternary complex. There are several ways in which RubisCO maybe converted to theactiveform.Various phosphorylated compounds have been shown to fully activate partially active RubisCO from plants, and the same metabolites serve as competitive inhibitors with respect to RuBP when the enzyme is in its fully

456 activated form (Jensen and Bahr, 1977). As pointed out earlier (Tabita, 1987), the work of Pelroy and Bassham (1972,1976) has shown that Synechocystis sp. strains PCC 6308 and PCC 6714 accumulate fairly high levels of these metabolites, particularly 6phosphogluconate, that cause an immediate cessation of fixation during light-dark transitions. Using permeabilized cells of Anabaena sp. strain CA, Synechocystis sp. strain PCC 6714, and Synechococcus sp. strain PCC 7002, it was shown that 6phosphogluconate and NADPH were most effective in regulating both the partially active and the fully activated enzyme in the cell (Tabita and Colletti, 1979). These whole-cell assays are relevant because RubisCO activity is measured in close approximation to the native cellular environment. Indeed, these in situ measurements indicated that intracellular RubisCO was more sensitive to effectors than the purified enzyme or freshly isolated crude preparations. Obviously, there is much to be learned about the factors which regulate the activity of RubisCO and photosynthetic assimilation in cyanobacteria. Various metabolites, such as the RubisCO inhibitor 2-carboxyarabinitol monophosphate (CA1P), are produced by certain plants in the dark (Gutteridge et al., 1986; Berry et al., 1987). Inhibitory compounds are also produced as a result of RuBP isomerization during catalysis (Zhu and Jensen, 1991a, b). Other phosphorylated metabolites, particularly RuBP itself, can block RubisCO catalysis under certain conditions. Thus, terrestrial plants have evolved a system to remove these high affinity phosphorylated inhibitor compounds from RubisCO. The enzyme RubisCO activase (Salvucci et al., 1985, 1986) catalyzes the removal of these inhibitory substances under conditions of active photosynthesis and thus represents at least one means to maximize RubisCO activity in vivo (Portis, 1990; 1992). Little is understood, however, concerning the factors that regulate RubisCO activity in cyanobacteria. Recently, the rca gene, encoding the RubisCO activase from the marine cyanobacterium Anabaena sp. strain CA (Li et al., 1993), was isolated sequenced and overexpressed; this is the first report describing a procaryotic rca gene. The role that RubisCO activase plays in cyanobacteria is a question that may be approached by both genetic (see Section IV), and biochemical studies. It is apparent that the Anabaena sp. strain CA RubisCO activase will have a different specificity and function than the activase of plants

F. Robert Tabita and green algae since noncarbamylated cyanobacterial RubisCO (E) does not bind RuBP (R) (Lee et al., 1991b). Since it is the E-R form of the enzyme that is used as a convenient substrate for plant RubisCO activase in vitro and that seems to be a substrate in vivo as well (Portis, 1990), cyanobacterial RubisCO activase must have some function other than catalyzing the ATP-dependent removal of RuBP. It is possible that RubisCO activase functions to facilitate the removal of inhibitor ligands such as 6phosphogluconate or CA1P from RubisCO, an important role for the enzyme in plants (Robinson and Portis, 1988). Moreover, so long as RuBP is in excess, cyanobacterial RubisCO does not exhibit the phenomenon of ‘fallover’ or decline in RubisCO activity in timed assays (Lee et al., 1991b; Read and Tabita, 1992b). Thus, the cyanobacterial RubisCO activase obviously is not required to remove fallover products from RubisCO as; it is in higher plants (Robinson and Portis, 1989). At this time, the most apparent and logical role for RubisCO activase in cyanobacteria would be as an agent to influence the carbamylation of RubisCO in vivo (Portis et al., 1986) or to remove important inhibitors such as CA1P and other ligands that accumulate in the dark. Experiments to resolve these issues are currently in progress (L. -A. Li and F. R. Tabita, unpublished results). In free-living cultures of both Anabaena sp. strain CA and Synechococcus sp. strain RF-1, an organism that shows a Circadian rhythm relative to nitrogenase activity and nif transcription (Huang et al., 1990), light-dark transitions do not have an effect on RubisCO activity in vitro (T.-J. Chow and F.R. Tabita, unpublished results). Since it is well established that fixation stops in the dark (Ihlenfeldt and Gibson, 1975; Joset-Espardellier et al., 1978), attempts to implicate RubisCO as a site for regulation must consider problems inherent to the cyanobacterial enzyme, particularly its low capacity to bind some of the classic inhibitors (see above) and potential changes that occur upon cell breakage. In this connection, prior studies with chemostat cultures of Synechococcus sp. strain PCC 6301 indicate that RubisCO activity increased substantially (Slater, 1975; Karagouni and Slater, 1979), suggesting that there are indeed growth conditions which influence RubisCO activity. In further support of the potential to regulate fixation at the level of RubisCO, Steinberg and Meeks (1989) showed that lightdependent fixation of Nostoc sp. strain UCD

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Metabolism

7801, immediately after separationfromits symbiotic partner Anthoceros punctatus, was eight-fold lower than the rate exhibited by a free-living culture of Nostoc sp. strain UCD 7801. This was found to be due to an eight-fold reduction of RubisCO activity, yet the levels of RubisCO protein were maintained at 4.3 and 5.2% of the total protein of Nostoc sp. strain UCD 7801 grown in symbiotic or free-living states, respectively, Furthermore, this apparent posttranslational modification of RubisCO did not appear to affect the ability ofthe inactivated enzyme to bind its substrate, RuBP. Although, it is difficult to predict the precise mechanism involved in the inactivation ofRubisCO in this system, it would be interesting to determine the role of RubisCO activase (rca has been detected in Nostoc sp. strain MAC; Li et al., 1993) as well as isolate and examine preparations of purified inactivated RubisCO. Other potential loci for light-dependent regulation of fixation in vivo are the thioredoxin-activated enzymes of the Calvin cycle and other enzymes of intermediary metabolism (see Chapter 24 and Crawford et al., 1984; Marsden and Codd, 1984; Serra et al., 1989). Obviously, a mutation in PRK, such that Ser-222 is converted to a phenylalanine (Su and Bogorad, 1991), has a profound effect on in vivo fixation and growth. IV. Organization of Reductive Pentose Phosphate Cycle Genes In all the cyanobacteria thus far examined, there appears to be a single type I RubisCO enzyme, comprised of eight L and eight S subunits in the familiar arrangement (Tabita, 1987). Without exception, the genes encoding the L and S subunits, rbcL and rbcS, respectively, are found to be closely associated and are part of a single transcriptional

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unit. In Synechococcus sp. strain PCC 6301 (Shinozaki and Sugiura, 1983), the genes are separated by a 93 bp spacer region, while inAnabaena sp. strain PCC 7120 (Nierzwicki-Bauer et al., 1985) and Anabaena sp. strain CA (L.-A. Li and F. R. Tabita, unpublished results) the spacer region is considerably larger, 545 and 542 bp, respectively. The significance ofthe spacer region and its different length in the two types ofcyanobacteria is not known at this time. However, it is clear that the rbcLrbcS genes from unicellular and heterocystous strains are flanked by completely different open reading frames. In Synechococcus sp. strain PCC 7942, there is an open reading frame (ORF) 5' to rbcL that is required for growth at low levels of (Friedberg et al., 1989). Likewise, 3' to rbcS, several ORF’s are present that appear to function in the transport of (Schwarz et al., 1992; also see Chapter 15). In Anabaena sp. strain PCC 7120 (Golden et al., 1988) and A. variabilis strain ATCC 29413 (Herrero and Wolk, 1986) the rbcLrbcS genes are found somewhat downstream from the nif structural genes and more recent studies with Anabaena sp. strain CA indicate that sequences in the immediate vicinity of the rbcLrbcS genes differ substantially from Synechococcus sp. strain PCC 7942 (Li et al., 1993). Specifically, an ORF encoding the enzyme RubisCO activase (rca) was found about 2 kb downstream from rbcS. Two unknown ORF’s (ORF’s 1 and 2) were found to be juxtaposed between rbcS and the rca gene and the whole region was sequenced and mapped (Fig. 9). The deduced amino acid sequence of the Anabaena sp. strain CA RubisCO activase showed both similarities and differences to the plant and algal activases (Li et al., 1993). The two ATP binding domains identified in plant RubisCO activase enzymes (Werneke et al., 1988) are highly conserved in the Anabaena sp. strain CA enzyme (amino acid residues 37 to 44 and 93 to 102) and the lysine

458 residue (residue 43) essential for ATP binding in the G--G-GKS/T glycine-rich consensus domain (Shen et al., 1991) is also conserved (Fig. 10). By contrast, the amino acid sequence similarity sharply differs at the carboxy-terminus. The Anabaena sp. strain CA RubisCO activase possesses additional residues at the carboxy-terminus and lacks much of the aminoterminal region present in plant and algal RubisCO activase. Thus, the additional carboxy-terminal sequence compensates for the truncated aminoterminus, yielding a molecular weight similar to those of the mature plant and algal enzymes. Besides specificity for different RubisCO enzymes, differences in the primary structure of RubisCO activase might reflect divergent regulation of fixation, since cyanobacterial RubisCO does not appear to form a noncarbamylated enzyme-RuBP (E-R) complex (Andrews and Abel, 1981; Badger, 1980; Lee et al., 1991b). Current studies are thus directed at elucidating the physiological role of cyanobacterial RubisCO activase by preparing rca null strains. In addition, nucleotide sequence analyses revealed that rbcLrbcS, rca, as well as the two intergenic unknown open reading frames of Anabaena sp. strain CA were all in the same transcriptional orientation. It is not clear whether all these genes are members of the same operon; however, preliminary studies indicate that the rbcLrbcS genes and rca may not be cotranscribed (L.-A. Li and F. R. Tabita, unpublished results). Because cyanobacteria, algae, and plants exhibit similar mechanisms to generate energy through photosynthesis and Anabaena sp. strain CA was found to contain homologous RubisCO activase, it was expected that the rca gene might be present in all cyanobacteria. Surprisingly, Southern blot hybridization analysis did not support this hypothesis. Indeed, the hybridization results indicated that genomic DNAs from a variety of unicellular and nonheterocystous filamentous cyanobacteria did not readily recognize the Anabaena sp. strain CA rca probe under the conditions of high and low stringency that were employed (Li et al., 1993). Certainly these results do not exclude the possibility of some form of RubisCO activase that is only poorly homologous to plant and heterocytous cyanobacterial rca, however, as indicated above, experiments with Synechococcus sp. strain 7942 have not uncovered any rca-like genetic information downstream from rbcS. Cyanobacteria have long been suggested to be the endosymbiotic progenitors of chloroplasts (Margulis,

F. Robert Tabita 1981; see Chapter 5). Since fixation is an essential function for cyanobacteria and chloroplasts, RubisCO activase may serve as an alternative to understand the phylogenetic relationship between cyanobacteria and chloroplasts; although the rca gene is nuclearencoded in plants, there is evidence for gene transfer from chloroplast to nucleus (Baldauf and Palmer, 1990). According to the hybridization analyses, heterocystous cyanobacteria seem to be phylogenetically closer to chloroplasts than are unicellular and nonheterocystous filamentous cyanobacteria. Although we have examined only a limited number of cyanobacterial strains at this time, our results are consistent with hypotheses suggesting that chloroplasts might be derived from an ancient Nostoc species (Bryant and Stirewalt, 1990). The results that have been collected at this time, with a limited selection of cyanobacterial strains, certainly suggests that additional studies with a more complete array of species is warranted. There are many questions that remain unanswered. Paramount among these is elucidation of the actual role RubisCO activase plays in Anabaena sp., particularly since unactivated cyanobacterial RubisCO does not bind RuBP tightly. It may be that RubisCO activase in heterocystous cyanobacteria catalyzes an enzyme-mediated removal of some phosphate inhibitor (Robinson and Portis, 1988, 1989). Biochemical studies, currently in progress with purified recombinant Anabaena sp. strain CA RubisCO activase, should help to define its role in vitro and facilitate an approach to learn its role in vivo. Finally, it will be necessary to determine the role of the two unknown open reading frames upstream from rca and downstream from rbcS and whether they are involved in the regulation of fixation. In various phototrophic and chemoautotrophic bacteria, the structural genes of the Calvin reductive pentose phosphate pathway are arranged in discrete operons, usually controlled by a single transcriptional activator gene (Tabita et al., 1992; 1993). These studies prompted us to examine our cyanobacterial rbcLrbcS clones for prk-containing sequences, using both spinach and bacterial heterologous probes. In Anabaena sp. strain CA it was found that chromosomal DNA strongly hybridized to a spinach prk probe but not to cosmid clones containing the rbcLrbcS genes (L.-A. Li and F. R. Tabita, unpublished results). The strongly hybridizing fragment was cloned and sequenced and found not to encode prk;

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Metabolism

the strong hybridization that was observed was due to the nucleotide sequence: ATGTTTCTGGTTTGTTGG, found in both spinach prk and the Anabaena sp.

459

clone. Moreover, after sequencing upstream and downstream from the Anabaena sp. strain CA rbcLrbcS genes, it became apparent that there was no prk gene associated with the rbcLrbcS and rca genes of this organism, suggesting that other Calvin

460 cycle genes may not be clustered in cyanobacteria. Certainly there was no indication that the prk clone of Su and Bogorad (1991) contained rbcLrbcS sequences. Thus, there is apparently no organized Calvin cycle structural gene operon in Synechococcus sp., and Synechococcus sp. fragments that contain the rbcLrbcS genes also do not contain prk. Curiously, Synechococcus sp. strain PCC 7942 DNA was found to complement Escherichia coli recA mutants to resistance to methyl-methane sulfonate. This complementing DNA, when sequenced, was found to encode a protein that showed great homology to spinach PRK (Borrias and de Vrieze, 1991). At this juncture, it is not clear how PRK confers methyl methane sulfonate resistance to E. coli (M. Borrias, personal communication). V. Regulation of Expression of Reductive Pentose Phosphate Cycle Genes

A. RubisCO Genes There is a remarkable scarcity of information relative to transcriptional regulation of the rbcLrbcS genes of cyanobacteria. There are indications that transcription is basically constitutive in the light or the dark, although this has not been thoroughly examined. In both Anabaena sp. strain PCC 7120 (Nierzwicki-Bauer et al., 1984) and Synechococcus sp. strain PCC 6301 (Shinozaki and Sugiura, 1985), the rbcL and rbcS genes were found to be part of a single transcriptional unit, as are the genes of the cyanobacterial-like endosymbiont (cyanelle) of Cyanophora paradoxa (Heinhorst and Shively, 1983). More recent studies indicate that the rbcLrbcS genes of Anabaena sp. strain CA are also co-transcribed, but may not be part of the same transcript as the closely associated rca gene (L.-A. Li and F. R. Tabita,

F. Robert Tabita unpublished results). There is an inverted repeat 31 base pairs 3' to the stop codon of rbcS in the two Anabaena sp. strains, 51 base pairs 3' to rbcS of Synechococcus sp. strain PCC 6301 (Fig. 11), suggestive of a hair-pin like structure which might function as a transcription terminator. This potential secondary structure is compatible with our preliminary indications that the rbcLrbcS and rca genes are found on separate transcripts. In plants, transcription of rbcS is light-regulated (Broglie et al., 1984). However, Nierzwicki-Bauer et al. (1984) reported that there was no difference in the levels of rbcS-specific mRNA of cultures of Anabaena azollae grown photoautotrophically in the light or photoheterotrophically with fructose as carbon source in the dark. However, subsequent experiments with both free-living Anabaena azolla and symbiotic Anabaena azollae-Azolla caroliniana indicated that there was a five to ten-fold decrease in the levels of rbcLrbcS transcripts in the symbiont compared to free-living A. azollae. Thus, transcription is presumably regulated by the metabolism of sugars by the symbiont (Nierzwicki-Bauer and Haselkorn, 1986). At variance with these results, Steinberg and Meeks (1989) found that the amount of RubisCO protein remained relatively constant in free-living Nostoc sp. strain UCD 7801 and symbiotic Nostoc sp. strain UCD 7801-Anthoceros punctatus cultures. Both in vitro RubisCO activity and light-dependent fixation, however, were about eight-fold lower in the symbiont compared to free-living Nostoc sp. strain UCD 7801. Clear indication of transcriptional control was obtained in nitrogen-starved Anabaena sp. strain PCC 7120 cells; rbcL-specific message was readily demonstrable in ammonia-grown cells but could not be detected in cells placed in a medium lacking fixed nitrogen containing DCMU, under an argon atmosphere (Haselkorn et al., 1983). These nitrogen-starvation conditions are known to result in

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Metabolism

decreased light-dependent fixation (Bradley and Carr, 1976) and loss of RubisCO and phosphoribulokinase activity (Bradley and Carr, 1977). Thus rbcLrbcS message stability or transcription appears to be under nitrogen control in Anabaena sp. cells starved for nitrogen under argon. When ammoniagrown cells were transferred to a medium free of fixed-N under aerobic conditions (in the presence of such that heterocyst differentiation is fully induced, the level of rbcLrbcS transcript remains at a constant level in the ammonia-grown and aerobic Nfree incubated cells (Lang and Haselkorn, 1989). The levels of rbcLrbcS transcript are obviously regulated by the nitrogen status of the cells, particularly if subsequent experiments show that anaerobiosis itself does not trigger the changes in rbcLrbcS transcript observed. In an effort to examine further the regulation of genes important for fixation, studies have been initiated with several strains, in particular Synechococcus sp. strains RF-1, Anabaena sp. strain CA, and other marine cyanobacteria. Synechococcus sp. strain RF-1 is a freshwater strain that shows an interesting diel response relative to nif transcription and nitrogenase activity under alternating light-dark conditions, reminiscent of a Circadian rhythm (Huang et al., 1988; 1990). This organism catalyzes active oxygen-evolving photosynthesis in the light but has the inherent ‘intelligence’ to catalyze oxygensensitive nif transcription, nitrogenase biosynthesis, and nitrogenase activity in the dark, i.e., under growth conditions where there is no oxygen produced. Thus, studies are in progress to determine whether there is a reciprocal dark-induced mechanism to turn off rbc transcription in Synechococcus sp. strain RF-1, particularly since rbc transcription does not appear to be under light-dark regulation in Anabaena sp. (Nierzwicki-Bauer et al., 1984) but may be in Synechocystis sp. strain PCC 6803 (Mohamed and Jansson, 1989).

B. prk and Other Genes Important for Metabolism As discussed earlier, the prk gene has only recently been cloned from Synechocystis sp. strain PCC 6803 (Su and Bogorad, 1991) and Synechococcus sp. strain PCC 7942 (Borrias and de Vrieze, 1991). In Synechocystis sp. strain PCC 6803, the mutation that renders cells sensitive to bright light (BRLS strain) maps to prk and there was no reported difference in

461

PRK mRNA levels in wild-type and BRLS cells exposed to dim light or bright light. It is not known if other mechanisms or environmental factors control prk expression. The gene encoding PEP carboxylase (ppc) has been isolated from four cyanobacterial strains: Synechococcus sp. strain PCC 6301 (Kodaki et al., 1985), Synechococcus sp. strain PCC 7942 (Luinenburg and Coleman, 1990), Anabaena sp. strain PCC 7120 (Luinenburg and Coleman, 1992) andAnabaena variabilis (Harrington et al., 1986). Attempts to prepare ppc null strains in Synechococcus sp. strain PCC 7942 have not been successful, presumably because of the inability to segregate the wild-type ppc gene from merodiploid strains containing copies of the inactivated and wild-type gene (Luinenburg and Coleman, 1990). Although the authors interpret these results to imply that active PEP carboxylase is obligatory for growth, it may be that strains containing only the inactivated ppc gene may be isolated after further efforts. In any case, additional experiments on the regulation and function of ppc and PEP carboxylase in cyanobacterial carbon metabolism are awaited with great interest. Comparisons of the deduced amino acid sequences of the Synechococcus sp. strain PCC 6301 and Anabaena sp. strain PCC 7120 enzymes show about 65% identity (Luinenburg and Coleman, 1992). There is also evidence that the Synechococcus sp. strain PCC 6301 ppc promoter may be functional in E. coli (Kodaki et al., 1985).

C. Environment

Gene Expression in the

Paul et al. (1990) used the polymerase chain reaction (PCR) to amplify rbcL from aquatic environments using oligonucleotide primers that bracketed conserved regions of the gene. These studies showed that this key gene could be detected in naturally occurring dissolved DNA of the water column, as well as in planktonic samples. This technology was correlated with the appearance of an Anabaenopsis sp. bloom and could be of potential value to predict the onset of cyanobacteria or algal intoxications. More recently Pichard and Paul (1991) have extended these studies to the detection of specific RubisCO mRNA in the marine environment. Using antisense and sense RNA probes, specific mRNA could easily be distinguished from contaminating target DNA. This approach thus provides a highly sensitive method to determine active transcription of an important

462 gene in the environment, and the sense and antisense protocols control for nonspecific hybridization. Indeed, the initial studies suggested light regulation of rbcL transcription in natural phytoplankton of the Dry Tortugas. Since tropical oligotrophic waters are usually dominated by Synechococcus sp.-like picoplankton, the laboratories of Paul and Tabita are collaborating to investigate rbc regulation of laboratory cultures and natural samples of oligotrophic marine cyanobacteria. VI. Conclusion In a prior review, it was predicted that advances in the biochemistry and regulation of fixation would be forthcoming (Tabita, 1987). The quality of the recent work and the totality of the various advances in this field have exceeded even the optimistic projections of one who actively participates in such studies. Although it is probable that additional breakthroughs will be made pertaining to cyanobacterial metabolism per se, it is also certain that these organisms will continue to serve as an excellent model system for investigations of fixation in more complex oxygen-evolving organisms. Acknowledgments Recent studies from our laboratory were supported by grants from the National Institutes of Health, the National Science Foundation, and the Department of Agriculture. The contributions of my students, postdoctoral associates, and my colleagues are gratefully acknowledged. I would particularly like to thank Janet Newman and Steve Gutteridge for the generous gift of computer graphics generated photographs of the Synechococcus sp. strain PCC 6301 RubisCO. References Andersen K and Caton J (1987) Sequence analysis of Alcaligenes eutrophus chromosonally encoded ribulose bisphosphate carboxylase large and small subunit genes and their gene products. J Bacteriol 169: 4547–4558 Andersson I, Knight S, Schneider G, Lindqvist Y, Lundqvist T, Brändén C-I and Lorimer GH (1989) Crystal structure of the active site of ribulose-bisphosphate carboxylase. Nature 337: 229–234

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F. Robert Tabita reconstitution ofribulose-1,5-bisphosphate carboxylase from Chromatium vinosum. Arch Biochem Biophys. 236:487–496. Jordan DB and Ogren WL (1981) Species variation in the specificity of ribulose bisphosphate carboxylase/oxygenase. Nature 291: 513–515 Jordan DB, Chollet R and Ogren WL (1983) Binding of phosphorylated effectors by active and inactive forms of ribulose-1,5-bisphosphate carboxylase. Biochemistry 22:3410– 3418. Joset-Espardellier F, Astier C, Evans EH and Carr NG (1978) Cyanobacteria grown under photoautotrophic, photoheterotrophic, and heterotrophic regimes: sugar metabolism and carbon dioxide fixation. FEMS Microbiol Lett 4: 261–264 Karagouni AD and Slater JH (1979) Enzymes ofthe Calvin cycle and intermediary metabolism in the cyanobacterium Anacystis nidulans grown in chemostat culture. J Gen Microbiol. 115:369– 376 Katagiri F, Kodaki T, Fujita N, Izui K and Katsuki H (1985) Nucleotide sequence ofthe phosphoenolpyruvate carboxylase gene of the cyanobacterium Anacystis nidulans. Gene 38: 265–269 Kettleborough CA, Parry MAJ, Burton S, Gutteridge S, Keys AJ and Phillips AL (1987) The role ofthe N-terminus ofthe large subunit of ribulose-bisphosphate carboxylase investigated by construction and expression of chimaeric genes. Eur J Biochem 170: 335–342 Knight S, Andersson I and Branden C-I (1989) Reexamination of the three-dimensional structure of the small subunit of rubisCO from higher plants. Science 244: 702–705 Knight S, Andersson I, and Branden C-I (1990) Crystallographic analysis of ribulose 1,5-bisphosphate carboxylase from spinach at 2.4 Å resolution. J Mol Biol 215: 113–160 Kodaki T, Katagiri F, Asano M, Izui K and Katsuki H (1985) Cloning of phosphoenolpyruvate carboxylase gene from a cyanobacterium, Anacystis nidulans, in Escherichia coli. J Biochem 97: 533–539 Kossman J, Klintworth R and Bowien B (1989) Sequence analysis of the chromosomal and plasmid genes encoding phosphoribulokinase from Alcaligenes eutrophus. Gene 85: 247–252 Krieger TJ and Miziorko HM (1986) Affinity labeling and purification of spinach leaf ribulose-5-phosphate kinase. Biochemistry 25: 3496–3501 Lanaras T and Codd GA (1982) Variations in ribuiose 1,5bisphosphate carboxylase protein levels, activities and subcellular distribution during photoautotrophic batch culture of Chlorogloeopsis fritschii. Planta 154: 284–288 Lang JD and Haselkorn R (1989) Isolation, sequence and transcription of the gene encoding the photosystem II chlorophyll-binding protein, CP-47, in the cyanobacterium Anabaena 7120. Plant Mol Biol 13: 441–456 Lee B and Tabita FR (1990) Purification ofrecombinant ribulose1,5-bisphosphate carboxylase/oxygenaselarge subunits suitable for reconstitution and assembly of active enzyme. Biochemistry 29: 9325–9357 Lee B, Berka RM and Tabita FR (1991 a) Mutations in the small subunit of cyanobacterial ribulose-bisphosphate carboxylase/ oxygenase that modulate interactions with large subunits. J Biol Chem 266: 7417–7422 Lee B, Read BA and Tabita FR (1991b) Catalytic properties of recombinant octameric, hexadecameric, and heterologous cyanobacterial/bacterial ribulose-1,5-bisphosphate carboxy-

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465 Mogel SN and McFadden BA (1989) Photomodification of a serine at the active site of ribulose-1,5-bisphosphate carboxylase/oxygenase by vanadate. Biochemistry 28: 5428– 5431 Mohamed A and Jansson C (1989) Influence of light on accumulation of photosynthesis-specific transcripts in the cyanobacterium Synechocystis 6803. Plant Mol Biol 13: 693– 700 Morell M, Kane HJ, Hudson GS and Andrews TJ (1992) Effects of mutations at residue 309 of the large subunit of ribulosebisphosphate carboxylase from Synechococcus PCC 6301. Arch Biochem Biophys 299:295–301 Newman J and Gutteridge S (1990) The purification and preliminary X-ray diffraction studies of recombinant Synechococcus ribulose-1,5-bisphosphate carboxylase/ oxygenase from Escherichia coli. J Biol Chem 265: 15154– 15159 Newman J and Gutteridge S (1993) The X-ray structure of Synechococcus ribulose-bisphosphate carboxylase/oxygenaseactivated complex at 2.2 Å resolution. J Biol Chem 268: 25876–25886 Nierzwicki-Bauer SA and Haselkorn R (1986) Differences in mRNA levels in Anabaena living freely or in symbiotic association with Azolla. EMBO J 5: 29–35 Nierzwicki-Bauer SA, Curtis SE and Haselkorn R (1984) Cotranscription of genes encoding the small and large subunits of ribulose-1,5-bisphosphate carboxylase in the cyanobacterium Anabaena 7120. Proc Natl Acad Sci USA 81: 5961–5965 Norman EG and Colman B (1991) Purification and characterization of phosphoglycolate phosphatase from the cyanobacterium Coccochloris peniocystis. Plant Physiol 95: 693–698 Norris L, Norris RE and Calvin M (1955) A survey of the rates and products of short-term photosynthesis in plants of nine phyla. J Exp Bot 6: 64–74 Owttrim GW and Colman B (1986) Purification and characterization of phosphoenolpyruvate carboxylase from a cyanobacterium. J Bacteriol 168: 207–212 Padan E (1979) Facultative anoxygenic photosynthesis in cyanobacteria. Annu Rev Plant Physiol 30: 27–40 Pate JS, Lindblad P and Atkins CA (1988) Pathways of assimilation and transfer of fixed nitrogen in coralloid roots of cycad-Nostoc symbioses. Planta 176: 461–471 Parry MAJ, Madgwick P, Parmer S, Cornelius MJ and Keys AJ (1992) Mutations in loop six of the large subunit of ribulose1,5-bisphosphate carboxylase affect substrate specificity. Planta 187: 109–112 Paul JH, Cazares L and Thurmond J (1990) Amplification of the rbcL gene from dissolved and particulate DNA from aquatic environments. Appl Environ Microbiol 56: 1963–1966 Paul K, Morell MK and Andrews TJ (1991) Mutations in the small subunit of ribulosebisphosphate carboxylase affect subunit binding and catalysis. Biochemistry 30:10019–10026 Pelroy RA and Bassham JA (1972) Photosynthetic and dark carbon metabolism in unicellular blue-green algae. Arch Mikrobiol 86: 25–38 Pelroy PA and Bassham JA (1976) Kinetics of light-dark fixation and glucose assimilation by Aphanocapsa. J Bacteriol 128: 633–643 Pichard SL and Paul JH (1991) Detection of gene expression in genetically engineered microorganisms and natural phyto-

466 plankton populations in the marine environment by mRNA analysis. Appl Environ Microbiol 57: 1721–1727 Pierce J, Carlson TJ and Williams JGK (1989) A cyanobacterial mutant requiring the expression of ribulose bisphosphate carboxylase from a photosynthetic anaerobe. Proc Natl Acad Sci USA 86: 5753–5757 Portis AR, Jr (1990) Rubisco activase. Biochim Biophys Acta 1015: 15–28 Portis AR, Jr (1992) Regulation of ribulose 1,5-bisphosphate carboxylase/oxygenase activity. Ann Rev Plant Physiol Plant Mol Biol 43: 415–437 Portis AR, Jr, Salvucci ME and Ogren WL (1986) Activation of ribulosebisphosphate carboxylase/oxygenase at physiological and ribulosebisphosphate concentrations by rubisco activase. Plant Physiol 82: 967–971 Ramage RT and Bohnert HJ (1989) Identification of an assembly domain in the small subunit of ribulose-l,5-bisphosphate carboxylase. Proc Natl Acad Sci USA 86: 1198–1202 Read BA and Tabita RF (1992a) Amino acid substitutions in the small subunit of ribulose-1,5-bisphosphate carboxylase/ oxygenase that influence catalytic activity of the holoenzyme. Biochemistry 31: 519–525 Read BA and Tabita FR (1992b) A hybrid ribulosebisphosphate carboxylase/oxygenase enzyme exhibiting a substantial increase in substrate specificity factor. Biochemistry 31:5554–5560 Read BA and Tabita FR (1992c) Catalytic properties of a hybrid ribulose bisphosphate carboxylase/oxygenase enzyme containing cyanobacterial large subunits and diatom small subunits. FASEB J 6: A209 Read BA and Tabita FR (1994) High substrate specificity factor ribulose bisphosphate carboxylase/oxygenase from eukaryotic marine algae and properties of recombinant cyanobacterial rubisco containing ‘algal’ residue modifications. Arch Biochem Biophys, in press Reichelt BY and Delaney SF (1983) The nucleotide sequence for the large subunit of ribulose 1,5-bisphosphate carboxylase from a unicellular cyanobacterium, Synechococcus PCC 6301. DNA 2: 121–129 Renström E and Bergman B (1989) Glycolate metabolism in cyanobacteria. I. Glycolate excretion and phosphoglycolate phosphatase activity. Physiol Plant 75: 137–143 Robinson SP and Portis AR Jr (1988) Involvement of stromal ATP in the light activation of ribulose-1,5-bisphosphate carboxylase/oxygenase in intact isolated chloroplasts. Plant Physiol 86: 293–298 Robinson SP and Portis AR Jr (1989) Ribulose-1,5-bisphosphate carboxylase/oxygenase activase protein prevents the in vitro decline in activity of ribulose-1,5-bisphosphate carboxylase/ oxygenase. Plant Physiol 90: 968–971 Roesler KR and Ogren WL (1990) Primary structure of Chlamydomonas reinhardtii ribulose 1,5-bisphosphate carboxylase/oxygenase activase and evidence for a single polypeptide. Plant Physiol 94: 1837–1841 Roy H and Cannon S (1988) Ribulose bisphosphate carboxylase assembly: what is the role of the large subunit binding protein? Trends Biochem Sci 13: 163–165 Roy H, Bloom M, Milos P and Monroe M (1982) Studies on the assembly of large subunits of ribulose bisphosphate carboxylase in isolated pea chloroplasts. J Cell Biol 94: 20–27 Salvucci ME, Portis AR Jr and Ogren WE (1985) A soluble chloroplast protein catalyzes ribulosebisphosphate carboxylase/

F. Robert Tabita oxygenase activation in vivo. Photosynth Res 7: 193–200 Salvucci ME, Portis AR, Jr and Ogren WE (1986) Light and response of ribulose-1,5-bisphosphate carboxyiase/oxygenase activation in Arabidopsis leaves. Plant Physiol 80: 655–659 Schneider G, Knight S, Andersson I, Branden C-Y, Lindqvist Y, and Lundqvist T (1990a) Comparison of the crystal structures of and rubisco suggests a functional role for the small subunit. EMBO J 9: 2045–2050 Schneider G, Lindqvist Y and Lundqvist T (1990b) Crystallographic refinement and structure of ribulose-15,-bisphosphate carboxylase from Rhodospirillum rubrum at 1.7 Å resolution. J Mol Biol 211: 989–1008 Schwarz R, Lieman-Hurwitz J, Hassidim M and Kaplan A (1992) Phenotypic complementation of high mutants of the cyanobacterium Synechococcus sp. strain PCC 7942 by inosine 5'-monophosphate. Plant Physiol 100: 1987– 1993 Serra JL, Llama MJ, Rowell P and Stewart WDP (1989) Purification and characterization of phosphoribulokinase from the cyanobacterium Anabaena cylindrica. Plant Science 59: 1–9 Shen JB, Orozco EM Jr and Ogren WL (1991) Expression of the two isoforms of spinach ribulose 1,5-bisphosphate carboxylase activase and essentiality of the conserved lysine in the consensus nucleotide-binding domain. J Biol Chem 266: 8963–8968 Shinozaki K and Sugiura M (1983) The gene for the small subunit of ribulose-1,5-bisphosphate carboxyiase/oxygenase is located close to the gene for the large subunit in the cyanobacterium Anacystis nidulans 6301. Nucl Acids Res 11: 6957–6964 Shinozaki K and Sugiura M (1985) Genes from the large and small subunits of ribulose-1,5-bisphosphate carboxylase/ oxygenase constitute a single operon on a cyanobacterium Anacystis nidulans 6301. Mol Gen Genet 200: 27–32 Shinozaki K, Yamada C, Takahata N and Sugiura M (1983) Molecular cloning and sequence analysis of the cyanobacterial gene for the large subunit of ribulose-1,5-bisphosphate carboxyiase/oxygenase. Proc Natl Acad Sci USA 80: 4050– 4054 Slater JH (1975) The control of carbon dioxide assimilation and ribulose 1,5-diphosphate carboxylase activity in Anacystis nidulans grown in a light-limited chemostat. Arch Microbiol 103: 45–49 Smith AJ (1982) Modes of cyanobacterial carbon metabolism. In: Carr NG and Whitton BA (eds) The Biology of Cyanobacteria, pp 47–85. Blackwell, Oxford Smrcka AV, Bohnert HJ and Jensen RG (1991a) Modulation of the tight binding ofcarboxyarabinitol 1,5-bisphosphate to the large subunit of ribulose 1,5-bisphosphate carboxylase/ oxygenase. Arch Biochem Biophys 286: 14–19 Smrcka AV, Ramage RT, Bohnert HJ and Jensen RG (1991b) Purification and characterization of large and small subunits of ribulose 1,5-bisphosphate carboxylase expressed separately in Escherichia coli. Arch Biochem Biophys 286: 6–13 Steinberg NA and Meeks JC (1989) Photosynthetic fixation and ribulose bisphosphate carboxyiase/oxygenase activity of Nostoc sp. strain UCD 7801 in symbiotic association with Anthoceros punctatus. J Bacteriol 171: 6227–6233 Su X and Bogorad L (1991) A residue substitution in phosphoribulokinase of Synechocystis PCC 6803 renders the mutant light-sensitive. J Biol Chem 266: 23698–23705

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Tabita FR (1980) Pyridine nucleotide control and subunit structure of phosphoribulokinase from photosynthetic bacteria. J Bacteriol 143: 1275–1280 Tabita FR (1981) Molecular regulation of carbon dioxide assimilation in autotrophic microorganisms. In: Dalton H (ed) Microbial Growth on Compounds, pp 70–81. Heydon and Sons, London Tabita FR (1987) Carbon dioxide fixation and its regulation in cyanobacteria. In: Fay, P and Van Baalen C (eds) The Cyanobacteria, pp 96–117. Elsevier, Amsterdam Tabita FR (1988) Molecular and cellular regulation of autotrophic carbon dioxide fixation in microorganisms. Microbiol Rev 52: 155–189 Tabita FR and Colletti C (1979) Carbon dioxide assimilation in cyanobacteria: regulation of ribulose 1,5-bisphosphate carboxylase. J Bacteriol 140: 452–458 Tabita FR and McFadden BA (1972) Regulation of ribulose-1,5diphosphate carboxylase by 6-phospho-D-gluconate. Biochem Biophys Res Commun 48: 1153–1159 Tabita FR and McFadden BA (1974a) D-ribulose 1,5-diphosphate carboxylase from Rhodospirillum rubrum. I. Levels, purification, and effect of metallic ions. J Biol Chem 249: 3453–3458 Tabita FR and McFadden BA (1974b) D-Ribulose 1,5diphosphate carboxylase from Rhodospirillum rubrum. II. Quaternary, structure, composition, catalytic and immunological properties. J Biol Chem 249: 3459–3464 Tabita FR and Small CL (1985) Expression and assembly of active cyanobacterial ribulose-1,5-bisphosphate carboxylase/ oxygenase in Escherichia coli containing stoichiometric amounts of large and small subunits. Proc Natl Acad Sci USA 82: 6100–6103 Tabita FR, Gibson JL, Bowien B, Dijkhuizen L and Meijer W (1992) Uniform designation for genes of the Calvin-BensonBassham reductive pentose phosphate pathway of bacteria. FEMS Microbiol Lett 99: 107–110

467 Tabita FR, Gibson JL, Falcone DL, Wang X, Li LA, Read BA, Terlesky KC and Paoli GC (1993) Current studies on the molecular biology and biochemistry of fixation in phototrophic bacteria. In: Murrell C and Kelly PD (eds) Microbial Growth on Compounds, pp 469–479. Intercept, Andover Voordouw G, DeVries PA, Van den Berg WAM and DeClerk EPJ (1987) Site-directed mutagenesis of the small subunit of ribulose-l,5-bisphosphate carboxylase/oxygenase from Anacystis nidulans. Eur J Biochem 163: 591–598 Wagner SJ, Thomas SP, Kaufman RI, Nixon BT and Stevens SE Jr (1993) The glnA gene of the cyanobacterium Agmenellum quadruplicatum PR-6 is nonessential for ammonium assimilation. J Bacteriol 175: 604–612 Wassmann CC, Ramage RT and Bohnert HJ (1989) Identification of an assembly domain in the small subunit of ribulose-1,5bisphosphate carboxylase. Proc Natl Acad Sci USA 86: 1198– 1202 Werneke JM and Ogren WL (1989) Structure of an Arabidopsis thaliana cDNA encoding rubisco activase. Nucl Acids Res 17: 2871 Werneke JM, Zielinski RE and Ogren WL (1988) Structure and expression of spinach leaf cDNA encoding ribulosebisphosphate carboxylase/oxygenase activase. Proc Natl Acad Sci USA 85: 787–791 Zeilstra-Ryalls J, Fayet O and Georgopoulos C (1991) The universally conserved GroE (Hsp60) chaperonins. Ann Rev Microbiol 45: 301–325 Zhu G and Jensen RG (199la) Xylulose 1,5-bisphosphate synthesized by ribulose 1,5-bisphosphate carboxylase/ oxygenase during catalysis binds to decarbamylated enzyme. Plant Physiol 97: 1348–13531 Zhu G and Jensen RG (1991b) Fallover of ribulose 1,5bisphosphate carboxylase/oxygenase activity. Plant Physiol 97:1354–1358

Chapter 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 Department of Botany, The Hebrew University of Jerusalem, 91904 Jerusalem, Israel Summary I. Introduction II. Adaptation to Changing Ambient Concentration and Gene Expression III. Mechanism of Inorganic Carbon Uptake A. A Primary Electrogenic Pump is Involved in Ci Uptake. B. The Ci Fluxes Associated with the CCM C. The Role of Photosynthetic Electron Transport D. Mutants Impaired in Ci Uptake IV. Role of Carboxysomes Concentration-Dependent Mutants and the Relevant Genomic Lesions V. Types of VI. Concluding remarks Acknowledgments References

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Summary The ability of cyanobacteria to adapt to a wide range of ambient concentrations involves modulation ofthe activity of an inorganic carbon-concentrating mechanism (CCM), as well as other changes at various cellular levels including the biosynthetic pathway of purines. Studies of mutants have identified several ofthe genes involved in the operation ofthe CCM and in the ability to grow under changing ambient concentration. In the case ofSynechococcus sp. strain PCC 7942 most ofthese genes have been mapped in the genomic region of the rbcLS operon. Higher levels of detectable transcripts originating from some of these genes have been observed after exposure of the cells to low concentration. Studies of mutants have confirmed quantitative models postulating crucial roles for carboxysomes and carboxysome-located carbonic anhydrase (CA) in cyanobacterial photosynthesis. A central role is also proposed for cytoplasmic-membraneassociated CA activity: CA may function to scavenge escaping by intracellular conversion to bicarbonate against the chemical potential.

I. Introduction Cyanobacteria show great flexibility with regard to the ambient concentrations under which they are able to grow. Their range extends from several percent in air to as low as one-tenth of the concentration of dissolved present in media at equilibrium with air. When cells aretransferredfrom D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 469–485. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

high to low concentration, they undergo a lightdependent adaptation process at various cellular levels (see Section II below). A prominent feature of this process, which has been intensively studied, is the induction of a Ci-concentrating mechanism (CCM) which is capable of building up an internal Ci concentration some 1000-fold than the external concentration. The operation ofthis mechanism is of

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particular significance in the lower range ofexternal concentrations. It enables the cells to overcome the more than 20-fold difference between the concentration of in their bulk medium and the of their ribulose 1,5-bisphosphate carboxylase/ oxygenase (RubisCO; see chapter 14). As a consequence the apparent photosynthetic affinity for external Ci is 10- to 20-fold higher in cells which have undergone the adaptation process (Kaplan et al., 1980). Furthermore, the elevated concentration of at the site of RubisCO (which is mainly located in polyhedral subcellular bodies, carboxysomes) reduces competition by and inhibits photorespiration. Though it has not been sufficiently investigated, it is very probable that the improved availability of consequent to the activity of the CCM results in reduced photoinhibitory damage to cells exposed to high light intensity. The scheme in Fig. 1 presents a tentative model of the CCM in cyanobacteria and its major components. It is discussed very briefly here but in greater detail in Sections III andIV,below. Inorganic carbon enters the cells from the surrounding unstirred layer and accumulates in the cytoplasm via energy-dependent system(s). Regardless of whether or is supplied, bicarbonate is the predominant species in the cytoplasm and the Ci species are not at chemical equilibrium there. Bicarbonate passes through the thylakoid region and enters the carboxysomes where localized carbonic anhydrase (CA) catalyzes the formation of Part of the generated is fixed at the closely adjacent carboxylation sites of RubisCO. Part may leak from the carboxysomes to the cytoplasm. Here a plasmalemma-localized CAlike activity converts it back to bicarbonate in an energy-dependent process and thus minimizes the wasteful leak of from the cells. The very large influx of bicarbonate associated with the CCM and subsequent fixation of would lead to the formation of ions within the cells. However, measurements of intracellular pH fail to reveal large changes in the internal pH consequent on the activity of the CCM (see Ogawa and Kaplan 1987; Kaplan et al., 1989). The mechanism(s) involved in maintenance and regulation of internal Abbreviations: CA – carbonic anhydrase; Ci – inorganic carbon; CCM – inorganic carbon- concentrating mechanism; Cm – chloramphenicol; DTT – dithiothreitol; bp – basepair; kbp – kilobasepair; –kanamycin resistance; ORF – open reading frame; RubisCO – ribulose 1,5-bisphosphate carboxylase/ oxygenase

pH in cyanobacteria are not fully understood (and are beyond the scope of the present chapter), but it is highly likely that the activity of the CCM is accompanied by large fluxes of protons across the cell envelope. The activity ofthe CCM may also play a significant role in determining the observed carbon isotopic fractionation i.e. the difference between the ratio in the atmosphere and that found in organic matter. This fractionation has been thought to result mainly from the activity of RubisCO which discriminates against the heavier isotope. The CCM may well contribute to the fractionation, due to the effect of its associated Ci fluxes on the isotopic composition of the internal Ci pool which is relatively closed. It has been noted that the higher the activity

Chapter 15 Ambient Inorganic Carbon Concentration Effects of the CCM, the smaller was the discrimination against the heavier isotope in Chlamydomonas (Berry, 1989). Natural variations in have been reported in marine organisms in different latitudes. It is likely (though yet to be experimentally established) that these differences reflect natural variations in the fluxes of Ci associated with the CCM activity. This chapter will focus on certain physiological and molecular aspects of the process of adaptation to changing ambient concentration, and of the operation of the CCM, both of which are currently under intensive investigation. For comprehensive reviews and literature citations, the reader is referred to Aizawa and Miyachi, 1986; Badger, 1987; Pierce and Omata, 1988; Kaplan et al., 1990; Miller et al., 1990; Coleman, 1991; Kaplan et al., 1991; Raven, 1991 and Badger and Price, 1992.

II. Adaptation to Changing Ambient Concentration and Gene Expression A syndrome of changes is observed at various cellular levels when cyanobacteria are transferred from high (1 –5% v/v in air) to low (air-level or lower) concentration. Noticeable changes include: modification of purine biosynthesis (Schwarz et al., 1992a); a rapid increase in the ability to accumulate Ci within the cells (half-time of about 1 h), and hence a substantially higher apparent photosynthetic affinity for extracellular Ci; a change in the amount of RubisCO (Mayo et al., 1989); an increase in number of carboxysomes (about 4-fold) and a change in their location from the cell-center towards its periphery (Turpin et al., 1984; Mckay et al., 1992); an alteration in thickness of some of the layers which comprise the cell envelope (Kaplan et al., 1990); accumulation of certain polypeptides in a cytoplasmic membrane enriched fraction, in particular a 42-kDa polypeptide (Omata and Ogawa, 1986); and a change in the pattern of phosphorylation of polypeptides (Bloye et al., 1992). The signal transduction pathway(s) involved in these responses to a single environmental factor (concentration of has not yet been elucidated. Evidence does not exist, therefore, demonstrating whether the changes associated with the adaptation process all result from a single perception process. A major experimental difficulty in the identification and analysis of the mechanisms involved is that most of these responses are

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quantitative rather than qualitative and thus demand very sensitive methods of assessment. Very little information is presently available on the nature of the signal which triggers adaptation. Badger (1987) has demonstrated that the ability to accumulate Ci internally depends on the ambient concentration of Ci rather than that of On the other hand, analyses of the time course of adaptation under various combinations of and levels have led to the suggestion that the cells respond to the relative activity of ribulose bisphosphate oxygenase (see Kaplan et al., 1990). Support for this suggestion may be seen in the conclusion that in Chlamydomonas reinhardtii the ratio of photosynthesis/ photorespiratory activities determines the extent of adaptation (Spalding and Ogren, 1982). A C. reinhardtii mutant impaired in phosphoglycolate phosphatase activity was unable to adapt to low level (see Marek and Spalding, 1991). It is not known whether the process of adaptation to ambient of cyanobacteria is related to the cell cycle and only occurs at a certain phase as is the case for C. reinhardtii (see Kaplan et al., 1990). A rapid decline in the ability to accumulate Ci was observed following the addition of bicarbonate to grown Synechocystis sp. strain PCC 6803. A decline following a similar time course was also observed when these cells were shifted from photoautotrophic to photoheterotrophic growth by the supply of glucose (Bloye et al., 1992). A significant increase in the extent of phosphorylation of several polypeptides was also noted when Synechocystis sp. strain PCC 6803 cells were supplied with either glucose or bicarbonate (Bloye et al., 1992). The nature of the phosphorylated polypeptides and their possible role in signal transduction is yet to be established. Further, it will be important to test whether the inhibition of the activity of the CCM by photoheterotrophic and high-Ci conditions occur via similar routes. A mutant impaired only in the ability to adapt to low would be expected to exhibit an apparent photosynthetic affinity to extracellular Ci similar to that of wild type but would not grow in the presence of low Several types of mutants have been isolated from cyanobacteria following chemical mutagenesis or insertional inactivation (see Section V and Fig. 6, below). Most of these mutants, however, exhibit an apparent photosynthetic affinity approximately two orders of magnitude lower than that of

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grown wild type. Thus, they do not qualify as ‘adaptation mutants’. Exceptions are mutants JR12, D4 and R14 of Synechococcus sp. strain PCC 7942 which were obtained following modifications ofthe genomic region downstream of the rbcLS operon (encoding the large and small subunits of RubisCO, see Fig. 6). Sequence analysis and metabolic complementation by inosine 5'-monophosphate (i.e., resumption of the ability to grow in the presence of low identified the open reading frame immediately downstream of rbcLS as the cyanobacterial purK gene (Fig. 6, Schwarz et al., 1992a). In eubacteria, purK encodes subunit II of phosphoribosyl aminoimidazole carboxylase which catalyzes the carboxylation step leading to the formation of inosine 5'-monophosphate in the purine biosynthetic pathway (Ebbole and Zalkin, 1987). This enzyme is composed of two subunits (encoded by the purEK operon). Subunit I (encoded by purE) carries the catalytic site and subunit II has been implicated in raising the affinity of the enzyme for Should this also be the case in cyanobacteria, the demand for exhibited by mutants impaired in purK would be explained (Schwarz et al., 1992). The cyanobacterial purE gene has not yet been identified, but the analysis of the sequences in the region of purK indicate that it must be located elsewhere in the genome and does not occur as a purEK operon as in other eubacteria. A transcript originating from the purK gene was not detectable in Synechococcus sp. strain PCC 7942 cells grown under high Under such conditions subunit I apparently suffices for purine biosynthesis. On the other hand, a 30 min exposure ofthese cells to low resulted in a detectable transcript originating from purK. Presumably, expression of purK confers the ability to synthesize purines in the presence of low (Schwarz et al., 1992a). It would be expected that, during the induction period under low the cellular purine level would transiently fall. It remains to be seen whether the (expected) transient lowering of the purine level is involved in the adaptation process. The observed responses to ambient stimulated the search for promoters. Several such promoters have been detected in Synechococcus sp. strain PCC 7942 with the aid oflacZ as a reporter gene (Scanlan et al., 1990). The nature of these promoters, the elements involved in their induction and the genes which they regulate, however, have yet

to be elucidated. An elevated level of transcripts following exposure to low has been noted in the cases of ndhB (encoding subunit II of NADH dehydrogenase, Marco et al., 1993), purK (Schwarz et al., 1992a), cmpA (encoding the cytoplasmicmembrane-located 42-kDa polypeptide) and rbcLS. Transformation of Synechococcus sp. strain PCC 7942 with a DNA construct obtained by fusing a 700-bp or 380-bp fragments containing the coding region upstream from cmpA with a promoter-less cat gene (encoding chloramphenicol acetyltransferase and capable of conferring resistance to chloramphenicol) resulted in strains capable of growing in the presence of chloramphenicol but only under low conditions. On the other hand, when the cmpA promoter region included only the first 150 bp upstream of the translational start codon, the transformant was able to grow under both high and low (see Fig. 2A). These data provide supporting evidence that the region between the nucleotides 150 to 380 bp upstream from translation start of cmpA contains regulatory elements involved in the response to ambient level. Interestingly, this is also the region that contains three of the four ‘boxes’ homologous to those identified in the genomic region upstream from the rbcLS operon (Fig. 2B; Kaplan et al., 1991). The role, if any, of these conserved sequences in conferring responsiveness to ambient level is yet to be established. Analysis of the polypeptide pattern in a cytoplasmic membrane-enriched fraction revealed that Synechococcus sp. strain PCC 7942 accumulates a 42-kDa polypeptide in large amount during adaptation to low (Omata and Ogawa, 1986). The higher level of mRNA originating from cmpA following exposure of cells to low concentration is consistent with this observation. Nevertheless, inactivation of cmpA by interposon mutagenesis did not affect the ability of the cells to adapt to low levels (Omata et al., 1990) suggesting that this polypeptide is most probably not directly involved in the adaptation process. Moreover, there are indications that this polypeptide is also not directly involved in Ci uptake. The mutant O221 of Synechococcus sp. strain PCC 7942 does not accumulate the 42-kDa polypeptide and is highbut is nevertheless able to accumulate Ci as efficiently as the wild type (Schwarz et al., 1988). The cmp genomic region contains four ORFs— cmpA, cmpB, cmpC and cmpD —and sequence

Chapter 15 Ambient Inorganic Carbon Concentration Effects

analyses have indicated that they are highly homologous to nrtA, nrtB, nrtC, and nrtD, respectively, involved in nitrate uptake (Omata, 1992). It should therefore be tested whether the ability to grow under low observed in the mutant where cmpA was inactivated, was due to functional compensation of the lesion by the gene product of nrtA. Should this be the case, the role of the 42-kDa CmpA polypeptide in Ci transport would have to be reexamined. It is interesting that the ability to accumulate the 42-kDa polypeptide is impaired in mutant O221 even though the mutation mapped within ccmN (fig. 6; Friedberg et al., 1989); Southern analysis indicated that cmpA must be located elsewhere. This may indicate the presence of transacting regulatory elements in the region containing the lesion in O221. Sequence analyses indicated the identity of cmpA and cbpA; the latter encodes a putative carotenoidbinding protein synthesized during exposure of Synechococcus sp. strain PCC 7942 to high light intensity (Reddy et al., 1989). These authors also demonstrated that, apart from being affected by light intensity and concentration, transcription of cbpA (cmpA) is also regulated by factors affecting DNA topology (Reddy et al., 1989).

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III. Mechanism of Inorganic Carbon Uptake Internal Ci concentrations as high as 50 mM or higher have been reported in cyanobacteria. When the and the membrane potential across the cytoplasmic membrane are taken into account, it is clear that Ci accumulation occurs against its electrochemical potential gradient (Kaplan et al., 1991; Badger and Price, 1992). Attempts to identify the physiological and molecular mechanisms involved are hampered by major difficulties such as: 1) Assessment of the accumulation ratio is complicated by lack of chemical equilibration between and within the cytoplasm and in the boundary layer surrounding the cells. Furthermore, the assessment of this ratio depends on accurate estimation of the pH at the respective sites. 2) The interconversion between the various Ci species and recycling (see Section D 2, below) complicates the determination of the kinetics of Ci transport during steady-state of photosynthesis.

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3) Isolated, sealed membrane vesicles are commonly used as a powerful tool to study unidirectional fluxes of ions and metabolites and to study the bioenergetics of transport. Attempts to isolate suitable sealed plasma membrane vesicles from cyanobacteria have not yet been successful. 4) Although mutants which require high for growth are obtained without difficulty, mutants impaired in Ci transport have rarely been reported. Nevertheless, progress has been made in clarifying the mechanisms involved in Ci uptake and concentration, and the following discussion briefly summarizes the evidence at hand.

A. A Primary Electrogenic Pump is Involved in Ci Uptake. Hyperpolarization of the membrane potential (immediate but transient) is triggered by the supply of bicarbonate to Ci-limited cells, suggesting the involvement of an electrogenic pump (see Kaplan et al., 1990). The pump might be a primary Ci pump directlyenergizedbyATP. However, aprimary pump has not been confirmed so far in any other organism. Furthermore, it might be difficult to distinguish between a primary pump (with subsequent utilization of in photosynthesis) and a primary pump. Particularly in the lower pH range (below pH 7.0), proton-bicarbonate symport driven by proton motive force could be envisaged. Here a stoichiometry of more than one proton per would have to be postulated (see Kaplan et al., 1990). Alternatively, the primary pump might establish a trans-membrane electrochemical potential gradient for some other ion which would than serve as the immediate source of energy for Ci uptake. For instance, dependence of uptake on the presence of (Section III B, below) suggests that Ci uptake might be driven by a secondary active symport (Reinhold et al., 1984). In this case the immediate source of energy for uptake would be a transmembrane Na+ gradient. Recent studies have provided direct indications for the presence in cyanobacteria of a light-dependent primary electrogenic sodium pump (Brown et al., 1990) that would establish such a gradient. However, conclusive evidence that cyanobacterial Ci uptake is driven by

symport secondary to a primary pump would depend on an unequivocal demonstration that the steady-state is large enough to drive the accumulation of bicarbonate. For example, when the accumulation ratio for Ci is as high as 1000 and the external sodium concentration is a few millimolar, the internal concentration should be in the low micromolar range. That sodium ions play a specific and significant role in cyanobacterial photosynthesis and growth is well established. (The reader is referred to Section III B below and Miller et al., 1990; Kaplan et al., 1990, for comprehensive reviews). However, this role may not necessarily be directly related to the primary pump involved in Ci transport. For instance, a antiporter might be involved in the maintenance and regulation of internal pH during bicarbonate uptake and subsequent fixation of (Kaplan et al., 1989).

B. The Ci Fluxes Associated with the CCM During steady-state photosynthesis, Ci influx should be equal to the sum of the rate of its intracellular utilization plus its rate of efflux. Kinetic parameters for the unidirectional Ci fluxes at steady-state are not available. Those parameters at hand derive from initial rates obtained in dis-equilibrium experiments. In such experiments the cells are allowed to utilize the ambient Ci until the oxygen compensation point is reached. They are then provided with various Ci concentrations in the form of or bicarbonate for a short period. Influx of Ci is followed with time by observing either the accumulation of within the cells, the removal of from the medium, or the initial rate of fluorescence quenching. The techniques used are filtering centrifugation, mass spectrometry or fluorimetry, respectively. The results obtained in these experiments have been interpreted as providing evidence that both and bicarbonate are taken up by the cells. They have therefore confirmed the earlier conclusion that bicarbonate ions are transported into the cell, based on the calculation that the rate of fixation is higher than the rate of formation from bicarbonate in the bulk medium (Miller et al., 1990). The saturation kinetics observed when internal Ci accumulation was measured as a function ofexternal concentration suggested that uptake is not merely the result of passive diffusion. Competition between andbicarbonate during uptake (Volokita et al., 1984) and between and (Espie et

Chapter 15 Ambient Inorganic Carbon Concentration Effects al., 1991) confirmed that a saturable mechanism is involved. Analysis of exchange by mass spectrometry demonstrated that Synechococcus sp. strain PCC 6301 cells are capable of efficiently removing fromthemediumalmosttocompletion (Espie et al., 1991). These authors concluded that the uptake system has a very high affinity for For some reason this apparent affinity is approximately 10-fold higher than that determined using the filtering centrifugationtechnique. Uptakeofbicarbonatealso displays saturation kinetics but direct measurements of the parameters are not possible by mass spectrometry and the available data were calculated from filtering centrifugation experiments in the range of 60–100 and Volokita et al., 1984). Bicarbonate uptake and uptake differ not only in their kinetic parameters but also in certain other characteristics. Bicarbonate uptake is associated with hyperpolarization of the plasma membrane—not observed for uptake. The presence ofsodium (in the millimolar range) is required for the uptake of but a very low concentration of sodium (in the micromolar range) is large enough to saturate the response of uptake to sodium. This differential effect has been used in an attempt to distinguish quantitatively between the uptake of the two Ci species (Miller and Canvin, 1985). Interestingly, growth of Synechococcus sp. strain PCC 6301 in a standing culture in the presence of a very low Ci concentration resulted in sodium-independent bicarbonate uptake (Espie and Kandasamy, 1992). It has been suggested that uptake of the two Ci species is mediated by two distinct mechanisms. The SC mutant ofSynechocystis sp. strain PCC 6803, that is impaired in uptake but not in that of bicarbonate, has provided support for this suggestion (Ogawa, 1993; see Fig. 3 and Section III D below). Regardless of which Ci species is supplied, bicarbonate is the species which accumulates at the inner face of the membrane, and the Ci species are not at chemical equilibrium in the cytoplasm. This has been deduced from experimental evidence indicating a high for the photosynthetic utilization of the dominant Ci species in the intracellular pool, obtained both with Anabaena variabilis (Volokita et al., 1984) and with Synechococcus sp. strain PCC 7942 (Schwarz et al., 1988). It was concluded that this dominant species is The conclusion that and are not at chemical

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equilibrium within the cytoplasm has been confirmed in experiments in which a human CA gene was expressed in Synechococcus sp. strain PCC 7942 (see Section IV below). These findings led to the suggestion that uptake of involves a vectorial CA-like moiety located in the plasma membrane which converts to in transit (Volokita et al., 1984). The system would have to depend on metabolic energy supply, since the release of bicarbonate in the cytoplasm would occur against its chemical potential gradient. The suggestion has recently been strengthened by the discovery of plasma membrane-associated CA activity in Synechocystis sp. strain PCC 6803 (Bedu et al., 1993). The presence of CA-like activity would also serve the important function of scavenging molecules in the cytoplasm and thus minimizing their leak to the medium (see Fig. 1). Moreover, this postulate obviates the earlier suggestion that uptake of in cyanobacteria is mediated by a membrane-located transport mechanism. Diffusion of across the cytoplasmic membrane, and subsequent energydependent conversion and release of bicarbonate, would account for the reported saturation kinetics and apparent uphill transport of as demonstrated by a quantitative model (L. Fridlyand, A. Kaplan and L. Reinhold, unpublished results). The complex interaction between the Ci species during transport (see Kaplan et al., 1990) might stem from competition between exogenous molecules and those produced from cytoplasmic (or carboxysomal)

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bicarbonate for the energy-dependent CA. The Ci species are not at chemical equilibrium in the boundary layer surrounding the cells. This has been deduced from the burst of observed when CA was supplied to Synechococcus sp. strain PCC 7942 cells that had been kept in a gas exchange system at steady-state in the light (Ogawa and Kaplan, 1987). The magnitude of this burst (resulting in a transient concentration higher than that in the gas stream) clearly indicated the presence of an external concentration higher than that at equilibrium with the level in the gas stream. Efflux of probably must have occurred during Ci uptake and it is likely that this efflux was mediated by the same carrier system which is involved in influx (slippage of the pump). An accurate estimate of Ci efflux at steady state photosynthesis is necessary in order to evaluate the efficiency of the CCM and its energetic cost but as pointed out above, such data are not available. Efflux of has been observed following darkening of a Synechococcus sp. strain PCC 7942 culture and has been used to estimate the steady-state Ci efflux in the light (Badger et al., 1985). It must be borne in mind, however, that darkening would cause acidification of the cytoplasm and thus stimulate the uncatalyzed formation of from bicarbonate. It would also alter the state of energization of the cells. Since active uptake depends on an energy-dependent difference in the kinetic parameters of the carrier at the inner and outer membrane surfaces, discontinuation of the energy supply on darkening will equalize the kinetic parameters and result in efflux. Moreover, the activity of the proposed energydependent scavenging system (see above) would depend on continuous supply of energy. A strong indication that efflux of Ci in the dark is faster than that in the light emerged from compartment analysis experiments performed on Anabaena variabilis (Marcus et al., 1982). A firm conclusion as to the relative roles of and bicarbonate in the steady-state uptake of Ci by any particular organism (there appears to be large species variation among cyanobacteria in this regard) would have to rest on detailed analysis of the uptake parameters at steady state for each Ci species. Moreover, interpretation of the results must take into account the mutual interaction between the Ci species and their relative abundance underthe specific growth and experimental conditions.

C. The Role of Photosynthetic Electron Transport Light energy is required for the energization of the Ci uptake system, and there is evidence that it is also involved in its activation (Kaplan et al., 1987). The electron transport chain, rather than fixation per se, is implicated as indicated by the observation that mutants display this effect on Ci uptake even at concentrations under which they fail to fix Studies by Ogawa and colleagues demonstrated that energization is most probably dependent on PS I activity (Ogawa et al., 1985). Activation of the Ci uptake, on the other hand, requires PS II activity but a very low level suffices. The activation by light is inhibited by DCMU but the active state can be restored by the addition of DTT. Since the activation by light is also inhibited by it is possible that some form ofreducing agent is involved (Kaplan et al., 1987). Recent studies (Miller et al., 1988; 1991; Espie et al., 1991) demonstrated a strong correlation between Ci uptake and fluorescence quenching when Synechococcus sp. strain PCC 6301 cells were provided with a range of Ci concentrations. The initial rate of fluorescence quenching and the initial declinein concentration in the medium showed similar dependence on the concentration of Ci supplied. These data provided supporting evidence for a link between Ci uptake and photosynthetic electron transport but the underlying mechanisms are not understood. Photosynthetic and respiratory electron transport in cyanobacteria share common constituents, although the interaction between these processes is not yet clear (see Peschek, (1987) and Chapter 13). In a scheme proposed by Nicholls et al (1992), the presence of two distinct electron carrier systems is postulated. NAD(P)H dehydrogenase is present on both the cytoplasmic and thylakoid membranes. Studies by Myers (1992) have indicated that exposure of Synechococcus sp. strain PCC 6301 to light of 680 nm (which is primarily absorbed by PS I) diverts the respiratory electron flow away from oxygen reduction to confirming mutual interaction between the electron carrier chains. Inactivation of ndhB, ndhK and ndhL, encoding various subunits of NADH dehydrogenase in Synechocystis sp. strain PCC 6803 (Ogawa, 1991), and ndhB in Synechococcus sp. strain PCC 7942 (Marco et al., 1993) resulted in mutants unable to accumulate Ci within the cell. On

Chapter 15 Ambient Inorganic Carbon Concentration Effects the other hand, mutants in which ndhC (Ogawa, 1992) and ndhF (Schluchter et al., 1993; Yu et al., 1992; Yu et al., 1993) were inactivated were able to grow in the presence of low The role ofNADH dehydrogenase in cyanobacterial photosynthetic electron transport has recently been investigated (see Mi et al., 1992; also see Chapters 10 and 13). The scheme presented by the latter author(s) involves donation of electrons by NADH dehydrogenase to the plastoquinone pool, providing a link between photosynthetic and respiratory electron transport. Theobservationthatphotosyntheticelectrontransport beyondthe site is severely inhibited in amutant in which ndhB is inactivated is in favor of this scheme (Marco et al., 1993). Importantly, conditions alleviated this inhibition in the mutant. This may have significance for the elucidation ofthe well known effect of bicarbonate on the activity of PS II (Eaton-Ray et al., 1986) and indicates possible involvement ofNADH dehydrogenase in this effect. The inability of the mutants impaired in various subunits of NADH dehydrogenase to accumulate inorganic carbon under air conditions may arise from disruption ofphotosynthetic electron transport.

D. Mutants Impaired in Ci Uptake Mutants defective in Ci uptake are highly desirable for resolving the mechanism of Ci transport and identifying the relevant genes. As indicated in the previous section, the mutants impaired in genes encoding various subunits ofNADH dehydrogenase have reduced ability to accumulate Ci but probably do not qualify as ‘transport mutants’ in a strict sense. Ogawa (1993) isolated a Synechocystis sp. strain PCC 6803 mutant, denoted SC, that is capable of growing like the wild type when exposed to 3% or 0.04% but unable to grow in the presence of 0.008% Fig. 3 shows that uptake of is severely depressed in the mutant, whereas bicarbonate uptake is apparently not affected. It is not known whether the gene impaired in mutant SC is directly involved in the uptake of Ci. For example, the observed differential effect of and bicarbonate uptake shown in the data in Fig. 3 could reflect a defect in the ability to convert to bicarbonate. On the other hand, five different Synechococcus sp. strain PCC 7942 mutants, that exhibit impaired ability to transport biocaronate but have normal uptake of have recently been

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isolated (M. Ronen-Tarazi and A. Kaplan, unpublished results). These mutants were obtained following transformation of the wild type with an inactivation plasmid library comprising small genomic fragments flanked with a cartridge encoding kanamycinresistance Single crossover recombination events resulted in inactivation ofvarious genes. Since the interposon mutated genes are tagged with the cartridge, the latter is being used to identify and map them. It is not yet know whether these genes are clustered, but they are not located in the genomic region of rbcLS. The availability of mutants impaired in either or bicarbonate uptake provides direct evidence for distinct uptake routes for the two Ci species.

IV. Role of Carboxysomes Carboxysomes are polyhedral bodies that are characteristic of cyanobacteria and that are also observed in certain bacteria (Codd, 1988). They contain most of the RubisCO in the cell but not some ofthe essential enzymes ofthe photosynthetic carbon reductive cycle. This has been deduced from fractionation experiments in which RubisCO activity appeared in the carboxysomal fraction but that of phosphoribulose kinase and phosphoglyceraldehyde dehydrogenase appeared in the supernatant. Further, immunogold labeling with the aid of antibodies has also clearly indicated that RubisCO is mainly located within the carboxysomes whereas phosphoribulose kinase is in the cytoplasm (McKay et al., 1992). These observations have provided strong support for the notion that the carboxysomes are the site of the carboxylation reaction in photosynthesis and not merely reserve bodies. The spacial separation between RubisCO and the other enzymes involved in carbon reduction indicates that fixation in cyanobacteria requires large fluxes of metabolites between the carboxysomes and the cytoplasm. The composition of carboxysomes in Synechococcus sp. strain PCC 7942 has recently been partly clarified following their isolation and analysis by SDS-PAGE (Price et al., 1992). Nevertheless, apart from the large and small subunits of RubisCO, the identities of the various polypeptides detected are not known. Immunogold labeling suggested the presence of RubisCO activase as well as chaperonin

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60 in Anabaena sp. strain PCC 7120 carboxysomes (Jager and Bergman, 1991). In Thiobacillus neopolitanus the carboxysomes are each surrounded by a protein layer which does not appear to contain any lipids (Holthijzen et al., 1986), and it is likely that this is also the case in cyanobacteria. Many of the mutants have the common feature that their carboxysomes are aberrant or completely absent (Pierce et al., 1988; Friedberg et al., 1989; Price and Badger, 1991, see Section V). Figure 4 presents an example of electron micrographs of Synechococcus sp. strain PCC 7942 and of one of its mutants showing aberrant carboxysomes in the latter. It is important to remember, however, that the biogenesis of carboxysomes has yet to be elucidated; the various forms of carboxysomes observed may be stages in their development so that the preponderance of any one form may represent retarded development. For example, the rod-shaped carboxysomes seen with high frequency in the Synechococcus sp. strain PCC 7942 mutants E1 and type I (Friedberg et al., 1989; Price and Badger, 1991) and the ‘empty’ inclusions within the carboxysomes (Fig. 4) are also observed, though rarely, in wild-type cells and may be stages in normal development (Orus et al., 1992). The apparent photosynthetic affinity for extracellular Ci in mutants which possess aberrant

carboxysomes is 2–3 orders of magnitude lower than that in the wild type. Their Ci uptake ability, however, does not differ significantly from that of the wild type (see Kaplan et al., 1991; Badger and Price 1992). Moreover, the phenotype of such mutants can not be accounted for by changes in the kinetic parameters of their RubisCO as shown in experiments with the in vitro-activated enzyme. These observations suggest the involvement of carboxysomes in the efficient photosynthetic utilization of the internal Ci pool, a conclusion supported by the fact that the number of carboxysomes increases during adaptation to low concentrations of (Turpin et al., 1984; McKay et al., 1992). A model for Ci fluxes and photosynthesis in cyanobacterial cells and carboxysomes has been proposed and modified which restricts the CAcatalyzed generation of to the latter bodies (Reinhold et al., 1989, Reinhold et al., 1991; see Fig. 1). The advantage of this model is that it obviates the need for assigning a high resistance to diffusion to the plasma membrane. Such a resistance has been regarded as necessary in order to limit back-diffusion of from the accumulated Ci pool and hence to lower the energy cost of the CCM. Attributing low permeability to the plasma membrane is inconsistent with the known properties of lipid bilayers and has been one of the major conceptual problems with

Chapter 15 Ambient Inorganic Carbon Concentration Effects regard to the CCM. The model assumes that the permeability of the plasma membrane in cyanobacteria is similar to that normally encountered in the boundary membranes of other organisms. In its original form the whole-cell model (Reinhold et al., 1989) transfers the diffusion barrier in the cell to the surface of the carboxysome. Themodified carboxysomal model doesawaywiththerequirementforadiffusionbarrier, since it places CA in the interior ofthe carboxysome and predicts that much ofthe formed at this site would be fixed as it diffuses outwards towards the carboxysome surface, past RubisCO sites located along the diffusion path (Reinhold et al., 1991). The model assumes that the predominant Ci species in the cytoplasm is always (see Section III C above). It furtherpostulates that conversion of to in the cytoplasm is uncatalyzed and that the Ci species do not reach chemical equilibrium. On arrival of the accumulated in the carboxysomes, is rapidly generated at CA sites in the immediate vicinity of RubisCO. The curve relating the rate of fixation to external Ci concentration predicted by the model agrees with experimental results remarkably well. The model predicts that, at 0.1 mM external Ci (the apparent for photosynthesis) a CA concentration that could accelerate the rate ofinterconversion of Ci species by a factor as low as 10-fold would be sufficientto allowphotosynthesistoproceedatalmost 90 per cent of maximum rate. This degree of catalysis, though very low, is nevertheless essential. When the interconversion factor is 1 (i.e., equivalent to the uncatalyzed rate) the predicted rate of photosynthesis drops to about 20 per cent ofthe maximum rate due to the limiting rate of formation. The model demonstrates that the ratio fixed to leaked is a function of the CA catalysis factor, and of the external Ci concentration, as well as of other factors such as carboxysome surface area. In other cyanobacterial species the ratio of fixed to leaked may vary owing to variations for these different parameters. The relative proportion of the formed in the carboxysomes that is fixed might affect the maintenance and regulation of the pH within these bodies, since the formation of from yields ions and the carboxylation reaction evolves protons (Fig. 1). The uptake mechanism may be depressed in vivo as internal Ci concentration rises above a certain value (i.e., a trans-inhibition mechanism), in which

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case the leakage predicted by the model would be an overestimation. Furthermore, the scavenging activity of the proposed energy-dependent CA (Section III B) would drastically reduce the leakage of This scavenging CA would also act as a trans-inhibition mechanism since the CA site would be accessible to both cytoplasmic and boundary layer-located molecules. Rising cytoplasmic concentration would compete with exogenous for the CA site. The predictions ofthe model that intracellular CA activity is confined to the carboxysomes (i.e., is absent from the cytoplasm) and that the Ci species are not at chemical equilibrium within the cytoplasm havebeen confirmed experimentally. CA activityhas been observed in cyanobacteria (Lanaras et al., 1985; Aizawa and Miyachi, 1986; Tu et al., 1987), though at a relatively low level (compared with green algae). On the basis of the CA and RubisCO activities of a carboxysome-enriched fraction, Price et al (1992) concluded that the intracellular activity of CA is associated with the carboxysomes. Further, expression ofa gene encoding human CA in Synechococcus sp. strain PCC 7942 resulted in a strain that demands high for growth due to a very large leakage of accumulated Ci (Price and Badger, 1989a). Mutants impaired in icfA (the gene encoding carboxysomal CA, located 20 kbp downstream ofthe rbcLS operon in Synechococcus sp. strain PCC 7942, Suzuki et al., 1991; Fukuzawa et al., 1992, see Fig. 6) were highin spite of the fact that the CA catalysis factor in mutant number 68 was large enough to drive the reaction at 250 times the uncatalyzed rate (i.e., 30-fold lower than in the wild type; Price et al., 1992). According to the model, this should be sufficient formaximal photosynthesis. Clearly, other factors such as correct organization and association of the CA and RubisCO within the carboxysomes are of critical importance (Reinhold et al., 1991). One of the CA-mutants, C3P-O, contains more carboxysomes than the wild-type strain but they appear to be smaller (Suzuki et al., 1991). Decisive experimental evidence as to the role ofcarboxysomal CA must await the isolation and characterization ofa mutant completely lacking carboxysomal CA activity. It is likely that RubisCO itself, as the major constituent of the carboxysomes might be involved in the correct organization of these bodies. Pierce et al. (1988) replaced the structural gene ofRubisCO in Synechocystis sp. strain PCC 6803 with that from the photosynthetic bacterium Rhodospirillum rubrum. The resulting organism, called ‘cyanorubrum’,

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expressed the gene encoding bacterial RubisCO, lacked visible carboxysomes, demanded high for growth and was very sensitive to oxygen. It also exhibited much higher photorespiratory activity as indicated by the Ci compensation point (Marcus et al., 1992). Modification of the small subunit in Synechococcus sp. strain PCC 7942 by the insertion of a gene conferring kanamycin resistance) near the 3' end of rbcS resulted in a Kmr, requiring mutant, EK6, in which the small subunit was 3 kDa larger (Lieman-Hurwitz et al., 1991) and in which the carboxysomes were aberrant (Orus et al., 1992). These observations strengthen the possibility that the small subunits have a role in the organization of the carboxysome. Adaptation of cyanobacteria to various ambient concentrations did not affect the performance of their RubisCO, that was assayed after activation in vitro. Furthermore, analyses ofthe kinetic parameters of the in vitro-activated enzyme did not reveal significant differences between those of the wild type and those of the mutants. On the other hand, the levels ofthe internal ribulose 1,5-bisphosphate pool were high in mutant cells when exposed to low but decreased to that observed in the wild type, on exposure to high (Fig. 5). These data suggest that RubisCO is in a low state of activation, in situ, in the high mutants EK.6, #68 and El, following pretreatment with low (Schwarz et al., 1992b). It is suggested that the phenotype in mutants

which possess defective carboxysomes stems from the inactivated state of RubisCO at low ambient This may well result from a low steady-state concentration at the RubisCO site in carboxysomes which are defective in their organization. V. Types of Concentration-Dependent Mutants and the Relevant Genomic Lesions As indicatedthroughout, mutants are currently serving as a major tool for elucidating the physiological and molecular basis of adaptation to changing levels and for elucidating the nature of the CCM (Marcus et al., 1986; Friedberg et al., 1989; Price and Badger, 1989b; Ogawa et al., 1987; Kaplan et al., 1991; Badger and Price, 1992; Ogawa, 1993). In Synechococcus sp. strain PCC 7942, all the mutations identified so far which resulted in highphenotypes have been mapped in the genomic region surrounding the rbcLS operon. This indicates that some genes involved in the ability to grow under changing ambient levels are clustered in this region. The scheme in Fig. 6 presents a partial restriction map of the rbcLS region of Synechococcus sp. strain PCC 7942, the location of the various ORFs and the sites ofmutations that affect the ability to grow at different ambient concentrations. The mutants that have been characterized can be divided into two main categories. Members of the first group show the expected

Chapter 15 Ambient Inorganic Carbon Concentration Effects

behavior of an ‘adaptation mutants’. They (D4 and R14) exhibit the same apparent photosynthetic affinity for extracellular Ci as observed in wild type but they are unable to grow under low They were isolated following modifications of purK (located immediately downstream of rbcS, Fig. 6), and the lesions affect their purine biosynthesis (see Section II). Therefore they cannot be regarded ‘adaptation mutants’ in the strict sense. The second group is heterogeneous and includes all the mutants in which the apparent photosynthetic affinity is approximately 100-fold lower than that of wild type. This group can be subdivided into three classes as described below. The first group contains mutants defective in their ability to accumulate Ci when either bicarbonate or is provided due either to lesions in genes encoding subunits ofNADH dehydrogenase (Section III C) or overexpress a foreign CA in the cytoplasm (Price and Badger, 1989a). In Synechococcus sp. strain PCC 7942, the extent of transcription of ndhB (located 12 kbp upstream of rbcL) appears to depend on the ambient level of (Marco et al., 1993). The second group of mutants are defective in the ability to accumulate Ci internally only when is the Ci species provided. The only example so far is mutant SC of Synechocystis sp. strain PCC 6803 (Ogawa, 1993; see Fig. 3). The third group of mutants is able to accumulate Ci internally only when but not bicarbonate, is provided (see Section III D above). The third fourth ofmutants are able to accumulate Ci internally at least as efficiently as the wild type but

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are defective in the ability to utilize the internal Ci pool photosynthetically. This group includes mutants with reduced carboxysomal-CA activity due to lesions in icfA (Fukuzawa et al., 1992; Price et al., 1992; see Section IV) and mutants that contain aberrant carboxysomes or lack these bodies. The latter were obtained following modifications of rbcLS (see Section IV; Pierce et al., 1988; Lieman-Hurwitz et al., 1991; Orus et al., 1992) as well as the region upstream of rbcL in the case of Synechococcus sp. strain PCC 7942 (Fig. 6). Inactivation, deletion or chemical mutagenesis of the five ORFs immediately upstream of rbcL resulted in phenotypes and the ORFs were therefore designated ccmK-ccmL-ccmM-ccmN-ccmO, respectively (Price et al., 1993). The ccmN and ccmO genes were earlier designated ORFI and ORFII, respectively (Friedberg et al., 1989). Insertion of a cartridge at the BclI site in the 3' region of the ccmO gene produced no detectable phenotype (Friedberg et al., 1989) but deletion ofa larger fragment from the 3' end ofccmO resulted in the mutant N1. The mutants obtained following modification of ccmK, ccmM and ccmN, and ccmO contained defective carboxysomes (see Section IV). It is not known whether normal carboxysomes are present in the mutants in which ccmL was inactivated. Insertion of a gene conferring kanamycin resistance at the EcoRI site downstream of ORF236 (Fig. 6) resulted in a mutant (M3) which also possess aberrant carboxysomes (Orus et al., 1992). The EcoRI site is located within ORF78 and also within the

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putative promoter region of ORF68 (not shown in Fig. 6) encoded by the reverse strand. In the case of mutant M3, it is not yet clear which inactivated ORF leads to the observed phenotype. In view of the observation that some of the mutants possess aberrant carboxysomes, it seems likely that the genes modified in the mutants encode carboxysome-related polypeptides. A detailed comparison of the polypeptide pattern in carboxysome-enriched fractions from the wild type and the mutants, however, is not yet available. Since the number of carboxysomes increases during adaptation to low it might be expected that transcription of some of these genes would depend on the ambient level of Studies by Ogawa (1993) located the mutation in the mutant G3 of Synechocystis sp. strain PCC 6803 within ORF535 whose product shows significant homology to CcmM of Synechococcus sp. strain PCC 7942 and some homology to RbcS. He also identified the product ORF242, encoded next to ORF535, as homologous to CcmN of Synechococcus sp. strain PCC 7942. It is not known whether normal carboxysomes are present in Synechocystis sp. strain PCC 6803 mutants impaired in ORF535 and ORF242 and whether these ORFs are located near the rbcLS operon in this organism. In addition to the mutants discussed above, Synechococcus sp. strain PCC 7942 mutants that only grow under low under certain circumstances have also been obtained. Insertional inactivation with a gene conferring resistance to kanamycin at the ClaI or SalI sites in ORF839 located downstream of ndhB, resulted in merodiploids (mutants Ml and Cl, respectively). The latter are able to grow under high or low in the absence of kanamycin but only under low in its presence. The product of ORF839 shows high sequence similarity to TopA, encoding topoisomerase I in E. coli, but the mechanism(s) leading to the expression of kanamycin resistance in the mutant exclusively under low requires further elucidation. Many of the genes involved in the ability to grow under changing levels are probably not yet recognized and might be located elsewhere. Moreover, Northern and sequence analyses of the region surrounding rbcLS of Synechococcus sp. strain PCC 7942 revealed several ORFs, whose inactivation did not result in selectable phenotypes under the experimental conditions used. ORF286 (formerly frxC, now chlL) and ORF466 (chlN) upstream of ccmK were identified as encoding two subunits of

protochlorophyllide reductase on the basis of homology with the corresponding genes in Synechocystis sp. strain PCC 6803 and liverwort (see Ogura et al., 1992, and Chapter 17), The product of ORF 145, that is highly homologous to rotA (encoding peptidylprolyl cis-trans isomerase) was identified downstream of ORF83 (Hassidim et al., 1992). Interposon mutagenesis of ORF 145 only resulted in merodiploids. Complete segregation of mutations in this gene appear to be lethal, and this suggests a crucial role for ORF 145 in protein folding. VI. Concluding remarks Clustering of the genes involved in certain physiological functions has been observed in a number of cases (e.g., see Chapter 21 and elsewhere in this book). Analyses of mutants has indicated that some ofthe relevant genes are clustered in the genomic region of the rbcLS operon, at least in the case of Synechococcus sp. strain PCC 7942. An interesting question, yet to be examined, is whether these clustered genes are coregulated. The possibility that DNA topology affects the transcription ofthe various ORFs in this region is another challenging problem for future research. Recognition of genes whose expression depends on level opens the way for clarification of the mechanism(s) conferring responsiveness of cyanobacteria to Experiments aimed at the identification ofthe relevant promoter regions, * and other components ofthe signal perception and transduction path will doubtless follow shortly. It should be borne in mind, however, that not all such genes are necessarily directly involved in the CCM or photosynthesis. Studies with mutants have indicated the importance of the correct structural organization of the carboxysomes. This probably involves the close association between CA and RubisCO envisaged by the theoretical models but still to be confirmed experimentally. The recent development ofimproved techniques for the isolation ofpurified carboxysomes is likely to result in the identification of their constituent polypeptides, and clarification of the manner by which carboxysomes fulfill their function in cyanobacterial photosynthesis. Furthermore, recognition of the constituent polypeptides should help to elucidate the intriguing problem of the mechanisms involved in the assembly and biogenesis

Chapter 15 Ambient Inorganic Carbon Concentration Effects of carboxysomes. Isolation of new mutants impaired in Ci transport will lead to identification of the bicarbonate carrier, that will then enable eventual reconstitution experiments. Such mutantsshould also help to verify whether an energy-dependent CA activity is in fact involved in the uptake and efficient recycling of

Acknowledgments Research in this laboratory was supported by grants from the U.S. A-Israel Binational Science Foundation (BSF); the National Council for Research and Development, Israel-GBF, Braunschweig, Germany; and the Ministry of Science of Niedersachsen, Germany.

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Ogawa T and Kaplan A (1987) The stoichiometry between and fluxes involved in the transport of inorganic carbon in cyanobacteria. Plant Physiol 83: 888–891 Ogawa T, Miyano A and Inoue Y (1985) Photosystem-I-driven inorganic carbon transport in the cyanobacterium, Anacystis nidulans. Biochim Biophys Acta 808: 77–84 Ogawa T, Kaneda T and Omata T (1987) A mutant of Synechococcus PCC 7942 incapable of adapting to low concentration. Plant Physiol 84: 711–715 Ogura Y, Takemura M, Oda K, Yamato K, Ohta H, Fukuzawa H and Ohyama K (1992) Cloning and nucleotide sequence of a frxC-ORF469 gene cluster of Synechocystis PCC6803: conservation with liverwort chloroplast frxC-ORF465 and nif operon. Biosci Biotechnol Biochem 56: 788–793 Omata T (1992) Characterization of the downstream region of cmpA: identification of a gene cluster encoding a putative permease of the cyanobacterium Synechococcus PCC7942. In: Murata N (ed) Research in Photosynthesis, Vol III, pp 807– 810. Kluwer, Dordrecht Omata T and Ogawa T (1986) Biosynthesis of a 42KD polypeptide in the cytoplasmic membrane of the cyanobacterium Anacystis nidulans strain R2 during adaptation to low concentration. Plant Physiol 80: 525–530 Omata T, Carlson TJ, Ogawa T and Pierce J (1990) Sequencing and modification of the gene encoding the 42-kilodalton protein in the cytoplasmic membrane of Synechococcus PCC 7942. Plant Physiol 93: 305–311 Orus MI, Martinez F, Rodriguez ML and Marco E (1992) Ultrastructural study of carboxysomes from requiring mutants of Synechococcus PCC 7942. In: Murata N (ed) Research in Photosynthesis, Vol III, pp 787–790. Kluwer, Dordrecht Peschek GA (1987) Respiratory and electron transport. In: Fay P and Van Baalen C (eds) The Cyanobacteria, pp 119–161. Elsevier Science Publishers, Amsterdam Pierce J and Omata T (1988) Uptake and utilization of inorganic carbon by cyanobacteria. Photosynth Res 16: 141–154 Pierce J, Carlson TJ and Williams JGK (1988) Anomalous oxygen sensitivity in a cyanobacterial mutant requiring the expression of ribulosebisphosphate carboxylase from a photosynthetic anaerobe. Proc Nat Acad Sci USA 86: 5753–5757 Price GD and Badger MR (1989a) Expression of human carbonic anhydrase in the cyanobacterium Synechococcus PCC 7942 creates a high requiring phenotype. Plant Physiol 91: 505–513 Price GD and Badger MR (1989b) Isolation and characterization of requiring mutants of the cyanobacterium Synechococcus PCC7942. Two phenotypes that accumulate inorganic carbon but are apparently unable to generate CO2 within the carboxysomes. Plant Physiol 91: 514–525 Price GD and Badger MR (1991) Evidence for the role of carboxysomes on cyanobacterial mechanism. Can J Bot 69: 963–973 Price GD, Coleman JR and Badger MR (1992) Association of carbonic anhydrase activity with carboxysomes from the cyanobacterium Synechococcus PCC7942. Plant Physiol 100: 784–793 Price GD, Howitt SM, Harrison K and Badger MR(1993) Analysis of a genomic DNA region from the cyanobacterium Synechococcus sp. strain PCC7942 involved in carboxysome

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Chapter 16 Assimilatory Nitrogen Metabolism and Its Regulation Enrique Flores and Antonia Herrero Instituto de Bioquímica Vegetal y Fotosíntesis, Universidad de Sevilla-CSIC, Facultad de Biología, Apartado 1113, E-41080 Sevilla, Spain Summary I. Introduction II. Nitrogen Fixation Fixation to Oxygen A. Relations of Cyanobacterial 1. Distribution of -Fixation Ability Among Cyanobacteria in Nonneterocystous Cyanobacteria 2. Protection of Nitrogenasefrom B. The Fixation Reaction Fixation System C. Genetic Structure of the D. Nitrogen Regulation of Fixation III. Nitrate and Nitrite Assimilation A. Nitrate and Nitrite Uptake B. Nitrate Reduction 1. Nitrate Reductase 2. Nitrite Reductase C. Regulation of Nitrate Assimilation 1. Regulation of the Activities of Nitrate and Nitrite Uptake 2. Regulation of the Expression of Proteins Involved in Nitrate Assimilation 3. Cotranscription of Some Genes Involved in Nitrate Assimilation IV. Assimilation of Organic Nitrogen A. Assimilation of Urea B. Assimilation of Amino Acids 1. Arginine 2. Glutamine V. Assimilation of Ammonium A. Ammonium Uptake B. Ammonium Assimilation Pathways 1. The Glutamine Synthetase/Glutamate Synthase Cycle 2. GS Enzyme and the glnA Gene 3. GOGAT 4. Other Ammonium Assimilating Enzymes VI. Distribution of Assimilated Nitrogen VII. Global Nitrogen Control Acknowledgments References

D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 487–517. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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Summary The element nitrogen (N) constitutes about 5–10% of the dry weight of a cyanobacterial cell. The purpose of this chapter is to review the assimilatory pathways which in free-living cyanobacteria lead from different extracellular N-sources to cellular N-containing components. Inorganic nitrogen in the form of ammonium is incorporated into glutamine and glutamate via the glutamine synthetase/glutamate synthase cycle. The glnA gene, encoding glutamine synthetase, has been characterized in a number of cyanobacteria. Glutamate (and glutamine) distribute N to other organic compounds by means of transaminases, and glutamate is itself a precursor of some other nitrogenous metabolites. Ammonium can be taken up from the external medium by the cyanobacterial cell, but it can also be derived from other nutrients, essentially nitrate and urea. Many cyanobacteria are able to fix under aerobic conditions. Strategies for protecting nitrogenase from in cyanobacteria include the temporal separation of nitrogenase activity and photosynthetic evolution, and in some filamentous cyanobacteria, the differentiation of heterocysts (cells specialized in fixation).A detailed characterization of nif genes has only been performed in a heterocyst-forming cyanobacterium. Nitrate reduction has been found to use photosynthetically reduced ferredoxin as an electron donor, and genes encoding nitrate transport and reduction proteins have been identified and shown to constitute an operon. Some amino acids like arginine and glutamine can also contribute N to some cyanobacteria; however, urea and amino acid utilization have been poorly investigated thus far. Pathways of N assimilation in cyanobacteria are induced upon ammonium deprivation, ammonium being the preferred N source. A gene, ntcA, encoding a transcriptional regulator required for expression of proteins subjected to nitrogen control has been identified. A major theme for future research is how information about the N status of the cell is sensed and transduced to the protein(s) effecting regulation of gene expression.

I. Introduction Nitrogen (N) can constitute as much as about 11 % of the dry weight of a cyanobacterial cell (Wolk, 1973). The cyanobacteria mainly use inorganic compounds (nitrate, dinitrogen, and ammonium) to fulfill their N requirements, but urea and other organic sources of N, such as some amino acids, can also be assimilated by some cyanobacteria. A general scheme representing major routes in N-assimilatory metabolism is depicted in Fig. 1. The fact that urea and inorganic N sources other than ammonium are first metabolized to ammonium to allow assimilation of their N atoms is emphasized in the left part of Fig. 1. Some amino acids, like arginine, are assimilated by the production of ammonium among their catabolic products. The central position played in cyanobacterial N metabolism by the glutamine synthetase/glutamate synthase (GS/GOGAT) cycle for ammonium assimilation is also emphasized in Fig. 1. Glutamate Abbreviations: ADH – alanine dehydrogenase; DCCD – N,N’dicyclohexylcarbodiimide; Fd – ferredoxin; GDH – glutamate dehydrogenase; GOGAT – glutamine(amide)-2-oxoglutarate aminotransferase (glutamate synthase); GS – glutamine synthetase; Ks – solute concentration that will give one half of the maximum rate of uptake; MSX – L-methionine-D,Lsulfoximine; Nar – nitrate reductase; Nir – nitrite reductase.

produced in this pathway is not only the major Ndonor for the biosynthesis of other N-containing metabolites but is itself a precursor for some amino acids and 5-aminolevulinate, the immediate precursor for porphyrin, phycobilin and chlorophyll biosynthesis (see Chapter 17). Glutamine also donates N to some metabolites. Finally, Fig. 1 shows that the amino acids arginine and aspartate together make up cyanophycin, a unique storage reservoir of N (and carbon) found in many cyanobacteria. Utilization of the different sources of N requires their passage through the cell wall. The cyanobacterial cell wall is like that of the Gram-negative bacteria that bear an outer membrane outside the peptidoglycan layer(s). The outer membrane contains a special type of proteins, known as ‘porins,’ which allow the passage of small molecules including amino acids and inorganic ions. There is evidence for the presence of such ‘porins’ in cyanobacteria (Resch and Gibson, 1983; Benz and Böhme, 1985). Therefore, it is the cytoplasmic membrane that represents the permeability barrier for the uptake of substances that are sources of N, and some of these substances are known to be taken up by the cyanobacterial cell with the aid of specialized transport systems. The convergence at the level of ammonium and

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the GS/GOGAT cycle of several N assimilation pathways (Fig. 1) allows the operation ofa regulatory network which helps the cyanobacterial cell to economize the process of N acquisition: the inhibition by ammonium ofthe activity or/and expression of proteins involved in the assimilation of alternative N sources. This system for N-control is currently a major theme of research in cyanobacteria.

II. Nitrogen Fixation Many cyanobacteria are able to grow at the expense of atmospheric under aerobic conditions and many more are able to perform fixation when anaerobic conditions are provided experimentally. Given the widespread distribution in nature of these microorganisms, it is believed that cyanobacteria

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contribute significantly to the process of biological fixation and thus participate in restoring to the soil combined-N lost through denitrification.

A. Relations of Cyanobacterial Oxygen

Fixation to

Nitrogenase, the enzymatic system for fixation, is irreversibly inactivated by oxygen when the enzyme is extracted from any organism (including cyanobacteria) so far tested. Therefore, those organisms able to perform fixation in aerobiosis have evolved mechanisms forprotecting their machinery from the deleterious effects of Because cyanobacteria are photoautotrophic organisms that perform oxygenic photosynthesis, they not only have to protect their machinery from

Enrique Flores and Antonia Herrero

490 atmospheric but also from intracellularly generated Cyanobacteria probably had to develop mechanisms to protect nitrogenase from intracellular photosynthetically evolved before they had to develop mechanisms to handle high levels of atmospheric The ability to fix was probably already present in cyanobacteria living in the primitive, basically anoxic atmosphere—long before cyanobacterial evolution led to the present-day levels of in the atmosphere.

1. Distribution of Cyanobacteria

Ability Among

The machinery for fixing has been shown to be present in all the currently recognized taxonomic groups of cyanobacteria (see Chapter 1). A phylogenetically-coherent group of filamentous cyanobacteria, that includesgenerasuchasAnabaena, Nostoc, and Fischerella of Sections IV and V of the taxonomic classification ofRippka et al. (1979), has developed a complex and efficient mechanism for performing fixation under aerobic conditions. This consists of the development of heterocysts, specialized cells that, under conditions ofaerobiosis and combined-N deprivation, differentiate from vegetative cells at semi-regular intervals in the filament and bear a series of modifications devoted to the protection of the apparatus from The structure, function and developmental biology of the heterocyst is dealt with in Chapter 27 of this book. Other cyanobacteria are able to use different strategies that permit fixation to be performed with a wide range of efficiencies under aerobic conditions. Finally, there are those cyanobacteria that are able to perform fixation only under anaerobic or microaerobic conditions. An experimental procedure has been established for screening nonheterocystous cyanobacteria for the ability to perform fixation in the complete absence of (Rippka and Waterbury, 1977). The procedure consists in subjecting cells to a period of N starvation under in the light (which would allow the cells to accumulate glycogen); the cells are subsequently treated with DCMU (to avoid photosynthetic oxygen evolution) under an atmosphere of Underthese conditions, nitrogenase activity supported by reducing power derived from the metabolism ofthe previously accumulated glycogen, is observable in many cyanobacteria. Alternatively, in the case of facultatively heterotrophic or

photoheterotrophic cyanobacteria, the initial period of N starvation can be substituted by the provision of an exogenous carbohydrate added at the onset of anaerobic induction. This method has revealed fixing capacity in 15% of the tested cyanobacterial strains belonging to Section I according to Rippka et al. (1979), including strains ofthe genera Gloeothece and Synechococcus; in 59% of the tested strains of Section II; and in 57% ofthe tested strains ofSection III, including strains of the genera Oscillatoria, Pseudanabaena, and the Lyngbya-PlectonemaPhormidium (LPP) group (Rippka and Waterbury, 1977). Some of these cyanobacteria had previously been described as aerobic This approach of searching for the potential to express nitrogenase activity has been complemented by searches, in a variety of nonheterocystous cyanobacteria, for DNA hybridizing to probes derived from the structural genes encoding nitrogenase in Anabaena sp. strain PCC 7120, the first cyanobacterium from which these genes were identified and cloned (Rice et al., 1982, see below). The presence of nif genes has been described in some strains of the genera Gloeothece (Kallas et al., 1983), Synechococcus, Cyanothece, the ‘LPP’ group, and Pseudanabaena (Kallas et al., 1985; Singh et al., 1987), Plectonema (Barnum and Gendel, 1985), and Trichodesmium (Zehr et al., 1991).

2. Protection of Nitrogenase from Nonheterocystous Cyanobacteria

in

For a more comprehensive review on strategies for protection ofthe cyanobacterial machinery from the reader should see Fay (1992). Among those nonheterocystous cyanobacteria that are able to perform fixation under aerobic conditions, the first to be described was the unicellular Gloeothece sp. strain PCC 6909 (previously known as Gloeocapsa sp.). This cyanobacterium is able to grow photoautotrophically with atmospheric as sole N-source (Wyatt and Silvey, 1969). A bellshaped curve results when the nitrogenase activity of suspensions of this cyanobacterium are plotted against concentration both in the dark and in the light (Maryan et al., 1986). Recently, it has been shown that in continuous cultures that were well-mixed and well-aerated, thus preventing supersaturation in the light, fixation occurs predominantly in the light (Ortega-Calvo and Stal, 1991). This means that more efficient provision of ATP and/or reductant

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can be supplied to nitrogenase in the light than in the dark. However, in batch cultures where higher concentrations of probably accumulate at least locally, fixation has been shown to be restricted to the dark period when the organism is subjected to light/dark diurnal cycles (Mullineaux et al., 1981); thus, fixation is temporally separated from photosynthetic production. These conditions probably mimic those of many natural communities of cyanobacteria that are usually characterized by supersaturation with during the day, whereas low or even anoxic conditionsprevail during the night (Ortega-Calvo and Stal, 1991). The occurrence of nitrogenase activity mainly in the dark period when a light/dark regime is imposed has also been described for some other aerobic, fixing cyanobacteria of the genera Synechococcus and Gloeothece (Huang and Chow, 1988) and Oscillatoria (Stal and Krumbein, 1985). In Synechococcus sp. strain RF-1, the rhythmic, diurnal fixation pattern parallels a similar pattern of nif mRNA expression, both persisting at least for some time after transferring synchronous cultures from the light/dark regime to continuous illumination. It has been suggested that the light/dark regime induces an endogenous rhythm for nitrogenase activity in this cyanobacterium (Huang and Chow, 1990). This rhythm might reflect fixation and photosynthesis taking place at different phases in the cell division cycle (Mitsui et al., 1986). This suggests that, even under continuous illumination, temporal separation of oxygenic photosynthesis and fixation can account for the ability of at least some nonheterocystous cyanobacteria to grow photoautotrophically under conditions. The organisms in which temporal separation of photosynthesis and fixation takes place should possess mechanisms for the provision of reductant and energy to nitrogenase while PS II is not functioning. These mechanisms probably rely upon the utilization of carbohydrate accumulated during a previous period ofphotosynthesis. Degradation and respiration in the dark of the accumulated carbohydrate not only can provide reductant and energy for nitrogenase, but can also contribute to lowering the levels of intracellular Planktonic Oscillatoria (Trichodesmium) spp. are considered to be the most important in marine pelagic environments (Carpenter and Romans, 1991). These microorganisms are found forming massive blooms under calm-water conditions. Based

491 on some data collected in the field, it was initially proposed that some cells, in the center of the bundleshaped colonies and in which photosynthetic evolution would not operate, could be specialized in fixation(CarpenterandPrice,1976).Nevertheless, spatial partitioning does not seem to be necessary for aerobic fixation in Trichodesmium spp. because in laboratory cultures colony formation does not seem to be required for fixation (Ohki and Fujita, 1988). Moreover, fixation in Trichodesmium spp. is essentially light-dependent. Natural populations of T. thiebautii have been described to exhibit a diel periodicity in nitrogenase activity which is ‘turned on’ near dawn and ‘turned off’ near dusk. This seems to result from de novo synthesis of nitrogenase each morning and inactivation and degradation of the enzyme in the late afternoon and night (Capone et al., 1990). When Trichodesmium spp. were grown under a light/dark regime, nitrogenase activity was almost exclusively found in the light (see Fay, 1992). Light-dependence of nitrogenase activity in Trichodesmium spp. could result from a requirement for a direct supply of photosynthetically generated ATP and reductant for fixation. An inability to provide reductant and energy when PS II is not working may also be the reason why another marine isolate of the genus Synechococcus is unable to fix in the dark (Spiller and Shanmugam, 1987). Respiration seems to contribute to the protection of nitrogenase from in these organisms (Bergman et al., 1993). The filamentous, nonheterocystous cyanobacterium Plectonema boryanum is an example of an organism requiring anaerobiosis for fixation (Stewart and Lex, 1970). The high sensitivity to of nitrogenase in vivo demonstrates the absence of mechanisms providing effective protection from in this cyanobacterium (see Fay, 1992; Rai et al., 1992). A similar situation seems to prevail in another filamentous, nonheterocystous cyanobacterium, Phormidium foveolarum (Weisshaar and Böger, 1983).

B. The

Fixation Reaction

The enzyme complex nitrogenase catalyzes the ATPdependent reduction of to two molecules of ammonium; this reaction requires the transference of six electrons to Nitrogenase has proven to be very similar in all organisms tested to date. It consists of two different protein components

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(Table 1): dinitrogenase (the Mo-Fe protein) and dinitrogenase reductase (the Fe protein). Dinitrogenase is an tetramer, and its and subunits are encoded by the nifD and nifK genes, respectively. This tetramer binds four [4Fe-4S] clusters, organized into two ‘P clusters’ and two Fe-Mo cofactors (see Fig. 3). Dinitrogenase reductase is a dimer of identical subunits, that are encoded by the nifH gene and that together bind one intersubunit 4Fe-4S center (see Figs. 2 and 3). Dinitrogenase reductase mediates the ATP-dependent tranference of electrons from external electron donors, such as

Enrique Flores and Antonia Herrero

ferredoxin or flavodoxin, to the P clusters of dinitrogenase. Dinitrogenase binds through its Mo-Fe cofactors and catalyzes the reduction of to ammonia. A recent major advance in nitrogenase research has been the determination by X-ray crystallography of the structures of the Azotobacter vinelandii dinitrogenase and dinitrogenase reductase proteins, as well as the formulation of models for their metal cofactors (Georgiadis et al, 1992; Kim and Rees, 1992, 1993, 1994; Chan et al., 1993; also see Dean et al., 1993). Figure 2 shows the alpha carbon backbone structure of an dimer of

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Nitrogen Metabolism

dinitrogensase; dinitrogenase reductase, whose structure was independently determined (Georgiadis et al., 1992), has been docked on to this dimer through a computer simulation-energy minimization procedure. The positions of the cofactors for the enzyme are also shown in Fig. 2; a more detailed scheme for the structures of the metal cofactors and the pathway of electron transport is shown in Fig. 3. Readers seeking more details concerning the structure

493 and function of nitrogenase should consult the review article of Dean et al. (1993) or the primary publications describing the X-ray structures of the nitrogenase of Azotobacter vinelandii (Georgiadis et al, 1992; Kim and Rees, 1992, 1993; Chan et al., 1993). Different studies show that nitrogenase from cyanobacteria is much like that of other bacteria, (i) Native electrophoresis showed that the enzyme from

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heterocysts of Anabaena variabilis is composed of two protein components, one containing non-heme iron and the other non-heme iron and molybdenum; by electrophoresis in the presence of SDS these components were resolved into subunits of similar sizes to those of nitrogenase from other organisms (Peterson and Wolk, 1978). (ii) The Mo-Fe protein of nitrogenase from Anabaena cylindrica has been shown to form an active complex with the Fe-protein of Clostridium pasteurianum (Tsai and Mortenson, 1978). Additionally, acidified extracts from Plectonema boryanum induced for nitrogenase complemented the defective dinitrogenase of a mutant of Azotobacter sp. impaired in the formation of the Fe-Mo cofactor. Dinitrogenase from eitherKlebsiella sp. orAzotobactersp. complementedcell-free extracts from Plectonema sp. cultures induced under molybdenum starvation (Nagatani and Haselkorn, 1978). These reconstitution experiments showed that cyanobacterial nitrogenase is analogous to those of other bacteria. (iii) The best proof of similarity between cyanobacterial nitrogenase and the enzyme from other procaryotes has been obtained through the identification and sequencing of the structural genes nifH, nifD, and nifK encoding the nitrogenase complex from a variety of cyanobacteria. Sequences of the protein products of these genes have proven very similar to those of the products of the corresponding nif genes of other bacteria (Dean and Jacobson, 1992).

Enrique Flores and Antonia Herrero

The reduction of to catalyzed by nitrogenase is a highly endergonic reaction requiring metabolic energy in the form ofATP. The biological reduction of by nitrogenase always occurs concomitantly with the reduction of protons to a wasteful reaction costing both energy and reductant that decreases the efficiency of the fixation process. Like many other organisms, cyanobacteria possess an enzyme, uptake hydrogenase, that returns electrons from at least a portion of the produced by nitrogenase to both respiratory and photosynthetic electron transport chains for the production of ATP and reductant (see Chapters 12 and 13). Besides recovering ATP and reducing power, the stimulated operation of respiration promoted by uptake hydrogenase contributes to the consumption of and could thus represent a mechanism for protecting the fixation system from (see e.g., Houchins, 1984; Spiller and Shanmugam, 1987). In the heterocyst, ferredoxin (Fd) is an electron donor to nitrogenase. This was shown by the ability of Fd to support nitrogenase activity in extracts from Anabaena sp. (Smith et al., 1971; Schrautemeier and Böhme, 1985). More recently, the identification and cloning of the gene fdxH, that encodes a Fd specific to the heterocyst (that serves as immediate electron donor to nitrogenase) has been achieved; the fdxH gene is present in the nif gene cluster ofthe Anabaena sp. strain PCC 7120 genome (Böhme and Haselkorn, 1988). The fdxH gene has also been identified in

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Nitrogen Metabolism

other heterocystous cyanobacteria, as well as in the filamentous, nonheterocystous, Plectonema sp. strain PCC 73110 (Schrautemeier and Böhme, 1992; Schrautemeier et al., 1994). In Anabaena sp. strain PCC 7120, a gene has recently been cloned that encodes a protein sharing 42% identity to pyruvate flavodoxin oxidoreductase, the product of gene nifJ, from Klebsiella pneumoniae (Bauer et al., 1993). Schmitz et al. (1993) have recently reported the isolation of nifJ homologs from Anabaena sp. strain PCC 7119 as well as Anabaena variabilis strain ATCC 29413. The Anabaena sp. strain PCC 7120 nifJ gene is expressed only under iron-limiting conditions and is required for growth only on media that is both iron- and N-limited. A gene encoding flavodoxin has been cloned from Anabaena sp., and flavodoxin mRNA has only been detected in filaments grown without iron (Leonhardt and Straus, 1989; Fillat et al., 1991). However, it has not been established that this particular flavodoxin can be an electron donor to nitrogenase in Anabaena sp. The pathways by which Fd could be reduced, as well as those providing ATP, to support fixation have mostly been studied in heterocysts (see Chapter 27 of this book). In nonheterocystous cyanobacteria, only a few physiological studies have been performed addressing the question of the metabolic processes providing ATP and reducing power for fixation (see e.g. Maryan et al., 1986; Spiller and Shanmugam, 1987). Reducing power could be provided photosynthetically in the light, or via degradation ofeither externally provided sugars (in the case of heterotrophic or photoheterotrophic cyanobacteria) or reserve-carbohydrates, both in the dark and in the light. ATP could be generated photosynthetically via cyclic or non-cyclic electron flow or by respiration of the reduced pyridine nucleotides generated in sugar oxidation pathways. Prevalence of some of those routes over the others probably depends on the environmental conditions to which the organism is confronted, as well as on the particular cyanobacterium being considered. More studies on this subject are evidently necessary.

C. Genetic Structure of the

Fixation System

A region of the chromosome of the heterocystforming cyanobacterium Anabaena sp. strain PCC 7120 containing the nifHDK genes, encoding the

495 three polypeptides of nitrogenase, was identified and cloned by means of hybridization with the Klebsiella pneumoniae nif genes (Mazur et al., 1980; Rice et al., 1982). In contrast to the arrangement in K. pneumoniae, in which the nifHDK genes are contiguous, in the identified region ofAnabaena sp. strain PCC 7120 (later demonstrated to correspond to the genome of vegetative cells) nifH and nifD were contiguous while nifK was separated from nifD by about 11 kb of DNA. The three genes were sequenced and found to be strongly similar to the corresponding genes ofother bacteria (see Haselkorn and Buikema, 1992). In the genome ofthe heterocyst, the structure of this nif region (Fig. 4) is somewhat different as the result of a DNA rearrangement—an excision event that takes place during heterocyst differentiation. The most striking consequences of this rearrangement are a change in the C-terminal end of nifD and expression of nifHDK as an operon (see Haselkorn and Buikema, 1992, and Chapter 27 of this book). A different region of the genome of Anabaena sp. strain PCC 7120 was cloned that contained DNA homologous to K. pneumoniae nifH but differing to some extent from the nifH gene present in the nif cluster referred to above (Rice et al., 1982). Besides nifHDK, other genes have been identified and characterized in the nif-region of Anabaena sp. strain PCC 7120 (Fig. 4). Upstream from nifH are: nifU, nifS, fdxN, and nifB (Mulligan et al., 1988; Mulligan and Haselkorn, 1989). The three nif genes were identified by their similarity to the corresponding genes of other bacteria. Although the actual function of the products of nifU and nifS are unknown, the nifB product is required for Mo-Fe cofactor synthesis (Shah et al., 1988). The fdxN gene codes for a bacterial-type Fd whose function in fixation has not yet been determined. A 55-kb DNA fragment which interrupts the coding region of fdxN in the chromosome ofvegetative cells is also excised during heterocyst development in many species. In the heterocysts, the four genes are probably transcribed as an operon from a promoter located upstream of nifB (Mulligan and Haselkorn, 1989; see Fig. 4). About 6–7 kb downstream from nifK, the fdxH gene encoding the heterocyst-specific Fd (see above) is found (Böhme and Haselkorn, 1988). Between fdxH and nifK there are seven ORFs, four of which have, on the basis of sequence similarity to genes of Klebsiella sp. or of Azotobacter sp., been identified

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as the genes nifW, nifX, nifN and nifE (Borthakur et al, 1990; Haselkorn, 1992; Haselkorn and Buikema, 1992). The genes nifV, encoding homocitrate synthetase required for Mo-Fe cofactor synthesis (Strieker et al., 1992) and nifJ, encoding pyruvate flavodoxin oxidoreductase (see above), have recently been identified. Apart from the regulatory genes that do not necessarily exist in Anabaena sp. strain PCC 7120 (since this organism can have a different scheme of regulation), nifM, required for maturation of dinitrogenase reductase, is the only essential K. pneumoniae nif gene not yet identified in Anabaena sp. strain PCC 7120. Whereas two heterocystous strains belonging to Section V of Rippka et al. (1979) have contiguous nifHDK genes in the vegetative cell chromosome (Saville et al., 1987; Singh and Stevens, 1992), the same disposition of the nifHDK genes present in Anabaena sp. strain PCC 7120, i.e. nifH and nifD contiguous and nifK separated from nifD, has normally been found in the vegetative cell chromosome of heterocystous cyanobacteria, that belong to Section IV of Rippka et al. (1979), studied to date (see Haselkorn and Buikema, 1992). In Anabaena variabilis strain ATCC 29413, the DNA insertion of 55 kb that in the vegetative cells of Anabaena sp. strain PCC 7120 interrupts the fdxN coding region does not seem to be present, since the rbcLS genes (encoding the two subunits of ribulose bisphosphate carboxylase) are separated from nifH by only about 10 kb in the vegetative cell chromosome (Herrero and Wolk, 1986); in Anabaena sp. strain PCC 7120 the distance between rbcLS and nifHDK is about 70 kb in vegetative cells and about 15 kb in heterocysts. Anabaena variabilis strain ATCC 29413 has recently been shown to carry a second nif gene cluster encoding a second molybdenum-dependent nitrogenase that appears to be expressed in vegetative cells under

Enrique Flores and Antonia Herrero

anaerobic conditions (Thiel et al., 1993) and that might correspond to that described by Hirschberg et al. (1985). The nifHDK genes have been identified in a number of nonheterocystous cyanobacteria including unicellular and filamentous strains (see Section II A, l), and their relative positions determined. In all the cases, nifHDK have a contiguous disposition (lacking intervening sequences between nifD and nifH) like that found in the heterocysts of the heterocyst-forming cyanobacteria. The nifH and nifD genes from Plectonema boryanum have been sequenced and found to encode polypeptides 84% and 76% identical, respectively, to those of Anabaena sp. strain PCC 7120 (Fujita et al., 1991). In P. boryanum, nifU is located upstream from nifH (Fujita et al., 1991). In Nostoc commune strain UTEX 584 a gene, namely glbN, not described in Anabaena sp. strain PCC 7120 is found between nifU and nifH (Potts et al., 1992). The glbN product (cyanoglobin) shows similarity with myoglobins of some ciliated protozoa. The fact that cyanoglobin is only expressed under conditions of N-deprivation, together with its location in the cluster of nif genes, suggests a possible scavenging function for cyanoglobin in cells. Anabaena variabilis strain ATCC 29413 has been reported to grow while fixing at about the same rate in the presence of molybdenum or of vanadium (Kentemich et al., 1988). This was an indication of the existence in this cyanobacterium of an alternative, V-dependent nitrogenase as has been found in other diazotrophs. The genes vnfDGK, encoding polypeptides of the V-dependent nitrogenase, have in fact been cloned and sequenced showing strong similarity to the corresponding genes of Azotobacter sp. (Thiel, 1993). Some indications of the existence

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in A. variabilis ofa third Fe nitrogenase, also present in some other diazotrophs, have been reported (Kentemich et al., 1991).

D. Nitrogen Regulation of

Fixation

Like other diazotrophs, cyanobacteria preferentially use fixed N (e.g., nitrate or ammonium), and express the system for fixation only in the absence of a suitable source of combined N (see e.g., Stewart and Rowell, 1975). The ability of a particular source of combined N to inhibit the expression of the fixation system depends upon the efficiency with which that nutrient is assimilated. In Anabaena sp. strain PCC 7120, mRNAs hybridizing to nifH, nifD, nifK (Rice et al., 1982), nifB (Mulligan and Haselkorn, 1989), and fdxH (Böhme and Haselkorn, 1988) have been found in filaments derepressed by incubation in N-free medium, but not in ammonium-grown cells. However, because heterocysts (the cells that express nitrogenase) do not develop in the presence of ammonium, absence of nif gene expression in ammonium-supplemented cultures might be a consequence of inhibition by ammonium of heterocyst development, with ammonium not having a direct effect on nif gene expression. Nonetheless, ammonium has also been shown to inhibit the synthesis of nitrogenase in cultures that bear fully developed heterocysts (Rowell et al., 1977; Ramos et al., 1985). Therefore, ammonium seems to negatively affect nitrogenase synthesis besides the effect it has on heterocyst development. A negative effect ofammonium on nitrogenase protein synthesis has alsobeendescribed in Gloeothecesp. (Mullineaux et al., 1983) as well as in Plectonema boryanum (Rai et al., 1992). The negative effect of ammonium on nitrogenase synthesis is not manifest if glutamine synthetase activity (GS, the main enzyme for ammonium assimilation, see Section V B) is depressed, e.g. by treatment with MSX (Stewart and Rowell, 1975; Ramos et al., 1985). Thus, ammonium assimilation via GS seems to be required for ammonium-promoted repression to operate. This is also supported by the phenotype exhibited by mutant strains isolated from Anabaena variabilis strain ATCC 29413 which are impaired in GS (Polukhina et al., 1982, Spiller et al., 1986; see also Hien et al., 1988). In contrast to what it is the case in the parental strain, these mutants

497 exhibit high levels ofnitrogenase in the presence of ammonium and excrete to the medium ammonium resulting from fixation. Nitrate has also been shown to repress nitrogenase synthesis in Anabaena sp. (Meeksetal., 1983;Ramos and Guerrero, 1983; Martin-Nieto et al., 1991), Plectonema boryanum (Nagatani and Haselkorn, 1978) and Synechococcus sp. strain RF-1 (Huang and Chou, 1991). Nitrate-promoted inhibition of nitrogenase synthesis operates at the level of nifHDK mRNA (Fujita et al., 1991; Huang and Chou, 1991; Martín-Nieto et al., 1991) and requires both nitrate reduction (Martín-Nieto et al., 1991) and further metabolism through GS of the ammonium resulting from nitrate reduction (Nagatani and Haselkorn, 1978; Ramos and Guerrero, 1983). The repressive effect of nitrate on nitrogenase seems thus to be of the same nature as the effect due to externally-added ammonium. In addition to regulating nitrogenase synthesis, ammonium, under certain conditions (pH 10 and ammonia concentrations in the range 0.5–1 mM), provokes an inactivation of nitrogenase through a modification of the Fe protein (dinitrogenase reductase) (see Böhm et al., 1992, and references therein).

III. Nitrate and Nitrite Assimilation Nitrate is probably the most abundant source of combined N for cyanobacterial nutrition. The assimilation of nitrate by cyanobacteria involves nitrate uptake and reduction of intracellular nitrate (via nitrite) to ammonium, which is the N form incorporated into organic compounds. Nitrite, which can also fulfill the N requirement ofcyanobacteria, is taken up into the cell and then reduced to the level of ammonium.

A. Nitrate and Nitrite Uptake In cyanobacteria, the uptake of nitrate takes place through a transport system that exhibits a high affinity for nitrate (Flores et al., 1983a), with values of having been determined in Synechococcus sp. and of in Anabaena sp. (see e.g., Meeks et al., 1983; Tischner and Schmidt, 1984; Rodríguez et al., 1992). In the unicellular cyanobacteria Synechococcus sp. strains PCC 6301

498 (Anacystis nidulans) and PCC 7942, intracellular accumulation of nitrate has been measured and positive free energy changes for the transport of nitrate have been calculated (Lara et al., 1987; Rodríguez et al., 1992). The endergonic nature ofthe transport process is also manifest in the observed sensitivity ofnitrate uptake to a protonophore and to the inhibitor of the DCCD (Ohmori et al., 1977; Flores et al., 1983a). A cytoplasmic-membrane protein of about 48 kDa was identified in Synechococcus sp. strain PCC 7942 which exhibited the same pattern of regulation as other components of the nitrate assimilation system (i.e., repression by ammonium, see below) and whose expression was altered in mutant strains pleiotropically impaired in nitrate assimilation (Madueño et al., 1988b). This suggested an involvement ofthat protein in nitrate transport. A similar protein has been described in the cytoplasmic membrane of Synechococcus sp. strain PCC 6301 (Sivak et al., 1989). The complete nrtA gene encoding the 48 kDa polypeptide involved in nitrate transport was cloned from a mutant of Synechococcus sp. PCC 7942 carrying an gene cassette inserted into nrtA (Omata, 1991). Interestingly, the nrtA gene has been located just downstream of nir, the gene encoding nitrite reductase (Luque et al., 1992). Downstream of nrtA three other genes (nrtBCD) whose predicted products show homology to proteins of bacterial transport systems dependent on binding proteins (‘multicomponent systems’) have been located (Omata et al., 1993). NrtA, although being membrane-bound, has been proposed to represent a substrate-binding protein. NrtB is homologous to Pro W, the integral membrane component of the glycine-betaine permease of E. coli. NrtD exhibits ATP-binding features and extensive homology to the ‘conserved component’ (the ATP-binding component) of the bacterial multicomponent transport systems. NrtC is a polypeptide that has a large Cterminal domain with structural similarity to NrtA (30% identity) and an N-terminal domain that has features ofan ATP-binding protein and is homologous (58% identity) to NrtD. The N-terminal domain of NrtC can provide a second ATP-binding site while a possible function for the C-terminal domain ofNrtC is currently unknown. Because the nrt genes are located in a cluster of genes encoding proteins involved in nitrate assimilation (downstream of nir and upstream from narB, see below; Luque et al.,

Enrique Flores and Antonia Herrero 1992; Omata et al., 1993), and because inactivation of any of the nrt genes renders mutant strains unable to grow on low concentrations (< 1 mM) of nitrate (Omata et al., 1989,1993), these genes are thought to encode the components of the transport system for nitrate in Synechococcus sp. strain PCC 7942. Bacterial multicomponent transport systems have recently been demonstrated to use ATP directly as the powering substrate (see Ames and Joshi, 1990). Thus, this should be the case for the multicomponent nitrate transport system of Synechococcus sp. However, based on the positive effect of on the level of accumulation ofnitrate in a nitrate reductase mutant of Synechococcus sp., the operation of a has been suggested (Rodríguez et al., 1992). Nonetheless, a strict requirement for of nitrate uptake has not been demonstrated, and the observed influence of on the extent of nitrate accumulation could be due to an effect of on membrane potential which in turn might affect a process distinct from nitrate inwards transport (for instance nitrate efflux). The occurrence of a high-affinity, concentrative transport system for nitrate in cyanobacteria permits the uptake by these organisms of the rather low concentrations of nitrate that may be found in their natural environments (e.g., in seawater and some freshwater lakes), and the intracellular accumulation of nitrate will allow nitrate reductase to function. However, interfering with nitrate transport by inactivation of nitrate transport genes does not preclude nitrate-dependent growth when nitrate is supplied at a high concentration, e.g. >10 mM (Omata et al., 1989, 1993). Entrance of nitrate into the cell under these conditions appears to take place by passive diffusion since, at nitrate concentrations higher than 1 mM, nitrate uptake increases linearly as the nitrate concentration in the external medium is raised (Omata et al., 1989). Nitrite uptake by cyanobacteria takes place in two different ways: (i) active transport, which is sensitive to DCCD and exhibits high affinity for nitrite 6– Flores et al., 1987; Madueño et al., 1987); and (ii) diffusion of nitrous acid whose contribution to the net uptake ofnitrite decreases as the pH of the medium is raised (Flores et al., 1987; Martín-Nieto et al., 1989). Competition for uptake observed between nitrate and nitrite suggests that the same system operates the active transport of both substrates (Madueño et al., 1987; Rodríguez et al., 1992). Disruption of the nrtD gene in Synechococcus sp.

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strain PCC 7942 not only impaired nitrate transport, but also abolished active nitrite transport, thus showing that nitrate and nitrite are transported by the NrtABCD permease (Luque et al., 1994a).

B. Nitrate Reduction Intracellular nitrate is reduced to nitrite in a twoelectron reaction catalyzedby nitrate reductase (Nar), theresulting nitritebeing thenreduced to ammonium in a six-electron reaction catalyzed by nitrite reductase (Nir). Both Nar and Nir use reduced Fd as the physiological electron donor (Manzano et al., 1976; Méndez et al., 1981; Arizmendi and Serra, 1990). In the reaction catalyzed by Nar, two moles of Fd are oxidized permoleof nitrate reducedtonitrite,whereas in that catalyzed by Nir six moles of Fd are oxidized permoleofnitritereducedtoammonium.Flavodoxin can serve as an electron donor alternative to Fd during growth under iron-limited conditions (Manzano, 1977). Both Nar andNir are conveniently assayed in vitro by using methyl viologen, chemically reduced with dithionite, as an electron donor (see e.g. Manzano et al., 1976; Martín-Nieto et al., 1989). In some cyanobacteria either Nar or both Nar and Nir have been found firmly bound to thylakoid membranes. This has permitted the demonstration of nitrate or nitrite photoreduction by subcellular preparations in a PS I-dependent reaction (Hattori and Myers, 1967; Manzano et al., 1976; Peschek, 1979). Alternatively, using water as a source of electrons, photoreduction of nitrate or nitrite can be shown in a reaction involving activity of both PS I and PS II (Candau et al., 1976; Ortega et al., 1976). This, together with the observed quenching of the chlorophyll fluorescence induced by nitrate and nitrite (Serrano et al., 1981) and in vivo determined stoichiometries of 2 moles of evolved per mole of nitrate reduced to ammonium and 1.5 moles of per mole of nitrite reduced to ammonium (Flores et al., 1983a), indicates that nitrate reduction in cyanobacteria is a genuine photosynthetic process, analogous to the photosynthetic reduction of NADP+.

499 Herrero and Guerrero, 1986; Martín-Nieto et al., 1990) which would be integrated into a Mo cofactor (Martín-Nieto et al., 1990), and non-heme iron and acid-labile sulfide (Candau, 1979; Mikami and Ida, 1984), which in Nar from Plectonema boryanum mightconstitutetwo clusters (Mikami and Ida, 1984). Nar from heterocyst-forming cyanobacteria exhibits a biphasic kinetic behavior with nitrate as the variable substrate that is not generally observed with Nar from nonheterocystous strains (MartínNieto etal., 1992). FromSynechococcussp. strainPCC7942,multiple mutants that are unable to grow at the expense of nitrate and that are impaired in Nar have been isolated (Stevens and Van Baalen, 1970; Kuhlemeier et al., 1984a, 1984b; Madueño et al., 1988a; Vega-Palas et al., 1990). By means of complementation ofsome of these mutants with gene banks ofwild-type DNA, or by means of marker-rescue of transposon-induced Nar deficient mutants, three genes, namely narA, narB, and narC, involved in the reduction of nitrate to nitrite were isolated (Kuhlemeier et al., 1984a, 1984b). These three genes are unlinked in the genome of Synechococcus sp. The narB gene has been sequenced (Andriesse et al., 1990). Based on the similarity of the predicted amino acid sequence of the NarB protein to that of the of E. coli Nar and of other molybdo-proteins, and on the similarity of its predicted amino acid composition to that of the Plectonema boryanum Nar (Mikami and Ida, 1984), it has been suggested that narB could represent the structural gene for nitrate reductase (Andriesse et al., 1990). Nothing is currently known about the actual function ofthe products ofthe genes narA or narC, although they could be involved in the biosynthesis of the molybdenum cofactor of Nar. With respect to molybdenum metabolism, the existence in Anabaena variabilis strain ATCC 29413 of both common genes for the synthesis of the Mo cofactor of Nar and the Fe-Mo cofactor of nitrogenase, and specific genes for the synthesis of the Mo cofactor of Nar, has been proposed (Martín-Nieto et al., 1990).

2. Nitrite Reductase 1. Nitrate Reductase Cyanobacterial nitrate reductases studied to date are proteins of about 75 kDa which appear to be constituted by a single polypeptide (Candau, 1979; Mikami and Ida, 1984; Martín-Nieto et al., 1992; see Table 1). They contain molybdenum (Peschek, 1979;

Cyanobacterial Nir consists of a single polypeptide of 52–68 kDa (Hattori and Uesugi, 1968; Manzano, 1977; Méndez and Vega, 1981; Yabuki et al., 1985; Arizmendi and Serra, 1990; Miyaji and Tamura, 1992). From spectrophotometric studies ofNir from several cyanobacteria, the presence of siroheme as a

500 prosthetic group has been proposed (Méndez and Vega, 1981; Arizmendi and Serra, 1990; Miyaji and Tamura, 1992). Kinetic parameters of Nir from some strains of cyanobacteria have been determined (see Table 1). Mutants unable to grow on nitrate or nitrite and impaired in Nir have been isolated (Stevens and Van Baalen, 1970; Madueño et al., 1988a; Vega-Palas et al., 1990). The nir gene, encoding Nir, from Synechococcus sp. strain PCC 7942 has been cloned after rescue in E.coli of a Tn901-carrying genomic fragment from a Tn901-induced Nir mutant (Luque et al., 1992). The nir gene encodes a protein with a molecular weight of 56,506 which is 54% and 50% identical to Nir from Zea mais (maize) and Spinacea oleracea (spinach), respectively (Luque et al., 1993). Nir from Synechococcus sp. also shows homology to the hemoprotein component of enterobacterial sulfite reductase, and bears the four conserved cysteine residues, as well as other conserved amino acids surrounding these cysteines, that have been attributed structural importance in the configuration of the active site of the sulfite reductase hemoprotein and that are also present in higher-plant nitrite reductase (Ostrowski et al., 1989). Synechococcus sp. strain

Enrique Flores and Antonia Herrero PCC 7942 Nir would therefore bear an active center much alike that present in these other reductases (Luque et al., 1993) in which an cluster is bound to the protein through the S atoms of the above-mentioned four conserved cysteine residues, one of which also binds one molecule of siroheme thus bridging together the two prosthetic groups (Ostrowski et al., 1989; see Fig. 5). The facts that Nir from a cyanobacterium and higher plants have been characterized as homologous proteins (Luque et al., 1993) and that Nir from higher plants is located within the chloroplast and encoded by a nuclear gene, provide support to the notion that the chloroplast originates from cyanobacterial-like ancestor(s), and that genes from the endosymbiont have been transferred to the nucleus of the host during the course of higher-plant evolution (see Chapters 4 and 5 of this book). Fig. 6 shows the structure of the Synechococcus sp. strain PCC 7942 genomic region that contains the nitrate-assimilation genes. A similar cluster of nitrateassimilation genes has recently been identified in Anabaena sp. strain PCC 7120 (J. E. Frías, E. Flores and A. Herrero, unpublished; Y. Cai and C. P. Wolk, personal communication).

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501

C. Regulation of Nitrate Assimilation

6301 exhibits a strict dependence upon the operation of fixation, and it has been proposed that some product(s) may have a regulatory, positive action on nitrate uptake (Flores et al., 1983c). Nitrate transport is freed from its requirement for products in cells that cannot assimilate ammonium, and has a positive effect on the MSX-promoted restoration ofnitrate uptake after its inhibition by ammonium (Flores et al., 1983c). These observations establish a link between the negative effect of ammonium assimilation and the positive effect of fixation on nitrate transport in Synechococcus sp., the proposal having been made that the products (putative positive effectors) could combine with the ammonium assimilation products (putative negative effectors) removing these latter compounds in the cell (Flores et al., 1983c). The actual effector(s), either positive or negative, ultimately interacting with the nitrate transport proteins remain to be identified. As is the case for the transport ofnitrate, the active transport of nitrite is inhibited by product(s) formed through ammonium assimilation via GS (Flores et al., 1987). Because nitrate and active nitrite uptake take place through the same transport system (see Section III A), it is expected that the same mechanism regulates nitrate and active nitrite transport.

When reduced N in the form of ammonium is available, it is preferentially used over other good N sources such as nitrate or nitrite. The negative effect of ammonium on nitrate assimilation is exerted at two different levels: (i)ammoniumaddition provokes an almost immediate cessation of the uptake of nitrate by cyanobacterial cells that are actively assimilating this nutrient; this inhibitory effect operates at the level of nitrate transport (Ohmori et al., 1977; Flores et al., 1980; Lara et al., 1987); (ii) exposure to ammonium results in repression of the synthesis of proteins involved in nitrate assimilation.

1. Regulation of the Activities of Nitrate and Nitrite Uptake The inhibition of nitrate uptake by ammonium requires that ammonium be incorporated into C skeletons through the GS/GOGAT cycle, as ammonium is unable to exert any negative effect on nitrate uptake in cells that bear an inactive GS (e.g., by treatment of the cells with MSX, see Section V B) or an inactive GOGAT (by treatment with azaserine; Flores et al., 1980). Although this inhibition by ammonium of nitrate uptake has been suggested to be due to a reduction of the cellular levels of ATP caused by ammonium assimilation through GS (Ohmori et al., 1977), this does not seem to be the case as only transient changes in the cellular levels of ATP are observed upon ammonium exposure, the original high levels being soon recovered even when inhibitory concentrations of ammonium are maintained in the external medium (Ohmori and Hattori, 1978). Itseems more likelythat ammonium inhibition of nitrate uptake operates through feed-back inhibition exerted by product(s) of ammonium assimilation via GS (Flores et al., 1980). The suggestion has been made that inhibition could be mediated by some nitrogenous metabolite(s) generated in reactions that involve glutamine as the amido N-donor (Romero et al., 1985). Nitrate uptake by Synechococcus sp. strain PCC

2. Regulation of the Expression of Proteins Involved in Nitrate Assimilation When cells of a variety of cyanobacteria are transferred from ammonium-containing media to media containing either nitrate or nitrite as the sole N-source, an increase in the cellular levels of Nar (Stevens and Van Baalen, 1974; Herrero et al., 1981, 1985; Martín-Nieto et al., 1989; Rai et al., 1992), Nir (Méndez et al., 1981; Herrero and Guerrero, 1986; Arizmendi et al., 1987; Martín-Nieto et al., 1989), and the 48 kDa cytoplasmic membrane protein that is part of the nitrate transporter (Madueño et al., 1988b; Sivak et al., 1989) is observed. For Nar and Nir, it has been shown that this increase in activity levels does not take place in the presence ofinhibitors

502 of transcription or translation such as rifampin, chloramphenicol or erythromycin (Stevens and Van Baalen, 1974; Herrero et al., 1981, 1985; Herrero and Guerrero, 1986; Arizmendi et al., 1987; Rai et al., 1992). Moreover, in Synechococcus sp. strain PCC 7942 (Suzuki et al., 1993; Luque et al., 1994b) and Anabaena sp. strain PCC 7120 (J. E. Frías, E. Flores and A. Herrero, unpublished results), mRNA of nitrate assimilation genes is not detected in the presence of ammonium. Thus, ammonium promotes inhibition of the synthesis of nitrateassimilation proteins. As is also the case for ammonium-promoted inhibition of nitrate and nitrite transport (see above), ammonium must be metabolized through GS in order to repress Nar and Nir, since treatment of the cells with MSX leads to Nar and Nir development even in the presence of ammonium (Herrero et al., 1981, 1985; Herrero and Guerrero, 1986). The presence of nitrate or nitrite generally has a positive effect on Nar and Nir levels and is strictly required for the expression of Nar and Nir activities in cyanobacteria (Herrero et al., 1981, 1985; Martín-Nieto et al., 1989; Rai et al., 1992). In those instances when nitrate or nitrite is required, ammonium has been established to promote a distinct inhibition of Nar and Nir synthesis that is not merely exerted through exclusion of the positive effector (that would be a consequence of inhibition by ammonium of nitrate transport and active nitrite transport). Under conditions in which nitrite mostly enters the cyanobacterial cell in a passive way (insensitive to ammonium inhibition), Nar and Nir do not develop in a medium containing nitrite and ammonium (Martín-Nieto et al., 1989). In Anabaena sp. strain PCC 7120, as is also the case for Synechococcus sp. strain PCC 7942 (Luque et al., 1994b), nitrate or nitrite is not required for transcription of nitrate assimilation genes (Cai and Wolk, 1983; J. E. Frías, E. Flores and A. Herrero, unpublished results). In Synechococcus sp., the stability of Nar is higher in media containing either nitrate or ammonium than in media lacking combined N (Herrero et al., 1984). This could contribute to maintaining higher cellular levels of Nar in media containing nitrate than in media lacking combined N. In Anabaena sp. strain PCC 7120, the basis for the requirement of nitrate or nitrite to obtain high levels of Nar and Nir is unknown.

Enrique Flores and Antonia Herrero

3. Cotranscription of Some Genes Involved in Nitrate Assimilation As mentioned above, the genes encoding Nir (nir), the nitrate transport system (nrtABCD), and the putative structural gene for Nar (narB) map close together in the genome of Synechococcus sp. strain PCC 7942 (Luque et al., 1992; Omata et al., 1993). Mutant strains unable to use nitrate as an N source and exhibiting pleiotropic phenotypes with respect to nitrate assimilation have been isolated from Synechococcus sp. strain PCC 7942. Strain FM16, isolated after Tn901 mutagenesis, is unable to grow on nitrate but grows on nitrite, and exhibits low but appreciable Nar levels, high Nir levels, and nondetectable nitrate transport activity. The low Nar activity of this strain is, in contrast to what it is found in the wild-type strain, constitutive, i.e., not subjected to ammonium-promoted repression (Madueño et al., 1988a). Strain FM2 bears Tn901 inserted within the nir gene (Luque et al., 1992). As a consequence of this, it is unable to grow on either nitrate or nitrite and exhibits non-detectable Nir activity, but it constitutively expresses nitrate transport and Nar activities (Madueño et al., 1988a). Additionally, mutant strains (CSI1 through CSI6) generated by insertion of a gene cassette at defined positions inside the nir gene exhibited undetectable Nir activity, irrespective of the orientation of the inserted cassette, and only basal levels of Nar (Luque et al., 1992). These polar effects of insertional mutations in the cluster of nitrate assimilation genes indicate that nitrate assimilation genes in that cluster are cotranscribed (Luque et al., 1992). Northern experiments using probes of the different genes from the cluster are consistent with these genes being cotranscribed (Suzuki et al., 1993; Luque et al., 1994b). IV. Assimilation of Organic Nitrogen In a careful study carried out by Neilson and Larsson (1980) which included seven cyanobacterial strains, the only organic compounds found to serve as sole N-sources for some cyanobacteria were: urea, a few amino acids (glutamine, asparagine, arginine and ornithine), and urate. On the other hand, the unicellular cyanobacterium Synechococcus sp. strain PCC 7002 has been reported to utilize as N-sources

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503

a wide number of organic compounds, including urea, most amino acids, and purines (specially hypoxanthine and xanthine; Kapp et al., 1975). In this section, we shall summarize information available on the assimilation of urea and some amino acids. Unfortunately, the pathways of purine (hypoxanthine, xanthine, and urate) assimilation in cyanobacteria are unknown.

been reported (see e. g., Hall and Jensen, 1980; Chapman and Meeks, 1983; Rawson, 1985; Flores and Muro-Pastor, 1990), but only in the case of aromatic amino acids the metabolic basis of inhibition has been elucidated (see e. g., Hall and Jensen, 1980).

A. Assimilation of Urea

Arginine was shown to be taken up and concentrated within the cells in Synechocystis sp. strain PCC 6803 and Anabaena sp. strain PCC 7120 by means of transport systems specific for basic amino acids rather than for arginine only (Labarre et al., 1987; Flores and Muro-Pastor, 1990; Herrero and Flores, 1990). Synechocystis sp. strain PCC 6803 cells grown on arginine exhibit a very high affinity for arginine and, based on the behavior of mutants impaired in basic amino acid transport, the basic amino acid transport system of this cyanobacterium has been suggested to be of the binding proteindependent type (Flores and Muro-Pastor, 1990). Two kinetic components are observed for arginine transport in Anabaena sp. strain PCC 7120 cells, with values of and 0.75 mM, respectively. One of these components corresponds to a highaffinity system that is missing from canavanine- or hydroxylysine-resistant mutants and catalyzes a concentrative (active) transport, and the other one corresponds to a low-affinity system that is still present in those mutants and mediates a nonconcentrative (passive) uptake (Herrero and Flores, 1990). The involvement of these transport systems in arginine-dependent growth is deduced from the behavior of transport mutants. Thus, Synechocystis sp. strain PCC 6803 mutants severely impaired in arginine transport are unable to grow using arginine (5 mM) as the sole N-source (Flores and MuroPastor, 1990), and Anabaena sp. strain PCC 7120 mutants lacking the high-affinity system show a much reduced arginine-dependent growth on low (0.5 mM) but not on high (5 mM) concentrations of arginine (Herrero and Flores, 1990). Whereas Synechocystis sp. strain PCC 6803 grows on either arginine or nitrate at similar rates (Flores and MuroPastor, 1990), arginine-dependent growth of Anabaena sp. strain PCC 7120 (studied in a mutant unable to fix as well as of Pseudanabaena sp. strain PCC 6903 (Neilson and Larsson, 1980), is

Urea is a good N-source for many cyanobacteria belonging to different taxonomic groups (see e. g., Kratz and Myers, 1955; Neilson and Larsson, 1980; Rawson, 1985). Since urea concentrations in natural environments (e. g., in seawater) can be similar to those of nitrate and ammonium, it follows that urea can be an important N-source for cyanobacteria. Urea appears to be taken up intact and actively by the cells and then metabolized intracellularly (Healey, 1977). Intracellular urea is degraded in cyanobacteria by the enzyme urease releasing one molecule of and two molecules of ammonium (Berns et al., 1966; Carvajal et al., 1982; Mackerras and Smith, 1986). Urease has been shown to require the presence of in the growth medium (Mackerras and Smith, 1986; Singh, 1990) and has been purified from Spirulina maxima (Carvajal et al., 1982; see Table 1) and Anabaena cylindrica (Argall et al., 1992). While the uptake system for urea might be subject to repression by ammonium (Healey, 1977; Singh and Ahmad, 1989), conflicting reports on repression by ammonium of urease have appeared (Mackerras and Smith, 1986; Singh, 1990). Further studies on the regulation and genetics of urea assimilation in cyanobacteria would be worthwhile.

B. Assimilation of Amino Acids Arginine (Kapp et al., 1975; Neilson and Larsson, 1980; Rawson, 1985; Flores and Muro-Pastor, 1990; Herrero and Flores, 1990; see also Wolk, 1973) as well as glutamine and asparagine (Kapp et al., 1975; Neilson and Larsson, 1980; Rawson, 1985; Thiel and Leone, 1986) have frequently been reported to serve as sources of N for cyanobacteria, and some information is available on the assimilation of arginine and glutamine. On the other hand, inhibition of cyanobacterial growth by some amino acids has

1. Arginine

504 slower than nitrate-dependent growth (Herrero and Flores, 1990). Arginine transport might represent a rate-limiting step for growth on arginine in these latter strains. Two common arginine-degrading enzymes in bacteria are arginase (which produces urea and ornithine) and arginine deiminase (which produces citrulline and ammonium). These two enzymes have been reported in cyanobacteria (Weathers et al., 1978; Gupta and Carr, 1981; Martel et al., 1993). Studies on the fate of exogenously supplied in Synechocystis sp. strain PCC 6803 suggest the operation of the arginase pathway (E. Flores, unpublished) which would release, among its final products, ammonium and glutamate.

2. Glutamine In cells of Anabaena sp., a high-affinity transport system for glutamine is evident (Rowell et al., 1977; Chapman and Meeks, 1983; Flores and Muro-Pastor, 1988). A second, lowaffinity glutamine transport activity 1.1 mM) is observed in Anabaena variabilis strain ATCC 29413 (Chapman and Meeks, 1983). The high-affinity system seems not to be specific for glutamine but, rather, it appears to be a broad-specificity system able to transport neutral amino acids and also glutamate (Chapman and Meeks, 1983; Flores and Muro-Pastor, 1988). Two kinetic components are also found for glutamine transport in cells of Synechocystis sp. strain PCC 6803 0.01 and 5 mM, respectively), the low affinity activity being attributable to the basic amino acid transport system (Labarre et al., 1987; Flores and Muro-Pastor, 1990). Nitrogen fixation mutants of A. variabilis strain ATCC 29413 have been reported to grow using glutamine as an N-source and, interestingly, a mutant of this strain which showed an increased activity of glutamine transport grew better than its parental fixation minus) strain on glutamine (Thiel and Leone, 1986). It should be noted, however, that because glutamine hydrolysis can take place spontaneously in the culture medium releasing ammonium, data on growth on high concentrations of glutamine should be taken with caution (Chapman and Meeks, 1983). Metabolism of exogenously supplied glutamine to glutamate has been shown to take place in Anabaena sp. (Rowell et al., 1977; Thiel and Leone, 1986). Although glutaminase activity has been detected (Haystead et al., 1973; Chen et al., 1987),

Enrique Flores and Antonia Herrero GOGAT appears to be the main enzyme converting exogenously supplied glutamine to glutamate, since azaserine (an inhibitor of glutamine-amido transferases, see Section V B, 1) has been shown to inhibit close to 80% the generation of from exogenously supplied (Rowell et al., 1977).

V. Assimilation of Ammonium As mentioned above, the first step in the assimilation of N-sources like nitrate and urea is their metabolism to render intracellular ammonium. Ammonium can also be directly taken up from the outer medium and cyanobacteria are in general able to grow with ammonium as an N-source. In this section, we shall first address the question of how ammonium is taken up from the extracellular medium by cyanobacteria; we shall then describe the pathways that operate for the incorporation of ammonium into C skeletons in these organisms.

A. Ammonium Uptake Cellular suspensions of several cyanobacteria have been shown to efficiently take up low concentrations (<0.25 mM) of ammonium from the outer medium (Healey, 1977; Ohmori et al., 1977; Flores et al., 1980; Boussiba et al., 1984b; Boussiba, 1989). Methylammonium can be used as a probe to study the ammonium transport system. In Synechococcus sp. strain PCC 7942 and Anabaena variabilis strain ATCC 29413, time-course experiments show a rapid accumulation of within the cells followed by the formation, which proceeds at a slower rate, of a product of metabolism via GS (Boussiba et al., 1984a; Rai et al., 1984). A gradient of of up to 200 between the cells and the medium has been observed, and accumulation is hampered by a number of metabolic inhibitors including some ionophores (Boussiba et al., 1984a; Rai et al., 1984). Available data suggests that uptake is mediated by a permease which catalyzes an active transport that is dependent on the membrane potential. of the cells for methylammonium is about (Boussiba et al., 1984a), but it seems to be lower for ammonium which is likely the natural substrate for the methylammonium transport system. The ammonium/methylammonium transport

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system is repressed in ammonium-supplemented cultures (Boussiba et al., 1984a; Vega-Palas et al., 1990) and, under these conditions, passive diffusion ofunprotonated ammonia may represent a mechanism for net uptake of ammonium. Nonetheless, the ammonium/methylammonium transport system can obviouslyhave arole inammoniumuptake innatural environments where ammonium can be found at low concentrations (e.g., in seawater). A role for this transport system in recapture of ammonium leaked from cells growing on nitrate or can also be considered (Boussiba et al., 1984a; Rai et al., 1984).

B. Ammonium Assimilation Pathways In general, there are two major pathways by which ammonium can be incorporated into organic constituents of the cell: directly into glutamate by means of glutamate dehydrogenase (GDH) or by a cycle of reactions catalyzed by GS and GOGAT (Fig. 1). GDH catalyzes the reductive amination of 2-oxoglutarate to glutamate, GS catalyzes the ATPdependent ligation of glutamate and ammonium rendering glutamine, and GOGAT transfers the amido group of glutamine to 2-oxoglutarate rendering two glutamate molecules. The concerted action of GS and GOGAT yields one molecule of glutamate per molecule of ammonium assimilated.

1. The Glutamine Synthetase/Glutamate Synthase Cycle The notion that ammonium assimilation in cyanobacteria takes place primarily via the GS/GOGAT cycle originated from the observation that, whereas GDH activity is very low or even undetectable in some cyanobacteria (Hoare et al., 1967; Neilson and Doudoroff, 1973), high activity levels of GS could be found in Anabaena cylindrica (Dharmawardene et al., 1972). The GS inhibitor MSX was then observed to depress severely the assimilation of newly fixed ammonium in cultures of Anabaena cylindrica (Stewart and Rowell, 1975), and GOGAT was detected in two heterocyst-forming cyanobacteria (Lea and Miflin, 1975). However, it was the experimental approach set up by Wolk and co-workers which firmly established the operation of the GS/GOGAT cycle as the major pathway for the assimilation of ammonium in cyanobacteria. Their experiments revealed that, in Anabaena cylindrica, after fixation of (which first

505 yielded or assimilation of exogenously supplied the first labeled organic compound is glutamine followed by glutamate (Wolk et al., 1976; Meeks et al., 1977). Similar patterns of distribution of or in glutamine and glutamate were found in other fixing cyanobacteria (Meeks et al., 1978). Glutamine is first labeled in its amido-N which, as shown by chase experiments, is then transferred to 2oxoglutarate to form That 2oxoglutarate is aminated by glutamine (the reaction catalyzed by GOGAT) rather than directly by ammonium (the reaction catalyzed by GDH) was corroborated by studying the effect of azaserine, an inhibitor of amide transfer from glutamine, that largely prevented the formation of but not of MSX, which acts by inhibiting GS, hampered the formation of tamine and, as a consequence, largely also the formation of (Wolk et al., 1976; Meeks et al., 1977). A major role ofthe GS/GOGAT cycle in ammonium assimilation in Anabaena cylindrica was also suggested from a study of incorporation of N into metabolites (Lawrie et al., 1976) and from the effect ofazaserine on lowering the glutamate pool of the cells (Rowell et al., 1977). Conversely, the absence of an effect of azaserine on the glutamate pool was taken to suggest that an ammonium-assimilating pathway alternative to the GS/GOGAT cycle might be operative in Anabaena sp. strain 1F (Chen et al., 1987). In Synechococcus sp. strain L-1402-1 (Flores et al., 1983b), Synechococcus sp. strain PCC 7942 (Boussiba et al., 1984b) and Spirulina platensis (Boussiba, 1989), but not in Synechococcus sp. strain UTEX 625 (Meeks et al., 1978), a significant inhibition by MSX of ammonium assimilation was observed. Moreover, in Synechococcus sp. strain L1402-1 and Synechocystis sp. strain PCC 6803, the cellular levels of glutamate were observed to be significantly depressed by treatment of the cells with azaserine (Flores et al., 1983b; Mérida et al., 1991). Therefore, the GS/GOGAT cycle appears to be a major pathway for ammonium assimilation not only in but alsoin cyanobacteria. In enterobacteria, the GS/GOGAT pathway is operative only when the organism is growing on Nsources poorer than ammonium or glutamine, or on a limiting concentration of ammonium (Magasanik and Neidhardt, 1987). In cyanobacteria, however, the major pathway for ammonium assimilation

506 appears to be the GS/GOGAT cycle independent of the N-source used for growth (Wolk et al., 1976; Meeks et al., 1977; Flores et al., 1983b). In agreement with this, GOGAT specific activity is similar in cells grown on ammonium, nitrate or (Meeks et al., 1977; Florencio et al., 1987) and, generally, GS activity in ammonium cultures is only reduced to half of the activity found in or nitrate-grown cultures (see e.g., Meeks et al., 1977; Rowell et al., 1977; Stacey et al., 1977; Vega-Palas et al., 1990).

2. GS Enzyme and the glnA Gene GS can account for about 0.5–2% of the total cell protein. The enzyme has been purified to electrophoretic homogeneity from a number of cyanobacteria (Stacey et al., 1977; Sampaio et al., 1979; Orr et al., 1981; Florencio and Ramos, 1985; Blanco et al., 1989; Mérida et al., 1990). Some molecular and kinetic properties of GS are summarized in Table 1. Each GS molecule is composed of 12 identical subunits arranged in two superimposed hexagonal rings (Sampaio et al., 1979; Orr et al., 1981; Mérida et al., 1990), a structure similar to that described for the GS protein from other procaryotes (Yamashita et al., 1989) (Fig. 7). The enzyme exhibits a requirement for a divalent cation that in general is most efficiently met by but also supports appreciable activity (Blanco et al., 1989; Mérida et al., 1990). The glnA gene from Anabaena sp. strain PCC 7120 encoding GS was cloned by hybridization using the glnA gene from Escherichia coli as a probe (Fisher et al., 1981) and sequenced (Tumer et al., 1983). Other cyanobacterial glnA genes have been cloned either by hybridization using the Anabaena sp. strain PCC 7120 glnA gene as a probe (Mérida et al., 1992; Elmorjani et al., 1992) or by complementation of E. coli glutamine auxotrophs (Riccardi et al., 1985; Wagner et al., 1993). The Calothrix sp. strain PCC 7601 (Elmorjani et al., 1992), Synechocystis sp. strain PCC 6803 (J. C. Reyes and F. J. Florencio, personal communication), and Synechococcus sp. strain PCC 7002 (Wagner et al., 1993) glnA genes encode polypeptides which are 89, 79 and 75% identical, respectively, to the Anabaena sp. strain PCC 7120 GS polypeptide. As expected from the structure of the GS protein, the polypeptide deduced from thecyanobacterialglnA geneisstrongly similar to other procaryotic (not only eubacterial) GS polypeptides and shows conserved regions which

Enrique Flores and Antonia Herrero are found in both procaryotic and eucaryotic GS polypeptides (see e. g., Elmorjani et al., 1992). In general, the cyanobacterial glnA gene is expressed in E. coli and complements E. coli glutamine auxotrophs. Expression in E. coli of the Anabaena sp. strain PCC 7120 glnA gene takes place from an ‘E. coli-like’ promoter which reads as follows: TTGTGC (–35 box) and TAATAT (–10 box; Tumer et al., 1983). In Anabaena, Calothrix and Synechocystis sp., an identical putative ribosome binding site (AGGAG) that can be operative in E. coli is located five nucleotides upstream from the glnA ATG start codon. In Anabaena, Calothrix and Synechococcus sp. a monocistronic mRNA is detected with a glnA probe. In Anabaena sp. strain PCC 7120 transcription of glnA from the ‘E. coli-like’ promoter (as well as from three other promoters) takes place in ammonium-grown cells (Tumer et al., 1983). More glnA mRNA is observed after N starvation, consistent with the increased GS activity found in as compared to ammonium-grown cultures. Under N starvation, promoter is activated, whereas no transcription takes place from the ‘E. coli-like’ promoter (Tumer et al., 1983). Which promoters are used in cells steadily growing on or nitrate is unknown. In Calothrix sp. strain PCC 7601 and Synechococcus sp. strain PCC 7002, transcription initiation of glnA may also occur from several different promoters whose expression is modulated by the N source (Elmorjani et al., 1992; Wagner et al., 1993). In Synechococcus sp. strain PCC 7942, transcription takes place from a single promoter that is severly repressed by ammonium (Cohen-Kupiec et al., 1993; Luque et al., 1994b). Mutants of Anabaena variabilis with an altered GS, but not completely devoid of GS activity, have been isolated after selection for resistance to MSX (see e.g., Chapman and Meeks, 1983; Spiller et al., 1986) or to ethylenediamine (Polukhina et al., 1982; see also Hien et al., 1988). A mutant of Synechococcus sp. strain PCC 7002 carrying a deletion of the glnA gene has been described which is still able to synthesize glutamine, suggesting that another GS would be present in this cyanobacterium (Wagner et al., 1993). The Synechocystis sp. strain PCC 6803 glnA gene has been insertionally inactivated in a recombinant strain carrying the Anabaena sp. strain PCC 7120 glnA gene in the Synechocystis sp. genome (Mérida et al., 1992) and, more recently, in the wildtype strain (Reyes and Florencio, 1994). In the latter

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507

Enrique Floras and Antonia Herrero

508 case, the resulting strain lacking a functional glnA gene product is not a glutamine auxotroph, and a second GS, encoded by the glnN gene, has been identified in this cyanobacterium (Reyes and Florencio, 1994). In enterobacteria, the GS protein is rapidly inactivated by adenylylation upon addition of ammonium to cultures containing a poor supply ofN (Magasanik and Neidhardt, 1987), but no evidence for regulation by adenylylation ofthe cyanobacterial enzyme has been found (Rowell et al., 1977; Stacey et al., 1979; Fisher et al., 1981). On the other hand, all the purified cyanobacterial GS proteins are susceptible to inhibition by some amino acids and nucleotides (Stacey et al., 1979; Orr and Haselkorn, 1981; Florencio and Ramos, 1985; Blanco et al., 1989; Mérida et al., 1990). Cumulative inhibition has been detected with a mixture of aspartate, serine and AMP, indicating independent binding sites for these three compounds (Stacey et al., 1979; however, see Blanco et al., 1989), while alanine and glycine share a binding site with serine (Orr and Haselkorn, 1981; see also Stacey et al., 1979). Feedback inhibition by aspartate and alanine (and serine and glycine), and by ADP and AMP, can be considered a feasible in vivo regulatory mechanism for cyanobacterial GS. The GS protein found in ammonium-grown cultures of Synechocystis sp. strain PCC 6803 is largely inactive (Merida et al., 1990), and the addition of ammonium to nitrate-grown cultures of this cyanobacterium provokes a short-term inactivation of GS (90–95% inactivation in 25 min, see Mérida et al., 1991). Interestingly, inactivation ofthe GS from Anabaena sp. strain PCC 7120 in response to ammonium has been observed in a Synechocystis sp. strain PCC 6803 recombinant strain bearing a mutated version ofits own glnA gene and carrying integrated into its genome the glnA gene from Anabaena sp. strain PCC 7120 (Mérida et al., 1992). Synechocystis sp. strain PCC 6803 appears to have a regulatory system for GS which is absent, or much less active, in other commonly studied cyanobacterial strains. Transfer of light-grown cells to darkness also results in a reversible inactivation of GS in some cyanobacteria (Rowell et al., 1979; Marqués et al., 1992b). Whether the ammonium- and dark-promoted inactivations of GS are mediated by the same mechanism, and whether the protein is involved in this mechanism, are open questions. is the product of the glnB gene which in the enterobacteria

is involved in the regulation of GS (Magasanik and Neidhardt, 1987). The glnB gene has recently been identified in Synechococcus sp. and the degree of phosphorylation of the protein has been found to respond to the light regime and N nutrition of the cells (Tsinoremas et al., 1991; Forchhammer and Tandeau de Marsac, 1994).

3. GOGAT Cyanobacterial GOGAT uses reduced Fd rather than pyridine nucleotides as an electron donor, thus being more similar to that of the higher plant chloroplasts than to the enterobacterial enzyme (Lea and Miflin, 1975). Only one cyanobacterial GOGAT has been purified to electrophoretic homogeneity—that from Synechococcus sp. strain PCC 6301 (Marqués et al., 1992a; see Table 1). It is a flavoprotein containing one molecule of FMN and consisting of only one polypeptide that may also contain non-heme iron. Feedback inhibition by the reaction product, glutamate, might be a regulatory mechanism for the enzyme.

4. Other Ammonium Assimilating Enzymes Nitrate-grown cells of Synechococcus sp. strain PCC 6301 treated with MSX or azaserine, which cannot assimilate ammonium via GS/GOGAT, and fed with nitrate release to the outer medium an amount of ammonium accounting for 85–90% of the nitrate-N taken up (Flores et al., 1980, 1983a). A similar inhibition (86–90%) of assimilation of is provoked by MSX in several filamentous cyanobacteria (Meeks et al., 1978). Because under the conditions used in those experiments MSX completely abolishes GS activity (Flores et al., 1983b), the low amount of ammonium still assimilated in the presence of MSX probably reflects the operation of alternative, minor pathways able to incorporate to some extent the ammonium accumulated in the MSXtreated cells (Wolk et al., 1976). is produced from in some cyanobacteria under conditions of inactivation of the GS/GOGAT pathway (Wolk et al., 1976; Meeks et al., 1977, 1978). This indicates direct amination of pyruvate catalyzed by alanine dehydrogenase (ADH) which has also been suggested from the rapid labeling of alanine observed on adding N to Anabaena cylindrica cells (Lawrie et al., 1976). ADH has actually been detected in many, but not all,

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of the cyanobacteria tested (Haystead et al., 1973; Neilson and Doudoroff, 1973; Batt and Brown, 1974; Flores et al., 1983b). The enzyme has been purified onlyfromAnabaenacylindrica (Rowell and Stewart, 1976) and some of its properties are summarized in Table 1. It is specific and its activity in the aminating reaction is significantly higher than activity in the deaminating reaction thus supporting its role in ammonium assimilation. It is of interest that alanine, glycine and serine markedly inhibit the aminating activity of the enzyme. In Anabaena cylindrica, is also formed from though to a limited extent, under conditions of inactivation of the GS/GOGAT pathway (Meeks et al., 1977) and, in Synechococcus sp. strain UTEX 625, labeling ofglutamate is not inhibited by inactivation of GS with MSX (Meeks et al., 1978). These observations suggest that, at least in some cyanobacteria, a direct amination of 2oxoglutarate catalyzed by GDH can take place. Although no GDH activity is detected in Synechococcus sp. strain PCC 6301 (Hoare et al., 1967; Flores et al., 1983b) or in some other strains (Neilson and Doudoroff, 1973), and GDH activities have been detected in some cyanobacteria, though generally at very low levels (Haystead et al., 1973; Neilson and Doudoroff, 1973; Batt and Brown, 1974; Florencio et al., 1987; Martínez-Bilbao et al., 1987; Chávez and Candau, 1991). has been purified from Synechocystis sp. strain PCC 6803 (Florencio et al., 1987) and Phormidium laminosum (Martínez-Bilbao et al., 1988), and from Synechocystis sp. strain PCC 6803 (Chávez and Candau, 1992). Some properties ofboth enzymes are summarized in Table 1. The two enzymes catalyze the amination of 2-oxoglutarate preferentially over the reverse deamin–ating reaction, which is consistent with them having a role in ammonium assimilation. The gdhA gene encoding hasrecentlybeencloned from Synechocystis sp. strain PCC 6803 by complementation of an E. coli glutamate auxotroph (S. Chávez and P. Candau, personal communication). Sequence analysis has shown that the gdhA -encoded polypeptide is homologous to GDH from other organisms. Synechocystis sp. strain PCC 6803 gdhA insertional mutants lacking any activity have been generated that are unaffected in their growth characteristics under standard conditions (S. Chávez and P. Candau, personal communication).

509 VI. Distribution of Assimilated Nitrogen Glutamate is a precursor for the biosynthesis of glutamine, arginine, proline, and 5-aminolevulinate. Indeed, proline and arginine are labeled after feeding Synechococcus sp. strain PCC 6301 with tamate (Smith et al., 1967) and arginine and citrulline (an intermediate in arginine biosynthesis) are labeled in Anabaena cylindrica shortly after assimilation of (Meeks et al., 1977). Arginine biosynthesis has been reported to take place in cyanobacteria via the so-called ‘cyclic pathway of ornithine synthesis’ in which the acetyl group of acetylornithine is transferred to glutamate to form N-acetylglutamate and ornithine in a reaction catalyzed by ornithine acetyltransferase (Hoare and Hoare, 1966). In bacteria, ornithine acetyltransferase is the product of the argJ gene whereas acetylornithinase, an alternative enzyme producing ornithine from acetylornithine, is encoded by argE. Although a DNA fragment able to complement an E. coli argE mutant has been isolated from Synechococcus sp. strain PCC 7002 (Porter et al., 1986), complementation might have been accomplished by an argJ gene product. Three enzymes catalyze the conversion of N-acetylglutamate into acetylornithine, and the genes encoding two of these enzymes have been cloned from Anabaena sp. strain PCC 7120 by complementation of Anabaena sp. arginine auxotrophs and sequenced. They are argC, encoding Nacetylglutamate semialdehyde dehydrogenase, and argD, encoding acetylornithine 5-aminotransferase (Floriano et al., 1992; and unpublished). Ornithine is converted into arginine by means of three enzymatic steps, the first of which, rendering citrulline, is catalyzed by ornithine transcarbamylase, an enzyme that shows a notably high activity in cyanobacteria (Holm-Hansen and Brown, 1963; Gupta and Carr, 1981; Chen et al., 1987). On the other hand, whereas the proline biosynthesis pathway has not been specifically studied in cyanobacteria, much work has recently been carried out on 5-aminolevulinate biosynthesis which is covered in Chapter 17 of this book. In addition to being a precursor of some metabolites, glutamate is considered the major source of N for cellular metabolism. Transamination reactions, by means of which glutamate can donate its amino group to a number of oxo acids, have been reported in cyanobacteria. Particularly high activity of glutamate-aspartate aminotransferase is detected

510 in Anabaena cylindrica (Batt and Brown, 1974; Rowell et al., 1977) that can account for the formation of after incorporation of into glutamine and glutamate (Meeks et al., 1977). Transaminases that should be able to transfer the amino group from glutamate to the 2-oxo acids of branched-chain and aromatic amino acids have been detected (Hoare et al., 1967; Romero et al., 1985). Additionally,aspartate-alanineaminotransferasehas also been reported in Anabaena cylindrica (Batt and Brown, 1974; Rowell and Stewart, 1976) and the possibility of transamination of the group of glutamine to oxaloacetate to form aspartate has also been considered (Meeks et al., 1977; Thomas et al., 1977). The amido-N of glutamine is also incorporated into a number ofmetabolites including some amino acids. Carbamyl phosphate synthetase (Lawrie, 1979; Chen et al., 1987) and anthranilate synthetase (Bottomley et al., 1980) have been shown to be able to use glutamine as an N-donor in Anabaena sp. Carbamyl phosphate is used in reactions catalyzed by ornithine transcarbamylase (rendering citrulline, see above) and aspartate transcarbamylase (the first enzyme of the uridylic acid biosynthesis pathway which has been shown to be operative in Anabaena variabilis; see Currier and Wolk, 1978), and anthranilate synthetase is the first key enzyme in tryptophan biosynthesis. Arginine and aspartate together make up cyanophycin or multi-L-arginyl-poly(L-aspartic acid), a reservoir of N found in many, but not all, cyanobacteria. For a review on cyanophycin biology the reader is referred to Simon (1987). Although the fate of the arginine and aspartate released after cyanophycin mobilization has not been investigated, the arginase pathway (see Section IV B,1) and transaminases able to use aspartate as a substrate (see above) might be involved in their further metabolism.

VII. Global Nitrogen Control The presence ofammonium in cyanobacterial culture media brings about repression of expression of proteins involved in the assimilation ofalternative N sources (like or nitrate) or of ammonium itself (see Sections II, III, and V). Ammonium metabolism through OS appears to be necessary for all those repressive effects to be manifest suggesting that a common mechanism might be involved in regulation

Enrique Flores and Antonia Herrero by ammonium ofproteins ofdifferent N assimilation pathways. The isolation from Synechococcus sp. strain PCC 7942 of mutants simultaneously impaired in the expression of all the proteins known to be subject to regulation by ammonium corroborated the existence of global N-regulatory gene(s) in a cyanobacterium (Vega-Palas et al., 1990). The gene ntcA, which was altered in one of those pleiotropic mutants and which would encode a positive-acting element, was cloned by complementation (VegaPalas et al., 1990). Sequencing of this gene has shown that it encodes a protein homologous to members of a family of bacterial transcriptional regulators ofwhich Crp (the cAMP-receptor protein) of E. coli is the best-known example (Vega-Palas et al., 1992). The ntcA gene is widespread in cyanobacteria and has also been cloned and sequenced from Synechocystis sp. strain PCC 6803 and Anabaena sp. strain PCC 7120 (Frías et al., 1993). As is the case for Crp and all the proteins in the family, NtcA is predicted to have a helix-turn-helix motif for binding to DNA near its carboxyl terminus. Amino acid sequences in the helix-turn-helix motif are identical in the three characterized NtcA proteins. Binding of Synechococcus sp. strain PCC 7942 NtcA to the promoter regions of Synechococcus sp. glnA and nir genes, as well as to that of the ntcA gene itself, has been recently observed in vitro, and the DNA target sequence for NtcA has been determined and found to include the motif which is located 22 nucleotides upstream from the putative –10 promoterbox of those genes (Luque et al., 1994b). Therefore, the promoter structure of NtcA-, N-regulated genes in Synechococcus sp. strain PCC 7942 is: of transcription. This structure is also found in some Nregulated genes of Anabaena sp. strain PCC 7120: e.g., in a promoter located upstream of nir (J. E. Frías, A. Herrero and E. Flores, unpublished results) and in promoter of glnA (see Tumer et al., 1983). An ntcA insertional mutant of Anabaena sp. strain PCC 7120 has recently been isolated and found to be unable to grow on nitrate or and to develop heterocysts (J. E. Frías, A. Herrero and E. Flores, unpublished results). Thus NtcA appears to be involved in regulation of fixation and heterocyst development. The bifA gene of Anabaena sp. strain PCC 7120, which encodes a DNA-binding protein that interacts with the promoters of the xisA, glnA, rbcL, and nifH promoters, is identical to the ntcA gene. Some DNA-binding sites for BifA have been

Chapter 16

Nitrogen Metabolism

determined, and the proposed consensus BifA binding site is similar to the NtcA binding site described above (Ramasubramanian et al., 1994). In enterobacteria, N-regulation at the transcriptional level is mediated by the two-component ntr system with the (GlnB) protein having a role in signal transduction from the N-status of the cell to the Ntr proteins (Magasanik and Neidhardt, 1987). The protein has been found in cyanobacteria (see Section V B, 2), but its precise role in signal transduction in these organisms is not yet known. NtcA is a novel regulator for the operation of Ncontrol in procarytotes, distinct from the widespread NtrB/NtrC system, that directly influences the levels of transcription of some genes involved in N assimilation. In turn, NtcA should itself be regulated in response to the N-regime of the cell, by modulation of its activity and/or its expression through mechanisms that are yet to be established.

Acknowledgments We thank P. Candau, F. J. Florencio, T. Omata, M. L. Peleato, T. Thiel, and Y. Cai for communicating results prior to publication. We would also like to thank Drs David Eisenberg (University of California, Los Angeles) and Douglas Rees (Cal Tech University) for contributing figures for this chapter. Work in the authors’ laboratory is currently supported by grants from DGICYT (grant no. PB90-0114), CICYT (grant no. BI093-0124) and Junta de Andalucía (research group no. 3057), Spain.

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516 Rowell P, Enticott S and Stewart WDP (1977) Glutamine synthetase and nitrogenase activity in the blue-green alga Anabaena cylindrica. New Phytol 79: 41–54 Rowell P, Sampaio MJAM, Ladha JK and Stewart WDP (1979) Alteration of cyanobacterial glutamine synthetase activity in vivo in response to light and Arch Microbiol 120: 195– 200 Sampaio MJAM, Rowell P and Stewart WDP (1979) Purification and some properties of glutamine synthetase from the nitrogenfixing cyanobacteria Anabaena cylindrica and a Nostoc sp. J Gen Microbiol 111: 181–191 Saville B, Straus N and Coleman JR (1987) Contiguous organization of nitrogenase genes in a heterocystous cyanobacterium. Plant Physiol 85: 26–29. Schmitz O, Kentemich T, Zimmer W, Hindeshagen B and Bothe H (1993) Identification of the nifJ gene coding for pyruvate:ferredoxin oxidoreductase in dinitrogen-fixing cyanobacteria. Arch Microbiol 160: 62–67 Schrautemeier B and Böhme H (1985) A distinct ferredoxin for nitrogen fixation isolated from heterocysts of the cyanobacterium Anabaena variabilis. FEBS Lett 184: 304–308 Schrautemeier B and Böhme H (1992) Coding sequence of a heterocyst ferredoxin gene (fdxH) isolated from the nitrogenfixing cyanobacterium Calothrix sp. PCC 7601. Plant Mol Biol 18: 1005–1006 Schrautemeier B Cassing A and Böhme H (1994) Characterization of the genome region encoding an FdxH-type ferredoxin and a new 2[4Fe-4S] ferredoxin from the nonheterocystous, nitrogenfixing cyanobacterium Plectonema boryanum PCC 73110. J Bacteriol 176: 1037–1046 Serrano A, Rivas J and Losada M (1981) Nitrate and nitrite as ‘ in vivo’ quenchers of chlorophyll fluorescence in blue-green algae. Photosynthesis Res 2: 175–184 Shah VK, Hoover TR, Imperial J, Paustian TD, Roberts GP and Ludden PW (1988) Role of nif gene products and homocitrate in the biosynthesis of iron-molybdenum cofactor. In: Bothe H, de Bruijn FJ and Newton WE (eds) Nitrogen Fixation: Hundred Years After, pp 115–120. Gustav Fisher, Stuttgart Simon RD (1987) Inclusion bodies in the cyanobacteria: Cyanophycin, polyphosphate, polyhedral bodies. In: Fay P and Van Baalen C (eds) The Cyanobacteria, pp 199–225. Elsevier, Amsterdam Singh RK and Stevens SE Jr (1992) Cloning of the nifHDK genes and their organisation in the heterocystous cyanobacterium Mastigocladus laminosus. FEMS Microbiol Lett 94: 227–234 Singh RK, Stevens SE Jr and Bryant DA (1987) Molecular cloning and physical mapping of the nitrogenase structural genes from the filamentous, non-heterocystous cyanobacterium Pseudanabaena PCC 7409. FEMS Microbiol Lett 48: 53–58 Singh S (1990) Regulation of urease activity in the cyanobacterium Anabaena doliolum. FEMS Microbiol Lett 67: 79–84 Singh S and Ahmad S (1989) Regulation of urea uptake by ammonia in the cyanobacterium Anabaena doliolum. FEMS Microbiol Lett 61: 199–202 Sivak MN, Lara C, Romero JM, Rodríguez R and Guerrero MG (1989) Relationship between a 47 kDa cytoplasmic membrane polypeptide and nitrate transport in Anacystis nidulans. Biochem Biophys Res Commun 158: 257–262 Smith AJ, London J and Stanier RY (1967) Biochemical basis of obligate autotrophy in blue-green algae and thiobacilli. J Bacteriol 94: 972–983

Enrique Flores and Antonia Herrero Smith RV, Noy RJ and Evans MCW (1971) Physiological electron donor systems to the nitrogenase of the blue-green alga Anabaena cylindrica. Biochim Biophys Acta 253: 104–109 Spiller H and Shanmugam KT (1987) Physiological conditions for nitrogen fixation in a unicellular cyanobacterium, Synechococcus sp. strain SF1. J Bacteriol 169: 5379–5384 Spiller H, Latorre C, Hassan ME and Shanmugam KT (1986) Isolation and characterization of nitrogenase-derepressed mutant strains of cyanobacterium Anabaena variabilis. J Bacteriol 165: 412–419 Stacey G, Tabita FR and Van Baalen C (1977) Nitrogen and ammonia assimilation in the cyanobacteria: Purification of glutamine synthetase from Anabaena sp. strain CA. J Bacteriol 132: 596–603 Stacey G, Van Baalen C and Tabita FR (1979) Nitrogen and ammonia assimilation in the cyanobacteria: Regulation of glutamine synthetase. Arch Biochem Biophys 194: 457–467 Stal LJ and Krumbein WE (1985) Nitrogenase activity in the non-heterocystous cyanobacterium Oscillatoria sp. grown under alternating light-dark cycles. Arch Microbiol 143: 67–71 Stevens SE and Van Baalen C (1970) Growth characteristics of selected mutants of a coccoid blue-green alga. Arch Microbiol 72: 1–8 Stevens SE and Van Baalen C (1974) Control of nitrate reductase in a blue–green alga. The effects of inhibitors, blue light, and ammonia. Arch Biochem Biophys 161: 146–152 Stewart WDP and Lex M (1970) Nitrogenase activity in the bluegreen alga Plectonema boryanum strain 594. Arch Microbiol 73: 250–260 Stewart WDP and Rowell P (1975) Effects of L-methionine-DLsulphoximine on the assimilation of newly fixed acetylene reduction and heterocyst production in Anabaena cylindrica. Biochem Biophys Res Commun 65: 846–856 Strieker O, Almon H, Monnerjahn U and Böhme H (1992) Identification and characterization of nifV in Anabaena sp. PCC 7120. Second European Workshop on the Molecular Biology of Cyanobacteria, Bristol, Abstracts p 57 Suzuki I, Sugiyama T and Omata T (1993) Primary structure and transcriptional regulation of the gene for nitrite reductase from the cyanobacterium Synechococcus PCC 7942. Plant Cell Physiol 34: 1311–1320 Thiel T (1993) Characterization of genes for an alternative nitrogenase in the cyanobacterium Anabaena variabilis. J Bacteriol 175: 6276–6286 Thiel T and Leone M (1986) Effect of glutamine on growth and heterocyst differentiation in the cyanobacterium Anabaena variabilis. J Bacteriol 168: 769–774 Thiel T, Erker J and Lyons E (1993) Characterization of alternative nitrogen fixation systems in Anabaena variabilis. Fourth Cyanobacterial Workshop in Molecular Genetics, 1993, Asilomar (Pacific Grove, CA), Abstracts p 92 Thomas J, Meeks JC, Wolk CP, Shaffer PW, Austin SM and Chien WS (1977) Formation of glutamine from and by heterocysts isolated from Anabaena cylindrica. J Bacteriol 129: 1545–1555 Tischner R and Schmidt A (1984) Light mediated regulation of nitrate assimilation in Synechococcus leopoliensis. Arch Microbiol 137: 151–154 Tsai LB and Mortenson LE (1978) Interaction of the nitrogenase components of Anabaena cylindrica with those of Clostridium pasteurianum. Biochem Biophys Res Commun 81: 280–287

Chapter 16

Nitrogen Metabolism

Tsinoremas NF, Castets AM, Harrison MA, Allen JF and Tandeau de Marsac N (1991) Photosynthetic electron transport controls nitrogen assimilation in cyanobacteria by means of posttranslational modification of the glnB gene product. Proc Natl Acad Sci USA 88: 4565–4569 Tumer NE, Robinson SJ and Haselkorn R (1983) Different promoters for the Anabaena glutamine synthetase gene during growth using molecular or fixed nitrogen. Nature 306: 337–342 Vega-Palas MA, Madueño F, Herrero A and Flores E (1990) Identification and cloning of a regulatory gene for nitrogen assimilation in the cyanobacterium Synechococcus sp. strain PCC 7942. J Bacteriol 172: 643–647 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 Wagner SJ, Thomas SP, Kaufman RI, Nixon BT and Stevens SE (1993) The glnA gene of the cyanobacterium Agmenellum quadruplicatum PR-6 is nonessential for ammonium assimilation. J Bacteriol 175: 604–612 Weathers PJ, Chee HL and Allen MM (1978) Arginine catabolism in Aphanocapsa 6308. Arch Microbiol 118: 1–6 Wei T-F, Ramasubramanian TS, Pu F and Golden JW (1993)

517 Anabaena sp. strain PCC 7120 bifA gene encoding a sequencespecific DNA binding protein cloned by in vivo transcriptional interference selection. J Bacteriol 175: 4025–4035 Weisshaar H and Böger P (1983) Nitrogenase activity of the nonheterocystous cyanobacterium Phormidium foveolarum. Arch Microbiol 136: 270–274 Wolk CP (1973) Physiology and cytological chemistry of bluegreen algae. Bacteriol Rev 37: 32–101 Wolk CP, Thomas J, Shaffer PW, Austin SM and Galonsky A (1976) Pathway of nitrogen metabolism after fixation of labeled nitrogen gas by the cyanobacterium, Anabaena cylindrica. J Biol Chem 251: 5027–5034 Wyatt JT and Sil vey JKG (1969) Nitrogen fixation by Gloeocapsa. Science 165: 908–909 Yabuki Y, Mori E and Tamura G (1985) Nitrite reductase in the cyanobacterium Spirulina platensis, Agric Biol Chem 49: 3061–3062 Yamashita MM, Almassy RJ, Janson CA, Casio D and Eisenberg D (1989) Refined atomic model of glutamine synthetase at 3.5 Å resolution. J Biol Chem 264: 17681–17690 Zehr JP, Ohki K and Fuj ita Y (1991) Arrangement of nitrogenase structural genes in an aerobic filamentous nonheterocystous cyanobacterium. J Bacteriol 173: 7055–7058

Chapter 17 Biosynthesis of Cyanobacterial Tetrapyrrole Pigments: Hemes, Chlorophylls, and Phycobilins Samuel I. Beale Division of Biology and Medicine, Brown University, Providence, Rl 02912, USA Summary I. Introduction II. Tetrapyrrole Precursor Biosynthesis A. ALA Formation 1. In Vivo Evidence for ALA and Tetrapyrrole Formation from Glutamate 2. Mechanism of ALA Formation from Glutamate a. tRNAGlu b. Glutamyl-tRNA Synthetase c. Glutamyl-tRNA Reductase d. GSA e. GSA Aminotransferase B. PBG Synthesis III. The Pathway from PBG to Uroporphyrinogen III A. Hydroxymethylbilane Synthase B. Uroporphyrinogen III Synthase IV. Steps Leading to Siroheme and Corrins V. Conversion of Uroporphyrinogen III to Protoporphyrin IX A. Uroporphyrinogen Decarboxylase B. Coproporphyrinogen Oxidase C. Protoporphyrinogen Oxidase VI. The Fe Branch A. Ferrochelatase B. Synthesis of Other Hemes C. Phycobilin Formation 1. Ferredoxin-Linked Heme Oxygenase 2. Conversion of Biliverdin to Phycobilins a. Phycobilin Ethylidine Isomerization b. Ferredoxin-Linked Biliverdin IXa Reduction c. Phycoerythrobilin Isomerization d. 15,16-Dihydrobiliverdin IXa 3. Protein Ligation VII. The Mg Branch A. Mg Chelatase B. Mg-Protoporphyrin Methyltransferase C. Isocyclic Ring Formation D. Vinyl Reduction E. Ring D Reduction 1. The Light-Dependent Process 2. The Light-Independent Process F. Phytylation G. Reaction Center Chlorophylls H. Chlorophyll b Formation

D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 519–558. © 1993 Kluwer Academic Publishers. Printed in The Netherlands.

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Summary Cyanobacteria are versatile tetrapyrrole synthesizers that are able to produce end products representing all major branches of the tetrapyrrole biosynthetic pathway: hemes, chlorophylls, phycobilins, and siroheme. Although tetrapyrrole biosynthesis has not been characterized as extensively in cyanobacteria as in plants and anoxygenic photosynthetic bacteria, recent studies ofthe biochemistry and molecular genetics ofthis pathway have begun to exploit the advantages of these oxygenic procaryotic organisms. The results of these studies are increasing our understanding of the biosynthetic enzyme reaction mechanisms, the physical properties of the enzymes, modes of metabolic regulation, the evolution of the pathway, and the phylogenetic relationships among cyanobacteria, other bacteria, algae, and plants. In this article, emphasis is placed on the individual enzymatic steps of tetrapyrrole biosynthesis in cyanobacteria, the natures of substrates, reaction intermediates, and products, the physical, kinetic, and regulatory properties of the enzymes, and the identification of genes that encode the enzymes. Because ofthe limited amount ofavailable information that has been directly derived from cyanobacteria, results obtained from other organisms is discussed wherever it is likely to be applicable to cyanobacteria. I. Introduction Tetrapyrroles are nearly ubiquitous biomolecules that have many essential biological functions including electron transfer, oxygen binding, and light absorption. Biologically important tetrapyrroles include hemes, chlorophylls, bilins, and corrins. Tetrapyrroles may be classified on the bases of the presence and identity of a chelated metal, positions and identities of pyrrole ring substituents, and oxidation state of the porphyrin nucleus. The biological tetrapyrroles may be arranged as branches of a single biosynthetic pathway (Fig. 1). The existence of the branched biosynthetic pathway indicates the need for a complex regulatory system to insure that the end products are synthesized in appropriate proportions under changing environmental and developmental conditions. Cyanobacteria are versatile tetrapyrrole synthesizers: along with the ability to produce hemes and chlorophylls, a property which they share with plants, the cyanobacteria (as well as rhodophytes and cryptophytes) also synthesize a variety of phycobilins. These linear tetrapyrroles, when covalently Abbreviations: ALA – acid; EGTA – ethylene ether) N, N, N’, N’tetraacetic acid; gabaculine – 3-amino-2,3-dihydrobenzoic acid; GSA – glutamate-1-semialdehyde; GSH – glutathione; PBG – porphobilinogen

linked to specific phycobiliproteins, function as lightharvesting pigments (see chapter 5). In addition to these major tetrapyrrole end products, cyanobacteria are probably capable of forming siroheme, the prosthetic group of assimilatory sulfite and nitrate reductases, as is indicated by their ability to use

Chapter 17 Biosynthesis of Hemes, Chlorophylls and Phycobilins sulfate and nitrate as nutrients. Cyanobacteria may also be able to synthesize the corrin nucleus of vitamin but there is no evidence on this point. Despite the importance of cyanobacteria and their versatility as tetrapyrrole producers, they have not been extensively used as model systems in biosynthetic studies. In this review, attention will be focused, wherever possible, on results that were obtained with cyanobacteria. However, much information about tetrapyrrole biosynthesis in cyanobacteria must be inferred from results obtained with other organisms. Cyanobacteria, like other phototrophic organisms, are able to regulate their tetrapyrrole content and composition, especially those which are components of their photosynthetic apparatus, in response to environmental signals such as light intensity and spectral composition, nutritional status, and developmental programs. These topics are covered in detail in Chapters 21 through 28. In the present chapter, discussion of regulation will be restricted to the regulatory properties of individual biosynthetic enzymes. Progress in the field of tetrapyrrole biosynthesis has accelerated enormously in the past few years as a result of the identification and characterization of genes from several procaryotic and eucaryotic sources that encode nearly all of the enzymes involved in heme and chlorophyll formation. Molecular genetic approaches have facilitated the prediction of enzyme properties from their encodinggene sequences; studies of the regulation of enzyme synthesis at the transcriptional level; production of sufficient quantities of enzymes for physical studies by overexpression of their encoding genes; and elucidation of phylogenetic relationships by sequence comparison. Although these approaches have not yet been widely applied to the cyanobacteria, the results obtained from other organisms are neverthelesshighly relevant to the cyanobacteria and will be described wherever possible. Table 1 provides a list of the of tetrapyrrole biosynthetic enzymes that are present or are likely to be present in cyanobacteria. EC numbers are listed where they have been assigned, and genes that encode the enzyme polypeptides are also listed for reference. Although it must be emphasized that only a minority of these enzymes and genes have been detected and characterized in cyanobacteria, the overall high degree of conservation of tetrapyrrole biosynthesis indicates that results with other organisms are highly predictive for cyanobacteria.

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II. Tetrapyrrole Precursor Biosynthesis

A. ALA Formation ALA can be considered to be the first universal, committed tetrapyrrole precursor. However, two different biosynthetic routes to ALA exist. Eucaryotes that do not contain plastids (e.g., animals, yeasts, fungi) and members of the subgroup of purple bacteria (which includes the genera Rhodobacter, Rhodospirillum, Agrobacterium, Rhizobium, and Bradyrhizobium), form ALA by the action of ALA synthase, which catalyzes the condensation of glycine with succinyl-coenzyme A. Plants, algae, and most groups of bacteria, including cyanobacteria, form ALA by a different route (Fig. 2), which begins with activation of glutamate by ligation to to form glutamyl-tRNA. Next, the activated glutamate is reduced at C-1 to form GSA. Finally, the amino group at C-2 of GSA is replaced by one at C-1, yielding ALA. The only known role of GSA is as a tetrapyrrole precursor, and therefore GSA formation can be considered to be the first committed step of the tetrapyrrole pathway in cyanobacteria. GSA supplies all of the C and N atoms of the tetrapyrrole nucleus. The succinate-glycine route of ALA formation, which does not appear to occur in cyanobacteria, will not be considered here, and readers are directed to recent review articles for further information about this reaction (Jordan, 1990; 1991).

1. In vivo Evidence for ALA and Tetrapyrrole Formation from Glutamate Until the early 1970’s, the only known route of ALA formation was via condensation of succinyl-CoA and glycine, catalyzed by ALA synthase, an enzyme that was characterized from animal mitochondria and Rhodobacter sphaeroides. The first evidence indicating an alternative biosynthetic route was the preferential incorporation of from exogenous five-carbon precursors, such as glutamate, and poorer incorporation of from glycine and succinate, into ALA by greening intact plant tissues that had been treated with the ALA analog, levulinic acid, which causes ALA to accumulate in vivo by inhibiting PBG synthase (Beale and Castelfranco, 1974a, b; Meller et al., 1975). Degradation studies indicated that the glutamate carbon skeleton was incorporated intact into ALA (Beale et al., 1975; Meller et al., 1975). Shortly after the initial reports on the plant studies

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appeared, similar results were reported for cyanobacteria (Meller and Harel, 1978; Avissar, 1980; Kipe-Nolt and Stevens, 1980). Later, was found to be superior to glycine or succinate in contributing label to phycocyanobilin in growing cultures of Synechococcus sp. strain PCC 6301 (Laycock and Wright, 1981). On the basis of the relative abilities of or acetate to contribute label to chlorophyll and glutamate in Synechococcus sp. strain PCC 6301, McKie et al. (1981) concluded that the five-carbon pathway operates exclusively in tetrapyrrole precursor formation in this cyanobacterial species.

Samuel I. Beale

Particulate-free cell extracts capable of converting glutamate to ALA were first obtained from plants (Gough and Kannangara, 1977) and algae (Wang et al., 1984; Weinstein and Beale, 1985a; Mayer et al., 1987; Breu and Dörnemann, 1988). Later, active cell-free preparations were obtained from cyanobacteria (Rieble and Beale, 1988; O’Neill et al., 1988) and the prochlorophyte Prochlorothrix hollandica (Rieble and Beale, 1988). Similar or identicalreactionmechanisms appear to operate in all cases, and reaction components from some heterologous sources can be mixed to reconstitute activity in fractionated systems.

Chapter 17 Biosynthesis of Hemes, Chlorophylls and Phycobilins

2. Mechanism of ALA Formation from Glutamate The currently accepted model for the transformation of glutamate to ALA requires three enzyme-catalyzed reactions (Fig. 2). In the first, glutamate is activated by ligation to tRNA in a reaction identical to the charging reaction in protein biosynthesis. Like aminoacyl-tRNA formation in general, this reaction requires ATP and Next, the tRNA-bound glutamate is converted to a reduced form in a reaction that requires a reduced pyridine nucleotide. The product of this reduction has been characterized as GSA (Hoober et al., 1988), the hydrated hemiacetal form of GSA (Hoober et al., 1988), or a cyclized form of GSA (Jordan et al., 1993). Finally, the positions of the nitrogen and oxo atoms of the reduced fivecarbon intermediate are interchanged to form ALA. Consistent with this reaction model, the ALA-forming systems extracted from plants (Bruyant and Kannangara, 1987), algae (Weinstein et al., 1987), and Synechocystis sp. strain PCC 6803 (Rieble and Beale, 1991b) have been separated into four

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macromolecular components, all of which must be present to catalyze in vitro ALA formation from glutamate. Three of these components are enzymes and the fourth is a low molecular weight RNA.

a. tRNAGlu ALA formation in extracts of plants (Kannangara et al., 1984) and algae (Breu and Dörnemann, 1988; Huang et al., 1984; Weinstein and Beale, 1985b)was blocked by preincubation of the extracts with RNase A. Addition of the RNaseinhibitor, RNasin, plus low molecular weight RNA from the same species restored activity. Similarly, RNA was required for ALA formation from glutamate in extracts of cyanobacteria and a prochlorophyte (Rieble and Beale, 1988). In several plant and algal species examined, the tRNA required for ALA formation was found to contain the UUC glutamate anticodon (Schön et al., 1986; Schneegurt and Beale, 1988). In some cases, the tRNA was purified by affinity purification using an affinity ligand directed against the UUC glutamate

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anticodon (Schneegurt and Beale, 1988). In Synechocystis sp. strain PCC 6803, the affinitypurified UUC anticodon-bearing tRNA was further fractionated into two components by mixed mode (anion exchange and hydrophobic interaction) HPLC (Schneegurt et al., 1988). Each homogeneous tRNA was tested for the ability to be charged with glutamate and to participate in ALA formation and protein synthesis in extracts derived from Synechocystis sp. strain PCC 6803. The results indicated that the same species of tRNA can participate in both processes. In vitro, the two tRNAs differ functionally only in their relative abilities to be charged with glutamate by Synechocystis sp. strain PCC 6803 extracts. Once charged, they both participate equally well in both ALA formation and protein synthesis (Schneegurt et al., 1988; O’Neill and Söll, 1990). The two have the same nucleotide sequence and differ by some unspecified anticodon base modification (O’Neill et al., 1988). Only one gene (trnE) is present in the Synechocystis sp. strain PCC 6803 genome (O’Neill and Söll, 1990). Barley chloroplast like that of Escherichia coli, has a highly modified base at the anticodon: the first base is 5-methylaminomethyl-2-thiouridine (Schön et al., 1986). In E. coli, the base modification is important for efficient charging with glutamate by glutamyl-tRNA synthetase (Sylvers et al., 1993). It has been proposed that the modified base may have regulatory significance, because oxidation of the 5-methylaminomethyl-2-thiouridine with inactivated the tRNA for both glutamate acceptance and ALA synthesis, and subsequent reduction with thiosulfate reactivated it (Kannangara et al., 1988). However, treatment of the tRNA derived from darkgrown barley leaves with thiosulfate did not increase its activity. It has been pointed out that from plant and algal chloroplasts and cyanobacteria contains an base pair while most other tRNAs have a conserved G·C pair at this position (Jahn et al., 1992). Although the base pair may conceivably be involved in recognition by the glutamyl-tRNA reductase enzyme, it is not a universal requirement, since lacks this feature in several nonphotosynthetic bacteria that form ALA from glutamate.

b. Glutamyl-tRNA Synthetase In chloroplasts and cyanobacteria, the same glutamyl-tRNA synthetase (EC 6.1.1.17) is used to

charge for both protein and ALA synthesis (Bruyant and Kannangara, 1987; Rieble and Beale, 1991 b). Synechocystis sp. strain PCC 6803 glutamyltRNA synthetase has an apparent native molecular weight of 63,000 (Rieble and Beale, 1991b). The aminoacylation reaction requires ATP and Although Chlamydomonas reinhardtii glutamyltRNA synthetase is inhibited by heme under some conditions (Chang et al., 1990), and the Scenedesmus obliquus enzyme is inhibited by protochlorophyllide (Dörnemann et al., 1989), the Synechocystis sp. strain PCC 6803 glutamyl-tRNA synthetase is insensitive to heme or protochlorophyllide at physiologically relevant concentrations (Rieble and Beale, 1991b). Moreover, it was determined that Synechocystis sp. strain PCC 6803 was always fully acylated in vivo, and that the cellular content of was a constant fraction of the total cellular tRNA population under growth conditions in which chlorophyll content was modulated over a 10-fold range (O’Neill and Söll, 1990).

c. Glutamyl-tRNA Reductase Glutamyl-tRNA reductase catalyzes NADPH-linked reduction of tRNA-activated glutamate to GSA. For technical reasons, glutamyl-tRN A reductase is usually measured in coupled enzyme assays where the substrate is generated in vitro from glutamate plus tRNA, and/or the product is converted in situ to ALA. However, the reductase enzyme that utilizes NADPH is physically separable from the other enzyme components by affinity chromatography (Weinstein et al., 1987; Rieble and Beale, 1991a, b). Affinitypurified glutamyl-tRNA reductase from several sources, including Synechocystis sp. strain PCC 6803, is active in GSA formation from glutamyl-tRNA in the absence of glutamyl-tRNA synthetase and GSA aminotransferase (Rieble and Beale, 1991 a). Affinitypurified glutamyl-tRNA reductase from Chlorella vulgaris and Synechocystis sp. strain PCC 6803 requires a divalent metal such as or for activity, with optimum activity occurring at 15 mM (Mayer et al., 1993b). It is of interest that glutamyl-tRNA reductase from a given source can use, as substrate, glutamyl-tRNA from some, but not other, sources. For example, the barley and C. vulgaris enzymes can use plant and algal, but not E. coli, yeast, or animal glutamyl-tRNA (Kannangara et al., 1984; Weinstein et al., 1986). On the other hand, the reductases from C. reinhardtii and

Chapter 17 Biosynthesis of Hemes, Chlorophylls and Phycobilins the green bacterium Chlorobium vibrioforme f. thiosulfatophilum NCIB 8327 can use E. coli glutamyl-tRNA (Huang and Wang, 1986; Rieble et al., 1989). The differences in the substrate acceptability of glutamyl-tRNA reductases from different species were used in the identification of the structural gene which encodes the enzyme (see below). Glutamyl-tRNA reductaseisolated from different sources appears to have widely divergent physical properties. The C. reinhardtii enzyme was reported to be a monomer with a molecular weight of 130,000 (Chen et al., 1990). In contrast, E. coli appears to contain two different glutamyl-tRNA reductases with molecular weights of 45,000 and 85,000 (Jahn et al., 1991). The Bacillus subtilis glutamyl-tRNA reductase-encoding hemA gene encodes a 50,800 molecular weight peptide (Petricek et al., 1990), but the native enzyme migrated as oligomer with an apparent molecular weight of 230,000 (Schröder et al., 1992). Rieble and Beale (1991b) described the purification to apparent homogeneity of glutamyl-tRNA reductase from Synechocystis sp. strain PCC 6803, and reported the enzyme to have a native molecular weight of 350,000 and to be composed of subunits of 39,000 molecular weight. However, these values were later found to be in error: the major component in the purified, apparently homogeneous enzyme preparation was not glutamyl-tRNA reductase, but another enzyme, acetohydroxyacid isomeroreductase, which is not involved in ALA biosynthesis (Rieble and Beale, 1992). Additional information about the structure of glutamyl-tRNA reductase has been obtained from studies of the gene that encodes the enzyme. Before the existence of the glutamate route of ALA biosynthesis was known, ALA auxotrophic mutants of E. coli were described. Because the ALA auxotrophy was complemented by plasmids carrying DNA from animals and Rhizobium meliloti that encode the enzyme ALA synthase (Schoenhaut and Curtis, 1986; Leong et al., 1982), it was assumed that wild-type E. coli uses the ALA synthase route for ALA formation. Later, it was shown that E. coli synthesizes ALA via the glutamate route rather than with ALA synthase (Li et al., 1989), and that the ALA auxotrophic mutants are defective in genes that encode glutamyltRNA reductase or GSA aminotransferase (Avissar and Beale, 1989a; Ilag et al., 1991). Mutation of the E. coli hemA gene results in a deficiency of glutamyltRNA reductase (Avissar and Beale, 1989a). The tRNA substrate specificity of glutamyl-tRNA reduc-

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tase in complemented hemA strains of E. coli resembles that of the species from which the complementing DNA was derived, indicating that the hemA gene encodes a structural component of glutamyl-tRNA reductase (Avissar and Beale, 1990; Majumdar et al., 1991). Synechocystis sp. strain PCC 6803 DNA that shares sequence similarity with the E. coli hemA gene complements the E. coli hemA mutation and encodes a protein that has a calculated molecular weight of 47,525 (Verkamp et al., 1992; Grimm, 1992). Although glutamyl-tRNA reductase can catalyze the reduction of free glutamyl-tRNA to GSA in the absence of glutamyl-tRNA synthetase, there is some evidence that the two enzymes may form an association. In the presence of glutamyl-tRNA, C. reinhardtii glutamyl-tRNA synthetase and glutamyltRNA reductase form a complex that migrates as a single entity on glycerol gradient centrifugation (Jahn, 1992). A complex between the two enzymes may facilitate the channeling of glutamyl-tRNA toward ALA biosynthesis and competition with the protein synthesizing apparatus for glutamyl-tRNA. Glutamyl-tRNA reductase from many sources is allosterically inhibited by heme. The heme inhibition is likely to be of physiological significance in regulating the rate of ALA formation in response to the cellular demand for end-product tetrapyrroles. Synechocystis sp. strain PCC 6803 glutamyl-tRNA reductase is inhibited 50% by heme (Rieble and Beale, 1991a). The sensitivity of C. vulgaris glutamyltRNA reductase to heme inhibition is increased several fold by physiologically relevant concentrations of GSH (Weinstein et al., 1993). In contrast, the Synechocystis sp. strain PCC 6803 enzyme is unaffected by GSH (Rieble and Beale, 1991a).

d. GSA GSA has been chemically synthesized by several methods for use as a substrate for the enzymecatalyzed conversion to ALA (Kannangara and Gough, 1978; Houen et al., 1983; Gough et al., 1989). Material identical to chemically synthesized GSA accumulates in greening leaves (Wang et al., 1981; Kannangara and Schouboe, 1985) and algal extracts (Breu and Dörnemann, 1988) that have been treated with gabaculine, a mechanism-based suicide inhibitor of (see below) that blocks chlorophyll synthesis. Jordan et al. (1993) have investigated the structure

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of GSA in aqueous solution by NMR and mass spectroscopy and concluded that it exists as the cyclic ester between the group and the hydrated aldehyde group, rather than the free aldehyde (Fig. 2). The cyclic structure does not contain free aldehyde or carboxylic acid functions, and is more compatible with previously reported properties of the chemically synthesized product (stability in aqueous solution, heat stability) than the free The cyclic compound, and not GSA, was proposed to be the product of glutamyltRNA reductase and the substrate of GSA aminotransferase. It seems likely that the cyclic and linear forms of GSA coexist in solution in an equilibrium ratio, in analogy with the aldose sugars.

e. GSA Aminotransferase GSA aminotransferase (EC 5.4.3.8) catalyzes the conversion of GSA to ALA. Enzymes capable of converting chemically synthesized GSA to ALA have been purified from several plant, algal, and bacterial sources including the cyanobacteria Synechococcus sp. strain PCC 7002 and Synechocystis sp. strain PCC 6803 (Grimm et al., 1989; Rieble and Beale, 1991b). The transamination reaction requires no added substrate other than GSA. The enzyme has a bound pyridoxal-P cofactor (Avissar and Beale, 1989b; Bull et al., 1990). GSA aminotransferase is inhibited by the mechanism-based suicide substrate analog gabaculine (Kannangara and Schouboe, 1985; Avissar and Beale, 1989b). Gabaculine reacts with enzyme-bound pyridoxal-P irreversibly to form a secondary amine. Gabaculine-treated C. vulgaris GSA aminotransferase can be reactivated by gel filtration (to remove gabaculine-pyridoxal adducts and excess gabaculine) followed by incubation with pyridoxal-P (Avissar and Beale, 1989b). A proposed reaction mechanism for GSA aminotransferase involves transfer of from enzymepyridoxamine to the terminal aldehyde carbon of GSA to form enzyme-pyridoxal and 4,5-diaminovaleric acid, followed by transfer of the at the 4-position of the intermediate back to the cofactor, thereby forming ALA and regenerating the pyridoxamine form of the cofactor. Implicit in this mechanism is that the N atom of ALA is derived from a different precursor molecule than the C atoms. This prediction was tested by the use of a mixture of and glutamate molecules as substrate for

SamueI I. Beale conversion to ALA by algal cell extracts. When the heavy isotope labels were present on separate substrate molecules, asignificantproportion of theALA product molecules contained two heavy atoms, indicating that the conversion occurs by intermolecular nitrogen transfer (Mau and Wang, 1988; Mayer et al., 1993a). This result supports the proposed reaction mechanism and indicates that the enzyme catalyzes an aminotransferase reaction, rather than an aminomutase reaction, even though the substrate and product have the same atomic composition. Whereas chemically synthesized GSA is racemic, GSA that is derived from L-glutamate would be predicted to have an L configuration at the C-2 asymmetric center. Friedmann et al. (1992) synthesized both L(S)- and S(R)-4,5-diaminovaleric acid and showed that the L-isomer is the preferred substrate for Synechococcus sp. strain PCC 6301 GSA aminotransferase. Recombinant Synechococcus sp. strain PCC 6301 GSA aminotransferase showed a nearly complete preference for L-GSA, although the D-isomer was able to bind at the enzyme active site and elicit spectral changes (Smith et al., 1992). The GSA aminotransferase-encoding hemL gene (named gsa in plants and algae) has been cloned and sequenced from several plants, algae, and bacteria including cyanobacteria. These genes encode highly conserved peptides that have recognizable similarity to other members of the aspartate aminotransferase enzyme family (Elliott et al., 1990). The peptide encoded by the Synechococcus sp. strain PCC 6301 hemL gene has a predicted molecular weight of 46,038 (Grimm et al., 1991a). All hemL-encoded peptides have a conserved putative active site containing an essential lysine (at position 265 in the Synechococcus sp. strain PCC 6301 enzyme). It is believed that the pyridoxal-phosphate cofactor binds to this lysine. Mutagenesis of this lysine inactivates the enzyme (Grimm et al., 1992; Ilag and Jahn, 1992). A Synechococcus sp. strain PCC 6301 mutant that was selected for resistance to gabaculine has a GSA aminotransferase with a lower specific activity than the wild-type enzyme. The mutation that confers gabaculine resistance is (Grimm et al., 1991 b). Most GSA aminotransferases contain a methionine at this position. However, the deduced amino acid sequence of GSA aminotransferase encoded by the hemL gene of Propionibacterium freudenreichii naturally contains a valine at this position (Murakami et al., 1993). The sensitivity of the P. freudenreichii

Chapter 17 Biosynthesis of Hemes, Chlorophylls and Phycobilins enzyme to gabaculine has not been reported. Native GSA aminotransferase from Synechocystis sp. strain PCC 6803 has a molecular weight of 99,000 (Rieble and Beale, 199 1b). Purified aminotransferase from Synechococcus sp. strain PCC 6301 has a molecular weight of 46,000 on denaturing SDSPAGE (Grimm et al., 1989). Therefore, the native enzyme appears to be a homodimer, like other members of the aspartate aminotransferase enzyme family.

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B. PBG Synthesis Formation of the first pyrrole in the pathway, PBG (Fig. 3), by asymmetric condensation of two ALA molecules is catalyzed by the enzyme PBG synthase (ALA dehydratase) (EC 4.2.1.24). Although PBG synthase from animals and R. sphaeroides has an octameric structure, the enzyme from plants and algae appears to be a homohexamer, with a native molecular weight of approximately 300,000 (Jordan, 1991). An interesting difference among PBG syn-

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thase enzymes from different sources is the metal requirement: the animal, yeast, and E. coli enzymes require for activity, the R. sphaeroides enzyme requires and the plant enzyme requires (Jordan, 1991). A recent study has concluded that PEG synthase from all sources has tightly-bound but that a second, weaker metal-binding site binds in the animal, yeast, and E. coli enzymes and in the plant enzyme (Mitchell and Jaffe, 1993). The E. coli and plant enzymes are proposed to have a third metal-binding site which stimulates activity approximately two-fold when is bound (Mitchell and Jaffe, 1993). A comparative study of the deduced peptide sequences of the PEG synthases encoded by the hemB genes of several species has revealed that the cysteine and histidine residues which form a putative binding site in many PBG synthases are replaced with other amino acids in the enzyme from plants (Boese et al., 1991; Schaumberg et al., 1992) and C. reinhardtii (G.L. Matters and S.I. Beale, unpublished results). Single turnover experiments (Jordan and Seehra, 1980) have established that in the reaction catalyzed by PBG synthase from R. sphaeroides, the first bound ALA molecule is the one that contributes the propionic acid side chain of the product. In the formation of PBG, removal of hydrogen to form the aromatic pyrrole ring must occur on the enzyme, as is indicated by the stereospecific retention the pro-S hydrogen atom derived from the hydrogens of ALA (Abboud and Akhtar, 1976). III. The Pathway from PBG to Uroporphyrinogen III Uroporphyrinogen III, the last common precursor of all end-product tetrapyrroles, is synthesized from PBG by two enzymes (Fig. 3). Although the steps leading from PBG to uroporphyrinogen III are thought to be identical in all organisms, few if any studies of these steps have been done on cyanobacteria. Therefore, the following summary is derived from results obtained with other organisms.

A. Hydroxymethylbilane Synthase Hydroxymethylbilane synthase (PBG deaminase) (EC 4.1.3.8) condenses four PBG molecules to form the first tetrapyrrole, uroporphyrinogen. The initial product of enzymic catalysis is the linear tetrapyrrole,

Samuel I. Beale hydroxymethylbilane (preuroporphyrinogen), which, in the absence of another enzyme, Uroporphyrinogen III synthase, spontaneously cyclizes to form uroporphyrinogen I. Biosynthesis of the biologically relevant isomer, uroporphyrinogen III, requires the action of uroporphyrinogen III synthase during or immediately after release of hydroxymethylbilane from PBG deaminase. Hydroxymethylbilane synthase from all sources is a monomer of approximately 40,000 molecular weight, and the enzyme activity does not require metal ions or other cofactors (Jordan, 1991). The hemC gene, which encodes hydroxymethylbilane synthase, has been cloned from E. coli (Thomas and Jordan, 1986), B. subtilis (Hansson et al., 1991), Euglena gracilis (Sharif et al., 1989), and pea (Witty et al. 1993), and the E. coli enzyme has been crystallized and its structure determined by X-ray crystallography (Louie et al., 1992). Hydroxymethylbilane synthase from R. sphaeroides and E. gracilis was used to establish that the order of assembly of the four PBG units is ABCD, as they appear in uroporphyrinogen (Fig. 3; Jordan and Seehra, 1979; Battersby et al., 1979a). The nascent monopyrrole- through tetrapyrrole-enzyme complexes are bound via a dipyrrole cofactor (Battersby et al., 1979b; Scott et al., 1980; Jordan and Berry, 1981; Battersby et al., 1983; Hart et al., 1987; Jordan and Warren, 1987; Warren and Jordan, 1988). In hydroxymethylbilane synthase from E. coli, the dipyrrole cofactor is attached to a cysteine residue of the E. coli enzyme) after formation of the apoprotein, and it remains permanently attached to the enzyme, while the link between the cofactor and the nascent oligopyrrole chain is severed after the hexapyrrole stage is reached.

B. Uroporphyrinogen III Synthase Hydroxymethylbilane is unstable in solution and it rapidly cyclizes to uroporphyrinogen I, a physiologically nonproductive end product. Uroporphyrinogen III synthase (EC 4.2.1.75) catalyzes closure of the tetrapyrrole macrocycle with inversion of ring D to form the type HI product. The mechanism of ring inversion has been the subject of intense investigation and is now generally believed to involve a spiro intermediate (Crockett et al., 1991). Uroporphyrinogen III synthase has been purified from E. gracilis (Hart and Battersby, 1985), spinach (Higuchi and Bogorad, 1975), and E. coli (Jordan et al., 1988).

Chapter 17 Biosynthesis of Hemes, Chlorophylls and Phycobilins The native enzyme from all sources is a monomer of approximately 30,000 molecular weight and contains no reversibly-bound cofactors or metal ions. The hemD gene, which encodes uroporphyrinogen III synthase, has been cloned from E. coli, sequenced, and overexpressed (Sasarman et al., 1987; Jordan et al., 1988; Crockett et al., 1991). Hydroxymethylbilane synthase and uroporphyrinogen III synthase may form a complex that facilitates transfer of hydroxymethylbilane between the two enzymes. The presence of E. gracilis uroporphyrinogen III synthase influences the of E. gracilis hydroxymethylbilane synthase for PBG (Battersby et al., 1979c). The sedimentation velocity of wheat germ hydroxymethylbilane synthase is also influenced by the presence of wheat germ uroporphyrinogen III synthase (Higuchi and Bogorad, 1975). The presence of R. sphaeroides uroporphyrinogen III synthase was reported to facilitate release of the tetrapyrrole product from hydroxymethylbilane synthase (Rosé et al., 1988). IV. Steps Leading to Siroheme and Corrins Although siroheme biosynthesis has not been described in cyanobacteria, the presence of this compound is implied by the ability of cyanobacteria to grow on media in which N and S are supplied as and respectively, because siroheme is the prosthetic group of the assimilatory nitrite and sulfite reductases that are required for full reduction of these nutrients to and (Siegel, 1978), which are the only forms that are incorporated into biomolecules, e.g., amino acids. Additional evidence for the existence of siroheme in cyanobacteria includes the characterization of a ferredoxin-linked sulfite reductase from Spirulina platensis which has an absorption spectrum indicative of a siroheme prosthetic group (Koguchi and Tamura, 1988), and reports of the cysI and nirA genes, which encode sulfite and nitrite reductases, respectively, in the genome of Synechococcus sp. strain PCC 7942 (Gisselmann et al., 1993; Omata, 1991; Luque et al., 1993). Formally, conversion of uroporphyrinogen III to siroheme requires: (a) methylation of the tetrapyrrole ring at positions 1 and 3 to form precorrin 2; (b) oxidation to the tetrahydroporphyrin (sirodihydrochlorin) by removal of two electrons; (c) insertion of An alternative fate for precorrin 2 is conversion

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to corrins (vitamin nucleus). Because there is no evidence that cyanobacteria synthesize corrins, this topic will not be covered here and the reader is directed to a recent review (Battersby, 1993). Chemical arguments suggest that the order of the steps of siroheme formation is probably that given above: first methylation, then oxidation, and finally chelation. Methylation of rings A and B of uroporphyrinogen III to produce precorrin 2 effectively limits subsequent oxidation beyond the tetrahydroporphyrin (dihydrochlorin) state. Oxidation of precorrin 2 to sirodihydrochlorin produces a compound that has the aromaticity and metal binding properties necessary for efficient binding of The methyl groups of siroheme (Siegel et al., 1977) and precorrin 2 (Warren et al., 1990) are derived from S-adenosyl-L-methionine and are transferred by uroporphyrinogen methyltransferase. Based on the structure of a singly-methylated (1-methyl) intermediate, the order of methylation is deduced to be first at the 1-position, and then at the 3-position (Deeg et al., 1977; Brunt et al., 1989). In Pseudomonas denitrificans, uroporphyrinogen methyltransferase is encoded by the cobA gene (Crouzet et al., 1990). A somewhat different methyltransferase is encoded by the cysG gene in E. coli (Warren et al., 1990) and Salmonella typhimurium (Goldman and Roth, 1993). It is of interest that cysG is the only known genetic locus specifically associated with siroheme synthesis in enteric bacteria. cysG encodes a 52,000 molecular weight peptide. Its COOH-terminal region is similar to the smaller, 29,200 molecular weight peptide encoded by the cobA gene. Because cysG is the only known gene that is involved in siroheme synthesis, and because only a portion of its encoded peptide is similar to the smaller cobA product, the CysG protein may be a multifunctional enzyme that catalyzes all steps of the conversion of uroporphyrinogen III to siroheme in the enteric bacteria (Goldman and Roth, 1993). In any case, it appears that insertion into sirodihydrochlorin is not catalyzed by the ferrochelatase that is responsible for protoheme formation (Powell et al., 1973; Hansson and Wachenfeldt, 1993). That enzyme is encoded by the hemH gene in E. coli (Frustaci and O’Brian, 1993), B. subtilis (Hansson andHederstedt, 1992),and Bradyrhizobium japonicum (Frustaci and O’Brian, 1992). Synechococcus sp. strain PCC 6301 contains a gene that more closely resembles the smaller cobA gene than cysG (A. G. Smith, personal communication). This fact

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suggests that, in contrast to the apparent situation in the enteric bacteria, uroporphyrinogen III methylation, precorrin 2 oxidation, and Fe chelation are catalyzed by separate enzymes in cyanobacteria. V. Conversion of Uroporphyrinogen III to Protoporphyrin IX

A. Uroporphyrinogen Decarboxylase Uroporphyrinogen decarboxylase (EC 4.1.1.37) catalyzes the decarboxylation of all four of the acetate residues on uroporphyrinogen to yield coproporphyrinogen, which contains methyls in their place. At physiological substrate concentrations, the decarboxylations occur in a specific sequence, beginning at ring D and proceeding clockwise around the macrocycle (Luo and Lim, 1993). At higher substrate concentrations, the decarboxylation sequence becomes random. The gene that encodes uroporphyrinogen decarboxylase has been cloned from several eucaryotes including yeast and mammals, as well as B. subtilis, in which it has been identified with the hemE locus (Hansson and Hederstedt, 1992). B. subtilis hemE encodes a 40,347 molecular weight peptide. A portion of an open reading frame has been reported in DNA from Synechococcus sp. strain PCC 7942 that encodes a peptide that is highly similar to the region of the B. subtilis protein (Kiel et al., 1990). Uroporphyrinogen decarboxylase activity was measured in leaf extracts of several plants and purified 72-fold from tobacco leaves (Chen and Miller, 1974). No metal requirements were detected, and EDTA or other metal chelating agents enhanced activity. Uroporphyrinogen III is a much better substrate than uroporphyrinogen I for the tobacco enzyme. A greaterthan-400-fold purified enzyme from E. gracilis has a molecular weight of 54,000, is not inhibited by EDTA, is stimulated by dithiothreitol, and is able to decarboxylate both uroporphyrinogen I and uroporphyrinogen III, the latter being a much better substrate (Juknat et al., 1989).

B. Coproporphyrinogen Oxidase Coproporphyrinogen oxidase (EC 1.3.3.3) oxidatively decarboxylates the propionate groups at positions 2 and 4 of coproporphyrinogen III to vinyls,

Samuel I. Beale thereby producing protoporphyrinogen IX. The enzyme is specific for the III isomer of coproporphyrinogen over the nonphysiological I isomer, although chemically synthesized coproporphyrinogen IV is also decarboxylated by the enzyme (Mombelli et al., 1976). Evidence indicating that the 2-propionate is converted before the 4-propionate includes characterization of a 2-monovinyl intermediate in rat liver preparations (Elder et al., 1978) and the preferential action of the E. gracilis enzyme on the chemically synthesized 2-monovinyl porphyrin compared to the ring 4-monovinyl porphyrin (Cavaleiro et al., 1974). Seehra et al. (1983) have proposed a reaction mechanism involving pyrrolic N-assisted removal of single protons as hydride ions from the of the propionate groups. The of the propionate groups do not appear to be involved: in the reaction catalyzed by an avian blood extract, both of the of both propionate groups were retained on the terminal carbon atoms of the protoporphyrinogen vinyl groups (Zaman and Akhtar, 1978). In aerobic organisms (presumably including cyanobacteria) coproporphyrinogen oxidase is an reaction. Extracts of anaerobically-grown R. sphaeroides cells can carry out the reaction anaerobically in the presence of ATP, oxidized pyridine nucleotide, and methionine (Tait, 1972). Similar requirements were reported for anaerobic yeast extracts (Poulson and Polglase, 1974). Although coproporphyrinogen oxidase is associated with the mitochondrial membranes of animal cells, the yeast enzyme is cytosolic (Camadro et al., 1986). Yeast coproporphyrinogen oxidase is a 70,000 molecular weight homodimer that contains two molecules of Fe (Camadro et al., 1986). A yeast gene (HEMI3) encoding coproporphyrinogen oxidase has been cloned and sequenced (Zagorec et al., 1988). In B. subtilis, mutations at the hemY locus cause accumulation of either coproporphyrinogen III alone or both coproporphyrinogen III and protoporphyrinogen IX (Hansson and Hederstedt, 1992). The B. subtilis hemY gene contains a single open reading frame that encodes a polypeptide with no recognizable similarity to the yeast HEM13 product. It was proposed that the B. subtilis hem Y product, which at 52,000 molecular weight is significantly larger than the 37,600 molecular weight yeast HEM13 product, may be a bifunctional enzyme that has both coproporphyrinogen oxidase and protoporphyrinogen oxidase activities (Hansson and Hederstedt, 1992).

Chapter 17 Biosynthesis of Hemes, Chlorophylls and Phycobilins

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C. Protoporphyrinogen Oxidase

VI. The Fe Branch

Protoporphyrinogen oxidase (EC 1.3.3.4) catalyzes the removal of six electrons from the tetrapyrrole macrocycle to form protoporphyrin IX, the last biosynthetic step that is common to hemes, bilins, and chlorophylls. Although protoporphyrinogen autoxidizes in solution in the presence of the oxidationin cells isenzymemediated. Mutantstrains of B. japonicum that are deficient in protoporphyrinogenoxidase donotformleghemoglobinheme and are devoid of cytochrome oxidase activity (O’Brian et al., 1987; Ramseier et al., 1989). Moreover, inhibition of protoporphyrinogen oxidase in plants by diphenyl ether herbicides causes photobleaching. The herbicidal mechanism involves inhibition of protoporphyrinogen oxidase by the herbicide, causing accumulation of its substrate, protoporphyrinogen IX. This compound diffuses to the cell membrane where it is oxidized to protoporphyrin IX, which is a photodynamic sensitizer that causes light-induced damage to the membrane (Matringe et al., 1989; Becerril and Duke, 1989). The paradoxical effect that an inhibitor of protoporphyrinogen oxidase raises the concentration of the product of the enzyme reaction is explained by the fact that protoporphyrinogen IX diffuses away from the normal site of its enzymic oxidation and the nonspecifically oxidized product accumulates in cellular regions where it is inaccessible to the normal biosynthetic enzymes. The deficiency of later intermediates, such as chlorophyll precursors and protoheme, may induce a compensatory release of feedback control over protoporphyrinogen synthesis and thereby cause an overproduction of this compound. Protoporphyrinogen oxidase from barley chloroplasts and mitochondria is a membrane-associated protein that has an apparent molecular weight of 210,000 and is composed of36,000 molecular weight subunits (Jacobs and Jacobs, 1987). In contrast, protoporphyrinogen oxidase from the anaerobe Desulfovibrio gigas was reported to have a native molecular weight of 148,000 and to contain three dissimilar subunits of 12,000, 18,500, and 57,000 molecular weight (Klemm and Barton, 1987). As discussed earlier, the B. subtilis hemY gene encodes a 57,000 molecular weight protein that may have both coproporphyrinogen oxidase and protoporphyrin oxidase activities (Hansson and Hederstedt, 1992).

In many groups of photosynthetic organisms such as plants, green algae, and anoxygenic photosynthetic bacteria, the Fe branch receives a lower quantity of precursors than the Mg branch, a situation that arises because of the quantitative preponderance of chlorophylls over hemes in these organisms. However, in cyanobacteria, the relative distribution ofprecursors into the Fe and Mg branches is more equal, or even reversed, because of the high cellular content of phycobiliproteins, which derive their phycobilin chromophores from heme.

A. Ferrochelatase Ferrochelatase (protoheme ferrolyase, EC 4.99.1.1) catalyzes the last step of protoheme formation: the insertion of into protoporphyrin IX. In addition to its physiological substrates, ferrochelatase can use and as the metal substrate and deuteroporphyrin IX, mesoporphyrin IX, and hematoporphyrin IX (in which the vinyl groups of protoporphyrin IX are replaced with H, ethyl, and groups, respectively) as the porphyrin substrate. Ferrochelatase is an intrinsic membrane protein that requires detergents or chaotropic agents for solubilization. An apparent exception is the B. subtilis ferrochelatase, which was reported to be a soluble enzyme (Hansson and Hederstedt, 1992). The hemH gene, which encodes ferrochelatase, has been cloned and sequenced from yeast and several animal sources, as well as from E. coli (Miyamoto et al., 1991; Frustaci and O’Brian, 1993), B. japonicum (Frustaci and O’Brian, 1992), S. typhimurium (Xu et al., 1992), and B. subtilis (Hansson and Hederstedt, 1992). The encoded bacterial polypeptides range from 34,000 to 38,000 molecular weight and are somewhat smaller than the polypeptides encoded by the animal and yeast genes. Although ferrochelatase has not been purified to homogeneity from anyplantorcyanobacterial source, activity has been measured in cell-free extracts of several plants and the enzyme has been partially purified from etiolated barley (Golden and Little, 1969). Ferrochelatase activity has also been characterized from spinach chloroplasts that were stripped of their outer envelope membranes (Jones, 1968).

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Brown et al. (1984a) measured ferrochelatase activity in extracts of the unicellular rhodophyte, Cyanidium caldarium, in an assay using and deuteroporphyrin IXas substrates. In wild-type cells, which do not normally form pigment in the dark, the in vitro ferrochelatase level increased several fold within 72 h after the cells were transferred from darkness to light. The increase in ferrochelatase activity paralleled the accumulation ofphycocyanin, which suggests that the level of this enzyme may be an important rate-controlling factor in phycobilin synthesis. The intracellular localization of the measured ferrochelatase activity was not reported.

B. Synthesis of Other Hemes In addition to protoheme, cyanobacteria also contain heme c, the covalently bound prosthetic group of cytochromes c and f and heme a, which is an essential component of cytochrome type cytochrome oxidase (see Fig. 4). Apocytochromes of the c type are thought to incorporate protoheme by enzymecatalyzed ligation of the vinyl groups to cysteine residues on the protein. However, a specific ligating

Samuel I. Beale enzyme, named cytochrome c-heme lyase, has been described only in yeast and Neurospora crassa extracts (Taniuchi et al., 1983; Nicholson et al., 1987). The enzyme is localized in the inner mitochondrial membrane (Nargang et al., 1988; Enosawa and Ohashi, 1986) and requires reduced (ferro)protoheme as a substrate (Nicholson and Neupert, 1989). The gene for cytochrome c-heme lyase has been cloned and sequenced from yeast (CYC3) and N. crassa (cyt-2) (Dumont et al., 1987; Drygas et al., 1989). Although the two encoded peptides are 32% identical, the yeast peptide is considerably smaller than the N. crassa peptide (molecular weights of 29,600 and 38,000, respectively). Cytochrome oxidase (EC 1.9.3.1) of the type has been detected in cyanobacteria including the unicellular species Synechococcus sp. strain PCC 6301 (Molitor et al., 1987) and the filamentous nitrogen-fixing species, Anabaena variabilis (Houchins and Hind, 1984; Häfele et al., 1988). In A. variabilis, cytochrome oxidase is found in heterocysts, whereas in Synechococcus sp. strain PCC 6301, it is present in both the plasma membrane and thylakoid membranes. The heme a prosthetic group of cytochrome oxidase (Fig. 4) is thought to be formed from protoheme by ligation ofa farnesyl group to the 2-vinyl group and oxidation or oxygenation of the 8-methyl group to a formyl group. Although the enzymes responsible for these protoheme modifications have not been described, the precursor status of protoheme has been shown in B. subtilis. Strains that are defective in the later steps of protoheme formation (uroporphyrinogen decarboxylase, coproporphyrinogen oxidase, protoporphyrinogen oxidase, and ferrochelatase) could not form heme a unless protoheme was added to the medium, even though the same cells could form siroheme, presumably using a different ferrochelatase for its biosynthesis (Hansson and Wachenfeldt, 1993). Although the order of the two protoheme modifications is not known, it is of interest that heme o, the prosthetic group of cytochrome o in E. coli, has the structure of a possible heme a precursor in which the 2-farnesyl group has been added but the 8-methyl group has not been converted to a formyl group (Wu et al., 1992). The cyoE gene of E. coli, when overexpressed, causes conversion of protoheme to heme o, and is therefore thought to encode a farnesyltransferase (Saiki et al., 1992).

Chapter 17 Biosynthesis of Hemes, Chlorophylls and Phycobilins

C. Phycobilin Formation Phycobilins are ubiquitous light-harvesting photosynthetic accessory pigments in cyanobacteria, and it is the presence of a phycobilin, phycocyanobilin, that gives cyanobacteria their characteristic blue-green color. In their functional state, phycobilins are covalently linked to phycobiliproteins by one (or sometimes two) thioether bonds between protein cysteine residues and portions of the phycobilins that are derived from vinyl residues on bilin pyrrole rings

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A and D (Fig. 5). Phycobilins are released from some phycobiliproteins by treatment with refluxing methanol. The structures of methanol-liberated phycobilins are somewhat different from the proteinbound forms in that they contain an ethylidine group that is generated from scission of the bond. As will be discussed below, ethylidinecontaining phycobilins are also thought to be precursors to the protein-bound pigments. The first phycobilins for which the structure was determined are methanolysis-liberated phycocy-

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anobilin and phycoerythrobilin (Cole et al., 1967; Crespi et al., 1967; Chapman et al., 1967; Crespi and Katz, 1969) (Fig. 5). Structures of several additional phycobilins have been described, bringingthepresent number of distinct chromophores to eight (Glazer and Hixson, 1977; Bishop et al., 1987; Wedemayer et al., 1991, 1992). The pace of discovery of new phycobilins suggests that more structures will be uncovered as pigments of more organisms are examined in detail. Cyanobacteria are one of three groups of organisms that contain phycobilins; the others are rhodophyte and cryptophyte algae. Virtually all of the experimental work on phycobilin biosynthesis has been done with one rhodophyte species, Cyanidium caldarium., a unicellular organism that has only phycocyanobilin-containing phycobiliproteins. Therefore, much ofthe informationaboutthesynthesis of phycocyanobilin and other phycobilins in cyanobacteria is necessarily extrapolated from the results obtained with C. caldarium. Thefirst direct evidence indicatingthatphycobilins are derived from heme and biliverdin was the ability of exogenous protoheme and biliverdin to contribute label to phycocyanobilin in intact greening C. caldarium cells (Brown et al., 1981; 1984b; Schuster et al., 1983; Beale and Cornejo, 1983). These results were followed by in vitro detection of enzymes capable of converting protoheme to phycobilins, as described below.

1. Ferredoxin-Linked Heme Oxygenase Heme oxygenase (EC 1.14.99.3) is the enzyme that catalyzes the opening of the heme macrocycle to form biliverdin in heme degradation. Heme oxygenase activity was detected in extracts of C. caldarium (Beale and Cornejo, 1984b). Like the animal system, the unfractionated algal heme oxygenase system requires reduced pyridine nucleotide and and activity is powerfully inhibited by the competitive inhibitor, Sn-protoporphyrin IX. Ascorbate and other moderately strong reductants stimulate the reaction in unfractionated cell extracts, and are required after removal of low molecular weight materials from the enzyme system by gel filtration or dialysis. Products ofthe reaction with mesoheme and protoheme as substrate were identified as the isomers of mesobiliverdin and biliverdin, respectively, indicating that the enzymatic reaction specifically opens the macrocyclic ring at the same

Samuel I. Beale bridge carbon as does animal heme oxygenase (Beale and Cornejo, 1984b; Cornejo and Beale, 1988). The algal heme oxygenase system differs from the animal cell-derived microsomal system in that it is soluble, with virtually all activity appearing in the supernatant fraction of high speed-centrifuged cell homogenates (Beale and Cornejo, 1984b). Microsomal heme oxygenase from animal cells requires two protein components for activity: heme oxygenase and NADPH:ferrihemoprotein reductase (EC 1.6.2.4). In contrast, the soluble heme oxygenase system from C. caldarium can be fractionated into three required protein components: one has ferredoxin-dependent NADPH:cytochrome c reductase activity; the second is a small Fe-S protein that appears to be ferredoxin; and the third is a hemebinding oxygenase enzyme (Cornejo and Beale, 1988). Reconstitution of heme oxygenase activity in vitro required all three protein components. The ferredoxin-dependent NADPH:cytochrome c reductase-containing fraction could be replaced by spinach ferredoxin: reductase (EC 1.18.1.2), and the ferredoxin-containing fraction could be replaced by commercial ferredoxin derived from spinach or the red alga, Porphyra umbilicalis (Cornejo and Beale, 1988; Rhie and Beale, 1992). In the absence of NADPH and ferredoxin: reductase, algal heme oxygenase activity was supported by a lightdriven ferredoxin reduction system derived from a partially purified spinach leaf Photosystem I preparation (Rhie and Beale, 1992). In this mixed reconstitution assay, heme oxygenase activity was light dependent. In the dark, no activity was detected unless NADPH and ferredoxin: reductase were added to incubation mixtures. These results indicate that the sole, essential, role of NADPH and the ferredoxin-dependent NADPH:cytochrome c reductase(ferredoxin: reductase) in the algal heme oxygenase system is to reduce ferredoxin, and that ferredoxin is the direct electron source for algal heme oxygenase. The heme-binding component of the algal heme oxygenase system has an apparent native molecular weight of approximately 38,000 and is resistant to inactivation by p-hydroxymercuribenzoate (Cornejo and Beale, 1988). The enzyme is inactivated by diethylpyrocarbonate, and this inactivation isblocked by heme. In the reconstituted heme oxygenase system, the heme-binding component was the rate limiting one: addition of this component to unfractionated cell extracts increased the yield of biliverdin, whereas

Chapter 17 Biosynthesis of Hemes, Chlorophylls and Phycobilins addition of ferredoxin or ferredoxin: reductase did not. A gene that encodes a protein with strong similarity to animal microsomal heme oxygenase has been located in the chloroplast genome ofthe rhodophyte, Porphyra purpurea (Reith and Munholland, 1993). Preliminary attempts to detect heme oxygenase activity in extracts of E. coli that was transformed with plasmids carrying this gene have not been successful (G. Rhie and S. I. Beale, unpublished results). Because heme oxygenase catalyzes the first committed step of phycobilin synthesis, it would be expected to have a regulatory role by controlling the entry of precursors into this branch of the pathway. Heme oxygenase activity was several-fold higher in extracts of light-grown wild-type C. caldarium cells (which require light for pigment accumulation) than in dark-grown cell extracts. Growth on D-glucose, which is known to suppress pigment content, lowered extractable heme oxygenase activity. Finally, addition of ALA to the growth medium increased the yield of heme oxygenase activity (Rhie and Beale, 1994).

2. Conversion of Biliverdin to Phycobilins Because nearly all phycobilins that have been described so far contain at least two more hydrogen atoms than biliverdin, some form of biochemical reduction must be necessary for the transformation of biliverdin into these phycobilins. Biliverdin was converted to free phycocyanobilin in extracts of C. caldarium (Beale and Cornejo, 1984a). In addition to biliverdin the reaction required reduced pyridine nucleotide, NADPH being more effective than NADH. Activity was retained in the supernatant fraction after high-speed centrifugation and eluted with the protein fraction on gel filtration.

a. Phycobilin Ethylidine Isomerization It was initially puzzling that incubation products included both the (3Z)- and the (3E)-ethylidine isomers of phycocyanobilin (Fig. 5) (Beale and Cornejo, 1984a). Interestingly, both ethylidine isomers of phycocyanobilin are also formed upon methanolytic cleavage of the phycocyanin chromophore from the protein moiety (Fu et al., 1979), but the Z isomer, being less stable (Weller and Gossauer, 1980), isomerizes to the E isomer at the high

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temperatures at which methanolysis is carried out, and the equilibrium isomer ratio strongly favors the E form. The Z- and E-ethylidine isomers of phycocyanobilin can be interconverted by heating in acetic acid-methanol mixtures (Beale and Cornejo, 1984a). Preliminary evidence indicates that enzymatic cistrans isomerization oftheethylidinegroup iscatalyzed by C. caldarium cell extracts in the presence of reduced GSH (Beale and Cornejo, 199la). After removal of low molecular weight material from cell homogenates by Sephadex G-25 gel filtration, the major product formed enzymatically was (3Z)-phycocyanobilin. Addition of GSH to incubations containing gel-filteredcell extractsrestored theability to form (3E)-phycocyanobilin. Moreover, purified (3Z)-phycocyanobilin was converted to (3E)-phycocyanobilin by cell extracts, and this conversion was greatly stimulated by GSH. The reverse conversion was not detected. In the absence of enzyme, GSH did not catalyze the conversion. These results indicate that extracts from C. caldarium cells contain a GSH-dependent enzyme activity that isomerizes the ethylidine group of (3Z)-phycocyanobilin to the E configuration. The existence of the (3Z)-phycobilins may be rationalized by proposing that the substrate for the pyrrole 2,3-reductase reaction is a bilin that has a 3 -vinyl group and the immediate product is a 3 -viny l2,3-dihydrobilin. Gossauer et al. (1989) have shown that synthetic 3-vinyl-2,3-dihydrobilins spontaneously isomerize to (3Z)-ethylidine-2,3-dihydrobilins. Thus, the enzymatic pyrrole reduction of biliverdin or another 3-vinylbilin would initially yield the unstable intermediate 3-vinyl-2,3-dihydrobilin, which would isomerize nonenzymatically to the (3Z)-ethylidine-2,3-dihydrobilin. However, if only the more stable (3E)-phycobilins are acceptable substrates for ligation of the chromophore to the apoprotein,there wouldbeaneed foraZ-E isomerase to transform the products of pyrrole ring reduction into the substrates for ligation.

b. Ferredoxin-Linked Biliverdin

Reduction

Activity fractionation experiments revealed that two of the protein components of the C. caldarium biliverdin reducing system are the same ones that are needed for the heme oxygenase system: ferredoxin and ferredoxin: reductase (Beale and Comejo, 1991a). These components could be replaced with commercial counterparts from spinach or a red alga

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and, as with heme oxygenase, the requirement for NADPH and ferredoxin: reductase could be supplanted by a light-driven ferredoxin-reducing system derived from spinach thylakoids (Rhie and Beale, 1992). These results indicate that reduced ferredoxin is the only reductant needed for reduction of biliverdin to phycobilins.

c. Phycoerythrobilin Isomerization In incubations containing partially fractionated C. caldarium extracts, bilin products in addition to phycocyanobilin accumulated. One of these bilins was identified as (3Z)-phycoerythrobilin (Fig. 5) (Beale and Cornejo, 1991b). The occurrence of (3Z)-phycoerythrobilin as an incubation product was unexpected, because C. caldarium does not contain phycoerythrin or other phycoerythrobilin-bearing phycobiliproteins. However, C. caldarium extracts contain an enzyme that converts (3Z)-phycoerythrobilin to (3Z)-phycocyanobilin (Beale and Cornejo, 1991b). The reverse reaction was not detected. The reaction does not require any substrate other than (3Z)-phycoerythrobilin. Because (3Z)-phycoerythrobilin and (3Z)-phycocyanobilin are isomeric, the enzyme can be considered to be an isomerase. In addition to being able to transform the (3Z) isomer, the protein fraction also catalyzed conversion of (3E)-phycoerythrobilin to (3E)-phycocyanobilin. The enzyme activity was named phycoerythrobilin-tophycocyanobilin (15, isomerase. A gel filtration fraction containing proteins in the 30,000 to 40,000 molecular weight range catalyzed the conversion ofbiliverdin to phycoerythrobilin but not to phycocyanobilin. The fraction containing proteins ofmolecular weight greater than 60,000 was inactive in biliverdin reduction, but catalyzed isomerization of phycoerythrobilin to phycocyanobilin (Beale and Cornejo, 1991b). The presence of phycoerythrobilin in an organism that does not use this bilin as a phycobiliprotein chromophore, and of an enzyme that converts phycoerythrobilin to phycocyanobilin, strongly suggest that phycoerythrobilin is an intermediate in the biosynthesis ofphycocyanobilin from biliverdin.

d. 15,16-Dihydrobiliverdin Partially fractionated C. caldarium protein extract, when incubated with biliverdin and reductant,

produced, in addition to phycocyanobilin and phycoerythrobilin, a third bilin that was identified as 15,16-dihydrobiliverdin (Fig. 5; Beale and Cornejo, 1991 c). Further fractionation of the proteins yielded afraction thatformed 15,16-dihydrobiliverdin as the sole product ofbiliverdin reduction. Purified 15,16-dihydrobiliverdin when incubated with another protein fraction, was converted to (3Z)-phycoerythrobilin and (3Z)-phycocyanobilin. This conversion, as well as the conversion ofbiliverdin to 15,16-dihydrobiliverdin required reductant in addition to the bilin substrate. These results suggest that 15,16-dihydrobiliverdin is a partially reduced intermediate in the biosynthesis of phycoerythrobilin from biliverdin This finding was followed by the discovery that 15,16-dihydrobiliverdin occurs as a phycobiliprotein chromophore in some cryptophyte algae (Wedemayer et al., 1992; Wemmer et al., 1993). The protein fractionation results indicate that reduction of biliverdin to phycoerythrobilin proceeds by two two-electron steps, each of which is catalyzed by a different enzyme (Fig. 5). One enzyme, when supplied with biliverdin plus a source ofreduced ferredoxin, produces only 15,16-dihydrobiliverdin (Beale and Cornejo, 1991c). Further reduction of the bilin to phycoerythrobilin requires other proteins. The enzyme that catalyzes the first reduction step can be separated from the other reductase by affinity chromatography on ferredoxin-Sepharose. As indicated above, protein fractionation indicates that the reduction steps are catalyzed by enzymes having apparent native molecular weights ofapproximately 30,000 to 40,000, and isomerization of phycoerythrobilin to phycocyanobilin is catalyzed by an enzyme with an apparent native molecular weight greater than 60,000. There is no experimental information available on the formation of phycobilins other than 15,16-dihydrobiliverdin phycoerythrobilin, and phycocyanobilin.

3. Protein Ligation Genetic and biochemical results support a role for specific proteins in phycobilin ligation (Zhou et al., 1992; Swanson et al., 1992a). At the cpc gene locus of Synechococcus sp. strain PCC 7002, there are, in addition to genes that code for structural components of phycobilisomes, two open reading frames designated cpcE and cpcF. Insertional mutagenesis of cpcE or cpcF caused the appearance ofyellow-green

Chapter 17 Biosynthesis of Hemes, Chlorophylls and Phycobilins cells that failed to accumulate normal amounts of phycocyanin. The cells contained normal phycocyanin subunits, but most subunits lacked chromophores. These results were interpreted to indicate that the ligation ofthe chromophore to the subunit requires the cpcE and cpcF gene products. Recombinant CpcE and CpcF proteins were overproduced in E. coli. Together, these proteins catalyzed in vitro ligation of phycocyanobilin to the subunit at the correct cysteine residue (Cys-84) (Fairchild et al., 1992). In addition to catalyzing ligation of free phycocyanobilin, CpcE/CpcF catalyzed exchange of bilin between the of phycocyanin and resin-bound apophycocyanin Phycocyanobilin binds tightly to CpcE/CpcF. The kinetics of the CpcE/CpcF-catalyzed ligation of phycocyanobilin to apophycocyanin showed a rapid initial bilin ligation phase followed by a slower linear phase. The kinetics were interpreted to indicate that CpcE/CpcF catalyzed the rapid reactionofaminor conformerorconfigurational isomer of phycocyanobilin with the apoprotein. The subsequent slow phase represents further ligation of this form of phycocyanobilin as it is produced by slow isomerization of the bulk of the remaining phycocyanobilin. A possible structure of the fasterreacting bilin is (3Z)-phycocyanobilin. This isomer, which can be detected as a minor component in the products of phycocyanin methanolysis (Beale and Cornejo, 1984a), is the major product of in vitro phycocyanobilin formation from biliverdin in C. caldarium cell extracts (Beale and Cornejo, 1984a, 1991 a). As described above, C. caldarium cell extracts contain an enzyme that catalyzes GSH-dependent cis-trans isomerization of the 3-ethylidine group of phycocyanobilin (Beale and Cornejo, 199la). It will be of interest to determine if the ethylidine cis-trans isomerase is encoded by the cpcE and/or cpcF genes. Genes resembling cpcE and cpcF have been found in the cpc gene clusters of other cyanobacteria (Belknap and Haselkorn, 1987; Conley et al., 1988; Mazel and Marliére, 1989; Bryantet al., 1991; Glauser et al., 1992; Dubbs and Bryant, 1993). Because the CpcE/CpcF proteins appear to affect phycocyanobilin attachment only to the apophycocyanin there are presumably other, so far undiscovered proteins that are required for phycobilin ligation to apophycocyanin and other apophycobiliproteins. An indication that there are specific proteins that mediate attachment ofother bilins is the

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presence of genes that are similar to cpcE and cpcF in the phycoerythrin-encoding gene cluster of the marine cyanobacterium Synechococcussp. WH 8020 (Wilbanks and Glazer, 1993) and in the phycoerythrocyanin-encoding gene cluster of Anabaena sp. strain PCC 7120 (Swanson et al., 1992b). Functional evidence for multiple ligating enzymes is provided by the spontaneous appearance ofa pseudorevertant in a culture of a cpcE mutant strain of Synechococcus sp. strain PCC 7002 (Swanson et al., 1992a). The pseudo-revertant has a amino acid substitution in the phycocyanin subunit that presumably allows it to serve as a substrate for chromophore ligating enzymes that normally function at other attachment sites. Although all reported instances ofcpcE/cpcF-like genes occur within co-transcribed gene clusters that also contain apophycobiliprotein genes, some indirect evidence suggests that such placement and cotranscription may not be universal. In C. caldarium cells that require light for chlorophyll and phycobiliprotein accumulation, administration of ALA to cells growing in the dark causes the induction ofboth apophycocyanin and phycocyanobilin formation, but the pigment and apoprotein remain unligated until the cells are exposed to light (Turner et al., 1992). These results suggest that, while ALA induces the expression of the genes encoding the apophycobiliproteins, light is required for expression of the genes encoding the proteins that catalyze chromophore ligation. It is unclear how the different chromophores and apoprotein ligation sites are recognized by the components responsible for attaching the various different chromophores at specific sites. The ligation process is further complicated by the recent discovery that at most sites, ligation results in the generation of an group with an R configuration, but at some sites, the thioethyl group has the S configuration (Duerring et al., 1991). Furthermore, it has become apparent that the thioether links do not invariably involve the of the bilin substituent: certain doubly-linked 15,16-dihydrobiliverdin chromophores of cryptophyte algal phycobiliproteins have been shown to be attached to the apoproteins at the of the bilin 3-position substituent and at the of the bilin 18-position substituent, i.e., the linkages are and (Wemmer et al., 1993).

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VII. The Mg Branch The pathway from protoporphyrin IX to chlorophyll a consists of six steps: Mg chelation, 6-propionate methylation, isocyclic ring formation, 4-vinyl reduction, ring D reduction, and esterification of the 7-propionate. In cyanobacteria, chlorophyll a is the principal end product of the Mg branch, but in the prochlorophytes, a considerable amount of chlorophyll b is also produced. Anoxygenic photosynthetic bacteria contain a large gene cluster, named the photosynthesis gene cluster (PGC), that contains many bch genes that encode enzymes catalyzing specific steps in the conversion of protoporphyrin IX to bacteriochlorophyll (Yang and Bauer, 1990; Coomber et al., 1990; Burke et al., 1992; 1993). Work is currently underway in a number of laboratories to characterize the bch genes and the proteins they encode by DNA sequencing, mutagenesis, and cloning and expressionin E. coli. Because several of the steps of bacteriochlorophyll biosynthesis are common to chlorophyll biosynthesis, homologs of some bch genes are likely to be present in cyanobacteria, and indeed, some homologs have already been identified (see below).

A. Mg Chelatase Early attempts to study Mg chelation were successful in detecting activity only in intact, permeabilized bacterial cells (Gorchein, 1972) and intact chloroplasts (Castelfranco et al., 1979). Chloroplasts that were broken by lysis in hypotonic medium yielded much less than 1% of the activity of intact plastids, and the residual activity could have been due to a small number of plastids that had escaped lysis (Richter and Rienits, 1982). Using intact cucumber cotyledon chloroplasts, the substrate requirements of Mg-chelatase were investigated (Walker and Weinstein, 199la). ATP was required for activity, and nonhydrolyzable ATP analogs failed to support activity. Activity was inhibited 50% by 1.8 mM AMP, even in the presence of 4 mM ATP, and by 0.6 mM 1,10-phenanthroline. Acceptable porphyrin substrates were protoporphyrin IX, deuteroporphyrin IX (vinyl groups replaced by H), mesoporphyrin IX (vinyl groups replaced by ethyls), and porphyrins with either one of the two vinyl groups replaced by an ethyl group. N-Methylprotoporphyrin IX and N-methylmesoporphyrin IX, which are potent inhibitors of ferrochelatase, inhibited Mg chelation at

Samuel I. Beale micromolar concentration. Inhibition by the sulfhydryl reagent, p-chloromercuribenzenesulfonate, which does not permeate into the chloroplasts, suggested that Mg chelatase was located in the envelope membranes, rather than the thylakoids. Pea plastids were the first to yield Mg-chelatase activity after disruption (Walker and Weinstein, 1991b). Activity required both a soluble protein fraction and a membrane-associated protein fraction, both of which were inactivated by heating. The combined protein fractions required ATP, and porphyrin for activity. Membrane fractionation indicated that the membrane-associated protein is localized in the nonpigmented, light membranes rather than the thylakoids (Walker et al., 1992). A continuous activity assay revealed that there is a fivemin lag phase before Mg chelation begins. The lag phase was shortened by pre-incubating the membraneassociated fraction with ATP, whereas preincubation of this fraction without ATP abolished activity, suggesting that ATP is involved in activating and stabilizing the enzyme system. Cucumber chloroplasts also yielded Mg chelatase activity (Lee et al., 1992). In contrast to the pea plastid system, all activity was associated with the membranes of the cucumber system, and a stroma fraction was neither required nor stimulatory. Protoporphyrin IX, as well as ATP, stabilized the cucumber plastid enzyme activity during isolation and, like the pea plastid system, activity required ATP and Because Mg chelation is the first committed step of the chlorophyll branch, it has been hypothesized to be a key regulatory step, and is has been expected that Mg chelation would be activated by light and inhibited by protochlorophyllide or other pigments. Mg chelatase activity in intact isolated cucumber plastids increased with increasing time of light exposure of the cotyledons before plastid isolation (Walker and Weinstein, 199la). However, activity was not inhibited by added protochlorophyllide, chlorophyllide, Mg-protoporphyrin IX, or heme. Mutations have been described in several plants and algae that cause protoporphyrin IX to accumulate and decrease or abolish chlorophyll accumulation. These mutations are presumed to be in genes that encode components of the Mg chelatase system. A pale-pigmented cs mutant of Arabidopsis thaliana was generated by T-DNA insertion (Koncz et al., 1990), that accumulates protoporphyrin IX in its tissues (T. Falbel, personal communication cited by

Chapter 17 Biosynthesis of Hemes, Chlorophylls and Phycobilins Koncz et al., 1992). The plant DNA boundaries of the T-DNA insert were used to isolate a cDNA that encodes a 46,251 molecular weight (including the transit sequence) chloroplast protein, which was suggested to be Mg chelatase. The physical properties of mature protein,deducedfrom the cDNA sequence, suggest that it is located in the stroma. The Rhodobacter capsulatus bchI gene, located in the bch gene cluster, encodes a homologous protein of 38,000 molecular weight (Armstrong et al., 1993). Disruption of this gene causes protoporphyrin IX to accumulate (Bollivar, 1993). Whereas the bchI-like gene is present in the nuclear genome of A. thaliana, similar genes have been located in the plastid genome of E. gracilis (Orsat et al., 1992), the rhodophyte P. purpurea (Reith and Munholland, 1993), and the cryptophyte Cryptomonas sp. (Douglas and Reith, 1993), and in the cyanelle genome of Cyanophora

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paradoxa (S. E. Douglas and M. Reith, personal communication). Two other R. capsulatus bch genes, bchD and bchO, were also reported to encodeproteins that are components of Mg chelatase (Burke et al., 1992). However, while disruption of the bchD gene caused protoporphyrin IX to accumulate (Bollivar, 1993), disruption of the bchO gene did not (D. W. Bollivar, personal communication). bchD encodes a 60,000 molecular weight protein (Burke et al., 1992). Recently, it was reported that mutations in two R. sphaeroides genes that are similar to R. capsulatus bchD and bchI abolishe Mg chelation as measured in a whole cell assay (Gorchein et al., 1993). It is possible that the bchD and bchI gene products correspond to the two plastid protein fractions needed for Mg chelatase activity, but this possibility awaits confirmation.

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B. Mg-Protoporphyrin Methyltransferase Methylation of the 6-propionic group of Mgprotoporphyrin IX (Fig. 6) is required prior to isocyclic ring formation, because an intermediate in the latter process contains a keto oxygen at a position to the carboxyl group, and unless esterified, the acid would readily decarboxylate (Bogorad, 1960). The enzyme that catalyzes the methylation, S-adenosylL-methionine:Mg-protoporphyrin IX methyltransferase (EC 2.1.1.11), has been isolated and characterized from several plant, algal, and bacterial sources, but not yet from cyanobacteria. The R. sphaeroides enzyme is tightly bound to membranous chromatophores (Hinchigeri et al., 1984), and the wheat methyltransferase is soluble in 0.5 M sucrose and 1.0 M NaCl (Ellsworth et al., 1974). Methyltransferase from different sources appears to have different kinetic reaction mechanisms: the wheat enzyme has a ping-pong mechanism, while the E. gracilis enzyme has a random mechanism and the R. sphaeroides enzyme has an ordered mechanism with Mg-protoporphyrin IX being bound before S-adenosyl-L-methionine (Richards et al., 1987). In R. capsulatus, the bchH gene has tentatively been identified as encoding the methyltransferase (Richards et al., 1991; Burke et al., 1992). The identification is based on the fact that strains carrying mutations of this gene are devoid of in vitro methyltransferase activity, whereas several other bacteriochlorophyll-deficient bch strains do have methyltransferase activity (Richards et al., 1991). However, bchH cells accumulate mainly protoporphyrin IX, and only small amounts of Mgprotoporphyrin IX (Biel and Marrs, 1983; Yang and Bauer, 1990). This anomaly has been explained as indicating that Mg chelatase and methyltransferase are functionally associated enzymes, and disruption of the methyltransferase also inhibits the chelatase reaction. Earlier results had also suggested that the Mg chelatase and methyltransferase steps are obligatorily linked. For example, Mg chelation by EDTApermeabilized R. sphaeroides cells required S-adenosyl-L-methionine and was inhibited by ethionine (Gorchein, 1972). Also, mutation ofthe R. sphaeroides bchH gene abolished not only methyltransferase activity, but also Mg chelation as measured in a whole cell assay (Gorchein et al., 1993). However, in vitro Mg chelatase from pea and cucumber plastids is active in the absence of S-adenosyl-L-methionine (Walker and Weinstein,

1991b; 1992; Lee et al., 1992). This discrepancy suggests that the degree of coupling of the two steps may differ in bacteria and plastids. The protein encoded by The R. capsulatus bchH gene has a molecular weight of 129,000 (Bollivar and Bauer, 1992; Burke et al., 1993).

C. Isocyclic Ring Formation Transformation of the methylated propionate at the 6-position to the isocyclic ring (Fig. 6) was originally postulated to proceed through acrylate, propionate, and intermediates, in analogy to offatty acids (Granick, 1950). The activated carbon of the pionate group then condenses with the bridge carbon of the macrocycle to form the isocyclic ring. This reaction creates a new asymmetric center with an R configuration at position 10 of the product, which is named Mg-2,4-divinylpheoporphyrin or, alternatively, divinylprotochlorophyllide. In contrast to the case of fatty acid where the keto oxygen atom is derived from via hydration of the acrylate intermediate, the keto oxygen in the isocyclic ring is derived from in plants and algae. This was shown by labeling for both protochlorophyllide in ALA-treated etiolated cucumber cotyledons (Walker et al., 1989) and chlorophylls a and b in greening C. vulgaris cells (Schneegurt and Beale, 1992). The fact that the isocyclic ring is formed in the absence of in anoxygenic photosynthetic bacteria indicates that the reaction must proceed by a somewhat different mechanism in these organisms. The location of the enzyme(s) within the intact chloroplast was investigated by the use of permeating and nonpermeating mercurial sulfhydryl reagents (Fuesler et al., 1984). Unlike Mg chelatase, the cyclase system in intact chloroplasts was insensitive to inhibition by nonpermeating reagent, p-chloromercuribenzenesulfonate, but it was inhibited by the permeating relative, p-chloromercuribenzoate, suggesting that the cyclase system is localized within the chloroplast. The cyclase system was active in broken cucumber cotyledon chloroplasts (Chereskin et al., 1982), and the activity was resolved into two required enzymic fractions, one of which is membrane bound and the other soluble (Wong and Castelfranco, 1984). Reconstitution of cyclase activity required both protein fractions, plus Mg-porphyrin substrate, and

Chapter 17 Biosynthesis of Hemes, Chlorophylls and Phycobilins reduced pyridine nucleotide. Activity with Mgprotoporphyrin IX monomethyl ester as substrate was stimulated by inclusion of S-adenosyl-L-methionine (presumably due to remethylation ofde-esterified substrate), and activity with Mg-protoporphyrin IX as substrate required S-adenosyl-L-methionine. The reconstituted cyclase system had an absolute requirement for a reduced pyridine nucleotide. The reconstituted cyclase system was used to test a variety of inhibitors and possible intermediates (Wong and Castelfranco, 1985; Wong et al., 1985; Walker et al., 1988). Although inhibitor studies with intact cucumber cotyledon plastids did not support the involvement of a hemoprotein in an ducing hydroxylation (Chereskin et al., 1982), the reconstituted system from broken plastids was inhibited by and but not by CO, suggesting the involvement of a ferric heme (Whyte and Castelfranco, 1993). The cyclase was inhibited by mercurial sulfhydryl reagents and the sulfhydryl alkylating agent, N-ethylmaleimide. Both the membranous and soluble fractions of the cyclase system were susceptible to inhibition by the latter reagent. The sulfhydryl-containingcompounds, dithiothreitol and mercaptoethanol, were also inhibitory, suggesting the presence of a sensitive disulfide. The chromatographic behavior of the soluble component on affinity columns suggested that this fraction binds the Mg-porphyrin substrate but not NADPH. The membrane-associated fraction appears to bind a required metal ion (Walker et al., 1991). The Mgporphyrin specificity was tested by administration of synthetic derivatives of Mg-protoporphyrin IX monomethyl ester, in which the side-chain at the 6 position of the macrocycle was modified. Both and derivatives (Fig. 6) were effective substrates for Mg-divinylpheoporphyrin formation. However, the acrylate derivative was ineffective. Only one enantiomer of the derivative was effective as substrate. With Mgprotoporphyrin IX monomethyl ester as substrate, an incubation product having the transient kinetic behavior of an intermediate was isolated and identified asthe derivative (Wong et al., 1985). This intermediate was not detected when the derivative was used as a substrate. With the derivative as the substrate, formation of Mg-divinylpheoporphyrin still required the presence of and NADPH as well as both the soluble and membranebound portions of the reconstituted cyclase system. The monovinyl (2-vinyl-4-ethyl) form of the

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derivative was four times more active than the 2,4derivative in the cyclization reaction (Walker et al., 1988). With Mg-protoporphyrin IX as substrate, the monovinyl and divinyl forms were equally effective. The cyclase reaction sequence depicted in Fig. 6 accounts for the known substrate requirements and acceptable in vitro substrates ofthe aerobic reaction, and predicts a stoichiometry of 3 moles each of and 3 NADPH consumed per mole of divinylprotochlorophyllide produced. Because iron deficiency or treatment with Fe chelators decreases the ability of plants to accumulate chlorophyll and causes Mg-protoporphyrin IX monomethyl ester to accumulate (Spiller et al., 1982), it was suggested that the cyclase requires Fe for activity. Although Fe chelators inhibited in vitro cyclase activity from cucumber cotyledon plastids, a dependence on Fe for activity was not directly demonstrated (Walker et al., 1991). A cyclase system was also characterized in isolated wheat etioplasts (Nasrulhaq-Boyce et al., 1987). Many of the properties of the wheat system are similar to those of the cucumber system, but the wheat system has an absolute requirement for organelle intactness and was inhibited by lipophilic Fe chelators and anaerobiosis. Greening ofetiolated tissue for 10 h did not increase or decrease the activity in subsequentlyisolated plastids. The products oftwo R. capsulatus genes within the bch gene cluster have been assigned roles in the cyclase enzyme system. Insertional disruption or point mutation of bchE causes accumulation of Mgprotoporphyrin IX monomethyl ester (Biel and Marrs, 1983; Bollivar, 1993), while disruption of the other gene, called bchM (Burke et al., 1992), causes accumulation ofMg-protoporphyrin IX monomethyl ester plus additional, unidentified pigments (Yang and Bauer, 1990; Bollivar, 1993). Interestingly, when bchE and bchM cells were co-cultured, the mixed culture was able to form bacteriochlorophyll (Bollivar, 1993). This result indicates that each mutant is blocked at a different point in the pathway and that an intermediate which is excreted into the medium by one strain can be transformed to bacteriochlorophyll by the other strain. R. capsulatus bchE and bchM encode proteins of 66,000 and 25,000 molecular weight, respectively (Burke et al., 1992). It is of interestthat the existence of twoR. capsulatus genes associated with the cyclase reaction correlates with the requirement for two plastid protein fractions for in vitro activity. However, it is important to stress

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that, because R. capsulatus is capable of anaerobic isocyclic ring formation, the enzymes catalyzing the reaction, and their encoding genes, may be different from those of plastids (and presumably those of cyanobacteria as well), which perform an oxygenation reaction. For example, as was noted above, the 6-acrylic derivative of Mg-protoporphyrin IX monomethyl ester is not a substrate for the requiring cyclase system from cucumber plastids, which apparently directly oxygenates the 6-propionate. However, the acrylate may turn out to be an intermediate in the anaerobic cyclase reaction, in which the keto oxygen could be derived from by hydration of the acrylic double bond.

D. Vinyl Reduction The step in the chlorophyll biosynthetic pathway at which reduction of the 4-vinyl group occurs has been the subject of extended controversy. The first reports, using etiolated wheat homogenates, indicated that Mg-protoporphyrin IX and its monomethyl ester were the preferred substrates for 4-vinyl reductase (Ellsworth and Hsing, 1973). A later report, using 4-vinyl reductase purified from wheat etioplasts, indicated that the enzyme could use Mg-protoporphyrin IX monomethyl ester, divinylprotochlorophyllide, or divinylchlorophyllide as substrates (Richards et al., 1987). More recently, a 4-vinyl reductase has been described from greening cucumber cotyledon plastids that is specific for divinylchlorophyllide, and no activity was detected with divinylprotochlorophyllide as substrate (Parham and Rebeiz, 1992). However, in that study, the plastids were manipulated by a light flash regime to increase the content of divinylprotochlorophyllide, and only endogenous, membrane-bound pigment was tested as substrate by the membrane-associated 4-vinyl reductase. The possibility exists that all of the protochlorophyllide molecules present in the membranes at the beginning of the experiment were tightly associated with the ring D reducing system, and were not accessible to the 4-vinyl reductase until they were released from the protochlorophyllide reductase after reduction of ring D. It appears that both monovinyl and divinyl protochlorophyllides are suitable substrates for the light-dependent protochlorophyllide reductase enzyme of plastids (Griffiths and Jones, 1975; Knaust et al., 1993), and that etiolated plastids of different

Samuel I. Beale plant species, at different developmental stages, accumulate monovinyl and divinyl protochlorophyllides in different ratios (Carey and Rebeiz, 1985; Carey et al., 1985). What is less clear at present is whether these differences have physiological significance, as has been proposed (Rebeiz et al., 1983). A recent study of the time course of monovinyland divinyl-protochlorophyllide accumulation after a brief light exposure in etiolated wheat leaves and cucumber cotyledons has convincingly shown that divinylprotochlorophyllide behaves as a precursor of monovinylprotochlorophyllide in vivo (Whyte and Griffiths, 1993). It was proposed that broad substrate specificity of 4-vinyl reductase and light-dependent protochlorophyllide reductase, combined with different relative activities of the two reactions in different species and at different developmental stages, can account for results that were previously interpreted in terms of a multi-branched pathway. The alternative routes from divinylprotochlorophyllide and chlorophyllide a are shown in Fig. 7. In cyanobacteria, both monovinyl and divinyl protochlorophyllides exist and are probably biosynthetic precursors of chlorophyll a. Synechococcus sp. strain PCC 6301 cells grown in light of greater than 650 nm wavelength accumulated both monovinyl and divinyl protochlorophyllide in approximately equal quantities, each comprising approximately 5% of total tetrapyrrole pigments, and the cells accumulated less chlorophyll a than did cells grown in white light (Myers et al., 1982). Protochlorophyllide was shown to be present in isolated Synechococcus sp. strain PCC 6301 cell membranes, but the published data do not permit evaluation of whether the monovinyl and/or divinyl forms are present (Peschek et al., 1989a). 4-Vinyl reductase was obtained as a membrane fraction from cucumber cotyledon plastids (Parham and Rebeiz, 1992). In contrast, a soluble enzyme from lysed wheat etioplasts had 4-vinyl reductase activity (Richards et al., 1987). The reaction catalyzed by the wheat enzyme specifically incorporated the (4R)-proton of NADPH into the product. Genetic studies have provided another approach to determining the in vivo substrate for 4-vinyl reductase. Mutation of the bchJ locus of R. capsulatus causes the cells to accumulate divinylprotochlorophyllide, and the bacteriochlorophyll content of the mutant cells is much lower than that of wild-type cells (J. Y. Suzuki, personal communication cited in Bollivar,

Chapter 17 Biosynthesis of Hemes, Chlorophylls and Phycobilins

1993). The most straightforward interpretation of these results is that bchJ encodes 4-vinyl reductase, and that monovinyl protochlorophyllide is the preferred substrate for the ring D reductase enzyme system, with divinylprotochlorophyllide being only poorly converted to divinylchlorophyllide. It has not been determined whether the bacteriochlorophyll present in bchJ cells has a vinyl or ethyl group at position 4 (D. W. Bollivar, personal communication). The R. capsulatus results also suggest that the bchJ gene product is responsible for all or nearly all 4-vinyl reductase activity in the cells. The peptide encoded by bchJ has a predicted molecular weight of 23,000 (Burke et al., 1992).

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E. Ring D Reduction Cyanobacteria, like anoxygenic photosynthetic bacteria and many algae and plants, but unlike angiosperm plants, are able to reduce the porphyrin ring of protochlorophyllide to a chlorin (chlorophyllide) without a requirement for light. Nevertheless, cyanobacteria have retained a second, lightrequiring protochlorophyllide reducing system similar to that of angiosperms. The existence of the lightdependent process in cyanobacteria was revealed in mutant strains in which the light-independent system had been inactivated (Fujita et al., 1992; 1993). Even though the two protochlorophyllide reducing systems have identical tetrapyrrole substrates and products,

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they differ substantially with respect to the identity and number of enzyme protein components.

1. The Light-Dependent Process Light-dependent protochlorophyllide reduction is catalyzed by NADPH:protochlorophyllide oxidoreductase (EC 1.6.99.1). Although the enzyme has not been characterized in cyanobacteria, its existence is inferred from the light requirement for protochlorophyllide reduction in mutant Plectonema boryanum cells in which the light-independent system has been inactivated (Fujita et al., 1992; 1993). NADPH:protochlorophyllide oxidoreductase has been characterized from a number of plant sources, and was originally isolated from the prolamellar bodies of etiolated plastids of grasses such as barley, oat, and rye. In fully mature spinach plastids, the enzyme is associated with the plastid envelope membranes (Joyard et al., 1990). As its name indicates, NADPH: protochlorophyllide oxidoreductase requires NADPH for activity, and it specifically transfers the (4S)-proton of NADPH to protochlorophyllide (Griffiths and Jones, 1975; Yee et al., 1989). As was discussed above, both monovinylprotochlorophyllide and divinylprotochlorophyllide are effective substrates (Griffiths and Jones, 1975; Belanger and Rebeiz, 1980), but protochlorophyll is not (Griffiths, 1974). The purified enzyme from etiolated oat seedlings is composed of 346 amino acids, contains one cysteine and no tryptophan residues, and has a molecular weight of 37,800 (Roper et al., 1987). Enzyme activity was inhibited by the flavin analog, quinacrine, and the activity in various preparations was proportional to the amount of extractable FAD, suggesting that the enzyme is a flavoprotein (Walker and Griffiths, 1988). The lpcr gene that encodes protochlorophyllide reductase has been cloned and sequenced from several plant species including gymnosperms (Spano et al., 1992), and the barley cDNA has been expressed in E. coli to yield active enzyme (Schultz et al., 1989). A C. reinhardtii nuclear gene, named pc-1, has also been identified that is associated with light-dependent chlorophyll synthesis. The mutation was originally produced and screened in y-1 mutant cells that are incapable of chlorophyll formation in the dark (Ford et al., 1981). The y-1, pc-1 double mutants accumulated protochlorophyllide and were unable to form chlorophyll in the light or dark. Membrane preparations from pc-1 cells did not have

Samuel I. Beale NADPH :protochlorophyllide oxidoreductase activity, whereas wild-type cell extracts did (Ford et al., 1983). Cells carrying only the pc-1 mutation formed approximately 36% as much chlorophyll as wild type cells in the light, which indicates that the lightindependent pathway operates in the light, at least in the pc-1 mutants. Surprisingly, pc-1 cells accumulated only 52% as much chlorophyll as wild-type cells in the dark. The depression in the dark chlorophyll level in the mutant suggests that the pc-1 gene product has additional regulatory functions over and above its biosynthetic role of catalyzing light-dependent protochlorophyllide reduction. The concentration of NADPH:protochlorophyllide oxidoreductase in plastids of etiolated angiosperm seedlings decreases precipitously after the seedlings are exposed to light (Mapleston and Griffiths, 1980; Santel and Apel, 1981;Dehesh et al., 1987). In grass seedlings (Apel, 1981; Batschauer and Apel, 1984), but not in seedlings of dicotyledonous plants (Kittsteiner et al., 1990; Benli et al., 1991), the level of mRNA that encodes the enzyme also falls rapidly after exposure of the seedlings to light. The low enzyme level present after the decrease has occurred is still sufficient to catalyze protochlorophyllide reduction at the high rates observed in greening tissues. The initial apparent overabundance of NADPH:protochlorophyllide oxidoreductase in etioplasts suggests that the enzyme may have some photoregulatory function in addition to its biosynthetic role in forming chlorophyllide. In gymnosperms, which contain both the light-dependent and light-independent protochlorophyllide reducing systems, the level of NADPH:protochlorophyllide oxidoreductase declines slowly over a period of 48 h after exposure of the seedlings to light, but the mRNA level remains constant (Spano et al., 1992).

2. The Light-Independent Process The first in vitro description of light-independent protochlorophyllide reduction by a cyanobacterial preparation was by Adamson et al. (1987) who used osmotically-lysed cells of Anabaena flos-aquae. Chlorophyll production was dependent on added protochlorophyllide, but dependence on NADPH was not observed. Peschek et al. (1989a, 1989b) reported lightindependent protochlorophyllide reduction in purified plasma membrane preparations from Synechococcus sp. strain PCC 6301. Activity in the mem-

Chapter 17 Biosynthesis of Hemes, Chlorophylls and Phycobilins branes was inactivated by heating. The reaction was insensitive to illumination, required NADPH, and was reversible in the presence of a high NADP/ NADPH ratio. The reaction demonstrated a requirement for and was inhibited by the -specific chelator EGTA. Recently, two genes were identified in P. boryanum that are required for light-independent protochlorophyllide reduction (Fujita et al., 1992; 1993). Disruption of either gene leads to the identical phenotype, in which the cells accumulate protochlorophyllide and are unable to form chlorophyll in the dark, but do form normal amounts ofchlorophyll in the light. The peptides encoded by these genes are significantly similar to two components of the nitrogenase enzyme system. One of the genes, frxC, bears a strong resemblance to the nifH gene which encodes the Fe-protein of nitrogenase, and the other gene (ORF467) has significant similarity to nifD and nifK, which encode the and subunits of the nitrogenase MoFe-protein. frxC and ORF467 are closely linked in P. boryanum.frxC-like and ORF467like genes have also been detected in the non-nitrogen fixing cyanobacterium, Synechocystis sp. strain PCC 6803 (Ogura et al., 1992). Three genes that are required for light-independent protochlorophyllide reduction have been characterized within the bch gene cluster of R. capsulatus and in the plastid genome of C. reinhardtii. R. capsulatus bchL (Yang and Bauer, 1990; Burke et al., 1992; 1993) and chlL (also called gidB) in the C. reinhardtii plastid genome (Suzuki and Bauer, 1992) are homologous to P. boryanum frxC, which is also present in the plastid genome of the liverwort, Marchantia polymorpha (Ohyama et al., 1986). R. capsulatus bchN and C. reinhardtii plastid chlN (also called gidA) (Choquet et al., 1992) are homologous to ORF465 of the M. polymorpha plastid genome (Ohyama et al., 1986) and P. boryanum ORF467. R. capsulatus bchB (Burke et al., 1993) and C. reinhardtii plastid chlB (Li et al., 1993; Liu et al., 1993) are homologous to the M. polymorpha plastid gene ORF513 (Ohyama et al., 1986), but a homologous gene has not yet been reported in cyanobacteria. chlL and chlN homologs have also been detected in the plastid genomes of conifers, but were not found in the plastid genomes of angiosperms (Lidholm and Gustafsson, 1991). Homologs ofchlB, chlL, and chlNhave also been found on the cyanelle genome of C. paradoxa (D. A. Bryant and V. L. Stirewalt, personalcommunication). In C. reinhardtii,

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as in R. capsulatus, disruption of any of the three genes (chlB, chlL, chlN) abolishes the ability to reduce protochlorophyllide in the dark. Mutations at several nuclear loci also abolish light-independent chlorophyll synthesis in C. reinhardtii, but their roles are unknown (Ford and Wang, 1980). None of the genes, or their encoded peptides that are involved in light-independent protochlorophyllide reduction, have detectable similarity to light-dependent NADPH :protochlorophyllide oxidoreductase. The R. capsulatus bchB, bchL, and bchN genes encode proteins with molecular weights of 57,000, 33,000, and 46,000, respectively (Burke et al., 1992). The fact that the products of the chlB, chlL, and chlN genes and their homologs resemble structural components ofthe nitrogenase system suggests that, like the nitrogen fixation process, light-independent protochlorophyllide reduction uses electrons that are derived from reduced ferredoxin. However, experimental evidence supporting an involvement of ferredoxin in the latter process has not been reported.

F. Phytylation As far as is known, all chlorophylls in cyanobacteria and prochlorophytes are esterifiedwiththe alcohol (2-E,7-R,1l-R)-phytol. Phytylation does not appear to have been studied in cyanobacteria, and the details ofthe process must be inferred from results obtained with plants and anoxygenic photosynthetic bacteria. Phytylation has been studied most extensively in plantplastids. In etioplasts, chlorophyllide is initially esterified with geranylgeraniol (Schoch et al., 1977), and the geranylgeranyl group is then reduced stepwise to dihydro- and tetrahydrogeranylgeranyl and finally to phytyl (hexahydrogeranylgeranyl) (Schoch and Schafer, 1978). The hydrogenation steps require NADPH (Benz et al., 1980) and (Schoch et al., 1980). The pyrophosphate ester of geranylgeraniol is the isoprene substrate of the etioplast isoprenyltransferase enzyme, called chlorophyll synthetase (Rüdiger et al., 1977; 1980). Chlorophyll synthetase was localized in both the prolamellar bodies and the prothylakoid membranes of oat etioplasts (Lütz et al., 1981). Both chlorophyllide a and chlorophyllide b are suitable substrates for chlorophyll synthetase obtained from etiolated oat plastids, but protochlorophyllide is not (Benz and Rüdiger, 1981; Helfrich and Rüdiger, 1992). The enzyme reaction mechanism for bacterio-

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chlorophyllide a phytylation was studied in R. sphaeroides and Rhodospirillum rubrum, where both the bridge and the nonbridge oxygen atoms of the ester were shown to be derived from the propionic acid of the chlorophyllide (Emery and Akhtar, 1985; Ajaz et al., 1985). This result could only occur if the isoprenyl alcohol is the activated group and is consistent with a isoprenyl pyrophosphate substrate. Fe deficiency, which is known to cause a decrease of chlorophyll content (chlorosis) in plants (Spiller et al., 1982), also induces chlorosis in cyanobacteria (Guikema and Sherman, 1983). Kittsteiner et al. (1991) found that treatment of etiolated cress (Lepidium sativum L.) seedlings with Fe chelators did not inhibit protochlorophyllide formation or its photoconversion to chlorophyllide, but it did inhibit the conversion of chlorophyllide to chlorophyll. It is not known whether the effect of the Fe chelators was to decrease the content of chlorophyll synthetase in the treated tissues, or whether the enzyme itself requires Fe for activity. At later stages of greening and in mature plastids, geranylgeranyl pyrophosphate appears to be reduced to phytyl pyrophosphate before it is ligated to chlorophyllide (Rüdiger, 1987). A mutant strain of S. obliquus that is deficient in vitamin E because it cannot form the phytol moiety of the vitamin is also unable to complete chlorophyll synthesis, and instead accumulates near normal amounts of chlorophylls a and b that contain geranylgeraniol instead of phytol (Bishop and Wong, 1974; Henry et al., 1986). Two genes have been identified in the R. capsulatus bch cluster that are involved with bacteriochlorophyllide phytylation. bchG mutants accumulate unesterified bacteriochlorophyllide a (Biel and Marrs, 1983; Bollivar, 1993), and bchP mutants accumulate bacteriochlorophyll a that contains geranylgeranyl instead of a phytyl group (Bollivar, 1993). bchG and bchP encode proteins with molecular weights of 33,400 and 43,000, respectively (Burke et al., 1992).

G. Reaction Center Chlorophylls Pheophytin a (chlorophyll a without the central Mg atom) is the primary electron acceptor of Photosystem II of plants and cyanobacteria (Kobayashi et al., 1988; Maeda and Watanabe, 1992). Because chlorophyll a readily loses its Mg atom under mild acid conditions, it has generally been assumed that

Samuel I. Beale

the pheophytin is derived from chlorophyll a by loss of Mg during incorporation into the Photosystem II reaction center. Chlorophyll a', the of chlorophyll a (Fig. 8), has been proposed as the primary electron donor of Photosystem I in plants and cyanobacteria (Kobayashi et al., 1988; Maeda et al., 1992). Redox state- and light-induced spectral changes were observed when chlorophyll a' was added to a 65,000 molecular weight Photosystem I apoprotein (Watanabe et al., 1987). Chlorophyll a is known to epimerize spontaneously in the presence of nucleophiles. As with the pheophytin in the Photosystem II reaction center, chlorophyll a' is assumed to be formed from chlorophyll a during incorporation into the Photosystem I reaction center.

H. Chlorophyll b Formation Although no cyanobacterial strain has yet been reported to contain chlorophyll b (Fig. 8), this pigment is present in prochlorophytes, which are thought to be related to cyanobacteria (see Chapter 21). Chlorophyll b is universally present in plants and green

Chapter 17 Biosynthesis of Hemes, Chlorophylls and Phycobilins algae and typically comprises approximately 25% of the total chlorophyll present in green plant tissues. In vivo results strongly suggest that chlorophyll b is derived from chlorophyll a or chlorophyllide a (reviewed in Beale and Weinstein, 1991). In vitro experiments attempting to show chlorophyll b formation have mostly yielded equivocal results. However, a preparation was obtained from a S. obliquus pigment mutant catalyzed conversion of chlorophyllide a and chlorophyll a to chlorophyllide b and chlorophyll b, respectively (Kotzabasis and Senger, 1989). Activity was associated with the membranesanddidnotrequirelight. Chlorophyllide b was detected in cell extracts, but not protochlorophyllide b. No further characterization of this in vitro chlorophyll b-forming system has been reported. Bednarik and Hoober (1985a, 1986) made the interesting observation that when degreened cells of C. reinhardtii strain y-1 are incubated in the dark at elevated temperatures (38°C) with o- or m-phenanthroline, a pigment rapidly accumulates and is excreted into the medium. The pigment was characterized as chlorophyllide b. Green cells treated the same way excreted a slightly different pigment that was characterized as divinylchlorophyllide b. Formation ofthese pigments required Cells incubated in the dark at elevated temperature without phenanthroline excreted protochlorophyllide. A membrane fraction was isolated from degreened cells which, when supplemented with phenanthroline, converted exogenous protochlorophyllide to chlorophyllide b (Bednarikand Hoober, 1985b). Although these results are consistent with the possibility that protochlorophyll(ide), rather than chlorophyll(ide) a, is the precursor to chlorophyll b, they are inconsistent with the results of Kotzabasis and Senger (1989), described above, which indicates that chlorophyll(ide) a is the precursor of chlorophyll(ide) b in S. obliquus. It therefore remains to be determined how generally applicable the C. reinhardtii observations are to chlorophyll b biosynthesis. All chlorophyll b-containing organisms are aerobic. This correlation suggests that the formyl oxygen atom ofchlorophyll b is derived from although its derivationfrom is also possible. In an experiment using the formyl oxygen atom of chlorophyll b was shown to be derived from in greening cells of C. vulgaris (Schneegurt and Beale, 1992). The result was confirmed for greening maize leaves (Porra et al., 1993).

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A marine prochlorophyte, Prochlorococcus marinus, was found to contain only the divinyl forms ofchlorophyll a and chlorophyll b (Fig. 8) instead of the normal monovinyl chlorophylls (Goericke and Repeta, 1992). Smaller amounts of a pigment that is probably divinylprotochlorophyllide were also detected. This is the first report ofthe replacement of chlorophylls by their divinyl analogs in “wild-type” organisms. However, a mutant strain ofmaize exists which contains only the divinyl forms ofchlorophylls a and b (Bazzaz et al., 1982; Brereton et al., 1983; Wu and Rebeiz, 1985). The mutant (OliveNecrotic 8147) is necrotic, its leaves contain 70% less chlorophyll than wild-type leaves, and its chlorophyll a-to-b ratio is three times higher, but it can carry out photosynthesis (Bazzaz et al., 1974). It is probable that both the mutant maize strain and P. marinus lack 4-vinyl reductase and that divinyl chlorophyll b is formed from the variant chlorophyll(ide) a.

Note Added in Proof The field of tetrapyrrole pigment biosynthesis is rapidly advancing, and several important new results have appeared after the literature review for this article was completed. Some ofthese new results are briefly described below. The assignment of genes encoding enzymes that catalyze the coproporphyrinogen oxidase and protoporphyrinogen oxidase reactions has recently been clarified. The Bacillus subtilis hemY gene, that was earlier proposed to encode a bifunctional enzyme that catalyzes both reactions, was expressed in Escherichia coli and its product was determined to have protoporphyrinogen oxidase activity but not coproporphyrinogen oxidase activity (Dailey et al., 1994). It was suggested that the Bacillus subtilis gene designation be changed to hemG to correspond with the designation for the Salmonella typhimurium gene encoding protoporphyrinogen oxidase (see below). Salmonella typhimurium contains bothan oxygendependent coproporphyrinogen oxidase, encoded by the hemF gene, and an oxygen-independent enzyme, encoded by the hemN gene (Xu et al., 1992; Xu and Elliott, 1993). The hemF gene product has apredicted molecular weight of 34,400 and is 44% identical to the yeast HEM13 product (Xu and Elliott, 1993). A soybean nuclear gene that encodes a 43,000 molecular weight protein complements a yeast HEM13 mutant (Madsen et al., 1993). The deduced soybean gene

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product is 50% similar to the yeast HEM13 gene product. The product of the S. typhimurium hemN gene has a deduced molecular weight of 52,800 and has no significant similarity to the S. typhimurium hemF product (Xu and Elliott, 1994), but it does have significant similarity to the product of a Rhodobacter sphaeroides gene, also named hemF, that has been proposed to encode an oxygen-independent coproporphyrinogen oxidase (Coomber et al., 1992). For the sake of conformity, it is suggested that if the Rb. sphaeroides hemF gene does in fact encode an oxygenindependent coproporphyrinogen oxidase, it should be renamed hemN. In S. typhimurium the hemG gene was suggested to encode protoporphyrinogen oxidase on the basis of the accumulation of protoporphyrin IX (as well as coproporphyrin III and uroporphyrin III) in hemG mutants (Xu et al., 1992). Protoporphyrinogen oxidase from B. subtilis differs from the enzyme from other sources in that it is a soluble enzyme and it is not inhibited by diphenyl ether herbicides (Dailey et al., 1994). Purified protoporphyrinogen oxidase from bovine liver mitochondria has a native molecular weight of 57,000 and appears to contain an FAD prosthetic group (Siepker et al., 1987). In addition to cytochrome c-heme lyase, yeast cells contain a second yeast heme lyase, cytochromelyase, which is responsible for assembly of cytochrome (Zollner et al., 1992). This protein is encoded by the CYT2 gene, that has been cloned and sequenced. The two yeast heme lyases are 35% identical. The yeast results suggest that each c-type cytochrome may require a different, specific heme lyase for its formation. The biosynthesis of heme a has been further elucidated. The overexpressed E. coli cyoE gene product catalyzes in vitro condensation of ferrous protoheme and farnesyl-pyrophosphate to form heme o (Saiki et al., 1993). In B. subtilis, heme a has been shown to be synthesized from protoheme by the sequential action of two enzymes that are the products of the ctaA and ctaB genes (Svensson et al., 1993). The ctaB gene complements E. coli cyoE mutants. B. subtilis ctaA mutants accumulate heme o instead of heme a. B. subtilis ctaB mutants accumulate neither heme o nor heme a. Interestingly, the B. subtilis ctaA gene, when expressed in E. coli, causes the accumulation of heme a, even though E. coli normally does not produce this heme. These results indicate that ctaB encodes a farnesyltransferase and that ctaA encodes a methyl oxidase or oxygenase, and suggest

that the farnesyl ligation step precedes formyl group formation in heme a biosynthesis. The functions of several genes involved in the steps between protoporphyrin IX and protochlorophyllide have been reassigned. The Rhodobacter capsulatus bchM gene, that was earlier reported to encode a component of the isocyclic ring-forming system, has instead been shown by heterologous expression in E. coli and enzyme assay to encode Sadenosyl-L-methionine:Mg-protoporphyrin IX methyltransferase (Bollivar et al., 1994). The Rb. capsulatus bchH gene, that was earlier considered to encode the methyltransferase, has significant sequence similarity to the cobN-encoded cobaltochelatase that functions in vitamin synthesis in Pseudomonas denitrificans (Debussche et al., 1992). A plant homolog of bchH, oli, is present in Antirrhinum majus, and its mutation results in partial disruption of chlorophyll biosynthesis (Hudson et al., 1993). It is now considered likely that oli and bchH encode a component of the Mg chelatase. In the cobaltochelatase reaction, CobN functions together with two other gene products, CobS and CobT (Cameron et al., 1991; Debussche et al., 1992).It will be of interest to determine whether CobS and CobT have structural or functional similarity to BchD and Bchl. With the reassignments of bchH and bchM, bchE is the only gene that is still assigned to isocyclic ring formation. Acknowledgments The author thanks D. W. Bollivar for critically reading the manuscript and making helpful suggestions, and several individuals who made research results available prior to publication. Research grant support from the National Science Foundation, the Department of Energy, and the U. S. Department of Agriculture is gratefully acknowledged. References Abboud MM and Akhtar M (1976) Stereochemistry of hydrogen elimination in the enzymatic formation of the C-2–C-3 double bond of porphobilinogen. J Chem Soc Chem Commun 1976: 1007–1008 Adamson H, Walker C, Bees A and Griffiths T (1987) Protochlorophyllide reduction in Anabaena. In: Biggins J (ed) Progress in Photosynthesis Research, Vol. IV pp 483-486 Martinus Nijhoff, Dordrecht Ajaz AA, Corina DL and Akhtar M (1985) The mechanism of the

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Samuel I. Beale (1983) Incorporation of haem into phycocyanobilin in levulinic acid treated Cyanidium caldarium. Plant Cell Rep 2: 85–87 Scott AI, Burton G, Jordan PM, Matsumoto H, Fagerness PE and Pryde LM (1980) N.M.R. spectroscopy as a probe for the study of enzyme-catalysed reactions: further observations of preuroporphyrinogen, a substrate for uroporphyrinogen III cosynthetase. J Chem Soc Chem Commun 1980: 384–387 Seehra JS, Jordan PM and Akhtar M (1983) Anaerobic and aerobic coproporpbyrinogen III oxidases of Rhodopseudomonas sphaeroides: mechanism and stereochemistry of vinyl group formation. Biochem J 209: 709–718 Sharif AL, Smith AC and Abell C (1989) Isolation and characterization of a cDNA clone for a chlorophyll synthesis enzyme from Euglena gracilis: the chloroplast enzyme hydroxymethylbilane synthase (porphobilinogen deaminase) is synthesized with a very long transit peptide in Euglena. Eur J Biochem 184: 353–359 Siegel LM (1978) Structure and function of siroheme and the siroheme enzymes. In: Singer TP and Ondarza RN (eds) Developmental Biochemistry, Vol. 1 pp 201–214 Elsevier, Amsterdam Siegel LM, Davis PS and Murphy MJ (1977) Incorporation of methionine-derived methyl groups into sirohaem by Escherichia coli. Biochem J 167: 669–674 Siepker LJ, Ford M, de Kock R and Kramer S (1987) Purification of bovine protoporphyrinogen oxidase: immunological crossreactivity and structural relationship to ferrochelatase. Biochim Biophys Acta 913: 349–358 Smith MA, Kannangara CG and Grimm B (1992) Glutamate 1-semialdehyde aminotransferase: anomalous enantiomeric reaction and enzyme mechanism. Biochemistry 31: 11249– 11254 Spano AJ, He Z and Timko MP (1992) NADPH:protochlorophyllide oxidoreductases in white pine (Pinus strobus) and loblolly pine (P. taeda): evidence for light and developmental regulation of expression and conservation in gene organization and protein structure between angiosperms and gymnosperms. Mol Gen Genet 236: 86–95 Spiller SC, Castelfranco AM and Castelfranco PA (1982) Effects of iron and oxygen of chlorophyll biosynthesis, I: in vivo observations on iron and oxygen-deficient plants. Plant Physiol 69: 107–111 Suzuki JY and Bauer CE (1992) Light-independent chlorophyll biosynthesis: involvement of the chloroplast gene chlL(frxC). Plant Cell 4: 929–940 Svensson B, Lübben M and Hederstedt L (1993) Bacillus subtilis CtaA and CtaB function in haem A biosynthesis. Mol Microbiol 10: 193–201 Swanson RV, Zhou J, Leary JA, Williams T, de Lorimier R, Bryant DA and Glazer AN (1992a) Characterization of phycocyanin produced by cpcE and cpcF mutants and identification of an intergenic suppressor of the defect in bilin attachment. JBiol Chem 267: 16146–16154 Swanson RV, de Lorimier R and Glazer AN (1992b) Genes encoding the phycobilisome rod substructure are clustered on the Anabaena chromosome: characterization of the phycoerythrocyanin operon. J Bacteriol 174: 2640–2547 Sylvers LA, Rogers KC, Shimizu M, Ohtsuka E and Söll D (1993) A 2-thiouridine derivative in is a positive determinant for aminoacylation by Escherichia coli glutamyl-tRNA synthetase. Biochemistry 32: 3836–3841

Chapter 17 Biosynthesis of Hemes, Chlorophylls and Phycobilins Tait GH (1972) Coproporphyrinogenase activities in extracts of Rhodopseudomonas sphaeroides and Chromatium strain D. Biochem J 128: 1159–1169 Taniuchi H, Basile G, Taniuchi M and Veloso D (1983) Evidence for formation of two thioether bonds to link heme to apocytochrome c by partially purified cytochrome c synthetase. J Biol Chem 258: 10963–10966 Thomas SD and Jordan PM (1986) Nucleotide sequence of the hemC locus encoding porphobilinogen deaminase of Escherichia coli K12. Nuc Acids Res 14: 6215–6226 Turner L, Houghton JD and Brown SB (1992) Isolation and partial purification of phycocyanin apoprotein and its role in studies of bilin-apoprotein attachment. Plant Physiol Biochem 30: 309–314 Verkamp E, Jahn M, Jahn D, Kumar AM and Söll D (1992) Glutamyl-tRN A reductase from Escherichia coli and Synechocystis 6803: gene structure and expression. J Biol Chem 267: 8275–8280 Walker CJ and Griffiths WT (1988) Protochlorophyllide reductase: a flavoprotein? FEBS Lett 239: 259–262 Walker CJ and Weinstein JD (1991a) Further characterization of the magnesium chelatase in isolated developing cucumber chloroplasts: substrate specificity, regulation, intactness, and ATP requirements. Plant Physiol 95: 1189–1196 Walker CJ and Weinstein JD (1991b) In vitro assay of the chlorophyll biosynthetic enzyme Mg-chelatase: resolution of the activity into soluble and membrane-bound fractions. Proc Natl Acad Sci USA 88: 5789–5793 Walker CJ, Mansfield KE, Rezzano IN, Hanamoto CH, Smith KM and Castelfranco PA (1988) The magnesium-protoporphyrin IX (oxidative) cyclase system: studies on the mechanism and specificity of the reaction sequence. Biochem J 255: 685–692 Walker CJ, Mansfield KE, Smith KM and Castelfranco PA (1989) Incorporation of atmospheric oxygen into the carbonyl functionality of the protochlorophyllide isocyclic ring. Biochem J 257: 599–602 Walker CJ, Castelfranco PA and Whyte BJ (1991) Synthesis of divinyl protochlorophyllide: enzymological properties of the Mg-protoporphyrin IX monomethyl ester oxidative cyclase system. Biochem J 276,: 691–697 Walker CJ, Hupp LR and Weinstein JD (1992) Activation and stabilization of Mg-chelatase activity by ATP as revealed by a novel in vitro continuous assay. Plant Physiol Biochem 30: 263–269 Wang W-Y, Gough SP and Kannangara CG (1981) Biosynthesis of in greening barley leaves, IV: Isolation of three soluble enzymes required for the conversion of glutamate to Carlsberg Res Commun 46: 243–257 Wang W-Y, Huang D-D, Stachon D, Gough SP and Kannangara CG (1984) Purification, characterization, and fractionation of the acid synthesizing enzymes from lightgrown Chlamydomonas reinhardtii cells. Plant Physiol 74: 569–575 Warren MJ and Jordan PM (1988) Further evidence for the involvement of a dipyrromethene cofactor at the active site of porphobilinogen deaminase. Biochem Soc Trans 16: 963–965 Warren MJ, Roessner CA, Santander PJ and Scott AI (1990) The Escherichia coli cysG gene encodes S-adenosylmethioninedependent uroporphyrinogen III methylase. Biochem J 265: 725–729

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Chapter 18 Carotenoids in Cyanobacteria Joseph Hirschberg and Daniel Chamovitz Department of Genetics, The Hebrew University of Jerusalem, Jerusalem, 91904 Israel

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 lX. Regulation of Carotenoid Biosynthesis and Accumulation References

559 560 560 562 564 565 567 571 572 574 575

Summary Carotenoids in cyanobacteria have two main functions: they serve as light-harvesting pigments in photosynthesis, and they protect against photooxidative damage. Carotenoids are generally hydrophobic isoprenoid compounds that are synthesized in membranes. They mostly accumulate in protein complexes in the photosynthetic membrane, in the cell membrane and in the cell wall. In addition to the prevailing and zeaxanthin, cyanobacteria contain unique ketocarotenoids such as echinenone and canthaxanthin. They do not synthesize and therefore contain but not or carotenes and their oxygenated forms. Glycosylated carotenoids are also very common in cyanobacteria. In the biosynthesis pathway of carotenoids, four enzymes convert geranylgeranyl pyrophosphate of the central isoprenoid pathway to It has been established that phytoene synthase carries out the two-step conversion of geranylgeranyl pyrophosphate to phytoene; phytoene desaturase catalyzes the dehydrogenation of phytoene and phytofluene to produce desaturase produces lycopene via neurosporene; and lycopene cyclase catalyzes two cyclization reactions that produce and Genes encoding several carotenogenic enzymes have been cloned from cyanobacteria. Their analysis has revealed some of the molecular mechanisms involved in carotenoid biosynthesis. The primary structures of the polypeptides are conserved with the homologous enzymes in algae and plants but are distinct from those of other microorganisms. Each of these enzymes is a single-gene produce that is functional in an autonomous manner in heterologous cells. The carotenoid biosynthesis pathway is a target to various ‘bleaching herbicides.’ Resistance to inhibitors of phytoene desaturase was found to be induced by mutations that lead to amino acid substitutions in the enzyme phytoene desaturase. Synthesis and accumulation of carotenoids is regulated by growth conditions and environmental signals. Very little is known about the molecular mechanisms that govern these processes.

D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 559–579. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

560 I. Introduction Carotenoids comprise the largest class of naturally occurring pigments in organisms. More than 640 carotenoids have been identified to date. They are responsible for most of the various shades of yellow, orange and red found in microorganisms, fungi, algae, plants and animals. Carotenoids are synthesized by all photosynthetic organisms as well as several nonphotosynthetic bacteria and fungi, however they are also widely distributed throughout the animal kingdom. Most carotenoids are composed of a hydrocarbon backbone, constructed from eight isoprenoid units and contain a series of conjugated double bonds. Carotenes do not contain oxygen atoms and are either linear or cyclized molecules containing one or two end rings. Xanthophylls are oxygenated derivatives of carotenes. Various glycosylated carotenoids and carotenoid esters have been identified. The backbone can be further extended to give or carotenoids, or shortened yielding apocarotenoids. Some nonphotosynthetic bacteria also synthesize carotenoids. General background on carotenoids can be found in (Goodwin, 1980; Goodwin and Britton, 1988). This chapter focuses on the biochemistry of carotenoids in cyanobacteria. Recent findings on carotenoid biosynthesis and function reveal striking similarities between cyanobacteria andplants, while significant differences exist between cyanobacteria and other microorganisms. Therefore, examples from algae and plants are described to illustrate certain points. II. Functions of Carotenoids In photosynthetic organisms carotenoid pigments serve two major functions: as accessory pigments for light harvesting and in prevention of photooxidative damage. As part of the light-harvesting antenna,

Abbreviations: CAR – carotenoid; CHL – chlorophyll; CPTA – 2-(4-chlorophenylthio)-triethylamine hydrochloride; DMAPP – dimethylallyl pyrophosphate; DMF – dimethylformamide; FPP – farnesyl pyrophosphate; GGPP – geranylgeranyl pyrophosphate; GPP – geranyl pyrophosphate; IPP – isopentenyl pyrophosphate; MPTA – 2-(4-methylphenoxy)-triethylamine hydrochloride; PPPP – prephytoene pyrophosphate; TLC–thinlayer chromatography; HPLC – high-pressure liquid chromatography; UV – ultraviolet

Joseph Hirschberg and Daniel Chamovitz carotenoids can absorb photons and transfer the energy to chlorophyll thus assisting in the harvesting of light in the range of 450–570 nm (Cogdell and Frank, 1987; Cogdell, 1988; Frank et al., 1991; Frank et al., 1992; Cogdell and Gardiner, 1993). Although carotenoids are integral constituents of the protein-pigment complexes of the light-harvesting antennae in photosynthetic organisms, they are also important components of the photosynthetic reaction centers. The identities of the photosynthetically active carotenoproteins and their precise location in lightharvesting systems are not known. Carotenoids in photochemically active chlorophyll-protein complexes of the thermophilic cyanobacterium Synechococcus sp. were investigated by linear dichroism spectroscopy of oriented samples (Breton and Kato, 1987). These complexes contained mainly the carotene pool absorbing around 505 and 470 nm, which is oriented close to the membrane plane. In photochemically inactive chlorophyll-protein complexes, the absorbs around 495 and 465 nm, and the molecules are orientedperpendicular to the membrane plane. Evidence that carotenoids are associated with cyanobacterial Photosystem II (PS II) has been described (Suzuki and Fujita, 1977; Newman and Sherman, 1978). There are two molecules in the reaction center core of PS II (Ohno et al., 1986; Gounaris et al., 1989; Newell et al., 1993) whose exact function(s) is still obscure (reviewed by Satoh, 1992). It was demonstrated that these two coupled carotene molecules protect chlorophyll P680 from photodamage in isolated PS II reaction centers (De Las Rivas et al., 1993), and this may be related to the protection against degradation of the D1 subunit of PS II (Sandmann et al., 1993). The light-harvesting pigments of a highly purified, oxygen-evolving PS II complex of the thermophilic cyanobacterium Synechococcus sp. consists of 50 chlorophyll a and 7 but no xanthophyll, molecules (Ohno et al., 1986). was shown to play a role in the assembly of an active PS II in green algae (Humbeck et al., 1989). Isolated complexes of Photosystem I (PS I) from Phormidium luridum, which contained 40 chlorophylls per P700, contained an average 1.3 molecules of (Thornber et al., 1976). In a preparation of PS I particles from Synechococcus sp. strain PCC 6301, which contained 130 ± 5 molecules of antenna chlorophylls per P700, 16 molecules of carotenoids

Chapter 18 Carotenoids in Cyanobacteria were detected (Lundell et al., 1985). A substantial content of and the xanthophylls cryptoxanthin and isocriptoxanthin were detected in PS I pigment-protein complexes of the thermophilic cyanobacterium Synechococcus elongatus (Coufal et al., 1989). A subunit protein-complex structure of PS I from the thermophilic cyanobacterium Synechococcus sp., which consisted of four polypeptides (of 62, 60, 14 and 10 kDa), contained approximately 10 molecules per P700 (Takahashi et al., 1985). This carotenoid is exclusively bound to the large polypeptides which carry the functional and antenna chlorophyll a. The fluorescence excitation spectrum of these complexes suggested that serves as an efficient antenna for PS I. An additional essential function of carotenoids is to protect against photooxidation processes in the photosynthetic apparatus that are caused by the excited triplet state of chlorophyll. Carotenoid molecules with conjugation of nine or more carbon-carbon double bonds can absorb tripletstate energy from chlorophyll and thus prevent the formation of harmful singlet-state oxygen radicals In Synechococcus sp. the triplet state of carotenoids, was monitored in closed PS II centers and its rise kinetics ofapproximately 25 ns is attributed to energy transfer from chlorophyll triplets in the antenna (Schlodder and Brettel, 1988). It is conceivable that this process, that has a lower yield compared to the yield of radical-pair formation, plays a role in protecting chlorophyll from damage due to over-excitation. The protective role ofcarotenoids in vivo has been elucidated through the use of bleaching herbicides, such as norflurazon (Sandoz 9879), that inhibit carotenoid biosynthesis in all organisms performing oxygenic photosynthesis (reviewed by Sandmann and Böger, 1989). Treatment with norflurazon in the light results in a decrease of both carotenoid and chlorophyll levels, while in the dark, chlorophyll levels alone are unaffected. Inhibition of photosynthetic efficiency in cells of Oscillatoria agardhii that were treated with the pyridinone herbicide, fluridone, was attributed to a decrease in the relative abundance of myxoxanthophyll, zeaxanthin and which in turn caused photooxidation of chlorophyll molecules (Canto de Loura et al., 1987). It has been demonstrated in plants that zeaxanthin is required to dissipate, in a nonradiative manner, the

561 excess excitation energy of the antenna chlorophyll (Demmig-Adams, 1990; Demmig-Adams and Adams, 1990). In algae and plants a light-induced deepoxidation of violaxanthin to yield zeaxanthin, is related to photoprotection processes (reviewed by Demmig-Adams and Adams, 1992). The lightinduced de-epoxidation of violaxanthin and the reverse reaction that takes place in the dark, are known as the ‘xanthophyll cycle’ (Stransky and Hager, 1970; Seifermann-Harms, 1977; Yamamoto, 1979; Hager, 1980; Demmig-Adams and Adams, 1992). Cyanobacterial lichens, that do not contain any zeaxanthin and that probably are incapable of radiationless energy dissipation, are sensitive to high light intensity; algal lichens that contain zeaxanthin are more resistant to high-light stress (DemmigAdams et al., 1990a,b). In contrast to algae and plants, cyanobacteria do not have a xanthophyll cycle. However, they do contain ample quantities of zeaxanthin and other xanthophylls that can support photoprotection of chlorophyll. Several other functions have been ascribed to carotenoids. The possibility that carotenoids protect against damaging species generated by near ultraviolet (UV) irradiation is suggested by results describing the accumulation of in a UVresistant mutant of the cyanobacterium Gloeocapsa alpicola (Buckley and Houghton, 1976). This has been demonstrated more elegantly in E. coli cells that produce carotenoids (Tuveson and Sandmann, 1993). Due to their ability to quench oxygen radical species, carotenoids are efficient anti-oxidants and thereby protect cells from oxidative damage. This function of carotenoids is important in virtually all organisms (Krinsky, 1989; Palozza and Krinsky, 1992). Other cellular functions could be affected by carotenoids, even if indirectly. Although carotenoids in cyanobacteria are not the major photoreceptors for phototaxis, an influence of carotenoids on phototactic reactions, that have been observed in Anabaena variabilis, was attributed to the removal of singlet oxygen radicals that may act as signal intermediates in this system (Nultsch and Schuchart, 1985). A variety carotenoids have important commercial uses. Since carotenoids are non-toxic, they are desirable as coloring agents in the food industry (Bauernfeind, 1981). Furthermore, the flesh, feathers or eggs of fish and birds assume the color of the dietary carotenoid provided, and thus carotenoids are frequently used in dietary additives for poultry

562 and in aquaculture farming. Certain cyanobacterial species, for example Spirulina sp. (Sommer et al., 1990), are cultivated in aquaculture for the production of animal and human food supplements. Consequently, the content of carotenoids, primarily of carotene, in these cyanobacteria has a major commercial implication in biotechnology.

III. Analytical Methods Carotenoids are readily characterized by their specific light absorption properties. Most of the common carotenoids have three major absorption maxima, whose wavelengths increase as the number of conjugated double bonds increases. Phytoene, with three conjugated double bonds (see Fig. 2), has an absorption peak at 289 nm, while the maximally desaturated lycopene, with eleven conjugated double bonds (see Fig. 2), absorbs at 472 nm. Cyclization and oxygenation also affect the absorption maxima. Absorption maxima of typical carotenoids are listed in Table 1. Carotenoids can be extracted from fresh or freezedried material. Methods for the extraction and analysis of carotenoids have been published (Liaaen-Jensen and Jensen, 1971; Davies, 1976; Britton, 1985). Carotenoids are very labile; both oxidation and isomerization can occur post-extraction by contact with acids, heat or light. Hence great care must be taken during extraction and analysis. Optimally all work should be done under an inert atmosphere, for example nitrogen or argon, and in a dimly lit room. Pigment extracts containing chlorophyll are more prone to carotenoid isomerization. Acetone has also been shown to induce carotenoid isomerization in the presence of light. For long storage carotenoids must be kept anaerobically in the dark. Additional precautionary steps are discussed by Britton (1985). In the past thin layer chromatography (TLC) was extensively used for carotenoid analysis. However, high-pressure liquid chromatography (HPLC) is now the method of choice. The advantages of HPLC analysis include higher sensitivity, better resolution, speed and reproducibility. Furthermore, artifactual results are minimized since the separation is achieved in a dark, oxygen-free environment. Numerous methods for carotenoid separation by HPLC have been published (Wright and Shearer, 1984; Ruddat and Will, 1985; Deleenheer and Nelis, 1992; Lesellier

Joseph Hirschberg and Daniel Chamovitz and Tchapla, 1993; Francis et al., 1993; for a recent review of published methods, see Craft, 1992). Application of any of these methods depends on the type of carotenoids that are to be analyzed and the available equipment. Basic analytical techniques, that have been utilized successfully for analyzing the most abundant carotenoids in cyanobacteria, will briefly be described here. For the determination of total colored carotenoid content, it suffices to extract pigments from a cell pellet with either acetone or dimethylformamide (DMF). The latter provides the advantage of being nonvolatile and thus reduces the risk oferrors due to solvent evaporation. Furthermore, carotenoid pigments are highly stable in DMF. Total carotenoids in a DMF extract can be estimated by the following formula: Carotenoid concentration (Chamovitz et al., 1993). An extinction coefficient of is used (Liaaen-Jensen and Jensen, 1971; Goodwin, 1980). Total chlorophyll concentration is measured from the DMF extract: Chlorophyll concentration × 11.92 (Moran, 1982). There are three advantages in using DMF for pigment extraction. Firstly, the DMF extract may be directly injected into the HPLC. Secondly, both carotenoids and chlorophyll a can be directly measured in the same analysis. Thirdly, this procedure avoids ether extractions, so there is little possibility of material loss due to low recovery. However, when using DMF great care must be taken in protecting the HPLC column from phycobiliproteins which may precipitate in the solvent. It is thus recommended to place a prefilter immediately following the injector. This filter must be replaced or washed with water whenever the system pressure rises too high. A disadvantage of injecting the DMF-pigment extract is that with certain columns and solvents the chlorophyll a peak may mask some carotenoids (e.g., lycopene, and some xanthophylls). The carotenoids can be separated from the chlorophyll by saponification. Total pigments are extracted from cyanobacteria with methanol or, if extracted with acetone, dried completely and dissolved in methanol, and treated with 6%

Chapter 18 Carotenoids in Cyanobacteria

563

564 methanolic KOH at 65 °C. An equal volume of saltsaturated water is added and the solution is partitioned against diethyl ether. Carotenoids in the ether phase are recovered by drying under a stream of The dried carotenoids are then dissolved in a small volume of acetone (or another solvent) and immediately injected into the HPLC apparatus. The type of column and elution conditions employed in the HPLC depend on the type of carotenoids that are to be analyzed. In normal-phase HPLC the polar sites of the carotenoid molecules compete with solvent modifiers for adsorption sites on the stationary phase and thus, the more hydrophobic carotenoids (carotenes) elute first, while polar carotenoids (xanthophylls) are retained longer. In reverse-phase HPLC the carotenoids partition between the nonpolar stationary phase and the polar mobile phase such that xanthophylls partition more effectively into the mobile phase and therefore elute first. Either isocratic or gradient elution can be used, and each will have a different separation profile. A sensitive isocratic system which allows the separation of up to fourteen carotenoids has been reported by (Granado et al., 1991). An isocratic system has also been used in analyzing carotenoids from Synechococcus sp. strain PCC 7942 and Synechocystis sp. strain PCC 6803 (Trick et al., 1988; Linden et al., 1990). The solvent acetonitrile: methanol: isopropanol (85:10:5), with a flow rate of in an ODS5 C18 reversed-phase column, provided an excellent separation of zeaxanthin and the two predominant carotenoids in these cyanobacteria. For cyanobacteria containing several types of xanthophylls, the use of a solvent gradient in the HPLC elution is recommended. Although these conditions require longer run times, better resolution and more accurate detection are achieved. Individual carotenoids can be identified by their characteristic absorption spectra and HPLC retention times; these can be determined by using purified standard compounds. IV. Carotenoid Composition in Cyanobacteria Cyanobacteria synthesize the same carotenes as do higher plants; however, they produce some unique types of xanthophylls, such as ketocarotenoids, 2hydroxy-derivatives and glycosides. is normally the only cyclic carotene present, since cyanobacteria do not synthesize end groups

Joseph Hirschberg and Daniel Chamovitz (Hertzberg and Liaaen-Jensen, 1971). The most common carotenoids in cyanobacteria are zeaxanthin, the ketocarotenoid echinenone, and the carotenoid-glycoside myxoxanthophyll (reviewed by Goodwin, 1980). The diketocarotenoid, canthaxanthin, is also widespread in cyanobacteria. While zeaxanthin is also found in all higher plants, echinenone is unique to cyanobacteria. Myxoxanthophyll is also found in some non-photosynthetic and photosynthetic bacteria. The molecular structures of some common cyanobacterial carotenoids are shown in Fig. 1. A list showing the carotenoid compositions of various cyanobacterial strains is given in Table 2. An accurate determination of carotenoid composition is highly dependent on the technique used in the analysis. This was illustrated in the study by Fresnedo et al. (1991), in which pigments were extracted from Phormidium laminosum either in a completely atmosphere or in air. Analysis by both TLC and HPLC of the pigments extracted in the absence of oxygen revealed only two carotenoids, carotene and nostaxanthin. However, exposure to an air-atmosphere during extraction led to the rapid appearance of at least three additional carotenoids: caloxanthin, zeaxanthin and (Fresnedo et al., 1991). This finding calls into question some of the accounts of carotenoid compositions that have been reported in the past. A distinct qualitative difference was found in the carotenoids extracted from fresh or desiccated samples of Nostoc commune (Olie and Potts, 1986). In the fresh samples and echinenone predominated, while in the desiccated samples large amount of canthaxanthin and and were found. In cultures of Spirulina platensis, zeaxanthin and 4-keto-3were found in natural culture from Lake Chad, while laboratory strains of the same species contained Surprisingly, this laboratory strain also produced The quantitative distribution of carotenoids in cyanobacteria varies considerably with the growth conditions, though the predominating pigments remain the same. During exponential-phase growth in a suspension culture, cells of Synechococcus sp. strain PCC 7942 were found to have a carotenoid distribution of 52% and 38% zeaxanthin, with minor amounts of caloxanthin, nostoxanthin and cryptoxanthin (Gombos and Vigh, 1986). A similar ratio was reported for the same strain by Chamovitz et al. (1993). In contrast,

Chapter 18 Carotenoids in Cyanobacteria

565 Synechococcus sp. strain PCC 7002 and Synechococcus sp. with altered carotenoid composition due to defects in carotenoid biosynthesis (van Baalen, 1965; Shestakov and Jevner, 1968; Asato and Folsome, 1969). Recently, mutants of Synechococcus sp. strain PCC 7942 which accumulate phytoene and have an altered carotene/xanthophyll ratio have been described (Linden et al., 1990; Chamovitz et al., 1993).

V. Carotenoproteins: The Cellular Location of Carotenoids

other studies reported zeaxanthin, with substantial amounts of echinenone and myxoxanthophyll, to be the dominant carotenoid in Synechococcus sp. strain PCC 7942 (Linden et al., 1990). Such differences in the carotenoid composition may be attributed to differences in growth conditions and growth stage of the cultures (see section on regulation of Carotenoids synthesis below), or could be the results of the methods of extraction and analysis. There have been several reports on mutants of

Carotenoids are often found in complexes with proteins (Lakshman and Okoh, 1993). Water-soluble (Holt and Krogmann, 1981) as well as detergentsoluble (Bullerjahn and Sherman, 1986) carotenoproteins have been isolated from cyanobacteria. Two membrane-intrinsic, xanthophyll-binding polypeptides with masses of 45 and 42 kDa were identified in Synechococcus sp. strain PCC 7942 (Bullerjahn and Sherman, 1986; Masamoto et al., 1987). The 45 kDa protein was detected in the outer membrane of the cell envelope and the 42 kDa protein was localized in the cytoplasmic membrane. The 42 kDa polypeptide accumulates during growth under high light intensity but is undetectable when grown at low light Masamoto et al., 1987). The cbpA gene, encoding this 42 kDa apoprotein, has been cloned and sequenced (Reddy et al., 1989). It was determined that high-light stress, most likely through the generation of oxygen radicals, induces transcription of cbpA (Reddy et al., 1993). Since novobiocin inhibits cbpA transcription, it is assumed that DNA supercoiling is involved in the regulation of this process. Interestingly, the cbpA gene was independently cloned by another group as a gene (cmpA) that codes for a cytoplasmic membrane polypeptide that is synthesized under carbon-limited growth conditions (Omata et al., 1990a,b). It was noted that this gene is homologous to another Synechococcus sp. strain PCC 7942 gene, nrtA, that encodes a 45 kDa cytoplasmic membrane polypeptide involved in nitrate uptake (Omata et al., 1989). Based on sequence similarity ofcmpA (= cbpA) to nrtA and its location in a cluster of other genes that is homologous to the nrt gene cluster, it was suggested (Omata, 1992) that the 42 kDa polypeptide is part of a transport system that is closely related to the nitrate transport system. Though the protein accumulates

566

under light-stress conditions, it is unlikely that it plays a major role in protecting against high-light damage, since a mutant strain of Synechococcus sp. strain PCC 7942 in which cmpA was inactivated grew quite well under high light intensity (Omata et al., 1989, 1990a). The role of the carotenoid moieties in this carotenoprotein is unknown. Another carotenoid protein has been identified in the aqueous extract of Synechococcus sp. strain PCC 7942 and purified to homogeneity (Diversé-Pierluissi and Krogmann, 1988). Based on absorption spectral properties, it was established that the main carotenoid component was zeaxanthin (83%). This carotenoidprotein complex is a dimer of two 23 kDa polypeptides and approximately 44 carotenoid molecules. A watersoluble, zeaxanthin-binding protein complex associated with the cell surface was purified from Prochlorothrix hollandica (Engle et al., 1991; Reddy et al., 1993). This complex accumulates in highlight-grown cells and is localized to the outer

Joseph Hirschberg and Daniel Chamovitz

membrane. In old suspension cultures the complex is released to the growth medium. The complex is acidic and is composed of two structurally similar polypeptides of 56 and 58 kDa. Its association with lipopolysaccharide in the cell surface indicates that it is a typical surface layer complex. Since it binds most of the zeaxanthin in the cells and is synthesized in high light, it is presumed that it plays a role in photoprotection. In Synechocystis sp. strain PCC 6714 carotenoids were found to be constituents of the outer membrane of the cell wall (Jürgens and Weckesser, 1985; Jürgens et al., 1985). Myxoxanthophyll and related carotenoid-glycosides, zeaxanthin, echinenone and carotene were identified as true constituents of the outer membrane. Using linear dichroism spectroscopy it was shown that, in Synechocystis sp. strain PCC 6714 and Synechococcus sp. strain PCC 6307, the predominant orientation of the carotenoids in the outer membrane is perpendicular to membrane plane

Chapter 18 Carotenoids in Cyanobacteria

(Jürgens and Mäntele, 1991). It was calculated that each carotenoid molecule spans one monolayer of the outer membrane, and the sugar moiety of the carotenoid-glycoside, myxoxanthophyll, serves as the polar head. Myxoxanthophyll and related carotenoid-glycosides,andzeaxanthin,arethemajor carotenoids of the cell wall in Synechocystis sp. strain PCC 6714; echinenone and are minor components (Jürgens and Weckesser, 1985). VI. Biosynthesis of Carotenoids The biosynthesis of carotenoids has been reviewed extensively in recent years (Spurgeon and Porter, 1980; Yokoyama et al., 1982; Britton, 1988; Kleinig, 1989; Goodwin, 1993). There are numerous

567

similarities in carotenogenesis of cyanobacteria and plants; hence, examples fromplants will be discussed as well. Carotenoids are produced from the general isoprenoid biosynthetic pathway (Fig. 2). While this pathway has been known for several decades, only recently, and mainly through the use of genetics and molecular biology, have some of the molecular mechanisms involved been elucidated. This is due to the fact that most of the enzymes which take part in the conversion of phytoene to carotenes and xanthophylls are labile, membrane-associated proteins that lose activity upon solubilization (Beyer et al., 1985; Bramley, 1985). However, solubilization of carotenogenic enzymes from Synechocystis sp. strain PCC 6714 that retain partial activity has been

568

reported (Bramley and Sandmann, 1987). There is no genuine in vitro system for carotenoid biosynthesis which enables a direct essay of enzymatic activities. A cell-free carotenogenic system has been developed (Clarke et al., 1982) and adapted for cyanobacteria (Sandmann and Bramley, 1985; Bramley and Sandmann, 1985). Reconstitution of phytoene

Joseph Hirschberg and Daniel Chamovitz

desaturase from Synechococcus sp. strain PCC 7942 in liposomes was achieved following purification of the polypeptide, that had been expressed in E. coli (Fraser et al., 1993). Carotenoids are synthesized from isoprenoid precursors. The central pathway of isoprenoid biosynthesis may be viewed as beginning with the

Chapter 18 Carotenoids in Cyanobacteria conversion of acetyl-CoA to mevalonic acid, isopentenyl pyrophosphate (IPP), a molecule, is formed from mevalonate and is the building block for all long-chain isoprenoids. Following isomerization of IPP to dimethylallyl pyrophosphate (DMAPP), three additional molecules of IPP are combined to yield the molecule, geranylgeranyl pyrophosphate (GGPP). These 1'-4 condensation reactions are catalyzed by prenyl transferases (Kleinig, 1989). There is evidence in plants that the same enzyme, GGPP synthase, carries out all the reactions from DMAPP to GGPP (Dogbo and Camara, 1987; Laferrière and Beyer, 1991). The first step that is specific for carotenoid biosynthesis is the head-to-head condensation of two molecules of GGPP to produce prephytoene pyrophosphate (PPPP). Following removal of the pyrophosphate, GGPP is converted to 15-cisphytoene, a colorless hydrocarbon molecule. This two-step reaction is catalyzed by the soluble enzyme, phytoene synthase, an enzyme encoded by single genes (crtB) in both cyanobacteria and plants (see Table 3; Chamovitz et al., 1992; Ray et al., 1992; for recent review on phytoene synthase, see Camara, 1993). All the subsequent steps in the pathway occur in membranes. Four desaturation (dehydrogenation) reactions convert phytoene to lycopene via phytofluene, and neurosporene. Each desaturation increases the number of conjugated double bonds by two such that the number of conjugated double bonds increases from three in phytoene to eleven in lycopene.

569 Relatively little is known about the molecular mechanism of the enzymatic dehydrogenation of phytoene (Jones and Porter, 1986; Beyer et al., 1989). It has been established that in cyanobacteria, algae and plants the first two desaturations, from 15-cisphytoene to are catalyzed by a single membrane-bound enzyme, phytoene desaturase (Jones and Porter, 1986; Beyer et al., 1989). Since the product is mostly in the all-trans configuration, a cis-trans isomerization is presumed at this desaturation step. The primary structure of the phytoene desaturase polypeptide in cyanobacteria is conserved (over 65% identical residues) with that of algae and plants (Pecker et al., 1992; Pecker et al., 1993). Moreover, the same inhibitors block phytoene desaturase in the two systems (Sandmann and Böger, 1989). Consequently, it is very likely that the enzymes catalyzing the desaturation of phytoene and phytofluene in cyanobacteria and plants have similar biochemical and molecular properties, that are distinct from those of phytoene desaturases in other microorganisms. One such a difference is that phytoene desaturases from Rhodobacter capsulatus, Erwinia sp. or fungi convert phytoene to neurosporene, lycopene, or 3,4-dehydrolycopene, respectively (Fig. 3). Desaturation of phytoene in daffodil chromoplasts (Beyer et al., 1989), as well as in a cell free system of Synechococcus sp. strain PCC 7942 (Sandmann and Kowalczyk, 1989), is dependent on molecular oxygen as a possible final electron acceptor, although oxygen is not directly involved in this reaction. A mechanism

570

of dehydrogenase-electron transferase was supported in cyanobacteria over dehydrogenation mechanism of dehydrogenase-monooxygenase (Sandmann and Kowalczyk, 1989). A conserved FAD-binding motif exists in all phytoene desaturases whose primary structures have been analyzed (Pecker et al., 1992; Pecker et al., 1993). The phytoene desaturase enzyme in pepper was shown to contain a protein-bound FAD (Hugueney et al., 1992). Since phytoene desaturase is located in the membrane, an additional, soluble redox component is predicted. This hypothetical component could employ as suggested (Mayer et al., 1992) or another electron and hydrogen

Joseph Hirschberg and Daniel Chamovitz

carrier, such as a quinone. The cellular location of phytoene desaturase in Synechocystis sp. strain PCC 6714 and Anabaena variabilis strain ATCC 29413 was determined with specific antibodies to be mainly (85%) in the photosynthetic thylakoid membranes (Serrano et al., 1990). In cyanobacteria algae and plants is converted to lycopene via neurosporene. Very little is known about the enzymatic mechanism, which is predicted to be carried out by a single enzyme (Linden et al., 1993). The deduced amino acid sequence of desaturase in Anabaena sp. strain PCC 7120 contains a dinucleotide-binding motif that is

Chapter 18 Carotenoids in Cyanobacteria similar to the one found in phytoene desaturase (Linden et al., 1994). Two cyclization reactions convert lycopene to carotene. Evidence has been obtained that in Synechococcus sp. strain PCC 7942 (Cunninghamet al., 1993), as well as in plants (Camara and Dogbo, 1986), these two cyclizations are catalyzed by a single enzyme, lycopene cyclase. This membranebound enzyme is inhibited by the triethylamine compounds, CPTA and MPTA (Sandmann and Böger, 1989). Cyanobacteria carry out only the and therefore do not contain and and their oxygenated derivatives. The is formed through the formation of a ‘ carbonium ion’ intermediate when the C-1,2 double bond at the end of the linear lycopene molecule is folded into the position of the C-5,6 double bond, followed by a loss of a proton from C-6. No cyclic carotene has been reported in which the 7,8 bond is not a double bond. Therefore, full desaturation as in lycopene, or desaturation of at least half-molecule as in neurosporene, is essential for the reaction. Cyclization of lycopene involves a dehydrogenation reaction that does not require oxygen. The cofactor for this reaction is unknown. A dinucleotide-binding domainwas found in the lycopene cyclase polypeptide of Synechococcus sp. strain PCC 7942 (F.X. Cunningham, personal communication), implicating NAD(P) orFAD as coenzymes with lycopene cyclase. The addition of various oxygen-containing side groups, such as hydroxy-, methoxy-, oxo-, epoxy-, aldehyde or carboxylic acidmoieties, form the various xanthophyll species. Little is known about the formation of xanthophylls. Hydroxylation of carotene requires molecular oxygen in a mixedfunction oxidase reaction.

VII. Molecular Characterization of Carotenoid Biosynthesis A molecular description of carotenogenesis in cyanobacteria and plants has been lacking for long time. This has probably been due to biochemical difficulties in obtaining purified enzymes that obstructed in turn the cloning of genes encoding carotenogenic enzymes by conventional approaches. Clusters of genes encoding the enzymes for the entire pathway have been cloned from the purple photosynthetic bacterium Rhodobacter capsulatus (Armstrong et al., 1989) and from the nonphoto-

571 synthetic bacteria Erwinia herbicola (Sandmann et al., 1990; Hundle et al., 1991; Schnurr et al., 1991) and Erwinia uredovora (Misawa et al., 1990). Two genes, al-3 for GGPP synthase (Nelson et al., 1989; Carattoli et al., 1991) and al-1 for phytoene desaturase (Schmidhauser et al., 1990) have been cloned from the fungus Neurospora crassa. However, attempts at using these genes as heterologous molecular probes to clone the corresponding genes from cyanobacteria or plants were unsuccessful due to lack of sufficient sequence similarity (see below). One report on using this approach to clone the phytoene desaturase gene from Synechocystis sp. strain PCC 6714 turned out to be in error (Schmidt and Sandmann, 1990). The first ‘plant-type’ genes for carotenoid synthesis enzyme were cloned from cyanobacteria using a molecular-genetics approach. In the first step towards cloning the gene for phytoene desaturase, a number of mutants that are resistant to the phytoenedesaturase-specific inhibitor, norflurazon, were isolated in Synechococcus sp. strain PCC 7942 (Linden et al., 1990). The gene conferring norflurazon-resistance was then cloned by transforming the wild-type strain to herbicide resistance (Chamovitz et al., 1990; Chamovitz et al., 1991). Several lines of evidence indicated that the cloned gene, formerly called pds and now named crtP (see Table 3), codes for phytoene desaturase. The most definitive one was the functional expression of phytoene desaturase activity in transformed E. coli cells (Linden et al., 1991; Pecker et al., 1992). The crtP gene was also cloned from Synechocystis sp. strain PCC 6803 by similar methods (Martinez-Ferez and Vioque, 1992). The cyanobacterial crtP gene was subsequently used as a molecular probe for cloning the homologous gene from an alga (Pecker et al., 1993) and higher plants (Hartley et al., 1991; Pecker et al., 1992). The phytoene desaturases in Synechococcus sp. strain PCC 7942 and Synechocystis sp. strain PCC 6803 consist of 474 and 467 amino acid residues, respectively, whose sequences are highly conserved (74% identities and 86% similarities). The calculated molecular mass is 51 kDa and, although it is slightly hydrophobic (hydropathy index –0.2), it does not include a hydrophobic region which is long enough to span a lipid bilayer membrane. The primary structure of the cyanobacterial phytoene desaturase is highly conserved with the enzyme from the green alga Dunalliela bardawil (61% identical and 81% similar; Pecker et al., 1993) and from tomato (Pecker et al., 1992), pepper (Hugueney et al., 1992) and

572 soybean (Bartley et al., 1991) (62–65% identical and ~79% similar; see Chamovitz, 1993). The eucaryotic phytoene desaturase polypeptides are larger (64 kDa); however, they are processed during import into the plastids to mature forms whose sizes are comparable to those of the cyanobacterial enzymes. There is a high degree of structural similarity in carotenoid enzymes of Rhodobacter capsulatus, Erwinia sp. and Neurospora crassa (reviewed in Armstrong et al., 1993), including in the crtI geneproduct, phytoene desaturase. As indicated above, a high degree of conservation of the primary structure of phytoene desaturases also exists among oxygenic photosynthetic organisms. However, there is little sequence similarity, except for the FAD binding sequences at the amino termini, between the ‘planttype’ crtP gene products and the ‘bacterial-type’ phytoene desaturases (crtI gene products; 19–23% identities and 42–47% similarities). It has been hypothesized that crtP and crtI are not derived from the same ancestral gene and that they originated independently through convergent evolution (Pecker et al., 1992). This hypothesis is supported by the different dehydrogenation sequences that are catalyzed by the two types of enzymes and by their different sensitivities to inhibitors. Although not as definite as in the case of phytoene desaturase, a similar distinction between cyanobacteria and plants on the one hand and other microorganisms is also seen in the structure of phytoene synthase. The crtB gene (formerly psy) encoding phytoene synthase was identified in the genome of Synechococcus sp. strain PCC 7942 adjacent to crtP and within the same operon (Bartley et al., 1991). This gene encodes a 36 kDa polypeptide of 307 amino acids with a hydrophobic index of –0.4. The deduced amino acid sequence of the cyanobacterial phytoene synthase is highly conserved with the tomato phytoene synthase (57% identical and 70% similar; Ray et al., 1987) but is less highly conserved with the CrtB sequences from other bacteria (29–32% identical and 48–50% similar with ten gaps in the alignment). Both types of enzymes contain two conserved sequence motifs also found in prenyl transferases from diverse organisms (Bartley et al., 1991; Carattoli et al., 1991; Armstrong et al., 1993; Math et al., 1992; Chamovitz, 1993). It is conceivable that these regions in the polypeptide are involved in the binding and/or removal of the pyrophosphate during the condensation of two GGPP molecules.

Joseph Hirschberg and Daniel Chamovitz The crtQ gene encoding desaturase (formerly zds) was cloned from Anabaena sp. strain PCC 7120 by screening an expression library of cyanobacterial genomic DNA in cells of E. coli carrying the Erwinia sp. crtB and crtE genes and the cyanobacterial crtP gene (Linden et al., 1993). Since these E. coli cells produce brownish-red pigmented colonies that produced lycopene could be identified on the yellowish background of cells producing The predicted desaturase from Anabaena sp. strain PCC 7120 is a 56 kDa polypeptide which consists of 499 amino acid residues. Surprisingly, its primary structure is not conserved with the ‘plant-type’ (crtP gene product) phytoene desaturases, but it has considerable sequence similarity to the bacterial-type enzyme (crtI gene product) (Sandmann, 1993). It is possible that the cyanobacterial crtQ gene and crtI gene of other microorganisms originated in evolution from a common ancestor. The crtL gene for lycopene cyclase (lcy) was cloned from Synechococcus sp. strain PCC 7942 utilizing essentially the same cloning strategy as for crtP . By using an inhibitor of lycopene cyclase, 2(4-methylphenoxy)-triethylamine hydrochloride (MPTA), the gene was isolated by transformation of the wild-type to herbicide-resistance (Cunningham et al., 1993). Lycopene cyclase is the product of a single gene product and catalyzes the double cyclization reaction of lycopene to The crtL gene product in Synechococcus sp. strain PCC 7942 is a 46 kDa polypeptide of 411 amino acid residues. It has no sequence similarity to the crtY gene product (lycopene cyclase) from Erwinia uredovora or E. herbicola (F.X. Cunningham, unpublished results). VIII. Inhibitors of Carotenoid Biosynthesis The carotenoid biosynthetic pathway is the target for a group of inhibitors known also as ‘bleaching herbicides’ (reviewed by (Sandmann and Böger, 1989). The most susceptible target in the pathway is phytoene desaturase, which is inhibited by a large number of chemically unrelated compounds. The oldest and most intensively studied are the phenylpyridazinones, as exemplified by norflurazon [SAN 9879 = 4-chloro-5-methylamino-2-(3trifluoromethylphenyl)-pyridazin-3(2H)one]. Others include: fluridone = 1-methyl-3-phenyl-5-(3-

Chapter 18 Carotenoids in Cyanobacteria trifluoromethylphenyl)-4(1H)-pyridone and fluorochloridone [R 40244= 1-(m-trifluoromethylphenyl)3-chloro-4-chloromethyl-2-pyrrolidone] (Bartels and Watson, 1992). These inhibitors have no effect on phytoene desaturation in organisms that do not perform oxygenic photosynthesis, such as Rhodobacter capsulatus or Erwinia uredovora. Early reports on the mode of action of norflurazon pointed out that the decrease in the levels of carotenoids and chlorophylls, together with the concurrent accumulation of phytoene is a common feature of this type of chlorosis-inducing herbicide (Urbach et al., 1976; Bartels and Watson, 1992). Through the use of cellfree assays using daffodil chromoplasts (Beyer et al., 1980) or photosynthetic membranes from the cyanobacterium Synechocystis sp. strain PCC 6803 (Clarke et al., 1982), it has been shown that phytoene accumulation originates from a direct interference of norflurazon with phytoene desaturase activity. The herbicidal effects of the other phytoene desaturase inhibitors are similar (Clarke et al., 1985; Wakabayashi et al., 1988; Sandmann et al., 1992; Kowalczyk-Schröder and Sandmann, 1992; Babczinski et al., 1992). Studies using membranes form the cyanobacterium Synechococcus sp. strain PCC 7942 have shown that norflurazon is a noncompetitive reversible inhibitor of phytoene desaturase (Clarke et al., 1985; Sandmann et al., 1989; Sandmann et al., 1992); however, its exact mode of interaction with the enzyme is unknown. Several mutations have been detected in crtP from herbicide-resistant strains of Synechococcus sp. strain PCC 7942 (Linden et al., 1990; Chamovitz et al., 1990; Chamovitz et al., 1993). Each of the mutant strains displayed a unique cross-resistance profile to a number of phytoene desaturase inhibitors. Most of them exhibited a considerable degree of crossresistance to the N-phenyl furone, fluorochloridone. For four mutants it was established that the resistance phenotype resulted from amino acid substitutions at positions 195, 320, 403 and 439 in phytoene desaturase, and that these changes in the polypeptide affected the enzyme-inhibitor interaction (Chamovitz et al., 1993). Since each of the mutations conferred resistance to herbicides with significantly different chemical structures, it was proposed that the different inhibitors have overlapping, but distinct binding sites on the enzyme. Interestingly, an altered amino acid at residue 195 of phytoene desaturase was also found in a norflurazon-resistant mutant of Synechocystis sp. strain PCC 6803 (Martinez-Ferez and Vioque, 1992).

573 In addition to conferring herbicide resistance, each of the point mutations detected in the gene crtP from Synechococcus sp. strain PCC 7942 also reduced, to varying extent, the enzymatic activity of phytoene desaturase. This reduction in enzymatic activity is expressed phenotypically in an increase of the steadystate levels of phytoene and a decrease in total colored carotenoid content, mostly expressed in reduction of xanthophylls (Chamovitz et al., 1993). Consequently, it was deduced that phytoene desaturation is a ratelimiting step in carotenogenesis in cyanobacteria. Since all of the mutations in crtP which lead to herbicide resistance also cause a reduction in the efficiency of the phytoene desaturation reaction, it was speculated that the herbicide binding sites and the catalytic site for desaturation of phytoene are either overlapping or in close proximity to each other on the phytoene desaturase enzyme. There are few inhibitors of desaturase. Two compounds, J 852 = 4-(3-methyl-propoxy)-2isopropylamino-6-methyl pyrimidine and LS 80707 = ethyl-cis-5-methyl-6-ethyl-2-phenyl-5,6-dihydropyran-4-one-3-carboxylate, inhibit the enzyme in Synechocystis sp. strain PCC 6714 (Sandmann et al., 1985; Sandmann and Böger, 1989). Direct interaction of the latter herbicide with the enzyme was demonstrated in a cell-free system from Synechocystis sp. strain PCC 6714 (Sandmann et al., 1985). A number of substituted triethylamine compounds inhibit the formation of cyclic carotenoids in plants, algae, and cyanobacteria and result in the accumulation of lycopene (reviewed by Sandmann and Böger, 1989). Particularly effective inhibitors ofthis class are the compounds CPTA = 2-(4-chlorophenylthio)-triethylamine hydrochloride and MPTA =2-(4-methylphenoxy)-triethylamine hydrochloride (Yokoyama et al., 1982; Cunningham, 1985). Experiments using a cell-free system from the cyanobacterium Synechocystis sp. strain PCC 6714 indicated that CPTA is an effective inhibitor of lycopene cyclization in vitro and that it inhibits the cyclization reaction in a noncompetitive manner (see Sandmann and Böger, 1989). The target site of MPTA and related compounds is the enzyme lycopene cyclase itself. This is demonstrated by the fact that a mutation in crtL caused MPTA-resistance in Synechococcus sp. strain PCC 7942 and that the enzyme was inhibited by MPTA when expressed in E. coli (Cunningham et al., 1993).

574

IX. Regulation of Carotenoid Biosynthesis and Accumulation Carotenoid biosynthesis is a regulated process. Various environmental and developmental factors control carotenoid synthesis and accumulation in procaryote as well as in eucaryotes (Bramley and Mackenzie, 1988). Changes in carotenoid content and composition, which occur in response to light intensity and light quality, are one of the adaptation mechanisms to high-light that exist in acclimatized cyanobacteria. The protective role of carotenoids in cyanobacteria against high light has been described (Kellar and Paerl, 1980; Codd, 1981). A strong indication to the importance of carotenoid content in photoprotection is provided by the finding that mutant cells of Synechococcus sp. strain PCC 7942, which contained less carotenoids, were more sensitive to photoinhibition (Sandmann et al., 1993). High-lightgrown cells of Spirulina platensis are more resistant to photoinhibition of photosynthesis, and have a better potential to recover from it under suitable reactivating conditions, than low-light grown cells (Shyam and Sane, 1989). Acclimation to high light, which reduces susceptibility to photoinhibition in Microcystis aeruginosa (Zevenboom and Mur, 1984), is associated with a decrease in the cell chlorophyll to carotenoid ratio (Raps et al., 1983), in addition to a decline in both chlorophyll and phycobiliprotein contents. In Synechococcus sp. strain PCC 7942 the proportion of the xanthophyll zeaxanthin increased with increasing light intensities (Goodall and Filipowicz, 1989), while in shade-adapted cells grown in low light, a reduced carotenoid content was detected (König, 1987). Different results were observed in Synechococcus sp. strain WH 7803 in which carotene decreased from 0.68 to with increasing light intensity and maintained a constant molar ratio with chlorophyll a (0.24 1 Chl a); on the other hand, the concentration of zeaxanthin remained unchanged at (Kana et al., 1988). An increase in the relative abundance of myxoxanthophyll and zeaxanthin, and a decrease in the relative abundance of echinenone and were observed in Oscillatoria agardhii following exposure to high intensity of white light (Millie et al., 1990). Maximum total carotenoid production of cell cultures of Spirulina platensis was measured under white light at an irradiance of which

Joseph Hirschberg and Daniel Chamovitz is the onset of light saturation for this organism (Olaizola and Duerr, 1990). However, the carotenoid content in the cells could increase at higher intensities of white light. When cells were grown under white light, and echinenone were most abundant at very low or very high irradiance levels, while myxoxanthophyll and zeaxanthin did not change. When cells were grown under red and blue light, the myxoxanthophyll content decreased and the levels of other carotenoids showed only small changes. Carotenoid composition in Synechococcus sp. strain WH 7803 cells is affected by light quality. The ratio of to chlorophyll a was constant when cells were grown in daylight, white or bluegreen light (Bidigare et al., 1989). However, zeaxanthin was two-fold higher in cells grown in blue-green light than in those grown under white light or daylight illumination. Other growth conditions may also affect carotenoid content. An increase in carotenoid content, with a concomitant depletion in phycocyanin has been observed in Pseudanabaena sp. strain M2 and Oscillatoria splendida following nitrogen deficiency during growth (Canto de Loura et al., 1987). In contrast, carotenoids were found to be relatively stable following nitrogen starvation or photobleaching that caused breakdown of chlorophyll and phycobiliproteins (Duke and Allen, 1990; Millie et al., 1990). A 42 kDa carotenoid protein in Synechocystis sp. strain PCC 6803, which is immunologically crossreactive with a similar size polypeptide in Synechococcus sp. strain PCC 7942, was conserved in membranes of nitrogen-starved cells (Duke and Allen, 1990). Changes of salinity from 0.012 M to 0.50 M NaCl in the growth medium altered carotenoid content in the freshwater cyanobacterium Synechococcus sp. strain PCC 6311 (Khomutov et al., 1990). Finally, a thermophilic Synechococcus sp., whose optimal temperature for growth is 58 °C, responded to a lowered growth temperature (38 °C) by increasing the proportion of unsaturated fatty acids in the membrane lipids as well as increased amounts of the carotenoid, myxoxanthophyll, that does not seem to be involved in energy transfer reactions (Miller et al., 1987). At present, the molecular mechanisms underlying the regulation of carotenoid biosynthesis remain obscure. The recent success in cloning genes for carotenogenic enzymes in cyanobacteria provide unique tools to examine these processes at the molecular level. Knowledge of the biochemistry of

Chapter 18 Carotenoids in Cyanobacteria carotenogenesis should now advance rapidly. The future availability of additional genes encoding enzymes involved in xanthophyll synthesis (e.g., enzymes for astaxanthin biosynthesis, important in coloring salmon flesh), should provide unique materials for biotechnological applications.

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Chapter 18 Carotenoids in Cyanobacteria Hugueney P, Römer S, Kuntz M and Camara B (1992) Characterization and molecular cloning of a flavoprotein catalyzing the synthesis of phytofluene and zeta-carotene in Capsicum chromoplasts. Eur J Biochem 209: 399–407 Humbeck K, Römer S and Senger H (1989) Evidence for the essential role of carotenoids in the assembly of an active Photosystem II. Planta 179: 242–250 Hundle BS, Beyer P, Kleinig H, Englert H and Hearst JE (1991) Carotenoids of Erwinia herbicola and an Escherichia coli HB101 strain carrying the Erwinia herbicola carotenoid gene cluster. Photochem Photobiol 54: 89–93 Jones BL and Porter JW (1986) Biosynthesis of carotenes in higher plants. CRC Crit Rev Plant Sci 3: 295–324 Jürgens UJ and Mäntele W (1991) Orientation of carotenoids in the outer membrane of Synechocystis PCC 6714 (cyanobacteria). Biochim Biophys Acta 1067: 208–212 Jürgens UJ and Weckesser J (1985) Carotenoid-containing outer membrane of Synechocystis sp. strain PCC6714. J Bacteriol 164: 384–389 Jürgens UJ, Golecki JR and Weckesser J (1985) Characterization of the cell wall of the unicellular cyanobacterium Synechocystis PCC 6714. Arch Microbiol 142: 168–174 Kana TM, Glibert PM, Goericke R and Welschmeyer NA (1988) Zeaxanthin and in Synechococcus WH7803 respond differently to irradiance. Limnol Oceanogr 33: 1623–1627 Kellar PE and Paerl HW (1980) Physiological adaptation in response to environmental stress during an Anabaena Bloom. Appl Environ Microbiol 40: 587–595 Khomutov G, Fry IV, Huflejt ME and Packer L (1990) Membrane lipid composition, fluidity and surface charge changes in response to growth of the fresh water cyanobacterium Synechococcus 6311 under high salinity. Arch Biochem Biophys 277: 263–267 Kleinig H (1989) The role of plastids in isoprenoid biosynthesis. Ann Rev Plant Physiol Plant Mol Biol 40: 39–59 Kowalczyk-Schröder S and Sandmann G (1992) Interference of fluridone with the desaturation of phytoene by membranes of the cyanobacterium Aphanocapsa. Pestic Biochem Physiol 42: 7–12 König F (1987) A role of the binding protein in the mechanism of cyanobacterial adaptation to light intensity? Z Naturforsch 42c: 727–732 Krinsky NI (1989) Antioxidant functions of carotenoids. Free Radical Biol Med 7: 617–635 Laferrière A and Beyer P (1991) Purification of geranylgeranyl diphosphate synthase from Sinapis alba etioplasts. Biochim Biophys Acta 216: 156–163 Lakshman MR and Okoh C (1993) Carotenoid - protein complexes. Meth Enzymol 214: 74–86 Lesellier E and Tchapla A (1993) Analysis of carotenoids by high-performance liquid chromatography and supercritical fluid chromatography. J Chromatogr 63: 9–23 Liaaen-Jensen S and Jensen A (1971) Quantitative determination of carotenoids in photosynthetic tissues. Meth Enzymol 23:586– 602 Linden H, Sandmann G, Chamovitz D, Hirschberg J and Böger P (1990) Biochemical characterization of Synechococcus mutants selected against the bleaching herbicide norflurazon. Pestic Biochem Physiol 36: 46–51 Linden H, Misawa N, Chamovitz D, Pecker I, Hirschberg J and Sandmann G (1991) Functional complementation in

577 Escherichia coli of different phytoene desaturase genes and analysis of accumulated carotenes. Z Naturforsch 46c: 1045– 1051 Linden H, Vioque A and Sandmann G (1993) Isolation of a carotenoid biosynthesis gene coding for zeta-carotene desaturase from Anabaena PCC 7120 by heterologous complementation. FEMS Microbiol Lett 106: 99–104 Linden H, Misawa N, Saito T and Sandmann G (1994) A novel carotenoid biosynthesis gene coding for desaturase: Functional expression, sequence and phylogenetic origin. Plant Mol Biol 24: 369–379 Lundell DJ, Glazer AN, Melis A and Malkin R (1985) Characterization of a cyanobacterial Photosystem I complex. J Biol Chem 260: 646–654 Martinez-Ferez IM and Vioque A (1992) Nucleotide sequence of the phytoene desaturase gene from Synechocystis sp. PCC 6803 and characterization of a new mutation which confers resistance to the herbicide norflurazon. Plant Mol Biol 18: 981–983 Masamoto K, Riethman HC and Sherman LA (1987) Isolation and characterization of a carotenoid-associated thylakoid protein from the cyanobacterium Anacystis nidulans R2. Plant Physiol 84: 633–639 Math SK, Hearst JE and Poulter CD (1992) The crtE gene in Erwinia herbicola encodes geranylgeranyl diphosphate synthase. Proc Natl Acad Sci USA 89: 6761–6764 Mayer MP, Nievelstein V and Beyer P (1992) Purification and characterization of a NADPH dependent oxidoreductase from chromoplasts of Narcissus pseudonarcissus–a redox-mediator possibly involved in carotene desaturation. Plant Physiol Biochem 30: 389–398 Miller M, Pedersen JZ and Cox RP (1987) Effect of growth temperature on membrane properties in a thermophilic cyanobacterium (Synechococcus sp.). In: Biggins J (ed) Progress in Photosynthesis Research, Vol II, pp 789–792. Martinus-Nijhoff, Dordrecht Millie DF, Ingram DA and Dionigi CP (1990) Pigment and photosynthetic response of Oscillatoria agardhii (Cyanophyta) to photon flux density and spectral quality. J Phycol 26: 660– 666 Misawa N, Nakagawa M, Kobayashi K, Yamano S, Izawa I, Nakamura K. and Harashima K (1990) Elucidation of the Erwinia uredovora carotenoid biosynthetic pathway by functional analysis of gene products in Escherichia coli. J Bacteriol 172: 6704–6712 Moran R (1982) Formulae for determination of chlorophyllous pigments extracted with N,N,-dimethylformamide. Plant Physiol 69: 1376–1381 Nelson MA, Morelli G, Carattoli A, Romano N and Macino G (1989) Molecular cloning of a Neurospora crassa carotenoid biosynthetic gene (albino-3) regulated by blue light and the products of the white collar genes. Mol Cell Biol 9:1271–1276 Newell RW, van Amerongen H, Barber J and van Grondelle R (1993) Spectroscopic characterisation of the reaction center of Photosystem II using polarised light: Evidence for excitons in PS II reaction centers. Biochim Biophys Acta 1057: 232–238. Newman PJ and Sherman LA (1978) Isolation and characterization of Photosystem I and II membrane particles from the bluegreen alga Synechococcus cedrorum. Biochim Biophys Acta 503: 343–361

578 Nultsch W and Schuchart H (1985) A model of the phototactic reaction chain of cyanobacterium Anabaena variabilis. Arch Microbiol 142: 180–184 Ohno T, Satoh K and Katoh S (1986) Chemical composition of purified oxygen-evolving complexes from the thermophilic cyanobacterium Synechococcus sp. Biochim Biophys Acta 852: 1–8 Olaizola M and Duerr EO (1990) Effects of light intensity and quality on the growth rate and photosynthetic pigment content of Spirulina platensis. J Applied Phycol 2: 97–104 Olie J and Potts M (1986) Purification and biochemical analysis of the cytoplasmic membrane from the desiccation-tolerant cyanobacterium Nostoc commune UTEX 584. Appl Environ Microbiol 52: 706–710 Omata T (1992) Characterization of the downstream region of cmpA: Identification of a gene cluster encoding a putative permease of the cyanobacterium Synechococcus PCC7942. In: Murata N (ed) Research in Photosynthesis, Vol. Ill, pp 807–810. Kluwer, Dordrecht Omata T, Ohmori M, Arai N and Ogawa T (1989) Genetically engineered mutant of the cyanobacterium Synechococcus PCC7942 defective in nitrate transport. Proc Natl Acad Sci USA 86: 6612–6616 Omata T, Carlson TJ, Ogawa T and Pierce J (1990a) Sequencing and modification of the gene encoding the 42-kilodalton protein in the cytoplasmic membrane of Synechococcus PCC7942. Plant Physiol 80: 525–530 Omata T, Ogawa T, Carlson TJ and Pierce J (1990b) Nucleotide sequence of the genes encoding the two major proteins in the cytoplasmic membrane of Synechococcus PCC7942. In: Baltscheffsky M (ed) Current Research in Photosynthesis, Vol. III, pp 525–528. Kluwer, Dordrecht Palozza P and Krinsky NI (1992) Antioxidant effects of carotenoids in vivo and in vitro–an overview. Meth Enzymol 213: 403–420 Pecker I, Chamovitz D, Linden H, Sandmann G and Hirschberg J (1992) A single polypeptide catalyzing the conversion of phytoene to is transcriptionally regulated during tomato fruit ripening. Proc Natl Acad Sci USA 89: 4962–4966 Pecker I, Chamovitz D, Mann V, Sandmann G, Böger P and Hirschberg J (1993) Molecular characterization of carotenoid biosynthesis in plants: the phytoene desaturase gene in tomato. In: Murata N (ed) Research in Photosynthesis, Vol III, pp 11– 18. Kluwer, Dordrecht Raps S, Wyman K, Siegelman HW and Falkowski P (1983) Adaptation of the cyanobacterium Microcystis aeruginosa to light intensity. Plant Physiol 72: 829–832 Ray JA, Bird CR, Maunders M, Grierson D and Schuch W (1987) Sequence of pTOM5, a ripening related cDNA from tomato. Nucl Acids Res 15: 10587–10588 Ray JA, Moureau P, Bird AS, Grierson D, Maunders M, Truesdale M, Bramley PM and Schuch W (1992) Cloning and characterization of a gene involved in phytoene synthesis from tomato. Plant Mol Biol 19: 401–404 Reddy KJ, Masamoto K, Sherman DM 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 Reddy KJ, Bullerjahn GS and Sherman LA (1993) Characteristics of membrane-associated carotenoid-binding proteins in

Joseph Hirschberg and Daniel Chamovitz cyanobacteria and prochlorophytes. Meth Enzymol 214: 390– 401 Ruddat M and Will OH, III (1985) High-performance liquid chromatography ofcarotenoids. Meth Enzymol 111:189–200 Sandmann G and Böger P (1989) Inhibition of carotenoid biosynthesis by herbicides. In: Böger P and Sandmann G (eds) Target Sites of Herbicide Action, pp 25–44. CRC Press, Boca Raton Sandmann G and Bramley PM (1985) Carotenoid biosynthesis by Aphanocapsa homogenates coupled to a phytoenegenerating system from Phycomyces blakesleeanus. Planta 164: 259–263 Sandmann G and Kowalczyk S (1989) In vitro carotenogenesis and characterization of the phytoene desaturase reaction in Anacystis. Biochem Biophys Res Com 163: 916–921 Sandmann G, Bramley PM and Böger P (1985) New herbicidal inhibitors of carotene biosynthesis. Pestic Sci 10: 19–24 Sandmann G, Linden H and Böger P (1989) Enzyme kinetic studies on the interaction of norflurazon with phytoene desaturase. Z Naturforsch 44c: 787–790 Sandmann G, Woods WS and Tuveson RW (1990) Identification of carotenoids in Erwinia herbicola and in transformed Escherichia coli strain. FEMS Microbiol Lett 71: 77–82 Sandmann G, Kowalczyk-Schröder S, Taylor HM and Böger P (1992) Quantitative structure-activity relationship of fluridone derivatives with phytoene desaturase. Pest Biochem Physiol 42: 1–6 Sandmann G, Kuhn M and Böger P (1993) Carotenoids in photosynthesis–protection of Dl degradation in the light. Photosynth Res 35: 185–190 Satoh K (1992) Structure and function of Photosy stem II reaction center. In: Murata N (ed) Research in Photosynthesis, Vol. II, pp. 3–12. Kluwer, Dordrecht Schlodder E and Brettel K (1988) Primary charge separation in closed Photosystem II with a lifetime of 11 ns. Flash-absorption spectroscopy with oxygen-evolving Photosystem II complexes from Synechococcus. Biochim Biophys Acta 933: 22–34 Schmidhauser TJ, Lauter FR, Russo VEA and Yanofsky C (1990) Cloning sequencing and photoregulation of al-1, a carotenoid biosynthetic gene ofNeurospora crassa. Mol Cell Biol 10: 5064–5070 Schmidt A and Sandmann G (1990) Cloning and nucleotide sequence of the crtI gene encoding phytoene dehydrogenase from the cyanobacterium Aphanocapsa PCC6714. Gene 91: 113–118 Schnurr G, Schmidt A and Sandmann G (1991) Mapping of a carotenogenic gene cluster from Erwinia herbicola and functional identification of six genes. FEMS Microbiol Lett 78: 157–162 Seifermann-Harms D (1977) The xanthophyll cycle in higher plants. In: Tevini M and Lichtenthaler HK (eds) Lipids and Lipid Polymers in Higher Plants, pp 218–230. Springer, Berlin Serrano A, Gimenez P, Schmidt A and Sandmann G (1990) Immunocytochemical localization and functional determination of phytoene desaturase in photoautotrophic prokaryotes. J Gen Microbiol 136: 2465–2469 Shestakov SV and Jevner VD (1968) Study of mutagenesis in blue-green alga Anacystis nidulans. Proc XII Int Congr Genet I: 84.(Abstract) Shyam R and Sane PV (1989) Photoinhibition of photosynthesis and its recovery in low and high light acclimatized blue green

Chapter 18 Carotenoids in Cyanobacteria algae (cyanobacteria) Spirulina platensis. Biochem Physiol Pflanz 185: 211–219 Sommer TR, Potts WT and Morrissy NM (1990) Recent progress in processed microalgae in aquaculture. Hydrobiologia 204/ 205: 435–443 Spurgeon SL and Porter JW (1980) Biosynthesis of carotenoids. In: Porter JW and Spurgeon SL (eds) Biochemistry of Isoprenoid Compounds, pp. 1–122. Wiley, New York Stransky H and Hager A (1970) Das carotenoid muster und die verbreitung des lichtinduzierten xanthophyll-cyclus in verschiedenen algenklassen. IV, Cyanophyceae und Rhodophyceae. Arch Microbiol 72: 84–96 Suzuki R and Fujita Y (1977) Carotenoid photobleaching induced by the action of photosynthetic reaction center. II: DCMU sensitivity. Plant Cell Physiol 18: 625–631 Takahashi Y, Hirota K and Katoh S (1985) Multiple forms of P700-chlorophyll a-protein complexes from Synechococcus sp.: the iron, quinone and carotenoid contents. Photosynth Res 6: 183–192 Thornber JP, Alberte RS, Hunter FA, Shiozawa JA and Kan K-S (1976) The organization of chlorophyll in the plant photosynthetic unit. Brookhaven Symp Biology 28: 132–148 Trick M, Dennis ES, Edwards KJR and Peacock WJ (1988) Molecular analysis of the alcohol dehydrogenase family of barley. Plant Mol Biol 11: 147–160 Tuveson RW and Sandmann G (1993) Protection by cloned

579 carotenoid genes expressed in Escherichia-coli against phototoxic molecules activated by near-ultraviolet light. Meth Enzymol 214: 323–330 Urbach D, Suchanka M and Urbach W (1976) Effect of substituted pyridazinone herbicides and of difunone (EMD-IT 5914) on carotenoid biosynthesis in green algae. Z Naturforsch 31c: 652–655 van Baalen C (1965) Mutation of the blue-green alga Anacystis nidulans. Science 149: 70 Wakabayashi K, Sandmann G, Ohata H and Böger P (1988) Peroxidizing herbicides: Comparison of dark and light effects. J Pesticid Sci 13: 461–471 Wright SW and Shearer JD (1984) Rapid extraction and highperformance liquid chromatography of chlorophylls and carotenoids from marine phytoplankton. J Chromatogr 294: 281–295 Yamamoto HY (1979) Biochemistry of the violaxanthin cycle in higher plants. Pure Appl Chem 51: 639–648 Yokoyama H, Hsu WJ, Poling SM and Hayman E (1982) Chemical regulation of carotenoid biosynthesis. In: Britton G and Goodwin TW (eds) Carotenoid Chemistry and Biochemistry, pp 371–385. Pergamon, Oxford Zevenboom W and Mur RL (1984) Growth and photosynthetic response of the cyanobacterium Microcytis aeruginosa in relation to photoperiodicity and irradiance. Arch Microbiol 139: 232–239

Chapter 19 Genetic Analysis of Cyanobacteria Teresa Thiel Department of Biology, University of Missouri - St. Louis, 8001 Natural Bridge Rd., St. Louis, MO 63121, USA Summary I. Introduction II. Gene Transfer A. Conjugation 1. General Aspects of Conjugation Systems 2. Mobilizable Vectors a. Replicating Shuttle Vectors b. Integrative Vectors c. Antibiotic Resistance Genes d. Positive Selection for Cloned DNA 3. Helper Plasmids 4. Applications of the Conjugation System to Anabaena sp. Strain PCC 7120 5. Applications of the Conjugation System to Other Filamentous Cyanobacteria B. Natural Transformation C. Electroporation III. Mutagenesis A. Chemical and UV Mutagenesis B. Targeted Mutagenesis by Gene Interruption C. Site-Directed Mutagenesis D. Transposon Mutagenesis E. Random Cartridge Mutagenesis F. Segregation of Mutants IV. Reporter Systems A. Introduction B. lacZ as a Reporter C. luxAB as a Reporter V. DNA Elements A. nif Elements B. Insertion Sequences C. Repetitive Sequences VI. Mapping VII. Expression of Foreign Genes in Cyanobacteria A. General Considerations B. Construction of Integration Platforms C. Promoters for Gene Expression VIII. Developing a Genetic System: Practical Problems and Possible Solutions A. The Need For Genetic Systems In Other Cyanobacteria B. Mechanism of Transfer and Initial Stability C. Maintenance of Donor DNA in the Host D. Expression of Donor Genes in the Host E. One Approach for Demonstrating Gene Transfer Acknowledgments References D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 581–611. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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Summary In recent years great strides have been made in developing genetic systems for the analysis of various aspects of cyanobacterial physiology and development. Transformation, electroporation and conjugation systems provide effective means for gene transfer in diverse cyanobacterial strains. Gene transfer, combined with the ability to clone and inactivate genes in cyanobacteria, has opened the door to advanced studies of photosynthesis, nitrogen fixation, heterocyst development and metabolism in these unique and important procaryotic microorganisms. This chapter reviews the advances of the last few years in the development and use of genetic systems in cyanobacteria. These include the gene transfer techniques of conjugation, transformation and electroporation as well as mutagenesis by conventional and recombinant DNA-based techniques. Reporter systems, particularly lacZ and luxAB, are used to measure gene expression and to identify novel genes that are expressed under specific physiological conditions. Transposons that have a reporter gene can simultaneously inactivate a gene and report expression of the interrupted gene. Such transposons provide a powerful tool for the identification of environmentally regulated genes. The technique ofpulsed-field gel electrophoresis, combined with Southern hybridization analysis using cloned cyanobacterial genes and transposon-tagged genes as probes, has allowed the construction of the first detailed maps of cyanobacterial chromosomes. The development of these genetic techniques has stimulated interest in cyanobacteria as hosts for the expression of foreign genes. A variety of host-vector systems combined with the ability to clone genes under the control of regulatable promoters, provide the tools required for expression of potentially beneficial genes in cyanobacteria. Because cyanobacteria grow in disparate habitats they may provide an ideal system for the expression and dissemination of environmentally useful gene products. I. Introduction The transfer of DNA to Synechococcus sp. by transformation, first reported over 20 years ago, heralded the beginning of genetic systems in cyanobacteria. Almost ten years ago the first conjugation system for cyanobacteria was described. As gene transfer techniques were being developed, cyanobacterial plasmids were isolated, vectors were constructed and the first cyanobacterial genes were cloned. By the late 1980’s the components were ready for assembly into powerful and sophisticated systems for genetic analysis of cyanobacteria. Much of the knowledge of photosynthesis, nitrogen metabolism, and heterocyst differentiation that is reported in this book is the result ofexperiments that relied upon these genetic systems. Many excellent reviews over the last few years have described the development of many aspects of genetic systems in cyanobacteria (Porter, 1986; Tandeau de Marsac and Houmard, 1987; Shestakov and Reaston 1987; Houmard and Tandeau de Marsac, 1988; Wolk, 1991; Buikema and Haselkorn, 1993) and several articles have reviewed and described methodologies (Thiel and Wolk, 1987; Elhai and Wolk, 1988b; Friedberg, 1988; Golden, 1988; Porter, 1988; Williams, 1988; Elhai et al., 1990; Haselkorn,

1991). Extensive tables compiled by Tandeau de Marsac and Houmard (1987) and Houmard and Tandeau de Marsac (1988) provide valuable information on strain designations, restriction endonucleases, and cyanobacterial plasmids and cloning vectors. This chapter will focus primarily on recent developments in genetic systems, their applications, and on some ofthe problems with such systems. For more detailed discussions of the application of genetic techniques to particular aspects of cyanobacterial metabolism or development, the reader is referred to other chapters herein that discuss those aspects in detail. II. Gene Transfer

A. Conjugation 1. General Aspects of Conjugation Systems Conjugation, which is DNA transfer mediated by cell-to-cell contact, is based upon the mobilization of DNA from one bacterium (usually Escherichia coli) to another bacterium by a broad-host-range conjugative plasmid. (For a review of the mechanisms of conjugative transfer see Ippen-Ihler and Minkley,

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1986). The conjugative apparatus, typically provided by an IncP conjugative plasmid, such as RP4 (also known as RK2), mediates DNA transfer across a broad spectrum of bacteria, including many cyanobacteria (Wolk et al., 1984; Flores and Wolk, 1985). Conjugation has been the method of choice for gene transfer in filamentous cyanobacteria, but is also useful for unicellular cyanobacteria (Wolk et al., 1984; Kreps et al., 1990; Sode et al., 1992; S. Golden,

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personal communication). The methodology, first described by Wolk et al. (1984), has been extensively reviewed (Thiel and Wolk, 1987; Elhai and Wolk, 1988b; Elhai et al., 1990). Three plasmids are typically involved in conjugative transfer of DNA to cyanobacteria (Fig. 1). The plasmid to be transferred to the cyanobacterial host (sometimes called the ‘cargo plasmid’) must have a site, called bom or oriT, that is nicked by an enzyme (the mob product) prior

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to transfer. The nicking enzyme is usually produced in trans by a second helper plasmid in the same donor cell. The nicked strand is mobilized from the donor cell to the recipient cell via transfer (tra) gene products provided by a third, mobilizing conjugal plasmid. The mobilizing plasmid may be maintained in the same donor cell as the other two plasmids, or it can be transferred from a separate E. coli cell to the cell containing the cargo and helper plasmids during conjugation. Maintenance of multiple plasmids in one strain requires that the plasmids be compatiblei.e., that they have different replicons. In triparental matings, the conjugative plasmid is in one E. coli strain, the cargo and helper plasmids are in a second E. coli strain, and the third partner is the recipient cyanobacterial cell (Fig. 1). The donor DNA is probably transferred as single-stranded DNA; however, a new DNA strand is synthesized immediately by the host cell, probably during transfer. The transferred plasmid can recircularize and replicate if the plasmid has a replicon that functions in the recipient cell. The transferred DNA may also recombine with homologous DNA in either the chromosome or in another plasmid in the recipient cell. Many common cloning vectors based on pBR322 (e.g., the pUC series of vectors beyond pUC7 and pBluescript) have lost the oriT (bom) site and cannot be used for conjugation; however, plasmids with a variety of useful features for conjugation to Anabaena sp. and other cyanobacteria have been created (Wolk et al., 1984; Elhai and Wolk, 1988a; Wolk et al., 1988; Buikema and Haselkorn, 1991b; see Table 1). Some of these vectors will be described in more detail in other sections of this chapter. There is no evidence that most E. coli replicons function in cyanobacteria. It appears, however, that some IncQ plasmids (pKT210 or pKT230) (Bagdesarian et al., 1981) can transfer to and replicate in some cyanobacteria after mobilization by the broadhost-range conjugal plasmid, RP4 (Kreps et al., 1990; Prosperi et al., 1991; Sode et al., 1992). The oriT of these plasmids can be nicked by the mob gene product of the IncP plasmid (e.g., RP4) that also provides the transfer functions. Plasmid pRL 153, which is based on pKT210 (Elhai and Wolk, 1988a), has been mobilized to and appears to replicate in several strains of cyanobacteria (e.g., Synechococcus sp. strain PCC 7002, Synechocystis sp. strain PCC 6803, Anabaena sp. strain PCC 7120, and Pseudanabaena sp. strain PCC 7409; J. Elhai and T. Thiel,

unpublished). A mobilizable plasmid based on pBR325 that contains the oriT from an IncP plasmid has been reported to transfer to and replicate in Plectonema boryanum strain UTEX 594 (Vachhani et al., 1993).

2. Mobilizable Vectors a. Replicating Shuttle Vectors Except for the IncQ plasmids described above, replicating shuttle vectors include two replicons: one that allows replication of the plasmid in E. coli, and one that allows replication in the host strain. These vectors have been constructed by cloning into a mobilizable E. coli plasmid (i.e., a plasmid with an oriT site) a segment of a cyanobacterial plasmid that includes the genes required for replication. A variety of shuttle vectors for transformation have been constructed using plasmids from unicellular cyanobacteria. These plasmids typically lack oriT and thus cannot be mobilized (Elhai and Wolk, 1988b; Houmard and Tandeau de Marsac, 1988); however, mobilizable shuttle vectors (see Table 1) that replicate in several unicellular or filamentous cyanobacteria are available (Wolk et al., 1984; Cobley, 1985; Elhai and Wolk, 1988b; Houmard and Tandeau de Marsac, 1988; Murry and Wolk, 1991; Chiang et al., 1992a). Vectors that include the replication origin from pDU1, a plasmid from Nostoc sp. strain PCC 7524 (Reaston et al., 1982; Schmetterer and Wolk, 1988; Walton et al., 1992), allow autonomous replication in several strains of cyanobacteria (Wolk et al, 1984; Flores and Wolk, 1985; Schmetterer and Wolk, 1988; Elhai and Wolk, 1988a).

b. Integrative Vectors Vectors designed to allow selection for recombination between a cloned gene of interest and a region of homology in the chromosome have also been constructed. These vectors, lacking a cyanobacterial replication origin, must have antibiotic resistance genes that can confer resistance when expressed from the chromosome as a low-copy-number gene (Elhai and Wolk, 1988a). These vectors are typically used either for gene inactivation by targeted mutagenesis (see Section III B) or for the integration of a second copy of a gene into the chromosome forming a merodiploid (Fig. 2). In addition, some of these vectors include the conditionally lethal sacB

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gene (Ried and Collmer, 1987; Cai and Wolk, 1990) that provides a selection for integration of the plasmid into the chromosome by double reciprocal recombination. Expression of the sacB gene (which encodes a levan sucrase) in the presence of sucrose kills many bacteria, including some cyanobacteria. Cyanobacterial cells in which the non-replicating plasmid has recombined into the chromosome by a singlecrossover event will retain the vector and the sacB gene. These cells are killed ifthey are grown on solid media containing 5% sucrose. Cells in which a double reciprocal recombination event has removed the vector (andthe sacB gene) from the chromosome can be selected because the non-replicating plasmid is lost during cell division and progeny cells grow in the presence of 5% sucrose. This selection system must be monitored carefully: Both the E. coli donor and the cyanobacterial host easily lose the selectable phenotype by mutation of sacB, often by the

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transposition of insertion elements (Cai, 1991). Some strains of cyanobacteria, e.g., Anabaena variabilis strain ATCC 29413, appear resistant to sacB selection (T. Thiel, unpublished results).

c. Antibiotic Resistance Genes Antibiotic resistance genes that are expressed well in cyanobacteria even in low copy (e.g., when integrated in the chromosome) have been incorporated into mobilizable vectors. One such gene is npt (aph, kan), from Tn5, encoding aminoglycoside 3'-phosphotransferase and conferring resistance to kanamycin and neomycin. In some vectors the native promoter has been replaced by the psbA promoter from Amaranthus hybridus (Elhai and Wolk, 1988a) or by the psbO (woxA) promoter of Anabaena sp. strain PCC 7120 (Borthakur et al., 1990). Other antibiotic resistance genes that function well in Anabaena sp.

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588 strain PCC 7120 are the fragment, that contains the aadA gene with its native promoter and that confers resistance to streptomycin and spectinomycin (Prentki and Krisch, 1984; Golden and Wiest, 1988), and streptomycin resistance (aadA, str) (with a six-base deletion that increases resistance) or bleomycin resistance (ble) from Tn5, both under the control of a psbA promoter (Wolk et al., 1991). Erythromycin resistance (erm) (combined with a cat gene encoding chloramphenicol acetyl transferase and conferring resistance to chloramphenicol for selection in E. coli) with its native promoter has been used primarily for selection of replicating shuttle vectors in Anabaena sp. strain PCC 7120 (Wolk et al., 1984), although it also functions well in the chromosome.

d. Positive Selection for Cloned DNA Positive selection vectors, based on the inviability of E. coli cells carrying long inverted repeats (palindromes) of DNA, have been constructed to facilitate the cloning of genes (Elhai and Wolk, 1988a). The palindromes in these vectors are separated by a short polylinker. Interruption of the inverted repeats by a fragment of DNA allows transformed E. coli cells to grow, while vectors lacking an insert kill the transformed host. The vector with the inverted repeat can be maintained in E. coli JC8111, a recB recC sbcB recF mutant strain. There are a variety of such vectors that incorporate different antibiotic resistance genes and various polylinkers (Elhai and Wolk, 1988). These vectors are generally useful for cloning experiments because they provide positive selection for an insert rather than screening for an insert, which is common in cloning vectors such as pUC 18 and pBluescript (Stratagene).

3. Helper Plasmids Restriction of foreign DNA by enzymes in the host cell reduces the number of exconjugants (Wolk et al., 1984; Elhai and Wolk, 1988b). Elhai and Wolk (1988b) have modified the original helper plasmid that provides the trans-acting mob gene product by the addition of genes that encoded AvaI methylase and Eco47II methylase (which modifies GGNCC sites and protects against restriction by Ava II; see Table 1). A derivative of this helper plasmid containing the methylase gene for EcoT22I (an isoschizomer of AvaIII) is also available (J. Elhai and C. P. Wolk, personal communication). Vectors destined for

Teresa Thiel cyanobacterial hosts are methylated in the E. coli donor carrying the helper plasmid and are thus protected from restriction by isoschizomers of AvaI, AvaII, or AvaIII that are present in many cyanobacteria (Houmard and Tandeau de Marsac, 1988). Anabaena sp. strain PCC 7120 has AvaI and AvaII activities (Duyvesteyn et al, 1983), and has recently been found to have AvaIII activity (A.M. Muro-Pastor and E. Flores, personal communication).

4. Applications of the Conjugation System to Anabaena sp. Strain PCC 7120 Conjugation has become a versatile tool for genetic manipulation of filamentous cyanobacteria, primarily Anabaena sp. strain PCC 7120. This technique has been used to obtain mutations in genes that have been cloned from Anabaena sp. strain PCC 7120 by replacement of a wild-type copy with a gene mutated in vitro (Golden and Wiest, 1988; Borthakur et al., 1990; Buikema and Haselkorn, 1991a; Brahamsha and Haselkorn, 1992; Muro-Pastor et al., 1992), for complementation of mutations (Wolk et al., 1988; Buikema and Haselkorn, 1991a, 1991b; Floriano et al., 1992), for promoter analysis (Elhai and Wolk, 1990; Lang and Haselkorn, 1991), and for the introduction of transposons for mutagenesis (Borthakur and Haselkorn, 1989; Wolk et al., 1991; Ernst et al., 1992). These applications in Anabaena sp. strain PCC 7120 are discussed in more detail below and in other chapters of this book (see, in particular, Chapters 16, 20 and 27).

5. Applications of the Conjugation System to Other Filamentous Cyanobacteria A. variabilis strain ATCC 29413 (also known as Anabaena sp. strain PCC 7937) has been the subject of many physiological studies. This organism grows vigorously under photoautotrophic conditions and chemoheterotrophically in the dark with fructose; thus, Photosystem I mutants are viable (Toelge et al., 1991; Mannan et al., 1991). This strain maintains its phenotype (fixes nitrogen in the presence of oxygen, see Ernst et al., 1992 for nomenclature) even when grown for many generations with a source of fixed nitrogen. In contrast, Anabaena sp. strain PCC 7120 loses its ability to grow aerobically under diazotrophic conditions in the absence of selection forthe phenotype. This may present a problem in that mutants of Anabaena sp. strain PCC

Chapter 19 Molecular Genetic Techniques for Analysis of Cyanobacteria 7120, which must be grown with fixed nitrogen, may eventually lose their ability to be complemented by the wild-type allele because ofone or more secondary mutations. A. variabilis also has an alternative vanadium-dependent nitrogenase system (Kentemich et al., 1988; Thiel, 1993) that may also contribute to its vigorous diazotrophic growth. In addition, A. variabilis has a second molybdenum-dependent nitrogenase system that functions only under anaerobic conditions (T. Thiel, unpublished). However, genetic systems currently available for this strain lack the flexibility of those for Anabaena sp. strain PCC 7120. The availability of cosmid (Herrero and Wolk, 1986) and lambda libraries (Hirschberg et al., 1985; Thiel, 1993) and the polymerase chain reaction have facilitated the cloning ofgenes from this organism (Hirschberg et al., 1985; Owttrim and Coleman, 1987; Johnson et al., 1988; van der Plas et al., 1989; Maldener et al., 1991; Toelge et al., 1991; Mannan et al., 1991;Bovy et al., 1992; Thiel, 1993). Shuttle vectors based on pDU1 do not readily transfer and replicate in A. variabilis; however, non-replicating plasmids with inserts of A. variabilis DNA can transfer and recombine with their homologues in the chromosome. Mutagenesis by gene replacement (see Section III B, ‘Targeted Mutagenesis’) has resulted in mutants in a protease gene (Maldener et al., 1991), in Photosystem I genes (Toelge et al., 1991; Mannan et al., 1991), in a nuclease gene (A.M. Muro-Pastor, personal communication) and in a vnf (vanadium-dependent nitrogenase) gene (Thiel, 1993). The replication region of the 41 -kb plasmid, pRDS 1, fromA. variabilis has been used to create hybrid plasmids that can serve as shuttle vectors between E, coli and either Anabaena sp. strain M-131 or A. variabilis (Murry and Wolk, 1991). One of these shuttle vectors is a cosmid vector that could facilitate the construction of a genomic library that may be useful for the type of complementation experiments that have been very successful in Anabaena sp. strain PCC 7120 (Wolk et al., 1988; Buikema and Haselkorn, 1991a, 1991b; Floriano et al., 1992). However, these vectors transfer poorly to A. variabilis and appear to require recombination with the resident plasmid in this strain for replication; thus, the frequency oftransfer is low and recovery of a complementing cosmid could be difficult. DNA transfer by conjugation and electroporation (see Section II C, ‘Electroporation’) has been successful in the cyanobacterium Calothrix sp. strain PCC 7601 (formerly Fremyella diplosiphon) (Cobley

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et al., 1987; Chiang et al., 1992a), a strain characterized by complementary chromatic adaptation. A shuttle vector for Calothrix sp. strain PCC 7601 was constructed using the replicon from an endogenous plasmid. A genomic library of Calothrix sp. DNA constructed in the shuttle vector was used to complement two phycobilisome regulatory mutants that give red or blue colonies (see Chapter 21 for a discussion ofthese mutants). The difference in color between the mutant and the wild-type strains provides a simple visual screen for exconjugants that have been complemented (Chiang et al., 1992a). A plasmid from a complemented red mutant was recovered from E. coli after transformation of E. coli with total DNA isolated from the complemented Calothrix sp. strain. A gene, rcaC, was shown to complement the original mutant and another independently isolated red mutant (Chiang et al., 1992b). Structural differences in the rcaC locus of the two red mutants complemented by the wild-type allele indicate that the mutations may have been caused by mobile insertion elements (see Chapter 21). Plasmids rescued from a blue mutant, after complementation of the mutant by conjugation ofthe genomic library, could not complement the blue mutant when introduced again (Chiang et al., 1992a). For a discussion of the open reading frames (ORFs) found on the plasmids that complemented the blue mutant, see Chapter 21. Vectors based on pDU1 have been transferred to three strains that are involved in symbiotic associations: Nostoc sp. strain ATCC 29133 and strain ATCC 27896 (Flores and Wolk, 1985), and Nostoc sp. strain PCC 8009 (strain ‘MAC,’ J. Meeks, personal communication). Plasmids based on ColE1 transfer poorly to these Nostoc sp. strains (Flores and Wolk, 1985); however a pDU1 derivative with oriV and oriT from RK2, pDUCA7 (Buikema and Haselkorn, 1991b), transfers well (J. Meeks, personal communication). The vector pDUCA7 was used to create a cosmid library for Nostoc sp. strain ATCC 29133, and that library was used to complement a UV-induced mutant (unable to fix aerobically) of that Nostoc sp. In addition, conjugation has been used to obtain transposon mutants (Wolk et al., 1991) of Nostoc sp. strain ATCC 29133 (J. Meeks, personal communication). Conjugation and electroporation (see Section II C, ‘Electroporation’) have been successful in the nonheterocystous filamentous cyanobacterium Plectonema boryanum (Fujita et al., 1992; Vachhani et al., 1993;Walton et al., 1993). A shuttle vector that is

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based on the endogenous plasmid pGL3 of Plectonema boryanum strain PCC 6306 has been constructed and was shown to replicate stably in Plectonema sp. and in Anabaena sp. (Walton et al., 1993). A mobilizable plasmid with only a pMB1 replicon also appears to replicate autonomously in Plectonema sp.; however, the actual origin of replication in the cyanobacterium is unknown (Vachhani et al., 1993).

B. Natural Transformation Transformation, which is the transfer of free DNA into cells, was first described for Synechococcus sp. strain PCC 7943 (Shestakov and Khuyen, 1970) many years ago and remains today the primary means for gene transfer in unicellular cyanobacteria. Transformation has been thoroughly reviewed (Porter, 1986; Shestakov and Reaston, 1987) and the methodology has also been described well (Golden and Sherman, 1984; Porter, 1988). In addition to the original transformable strain, two close relatives, Synechococcus sp. strain PCC 6301 (Herdman and Carr, 1971) and Synechococcus sp. strain PCC 7942 (Grigorieva and Shestakov, 1976), are also transformable. These three strains are genetically very similar (Wilmotte and Stam, 1984; Golden et al., 1989); however, transformation has been studied in more detail in Synechococcus sp. strain PCC 7942 because it is highly transformable (Golden and Sherman, 1984). Heterospecific transformation of Synechococcus sp. strain PCC 7942 by DNA from Synechococcus sp. strain PCC 6301 gives the same frequency as homologous transformation, again suggesting a very close relationship between these strains (Grigorieva, 1985). The photoheterotrophic strain (i.e., it grows in the absence of Photosystem II activity if it is provided with glycerol in the light), Synechococcus sp. strain PCC 7002, is also highly transformable (Stevens and Porter, 1980; Buzby et al., 1983; Porter, 1986). Other transformable unicellular strains include Synechocystis sp. strain PCC 6308 (Devilly and Houghton, 1977), Synechocystis sp. strain PCC 6714 (Astier and Espardellier, 1976), and Synechocystis sp. strain PCC 6803 (Grigorieva and Shestakov, 1982). Synechocystis sp. strain PCC 6803 is naturally a photoheterotroph; however, in the laboratory it has recently been successfully grown heterotrophically with glucose in the dark (Anderson and McIntosh, 1991) and has been used extensively for cyanobacterial studies of

Teresa Thiel photosynthetic reaction centers. The mechanism of transformation in cyanobacteria is poorly understood; however, most of these unicellular strains are naturally competent, and the mechanism may share some characteristics with other transformable bacteria (Porter, 1986). One strain in which transformation can be induced is Synechocystis sp. strain PCC 6308, which requires treatment for competency. The ability ofheterologous DNA to compete with homologous DNA for uptake in Synechococcus sp. strain PCC 7942 (Golden and Sherman, 1984) and in Synechococcus sp. strain PCC 7002 (Essich et al., 1990) may imply a mechanism similar to that of transformable Grampositive heterotrophic bacteria. Cyanobacteria appear to be competent during all phases of growth (Shestakov and Khuyen, 1970; Stevens and Porter, 1980; Grigorieva and Shestakov, 1982); however, cells are usually transformed during mid- to lateexponential growth (Porter, 1988). Transformation is dependent on DNA concentration; it shows singlehit kinetics, with full saturation at concentrations as low as in Synechococcus sp. strain PCC 7002 and as high as in Synechocystis sp. strainPCC6803(StevensandPorter, 1980;Grigorieva and Shestakov, 1982; Porter, 1986). While transformation frequencies are variable between different strains, and even between different experiments in the same strain, typical values are of DNA (Porter, 1986). Although both chromosomal and plasmid DNA can transform unicellular cyanobacteria, efforts in recent years have focused on transformation with plasmids containing cyanobacterial genes. There have been reports of replication of broad-host-range plasmids of the IncQ group in cyanobacteria (Kreps et al., 1990; Sode et al., 1992), but most plasmid transformation has used endogenous cyanobacterial plasmids as the source of cyanobacterial replicons. Many shuttle vectors carrying a variety ofselectable markers and capable of replication in both cyanobacteria and E. coli have been constructed (for an extensive list see Table IV in Houmard and Tandeau de Marsac, 1988). One problem is that these shuttle vectors recombine readily with the resident plasmid, often leading to loss of the shuttle vector (Porter, 1986). A strain of Synechococcus sp. strain PCC 7942 cured of the endogenous plasmid, pUH24, could stably maintain the donor plasmid, but the efficiency of transformation in that strain was much lower than in the wild-type strain (Chauvat et al.,

Chapter 19 Molecular Genetic Techniques for Analysis of Cyanobacteria 1983). The creation of stable merodiploids (i.e., strains containing two copies of a segment ofDNA) via replicating plasmids has been problematic. Chromosomal genes cloned in a shuttle vector and transformed into a cyanobacterial host recombine with their homologues in the genome. This can result in non-reciprocal recombination (also called gene conversion) in which there is conversion of a mutated copy ofa gene in a replicating plasmid to a wild-type copy (Porter et al., 1986). Thus, although the plasmid is maintained, the desired genotype may be lost. Vectors that cannot replicate in the cyanobacterial host allow selection for recombination of the plasmid into the chromosome (Williams and Szalay, 1983). These vectors have been useful for targeted mutagenesis (see Section III B), in which the gene in the chromosome is replaced by the mutated gene in the plasmid. In Synechococcus sp. strain PCC 7942, the frequency of replacement of the chromosomal gene is about 100-fold higher than the frequency of addition ofthe plasmid to the genome (Kolowsky et al., 1984), and in Synechocystis sp. strain PCC 6803 the frequency of replacement is about 1000-fold higher than addition (Williams, 1988; Labarre et al., 1989). Therefore, this is an effective means for creating mutants but is not efficient for creating merodiploids that have both a wild-type and mutant copy of the gene. If gene replacement occurs by double crossovers and merodiploid formation occurs by single crossovers, merodiploids should predominate. It has been suggested that the unusually high frequency ofapparent double-crossover events is the result of gene conversion (nonreciprocal recombination; Williams and Szalay, 1983; Kolowsky et al., 1984; Labarre et al., 1989). However, Porter (1986) argues that they can be better explained by a linearization of the plasmid during transformation. Linear plasmids could not readily recombine into the chromosome by a single-crossover event, but could efficiently yield gene replacement by doublereciprocal recombination with the chromosome. Gene replacement has provided a powerful tool for the construction of mutants for analysis of photosynthesis, a subject that is covered in depth in other chapters of this book.

C. Electroporation Electroporation has been used to introduce DNA into animal cells, plant cells, and many bacteria, including several cyanobacteria. Optimum conditions for

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electroporation of a replicating plasmid in Anabaena sp. strain M-131 were a field strength of and a time constant of 5 msec (Thiel and Poo, 1989). As is true for conjugation, restriction of DNA is a significant problem; a single unmodified AvaII site reduces transformation efficiency about 100-fold (Thiel and Poo, 1989). Electroporation has some advantages over conjugation: E. coli cells do not contaminate the transformants; vectors lacking a bom site (e.g. the pUC series) may serve as donors; and electroporation requires only DNA and washed host cells. The ability to methylate the donor DNA in vitro, to produce linear plasmids for transfer (yielding exclusively double recombinants after integration into the chromosome), and to use chromosomal DNA as the donor are potential advantages of electroporation that have not yet been explored. In studies of genes for iron-sulfur proteins in Plectonema boryanum strain IAM-M101 (Takahashi et al., 1991; Fujita et al., 1991, 1992), electroporation has been used to create targeted mutations of frxC (protochlorophyllide reductase) and nifH (dinitrogenase reductase)(Fujita et al., 1992). The cloned frxC and nifH genes were interrupted by a kanamycin/ neomycin resistance gene and were introduced into wild-type Plectonema boryanum strain by electroporation. Following electroporation, cells were grown for three days under non-selective conditions and then plated with kanamycin. The conditions for electroporation of P. boryanum were significantly different from those used previously (Thiel and Poo, 1989): The field strength, and the time constant, about 10 msec, are both much greater than is optimal for replicating plasmids introduced into Anabaena sp. strain M-131. In addition, the required DNA concentration was high. The number of transformants was low: There was one transformant for the nifH replacement, and it was a double recombinant. There were five transformants for frxC using DNA at two were double recombinants and three were single recombinants. Since the products of both single and double recombination events were found among the transformants, it appears that plasmids that enter the cell by electroporation may not be cleaved. In one experiment in which there was only 0.48 kb of genomic flanking sequence on one side and 1.05 kb on the other side of the antibiotic resistance gene no transformants were found. No efforts were made to pre-methylate the DNA or to determine whether restriction was a problem; thus, the low frequency

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may be at least partly attributable to restriction. DNA transfer by electroporation to Calothrix sp. strain PCC 7601 has also been successfully used to transform shuttle vectors and a genomic library, cloned in a replicative shuttle vector, into Calothrix sp. (Chiang et al., 1992a). The optimum conditions for electroporation of Calothrix sp. strain PCC 7601 are field strengths of with a time constant of 5 msec using DNA at (Chiang et al., 1992a). These conditions are intermediate between those reported for Anabaena sp. (Thiel and Poo, 1989) and for Plectonema boryanum (Fujita et al., 1992). Electroporation has been reported to be mutagenic in Calothrixsp. strainPCC 7601. Afterelectroporation pigmentation mutants were found at higher frequencies than could be accounted for by spontaneous mutation (Bruns et al., 1989). Although mutants have not been reported after electroporation of other bacteria, most organisms do not exhibit easily distinguishable mutant phenotypes. A high frequency ofspontaneous pigmentation mutants was reported in Calothrix sp. strain PCC 7601 by Tandeau de Marsac (1983) and this strain was subsequently show to harbor insertion sequences (Mazel et al., 1991). One possible mechanism of mutagenesis is that introduction of DNA by electroporation induces transposition of one or more of the IS elements present in Calothrix sp. Two red mutants of Calothrix sp. strain PCC 7601 have been found to have gross structural changes in the altered gene that may be the result of transposition of an insertion element (see Chapter 21). It is also possible that any entry of foreign DNA into a cell is mutagenic, perhaps by inducing the SOS response or by inducing insertion sequences to move. The effect may be more pronounced in electroporation than in conjugation because of the large amount of DNA that may enter the cell. III. Mutagenesis

A. Chemical and UV Mutagenesis A few techniques for chemical and ultraviolet (UV) light-induced mutagenesis of cyanobacteria have been well reviewed (Golden, 1988;Haselkorn, 1991). Chapman and Meeks (1987) described conditions for nitrosoguanidine mutagenesis that produce mutants

Teresa Thiel of A. variabilis strain ATCC 29413 at a frequency of about one in 104 viable cells. Auxotrophs of A. variabilis strain ATCC 29413 were obtained by nitrosoguanidine mutagenesis followed by penicillin enrichment (Currier et al., 1977). Diethyl sulfate mutagenesis (Chapman and Meeks, 1987; Haselkorn, 1991) or UV mutagenesis (Golden, 1988; Wolk et al., 1988) followed by penicillin enrichment have been used to obtain mutants of Anabaena sp. strain PCC 7120 unable tofixnitrogen (Buikema and Haselkorn, 1991 b). With the development of techniques for gene cloning, gene transfer, and transposition in cyanobacteria, both chemical and UV-light mutagenesis techniques have become less popular. However, it has been shown in Anabaena sp. strain PCC7120 (Wolk etal., 1988; Buikema and Haselkorn, 1991a, 1991b; Floriano et al., 1992) that the combination of chemical or UV light mutagenesis and complementation ofmutations by wild-type genes on plasmids can provide a powerful tool for the discovery and characterization of novel genes.

B. Targeted Mutagenesis by Gene Interruption In all cyanobacteria forwhich gene transfer is feasible, targeted inactivation of a gene by interruption of the cloned gene with a selectable marker has become the method of choice for mutagenesis (Fig. 2). Using cloned genes for the three copies of psbA in Synechococcus sp. strain PCC 7942, Golden et al. (1986) introduced an antibiotic resistance gene into each psbA gene. Plasmids carrying the interrupted cyanobacterial genes were transformed into wildtype Synechococcus sp. strain and mutants in psbA were selected by their antibiotic-resistant phenotype. More recently, targeted mutagenesis has been a valuable tool in the study of photosynthesis in Synechocystis sp. strain PCC 6803 (e.g., Williams, 1988; Pakrasi et al., 1988; Burnap and Sherman, 1991; Smart et al., 1991), Synechococcus sp. strain PCC 7002 (Bryant et al., 1990;Zhou et al., 1992),and in A. variabilis strain ATCC 29413 (Mannan et al., 1991; Toelge et al., 1991; Thiel, 1993), and for the study of a variety of genes in other cyanobacteria (Golden and Wiest, 1988; Borthakur et al., 1990; Buikema and Haselkorn, 1991a; Maldener et al., 1991; Brahamsha and Haselkorn, 1992; Fujita et al., 1992; Muro-Pastor et al., 1992) Transformed plasmid DNA in unicellular strains yields primarily recombinants that are the result of

Chapter 19 Molecular Genetic Techniques for Analysis of Cyanobacteria

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apparent double-reciprocal recombination, replacing the wild-type gene with the mutated copy. (See IIB, ‘Natural Transformation’, for a discussion of this phenomenon.) This is in contrast to Anabaena sp., in which recombination following conjugation yields primarily merodiploids, the result of single recombination (Cai and Wolk, 1990). Although the high frequency of double recombination in Synechococcus sp. strains may be attributable to the method of gene transfer, this may not be the only factor involved. Conjugal transfer of a non-replicating vector designed for targeted gene activation in Synechococcus sp. strain PCC 7942 also yields primarily double recombinants (S. Golden, personal communication). Even among different strains of filamentous cyanobacteria, the frequency ofdouble versus single recombinational events following conjugation is variable. Whereas double crossovers appear to be rare in Anabaena sp. strain PCC 7120, in A. variabilis strain ATCC 29413 double crossovers account for 10–30% of recombinants (T. Thiel, unpublished results). In Plectonema boryanum, two out of five recombinants obtained by electroporation with a plasmid carrying an inactivated copy of the frxC gene were double recombinants (Fujita et al., 1992); thus, double crossovers may also be common in that strain. The problem of a high frequency of merodiploids in Anabaena sp. strain PCC 7120 has been overcome by the development of a positive selection vector using sacB that facilitates selection of double recombinants (Cai and Wolk, 1990). (See Section II A, 2b; ‘Integrative Vectors.’)

provide a new restriction site for ease in identification of the mutated plasmid. The plasmid with the mutation is introducedintothe cyanobacteriumbygene transfer; replacement of the wild-type gene copy with the mutated form by double reciprocal recombination yields the desired mutant strain. Unless a mutation creates a selectable phenotype, replacement is most easily accomplished by including an antibiotic resistance gene near, but outside of, the coding region of the gene of interest (Golden et al., 1986). Selection for antibiotic resistance typically produces mutants that also carry the linked point mutation. In some cases gene conversion or a recombination event within the segment of DNA between the mutation and the antibiotic resistance gene will produce antibiotic-resistant strains that are not mutated at the site of interest. This problem can be overcome by first creating a cyanobacterial strain with a deletion ofthe gene of interest that precludes such a recombination event. After the plasmid with the site-directed mutation is transferred to the cyanobacterial deletion strain, antibiotic-resistant recombinants can be formed only by crossover events outside the gene ofinterest; thus, all such recombinants should contain the sitedirected mutation within that gene (Fig. 3; EatonRye and Vermaas, 1991). Alternatively, complementation in trans via a plasmid-borne copy of the gene being studied can be carried out in a strain harboring a chromosomal deletion of the gene of interest (e.g., Zhou et al., 1992).

C. Site-Directed Mutagenesis

Mutagenesis by transposition was first reported in Synechococcus sp. strain PCC 7942 when Tn901, introduced on a replicating plasmid, produced antibiotic-resistant colonies (Tandeau de Marsac et al., 1982). One transposon-containing strain was shown to be a methionine auxotroph. Transposon mutagenesis with Tn901 in plasmid pUH24 of Synechococcus sp. strain PCC 7942 (van den Hondel et al., 1980) has recently been used to identify a cluster of genes involved in nitrate assimilation (Madueño et al., 1988; Luque et al., 1992). The transposon was demonstrated to have integrated into the chromosome of Synechococcus sp. and the mutated region, carrying the transposon, was cloned in a plasmid which then served as a probe for cloning the wild-type genes (Luque et al., 1992). Transposon mutagenesis has been very successful in Anabaena

Site-directed mutagenesis, causing one or more specific changes in nucleotide sequence, has been used primarily in unicellular cyanobacteria for the study of photosynthesis (e.g., Vermaas et al., 1987 a,b; Metzetal., 1989; Pakrasi et al., 1991; Nixon and Diner, 1992; Ohad and Hirschberg, 1992; van der Bolt and Vermaas, 1992). The use of this technique for cyanobacteria was reviewed by Williams (1988). Various strategies and their requisite components, packaged as commercial kits, are available for creating the desired mutation in vitro. Site-directed mutagenesis requires that the sequence of the region of interest be known. Design of the mutagenic oligonucleotide typically includes one or more changes that alter a specific amino acid and may

D. Transposon Mutagenesis

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sp. strain PCC7120(Borthakur and Haselkorn, 1989; Wolk et al., 1991; Ernst et al., 1992).Mutagenesis of Anabaena sp. strain PCC 7120 minimally requires a transposon with an antibiotic resistance gene that is expressed well in Anabaena sp. on a mobilizable plasmid incapable of replication in that cyanobacterium. Tn5 is able to transpose, creating neomycin-resistant exconjugants in Anabaena sp. strain PCC 7120 (Borthakur and Haselkorn, 1989). Wolk and his coworkers have constructed several transposon vectors for cyanobacteria. These vectors have multiple antibiotic resistance genes with a strong plant promoter (from the psbA gene of Amaranthus hybridus), an origin of transfer for conjugation, an E. coli replication origin within the transposon that facilitates recovery of the mutated gene in E. coli, and, in some plasmids, promoterless luxAB genes as a transcriptional reporter of transposition (Table 1; Wolk et al., 1991; Ernst et al., 1992). Transposon mutants are obtained after conjugation of these vectors from E. coli to cyanobacteria by selecting for colonies resistant to an antibiotic to which the transposon encodes resistance. Insertion of transposons carrying luxAB into transcriptionally active genes results in luminescent colonies. A change in the luminescence

Teresa Thiel

of a colony that results from a change in its environment (such as a change in the composition of the medium, a change in temperature, or in light intensity) identifies that colony as potentially having a transposon in a gene that responds to the environmental change (Wolk et al., 1991). A mutant phenotype that appears concomitantly with transposition must be confirmed to be the result of the transposition event by recovery of the mutated fragment for reconstruction of the mutant (Ernst et al., 1992). A variety of mutants of Anabaena sp. strain PCC 7120 affected in their ability to fix nitrogen were isolated by transposon mutagenesis (Ernst et al., 1992). Transposons have not only facilitated the isolation of mutants and the identification of new genes, but have also aided in mapping new genes on the Anabaena sp. strain PCC 7120 chromosome (Kuritz et al., 1993) (see Section V, ‘Mapping’).

E. Random Cartridge Mutagenesis In unicellular cyanobacteria an alternative to transposon mutagenesis is the random insertion of an antibiotic resistance gene into the genome using restriction fragments ofgenomic DNA (Buzby et al.,

Chapter 19 Molecular Genetic Techniques for Analysis of Cyanobacteria 1985; Chauvat et al., 1989; Labarre et al., 1989). Chromosomal DNA from the cyanobacterium is digested with a restriction enzyme and randomly ligated to an antibiotic resistance gene that has compatible ends. The ligated fragments are then transformed into the cyanobacterial host and antibiotic-resistant transformants are selected. In Synechocystis sp. strain PCC6803, a high percentage of antibiotic-resistant transformants are defective in photosynthesis (Chauvat et al., 1989), Analysis of these mutants revealed that many are deletion mutants that map to a region of the genome containing the Photosystem II genes psbB, psbC, and psbDI. The transformants appear to result from integration of the two different chromosomal fragments, which flank the antibiotic resistance cassette, into the genome by either double reciprocal recombination or by gene conversion (via a heteroduplex hybrid DNA intermediate; Labarre et al., 1989). This results in insertion of the antibiotic resistance cassette into the genome with concomitant deletion of the region of the chromosome between the two DNA segments that flanked the antibiotic resistance cassette in the transforming DNA (Fig. 4). This technique yields a high frequency of mutation in Synechocystis sp.

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strain PCC 6803, and can cause large deletions (Chauvat et al., 1989; Labarre et al., 1989).

F. Segregation of Mutants All techniques for mutagenesis require consideration of genetic and morphological features of cyanobacteria that may differ from other bacteria. At least some cyanobacteria have as many as 10–15 copies of their haploid genome (Herdman et al., 1979; Labarre et al., 1989). Cells with a newly introduced mutation will, therefore, be heterozygous (i.e., having both wild-type and mutant genomes) until replication and subsequent segregation of these genomes produce a cell that is homozygous for the mutation. Strains with multiple copies of the genome will require several rounds of cell division to effect complete segregation. In filamentous cyanobacteria, even one cell in the filament with a wild type copy of a gene may confer a wild-type phenotype on the colony growing from the filament (Currier et al., 1977). These problems are overcome by allowing several generations of growth for segregation of mutant genomes before applying selection for the mutant, and by sonication of filaments to very short fragments to produce

596 colonies that are the progeny of a single cell (Wolk and Wojciuch, 1973).

IV. Reporter Systems

A. Introduction Reporter genes lacking a promoter have been fused to the promoter regions of genes of interest for the study ofgene expression in many biological systems. In cyanobacteria, the expression of the many genes that are environmentally regulated may be most easily studied through the use of reporters. In heterocystous cyanobacteria the differential expression of developmentally controlled genes in specialized cell types can best be studied by the use of reporter genes. The product of the reporter gene should be easily assayed and quantitated; however, for filamentous cyanobacteria, in situ localization of the product to particular cells may be equally important. Although the quantity of the product of the reporter gene is a convenient measure of promoter function, it is important to consider that the transcript of the reporter gene is not the same as the transcript of the gene that is replaced: Fusions may produce a transcript with different stability than the native transcript; hence, the quantity ofthe reporter product may not accurately reflect the amount of the native product of the gene transcribed from that promoter (Bustos and Golden, 1992). Moreover, the reporter itself may differ in stability in cells grown under different conditions. Techniques for the use ofseveral genes as reporters in cyanobacteria have been described by Friedberg (1988). Chloramphenicol acetyl transferase (cat) (Friedberg and Seijffers, 1986; Ferino and Chauvat, 1989; Lang and Haselkorn, 1991; see Section VI C, ‘Promoters for Gene Expression’, for further discussion of cat), (lacZ; Buzby et al., 1985; Schaefer and Golden, 1989; Elhai and Wolk, 1990; Bustos and Golden, 1991; 1992), glucuronidase (GUS; A. Grossman, personal communication) and luciferase (luxAB; Schmetterer et al., 1986; Elhai and Wolk, 1990; Wolk et al., 1991) have been shown to function in cyanobacteria. The cat gene, on a replicating shuttle vector, was used as a reporter for deletion analysis of the psbB promoter of Anabaena sp. strain PCC 7120 (Lang and Haselkorn, 1991). A novel reporter plasmid for promoters that are active in heterocysts has been constructed by C. Bauer and R. Haselkorn (personal

Teresa Thiel communication). The promoter of interest is cloned before a promoterless cat gene that is followed by the structural genes for nitrogenase (nifHDK, also without a promoter). The plasmid is transferred to a nifHdeletion strain of Anabaena sp. strain PCC 7120 (lacking nitrogenase activity; Golden et al., 1991) and exconjugants are selected as chloramphenicolresistant colonies. A promoter that functions in heterocysts drives the transcription of nitrogenase, which functions in heterocysts and confers a phenotype on the cell. Ifthe promoter functions only in vegetative cells, active nitrogenase will not be made and the cells will be When the psbB promoter region of Anabaena sp. strain PCC 7120 was cloned in this vector and transferred to the nifHdeletion strain, chloramphenicol-resistant exconjugants were suggesting that the psbB promoter may function in heterocysts (C. Bauer and R. Haselkorn, personal communication).

B. lacZ as a Reporter Expression of lacZ from a replicating plasmid was first demonstrated in Synechococcus sp. strain PCC 7002 (Buzby et al., 1985), and this reporter was subsequently used to monitor expression from the cpcBACDEF promoter of the same organism (Gasparich et al., 1987). The lacZ gene was used as a transcriptional reporter to measure expression of nifH in Anabaena sp. strain PCC 7120 (Elhai and Wolk, 1990). A replicating shuttle vector carrying lacZ under the control a nifH promoter was transferred by conjugation to Anabaena sp. strain PCC 7118, which cannot make heterocysts, and is, therefore, However, under anaerobic conditions, the expression of lacZ begins about 13 hours after removal of combined nitrogen from the medium, indicating that nif genes can be expressed anaerobically (Elhai and Wolk, 1990). The lacZ gene can be expressed in the cyanobacterium Synechococcus sp. strain PCC 7942 from a plasmid, or from a copy integrated into the chromosome where it was used to identify regulated promoters (Scanlan et al., 1990). The lacZ gene was also used in Synechococcus sp. strain PCC 7942 to study the light-regulated expression of the multicopy Photosystem II genes, psbA (Schaefer and Golden, 1989; see Chapter 23), and psbD (Bustos and Golden, 1992). The lacZ fusions were used to identify regions downstream from the transcriptional start of psbDII that are required for expression (Bustos and Golden, 1991). The lacZ fusions in Synechococcus

Chapter 19 Molecular Genetic Techniques for Analysis of Cyanobacteria sp. strain PCC 7942 were constructed in vitro and introduced into the cyanobacterial genome by transformation and recombination. Since recombination after transformation yields almost exclusively gene replacement (by double-recombination events or by gene conversion) rather than addition of the altered copy (by a single-recombination event), fusions to an essential gene would inactivate the normal copy and would be lethal (Bustos and Golden, 1992). Although the problem of gene replacement can be overcome by inserting the reporter fusion in a gene contained in a replicating plasmid, this poses a problem because the amount of product from the reporter gene will be determined in part by the copy number of the plasmid, which may vary under different growth conditions. A solution to these problems in Synechococcus sp. strain PCC 7942 was the construction of a plasmid bearing an integration platform, or ‘neutral site,’ to serve as the site for construction of the fusions. The integration platform allows recombination into the chromosome without disruption ofthe gene ofinterest (Fig. 5)(Bustos and Golden, 1992). For a more detailed explanation of

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integration platforms, including ‘neutral sites,’ the reader should see Section VI B, ‘Integration Platforms.’

C. luxAB as a Reporter The most elegant use of reporter genes has been the work from Wolk’s laboratory using the luxAB genes, encoding luciferase, as an in situ reporter of gene expression in vegetative cells and heterocysts of Anabaena sp. strain PCC 7120 (Elhai and Wolk, 1990). Light emitted from cells expressing the luxAB genes was captured by a highly sensitive camera and transformed into a pattern of light and dark areas. Superimposition of this image on a computergenerated skeleton of cells, obtained from a brightfield image of a filament, allows localization of luxAB expression to individual cells in the filament. The luciferase reporter was used to study the expression from nif H, rbcLS,glnA and hetR promoters during heterocyst differentiation (Elhai and Wolk, 1990; Black et al., 1993; see the Chapter 27 for further details on expression from these promoters).

598 The luciferase reporter system has recently been modified to amplifyexpressionfromweakpromoters using the bacteriophage T7 RNA polymerase gene. In this system, the promoter ofinterest is fused to the T7 RNA polymerase gene while luxAB is driven by a T7 promoter. Expression from the promoter of interest produces T7 RNA polymerase that then acts on the T7 promoter to transcribe luxAB (Wolk et al., 1993). Using the amplification by T7 polymerase, hepA (formerly hetA), a gene required for heterocyst formation (Holland and Wolk, 1990), was shown to be transcribed 7.5 hours after nitrate deprivation, hours before heterocysts are morphologically evident (Wolk et al., 1993). Luciferase has also been used as a reporter for identification of transposon-induced mutants that respond to changes in environment (Wolk et al., 1991). For more information, the reader should consult Section III D, ‘Transposon Mutagenesis,’ and Chapter 27.

V. DNA Elements

A. nif Elements The first elements to be characterized in cyanobacteria were the 11-kb insertion that interrupts nifD (Golden et al., 1985) and the 55-kb element in fdxN (Golden et al., 1988) in Anabaena sp. strain PCC 7120 that are excised by site-specific recombination (Golden et al., 1987). Little is known about the origin of these elements; however, the 11-kb element has some similarity to an insertion sequence IS892, which is discussed below. The 11-kb element is prevalent in cyanobacteria (Meeks et al., 1988), including A. variabilis strain ATCC 29413 (Brusca et al., 1989), but is missing in the heterocystous strain Fischerella sp. (Mastigocladus laminosum) (Saville et al., 1987; Singh and Stevens, 1992). The 55-kb element is present in Nostoc sp. strain PCC 8009 (strain ‘MAC’) (C. Carrasco and J. Golden, personal communication), but is not present in A. variabilis (Brusca et al., 1989). These elements are discussed in more detail in Chapter 27 by Wolk et al. and will not be discussed further here.

B. Insertion Sequences Bacterial insertion sequences (IS) are DNA elements, typically containing terminal inverted repeats, that are capable of transposition with concomitant

Teresa Thiel mutagenesis of interrupted genes. Transposition of an IS element typically causes a short duplication of the target sequence. A high frequency ofspontaneous mutations affecting pigmentation in Calothrix sp. strain PCC 7601 led to speculation that such mobile genetic elements were responsible for these mutations (Tandeau de Marsac, 1983). Several different IS elements have since been identified and characterized in cyanobacteria. Three IS elements, IS701, IS702, andIS703,havebeenfoundinthegenomeofCalothrix sp. strain PCC 7601 (Mazel et al., 1991). IS701 and IS 702 are typical bacterial IS elements: They have long terminal inverted repeats and their transposition causes a short duplication ofthe target DNA. Each IS element has an open reading frame (ORF) that may encode a transposase. IS701 is present in at least 15 copies and IS702 in nine copies in the Calothrix sp. strain PCC 7601 genome. These elements hybridize to the DNA ofother cyanobacteria, although there is wide variation in the number of hybridizing bands and in the restriction fragment patterns (Mazel, et al., 1991). The IS element IS891 was isolated from Anabaena sp. strain M-131. This element is atypical in that it lacks inverted terminal repeats and does not duplicate the target DNA site upon transposition (Bancroft and Wolk, 1989). Six IS elements were identified in Anabaena sp. strain PCC 7120. These were found during characterization of mutations that inactivated the conditionally lethal sacB gene that had been introduced into Anabaena sp. strain PCC 7120 on a replicating plasmid (Cai, 1991) (See Section II A, 2b, ‘Integrative Vectors’, for an explanation of sacB action). IS892 has terminal repeats and generates an 8-bp direct repeat in the target. It has two ORFs in which the codon usage is similar to that ofthe ORFs found in the excised 11-kb insertion element of nifD of Anabaena sp. strain PCC 7120 (Golden et al., 1985). In addition, the sequence of the inverted repeats in IS892 has some similarity to the 11-bp direct repeats that flank the 11-kb insertion in nifD (Cai, 1991). IS895 from Anabaena sp. strain PCC 7120 was identified by its proximity to psbAI (Alam et al., 1991), and, independently, as the cause of a mutation in sacB (Cai, 1991). IS895 has a 30-bp terminal inverted repeat, but does not duplicate the target site. Three other copies of IS895 from Anabaena sp. strain PCC 7120, designated A, B, and C, were also characterized (Alam et al., 1991). These three are very similar to IS895, differing only by small insertions or deletions that alter ORF I or ORF II in

Chapter 19 Molecular Genetic Techniques for Analysis of Cyanobacteria IS895B and IS895C. There is no evidence that these iso-IS895 elements transpose (Alam et al., 1991). IS892-like and IS895-like elements are also present in Anabaena sp. strain PCC 7118 and Anabaena sp. strain M-131 (Alam et al. 1991; Cai, 1991). IS892like elements were the type found most frequently among 15 mutant strains in which the sacB gene appeared to have been inactivated by insertion elements. When restriction digests of chromosomal DNA from Anabaena sp. strain PCC 7120 were hybridized to IS892, a pattern of hybridizing bands was initially observed; however, that pattern changed with increasing time of maintenance of the Anabaena sp. culture. DNA from a culture stored frozen for 8 years showed dramatic differences in hybridization pattern compared to the culture grown continuously during that period. In contrast, IS895 hybridization patterns didnot show significant changes. Cai suggests that IS892 is an actively transposing IS element, while others, like IS895, may be somewhat more stable (Cai, 1991). Elements IS893, IS894, IS897, and IS898 have not yet been characterized. The prevalence of IS elements in Anabaena sp. strain PCC 7120 may account for the instability of its phenotype: Cultures of this strain maintained on a medium containing a source of fixed nitrogen readily lose their phenotype (J Elhai, J. Golden, T. Thiel; independent unpublished results).

C. Repetitive Sequences In addition to insertion sequences, there are also reports of repetitive DNA sequences in cyanobacteria. Three families of tandemly repeated sequences, first identified in Calothrix sp. strain PCC 7601, have also been identified in many other filamentous cyanobacteria (Mazel et al., 1990). Anabaena sp. strain PCC 7120 has a variety of short, tandemly repeated sequences (Mulligan and Haselkorn, 1989; Haselkorn and Buikema, 1992). Most of these are heptamers, typically containing CCCC with other bases. While most of these sequences have been found in noncoding regions of the genome, the nifJ gene of Anabaena sp. strain PCC 7120 has five tandemly repeated heptamers of CCCCAGT within a coding region (Bauer et al., 1993).

VI. Mapping With the development oftechniques for pulsed-field

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gel electrophoresis, physical maps have been produced for many bacterial chromosomes. These maps allow the localization ofgenes to specific large restriction fragments of DNA, thus merging the genetic and physical maps into an accurate representation of the chromosome. Genetic and physical maps have been constructed for Anabaena sp. strain PCC 7120 (Fig. 6A)(Bancroft et al., 1989; Kuritz et al., 1993), Synechococcus sp. strain PCC 7002 (Fig. 6B)(Chen and Widger, 1993) and Synechocystis sp. strain PCC 6803 (Fig. 6C)(S. Shestakov, personal communication). In Anabaena sp. strain PCC 7120, restriction enzymes AvrII, PstI, and SalI, that cut the genome very rarely, produce 9–25 fragments of DNA in the 50–800 kb size range. These fragments can be separated and transferred to membranes for Southern hybridization to probes of known genes (Bancroft andWolk, 1988; Bancroft etal., 1989). In addition to locating genes to particular restriction fragments, Wolk and his coworkers (Kuritz et al., 1993) have devised a method to orient a gene on the physical map with respect to its direction of transcription. This requires that the gene to be oriented be cloned in a vector that contains restriction sites that are rare in the chromosome. The cloned gene in this vector is transferred to Anabaena sp. where antibiotic-resistant exconjugant cells will contain, by single-crossover recombination in the chromosome, a duplicated copy of the cloned region of the chromosome with the vector between the two copies (e.g., see Fig. 2B). Chromosomal DNA from wild-type or recombinant strains is digested with the enzymes that restrict the rare sites in the vector. Southern analysis of these fragments using gene specific and vector specific probes produces labeled bands that can be used to identify the orientation of the inserted plasmid in the genome by the size of the hybridizing fragments. Since the orientation of the gene of interest in the plasmid is known, the orientation of the gene in the chromosome can be inferred (Kuritz et al., 1993). A map of the chromosome of Anabaena sp. strain PCC 7120 is given in Fig.6A (from Kuritz et al., 1993; kindly provided by C. P. Wolk). Using this technique, cpcBA (C-phycocyanin) was mapped at 0.61 Mb and is transcribed clockwise. It is apparent from the map that neither genes for photosynthesis nor genes involved in heterocyst differentiation (e.g., hetR, hep A) or nitrogen fixation are tightly clustered. These mapping techniques are useful in the analysis of mutants and may help in the development ofa set of

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Chapter 19 Molecular Genetic Techniques for Analysis of Cyanobacteria overlapping clones representing the entire Anabaena sp. strain PCC 7120 genome (Kuritz et al., 1993). Such a set of clones would be very useful for the rapid localization ofnew genes or transposon mutants to a particular clone, which would then also indicate its approximate location on the genome. A map of the 2.7 Mb chromosome of Synechococcus sp. strain PCC 7002 has been constructed from AscI, NotI, and SfiI restriction fragments (Chen and Widger, 1993; Fig.6B). Twenty-onegenes orgene clusters have been localized to particular restriction fragments by Southern hybridization, thus providing information concerning the relative positions of many of these genes. A map of the 3.3 Mb chromosome of Synechocystis sp. strain PCC 6803 has also been constructed from NotI, Eco72I and MluI restriction fragments and 23 genes or gene clusters have been mapped on this genome (S. Shestakov, personal communication). There appears to be very little similarity in the organization of the genetic map among any of the three cyanobacterial genomes.

VII. Expression of Foreign Genes in Cyanobacteria

A. General Considerations The expression of foreign genes in cyanobacteria requires many of the genetic tools discussed in previous sections, including cloning vectors and gene transfer techniques. Expression of foreign genes also requires a promoter that functions in cyanobacteria and requires control of gene expression if the product is toxic to the cyanobacterial cell. Some promoters that may be useful for foreign gene expression are briefly described below; however, the reader is referred to Chapter 20 for a more complete discussion of promoters in cyanobacteria. Some genes that have been expressed in cyanobacteria are of cyanobacterial origin, but others are noncyanobacterial genes that may have practical applications. The apcAB genes, encoding the and subunits of allophycocyanin from Cyanophora paradoxa, were expressed from a replicating plasmid in Synechococcus sp. strain PCC 7002 (De Lorimier et al., 1987). Tandeau de Marsac et al. (1987) first expressed a Bacillus sphaericus entomocidal toxin gene in Synechococcus sp. strain PCC 7942 on a replicating shuttle vector. The toxin gene was expressed in Synechococcus sp. strain PCC 7942 and the protein

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killed mosquito larvae. Recently, Murphy and Stevens (1992) have translationally fused the cryIVD gene from Bacillus thuringiensis to the promoter region of cpcB gene of Synechococcus sp. strain PCC 7002 in a replicating shuttle vector for expression in that cyanobacterial strain. The cpcB-cryIVD fusion protein is expressed at high levels in Synechococcus sp. and is not degraded. The protein was toxic to mosquito larvae that ingested the transformed cyanobacteria (Murphy and Stevens, 1992).

B. Construction of Integration Platforms While some genes have been expressed from a replicating plasmid, other genes have been introduced into a cyanobacterial chromosome. Integration platforms and the conceptually similar ‘neutral sites’ (Bustos and Golden, 1992) are fragments of cyanobacterial chromosomal DNA that are cloned in a non-replicating vector. An antibiotic resistance gene that is expressed in the cyanobacterial host is cloned within the chromosomal fragment. The platform is used as a region for insertion, in vitro, of any gene (cyanobacterial or foreign) that is destined for the cyanobacterial chromosome. The plasmid is then transferred to the cyanobacterial host with antibiotic selection for integration of the platform (with the inserted gene) at its homologous site in the chromosome. The gene inserted within the platform is, therefore, also integrated in the chromosome. Such platforms are particularly useful in unicellular Cyanobacteria where single recombination, forming merodiploids, is rare (Williams, 1988). The first such vectors contained fragments of Synechocystis sp. strain PCC 6803 DNA, interrupted by a kanamycin resistance gene, cloned into pUC9 (Williams, 1988). This type of vector was used to integrate the psbA gene of the higher plant Poa annua into the genome of a strain of Synechocystis sp. strain PCC 6803, deleted of all its own psbA genes, and the gene was expressed (Nixon et al., 1991). This same vector was used by Merida et al. (1992) to insert a copy of the Anabaena sp. strain PCC 7120 glnA gene into the genome of Synechocystis sp. strain PCC 6803, creating a heterologous glnA merodiploid. The Synechocystis sp. strain PCC 6803 glnA gene in the merodiploid was mutated by targeted inactivation of the gene; however, the Synechocystis sp. strain PCC 6803 glnA mutant grew, using the Anabaena sp. strain PCC 7120 gene (Merida et al., 1992). A plasmid carrying a 2.8 BamHI chromosomal

602 fragment of Synechococcus sp. strain PCC 7942 was constructed to serve as a ‘neutral site’ locus for integration of fusions of psbD to lacZ (Bustos and Golden,1992). Inthiscase,integration ofthefusions at the psbD locus would have been lethal, since this gene is essential in Synechococcus sp. strain PCC 7942. The neutral site vector was constructed by inserting size selected (2–4 kb) chromosomal DNA from Synechococcus sp. strain PCC 7942 into the tetracycline resistance gene of pBR328. Plasmids obtained from this ligation were screened for a plasmid that had a restriction site that appeared once in the insert (near the center) but not in the vector. A spectinomycin resistance gene was inserted in the chromosomal fragment in the plasmid, transferred to Synechococcus sp. strain PCC 7942 and spectinomycin-resistant recombinants were obtained. A plasmid that integrated and expressed spectinomycin resistance, but that had no deleterious effects on growth became a neutral site vector (Fig. 5) (S. Golden, personal communication). While the original vector lacks the oriT site required for conjugation, later derivatives constructed in pBR322 are mobilizable by conjugation to Synechococcus sp. strain PCC 7942 with very high efficiency (S. Golden, personal communication). A platform vector has been constructed for Synechococcus sp. strain PCC 7002 using a fragment of DNA from that strain that complements an arg3 mutant of E. coli. A polylinker in the cyanobacterial DNA allows insertion of a foreign gene and a kanamycin/neomycin resistance gene within the cyanobacterial DNA allows selection for recombinants that have integrated in the chromosome. Unlike the vectors described above, the vector portion of this platform plasmid can be easily removed by digestion ofthe plasmid with NotI (sites for Not I are rare in cyanobacterial DNA) prior to transformation of Synechococcus sp. strain PCC 7002 (W. H. Muñiz, and R. D. Porter, personal communication). A somewhat different platform based on pBR322 was constructed in Synechococcus sp. strain PCC 7942 for the transfer and integration of the plastocyanin gene of A. variabilis strain ATCC 29413 into Synechococcus sp. strain PCC 7942, and for the integration of an extra copy of the petFI gene from Synechococcus sp. strain PCC 7942 itself (van der Plas et al., 1990). The platform consists of the oriV region of pBR322 and a segment of the ampicillin resistance gene (providing homology topBR322, but conferring ampicillin sensitivity on the host) separated

Teresa Thiel by a kanamycin resistance gene. This platform, integrated in the Synechococcus sp. strain PCC 7942 genome at the metF locus (hence, the strain is provides homology for the insertion of any gene cloned in pBR322 (or derivatives such as the pUC series of cloning vectors) and renders the strain resistant to kanamycin (see structure of R2-PIM9 in Fig. 7). Recombination between the homologous pBR322 sequences in the genome and those in the vector yields ampicillin-resistant, kanamycinsensitive transformants in which the kanamycin resistance gene is replaced by the gene cloned in pBR322 (van der Plas et al., 1990). The presence of oriV in the platform allows rescue of the integrated fragment as a replicating plasmid. An example of the use of this platform to insert the apcE gene of Calothrix sp. strain PCC 7601 in the platform of R2PIM9 is provided in Fig. 7 (modified from Capuano et al., 1993, and very kindly provided by J. Houmard). The apcE gene of Calothrix sp. strain PCC 7601 is expressed in the heterologous host; however, it could not substitute for the endogenous apcE gene, which is essential for growth of Synechococcus sp. strain PCC 7942 (Capuano et al., 1993).

C. Promoters for Gene Expression Whether integrated in the chromosome or on a replicating plasmid, expression of a foreign gene in a cyanobacterium requires a promoter that can be recognized in the host. Analysis of sequences upstream of transcriptional start sites has failed to identify a consensus promoter in cyanobacteria (see Chapter 20). In vitro experiments suggest that an E. coli consensus promoter should be an excellent cyanobacterial promoter (Schneider et al., 1991). In a derivative of Synechococcus sp. strain PCC 7942 (Anacystis R2K), the lambda promoter with a temperature-sensitive lambda cI represser gene (cI857) was fused to the cat gene. At 30 °C the strain produces an active cI repressor that somewhat inhibits transcription of cat. At 42 °C the repressor is inactivated and more cat is synthesizd (Friedberg, 1988). Expression of cat was about 20-fold higher after 15 hours at 42 °C compared to 30 °C. In Synechocystis sp. strain PCC 6803, the lambda promoter with the repressor gene cI857 was inserted upstream of cat in a plasmid that was then integrated into an endogenous plasmid in the cyanobacterium. The cat gene was not expressed at 30 °C, and was induced at least 2000fold at 39 °C (Ferino and Chauvat, 1989). Thus, the

Chapter 19 Molecular Genetic Techniques for Analysis of Cyanobacteria

lambda promoter appears to be a tightly regulated promoter in Synechocystis sp. strain PCC 6803. Unfortunately, few cyanobacteria grow well at temperatures of 39– 42 °C, thus limiting the potential use ofthis promoter in many cyanobacteria. The tac promoter of E. coli (a trp-lac hybrid promoter), without the lacI repressor gene, was also able to promote high levels of transcription of the cat gene in Synechocystis sp. strain PCC 6803 (Ferino and Chauvat, 1989); however, the tac promoter is poorly repressed, even in E. coli, by the lacI repressor gene (Brosius, 1988). The recA promoter of E. coli that is inducible by agents that activate the SOS response, such as UV light or mitomycin C, failed to express the cat gene in Synechocystis sp. strain PCC 6803, even in the presence of possible inducing agents (Ferino and Chauvat, 1989). Cyanobacterial genes may also be a source of regulatable promoters. Expression of the phycocyanin genes, cpcB2A2, of Calothrix PCC 7601 is regulated by the color of light (Conley et al., 1988; se Chapter 21). Red light increases expression of the cpcB2A2driven GUS reporter gene 30- to 50-fold (E. Casey and A. Grossman, personal communication). Genes that are induced by nutrient

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stress are also potential sources of regulatable promoters. When the promoter region of the phoA (phosphatase) gene of Synechococcus sp. strain PCC 7942 (Ray et al., 1991) was fused to GUS, phosphate starvation led to a 10- to 20-fold increase in enzyme activity (E. Casey and A. Grossman, personal communication). Similarly, the promoter for a sulfurstress gene, rhdA (Laudenbachetal., 1991), is induced 50- to 100-fold by sulfur starvation (D. Laudenbach and A. Grossman, personal communication). These promoters show clear potential for expression of toxic genes in cyanobacteria. J. Elhai and C. P. Wolk (personal communication) have shown that psbA (from Amaranthus hybridus), rbcLS ( from Anabaena sp. strain PCC 7120) and tac are strong promoters in Anabaena sp. strain PCC 7120. The rbcLS promoter and the tac promoter give similar levels of activity when fused to luciferase; the psbA promoter is about twice as strong (J. Elhai, personal communication). The tac promoter (with is induced 4- to 5-fold by the addition of isopropyl thio-galactoside (IPTG; J. Elhai, personal communication). We have recently cloned the 30 kDa movement protein of tobacco mosaic virus (Deom et al., 1987) in a replicating shuttle vector in Anabaena

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sp. strain PCC 7120, under the control of the tac promoter (Zahalak and Thiel, unpublished). The protein is produced in large amounts in Anabaena sp. strain PCC 7120, and IPTG increased the amount of product 3- to 5-fold. As is true in E. coli, expression from the tac promoter is not repressed well by the lac repressor in Anabaena sp. strain PCC 7120. VIII. Developing a Genetic System: Practical Problems and Possible Solutions

A. The Need For Genetic Systems In Other Cyanobacteria In the last ten years great progress has been made in developing genetic systems for cyanobacteria. Transformation and conjugation systems combined with the ability to clone and inactivate genes in cyanobacteria have opened the door to advanced studies of photosynthesis, nitrogen fixation, heterocyst development and metabolism. And yet relatively few organisms have very well developed genetic systems. There has been a trend in the last few years toward defining a few cyanobacterial strains as the paradigms for study. Synechocystis sp. strain PCC 6803 is one model for photosynthesis. Anabaena sp. strain PCC 7120 is the choice for nitrogen fixation and heterocyst development. This situation is at least partly attributable to the successes, and even more attributable to the failures, in the development of genetic systems. Clearly there are many fascinating aspects of cyanobacterial structure, function, growth, and metabolism yet to be explored. Hormogonia, akinetes, motility, chromatic adaptation, and symbiotic associations are still poorly understood. Have we learned anything in developing genetics to its current state that can accelerate the pace of development of genetic systems in cyanobacteria that are not currently in the mainstream? Are there simply some strains that are amenable to genetic manipulation and others that are not? Clearly there are more questions than answers, but there has been information gained in the development of gene transfer systems that may help to guide others in developing systems in other cyanobacteria. Three separate aspects of gene transfer must be considered in attempts to develop a system for a new organism: 1) mechanism of transfer and initial stability of the donor DNA in the recipient, 2) maintenance of the donor DNA in the

Teresa Thiel host, and 3) expression of desired genes in the host.

B. Mechanism of Transfer and Initial Stability Natural competence appears to be a trait of only a few species of bacteria, and conditions for the induction of artificial competence must be determined empirically. After many years of attempts with a variety of cyanobacteria, transformation is still largely restricted to those few strains described above (see Section II B, ‘Natural Transformation’). Conjugation and electroporation do not depend on competence and could be successful, potentially, for a broad spectrum of host strains. The fact that both conjugation and electroporation are successful in the same strains (and that neither is successful in many other strains; Thiel and Poo, 1989; Chiang et al., 1992a; J. Meeks, personal communication) suggests that both mechanisms are effective in the transfer of genes to virtually any organism, but that some organisms have barriers beyond the stage of transfer that prevent successful establishment or expression of the donor DNA. The large differences in frequency of transfer to different organisms by either conjugation or electroporation (Flores and Wolk, 1985; Thiel and Poo, 1989) supports the argument that the barrier is not transfer. Thus, for any new organism, one of these transfer mechanisms would be the obvious choice. A variety of vectors and helper plasmids for cyanobacteria that can be used for either method are available (Table l)(Elhai and Wolk, 1988b; Elhai et al., 1990). One known barrier to gene transfer is restriction of the donor DNA by the many endonucleases present in cyanobacteria (see Houmard and Tandeau de Marsac, 1988). Conjugation efficiencies are low when unmodified plasmids are transferred from E. coli to Anabaena sp. strain PCC 7120 (Wolk et al., 1984). Elhai and Wolk (1988b) found that conjugation efficiency decreases 5- to 13-fold for each unmethylated AvaII site, and 25- to 50-fold for each unmethylated AvaI site in the donor DNA. Even one unmodified AvaII site in a plasmid decreases electroporation efficiency 100-fold (Thiel and Poo, 1989). Similarly, a single AquI (an isoschizomer of AvaI) site in Synechococcus PCC 7002 redces transformation efficiency as much as 100-fold (Buzby et al., 1983; Porter, 1986). It is possible to use crude extracts of sonicated cyanobacterial cells with known plasmids or with bacteriophage lambda DNA to determine restriction enzyme sites by comparison of

Chapter 19 Molecular Genetic Techniques for Analysis of Cyanobacteria the restriction pattern with known enzymes (D. Bryant, personal communication). Many cyanobacterial restriction endonucleases are isoschizomers of enzymes for which modification enzymes are available. For DNA that is to be used for transformation or electroporation, modification can be done in vitro. For restriction enzyme activities for which no methylase is commercially available, it may be possible to use a crude extract of the cyanobacterial host with S-adenosyl methionine for modification of the donor DNA (Swanson et al., 1992). Alternatively, once a vector is established in the desired cyanobacterial host, the vector can be reisolated from that host (where it will be modified), manipulated in vitro (e.g., insert a cloned gene) and returned to that host (J. Meeks, personal communication). For conjugation, the plasmid must be modified in vivo, typically by expression of the gene for the appropriate methylase on a helper plasmid in the E. coli donor strain (Elhai and Wolk, 1988b). One potential problem has been the presence of nonspecific nucleases in many cyanobacteria (Wolk and Kraus, 1982; Muro-Pastor et al., 1992). It is not clear how much they affect establishment of donor DNA. Although both Anabaena sp. strain PCC 7120 and A. variabilis strain ATCC 29413 have a sugarnonspecific nuclease (Muro-Pastor et al., 1992; A.M. Muro-Pastor, personal communication), the former strain is much more amenable to genetic manipulation than the latter. Inactivation of this nuclease in either strain does not enhance its capabilities for gene transfer (A.M. Muro-Pastor, personal communication). Type I restriction systems, which may also provide a barrier to gene transfer, have not been characterized in cyanobacteria; however, these sites may be modified by treatment of DNA in vitro with extracts of the host cell (Swanson et al., 1992).

C. Maintenance of Donor DNA in the Host Establishment of a plasmid as an independent replicon in the host strain requires a replication origin that functions in the host. Many shuttle vectors based on endogenous cyanobacterial plasmids are available (Table 1 )(also, see Houmard and Tandeau de Marsac, 1988), but little is known concerning the species specificity of the replicons. While endogenous plasmids can be used to construct shuttle vectors, these shuttle vectors may integrate into the endogenous plasmid making recovery of large plasmids difficult. The availability of cloned genes in

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many cyanobacteria allows the transfer and integration of plasmids into the chromosome using the homology of the cloned gene to direct the plasmid to a particular site (Fig. 2). This has the advantage that no cyanobacterial replicon is required; thus, currently available plasmids can be used. Although a replicating shuttle vector is sometimes desirable, genetics in cyanobacteria isfeasible with only integrativevectors.

D. Expression of Donor Genes in the Host Successful transfer and replication or integration of a vector are not sufficient if the selectable marker is not expressed well in the cyanobacterial host. Although many antibiotic resistance genes appear to function in hosts for which gene transfer systems are well established, it cannot be assumed that antibiotic resistance genes with their native promoters will function in all cyanobacteria. For example, the kanamycin/neomycin resistance gene driven by a psbA promoter (Elhai and Wolk, 1988a) functions very well in both Anabaena sp. strain PCC 7120 and A. variabilis strain ATCC 29413 (Maldener et al., 1991; Toelge et al., 1991; Mannan et al., 1991). However, while the streptomycin/spectinomycin resistance gene with its native promoter can be used for gene targeted mutagenesis in Anabaena sp. strain PCC 7120 (Golden and Wiest, 1988), that gene has not worked well for that purpose in A. variabilis strain ATCC 29413 (T. Thiel, unpublished results). For some strains, antibiotic resistance cassettes with strong promoters that function well in cyanobacteria, such as the promoter from the psbA gene of Amaranthus hybridus, may be required.

E. One Approach for Demonstrating Gene Transfer While the road to efficient gene transfer is clearly strewn with potential roadblocks, the knowledge that roadblocks exist provides the basis for rational approaches to circumvent them. One reasonable approach would be to begin with a gene or fragment of DNA cloned from the organism of interest. It should be cloned in a plasmid like pRL447 (Elhai and Wolk, 1988a) which is mobilizable, and carries kanamycin/neomycin resistance with a psbA promoter. This vector could be transferred by conjugation or by electroporation to the host of interest, for Anabaena sp. neomycin at provides good selection, although effective drug

606 concentrations for other strains may have to be determined empirically. Plasmids such as pRL528, which encodes methylases for AvaI and AvaII sites (Elhai and Wolk, 1988b), or pRL623, which also protects AvaIII sites (J. Elhai and C. P. Wolk, personal communication), should be used for in vivo methylation of these type II potential restriction sites that may lead to restriction in the cyanobacterial host. In vitro methylation using a host-cell extract may also be used to protect DNA that is to be transferred by electroporation or by transformation. After selection for neomycin-resistant colonies, the transfer and integration of the plasmid in the recipient cell can be easily verified by Southern hybridization. Antibiotic resistance genes can also be inserted into cloned genes, allowing mutant isolation (Fig. 2B; see Section III B, ‘Targeted Mutagenesis’). It should be noted that a relatively large segment of homologous DNA (at least one kb, preferably two or more kb) facilitates recombination, particularly when integration by double recombination is required. In summary, a genetic system that functions at even minimal efficiency allows further empirical determination of optimum conditions for electroporation or conjugation with subsequent refinement of vectors for diverse applications.

Acknowledgments The author is grateful to C. Bauer, D. Bryant, E. Casey, J. Elhai, E. Flores, S. Golden, A. Grossman, R. Haselkorn, J. Houmard, A. Muro-Pastor, J. Meeks, W. Muñiz, R. Porter, S. Shestakov, and C. P. Wolk for providing unpublished information, and to J. Houmard, S. Shestakov, W. Widger, and C. P. Wolk for providing figures.

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Teresa Thiel van den Hondel CAMJJ, Verbeek S, van der Ende A, Weisbeek PJ, Borrias WE and van Arkel GA (1980) Introduction of transposon Tn901 into a plasmid of Anacystis nidulans: Preparation for cloning in cyanobacteria. Proc Natl Acad Sci USA 77: 1570–1574 van der Bolt F and Vermaas W (1992) Photoinactivation of Photosystem II as studied with site directed D2 mutants of the cyanobacterium Synechocystis sp. PCC 6803. Biochem et Biophys Acta 1098: 247–254 van der Plas J, Bovy A, Kruyt F, DeVrieze G, Dassen E, Klein B and Weisbeek P (1989) The gene for the precursor of plastocyanin from the cyanobacterium Anabaena sp. PCC 7937: isolation, sequence and regulation. Mol Microbiol 3: 275–284 van der Plas J, Hegeman H, deVrieze G, Tuyl M, Borrias M and (1990) Genomic integration system based on pBR322 sequences for the cyanobacterium Synechococcus sp. PCC 7942: transfer of genes encoding plastocyanin and ferredoxin. Gene 95: 39– 48 van Haute E, Joos H, Maes M, Warren G, van Montagu M and Schell J. (1983) Intergeneric transfer and exchange recombination of restriction fragment clones in pBR322: a novel strategy for the reversed genetics of the Ti plasmids of Agrobacterium tumefaciens. EMBO J 2: 411–417 Vermaas WJF, Williams JGK and Arntzen CJ (1987a) Sequencing and modification of psbB, the gene encoding CP-47 protein of Photosystem II in the cyanobacterium Synechocystis 6803. Plant Mol Biol 8: 317–326 Vermaas WJF, Williams JGK and Arntzen CJ (1987b) Sitedirected mutation in two histidine residues in the D2 protein inactivate and destabilize Photosystem II in the cyanobacterium Synechocystis 6803. Z Naturforsch 42c: 762–768 Walton DK, Gendel SM and Atherly AG (1992) Nucleotide sequence of the replication region of the Nostoc PCC 7524 plasmid pDUl. Nucleic Acids Res 20: 4660 Walton DK, Gendel SM and Atherly AG (1993) DNA sequence and shuttle vector construction of plasmid pGL3 from Plectonema boryanum PCC 6306. Nucleic Acids Res 21: 746 Williams JG (1988) Construction of specific mutations in Photosystem II photosynthetic reaction center by genetic engineering methods in Synechocystis 6803. Meth Enzymol 167:766–778 Williams JGK and Szalay AA (1983) Stable integration of foreign DNA into the chromosome of the cyanobacterium Synechococcus R2. Gene 24: 37–51 Wilmotte AMR and Stam WT (1984) Genetic relationships among cyanobacterial strains originally designated as ‘Anacystis nidulans’ and some other Synechococcus strains. J Gen Microbiol 103: 2737–2740 Wolk CP (1991) Genetic analysis of cyanobacterial development. Curr Opin Genet Dev 1: 336–341 Wolk CP and Wojciuch E (1973) Simple methods for plating single vegetative cells of, and for replica-plating, filamentous blue-green algae. Archiv Fur Mikrobiologie 91: 91–95 Wolk CP and Kraus J. (1982) Two approaches to obtaining low extracellular deoxyribonuclease activity in cultures of heterocyst-forming cyanobacteria. Arch Microbiol 131: 302– 307 Wolk CP, Vonshak A, Kehoe P and Elhai J (1984) Construction of shuttle vectors capable of conjugative transfer from

Chapter 19 Molecular Genetic Techniques for Analysis of Cyanobacteria Escherichia coli to nitrogen-fixing filamentous cyanobacteria. Proc Natl Acad Sci USA 81: 156l–l565 Wolk CP, Cai Y, Cardemil L, Flores E, Hohn B, Murry M, Schmetterer G, Schrautemeier B and Wilson R (1988) Isolation and complementation of mutants of Anabaena sp. strain PCC 7120 unable to grow aerobically on dinitrogen. J Bacteriol 170: 1239–1244 Wolk CP, Cai Y and Panoff J-M (1991) Use of a transposon with luciferase as a reporter to identify environmentally responsive

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genes in a cyanobacterium. Proc Natl Acad Sci USA 88:5355– 5359 Wolk CP, Elhai J, Kuritz T and Holland D (1993) Amplified expression of a transcriptional pattern formed during development of Anabaena Mol Microbiol 7: 441–445 Zhou JH, Gasparich GE, Stirewalt VL, Delorimier R and Bryant DA (1992) The cpcE and cpcF genes of Synechococcus sp. PCC 7002 – construction and phenotypic characterization of interposon mutants. J Biol Chem 267: 16138–16145

Chapter 20 The Transcription Apparatus and the Regulation of Transcription Initiation Stephanie E. Curtis Department of Genetics, Box 7614, North Carolina State University, Raleigh, NC 27695-7614, USA

James A. Martin Department of Biology, University of Iowa, Iowa City, Iowa 52242, USA

Summary I. Introduction II. Transcription in E. coli: Paradigms for Eubacteria A. RNA Polymerase Structure and Function B. Alternate Sigma Factors C. Sigma Factor Structure 1. The Family 2. The Family D. Structure and Function of Bacterial Promoters 1. Promoters a. Basal Promoters b. Regulated Promoters Promoters 2. III. Transcription in Cyanobacteria A. The Transcriptional Apparatus 1. RNA Polymerase Core Structure 2. Sigma Factors B. Cyanobacterial Promoters 1. Sequence Comparisons 2. In vitro Assays of Promoter Activity 3. In vivo Assays of Promoter Activity a. Verification of Transcription Initiation Sites and Identification of Minimal Basal Promoters b. Mutations in Basal Elements: The rbcL Promoter c. Dissection of Regulatory Elements: The psbDII Promoter C. Regulation of Transcription Initiation 1. Accessory Factors 2. Promoter Switching? D. Conclusions and Future Directions Acknowledgments References

D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 613–639. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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Summary Our knowledge of transcription in cyanobacteria has increased dramatically during the last decade. The aspects of transcription for which most is known include the properties of the transcription apparatus and the regulation of transcription initiation. The structure of the RNA polymerase of Escherichia coli was believed to be universal among eubacteria, and the finding that cyanobacteria possess a transcriptional apparatus of unique structure was unexpected. Cyanobacterial RNA polymerases contain equivalents of the and subunits of the E. coli enzyme, but differ in having two subunits that correspond to the subunit of E. coli. The genes encoding the sigma factors for the major form of RNA polymerase, as well as additional sigma factor genes have been identified in several cyanobacteria. All of the genes encode sigma factors with features of the principal sigma factors of other eubacteria. The roles of these multiple sigma factors remain to be elucidated, and factors of the alternative sigma class have not yet been identified. Comparisons of promoter sequences, the analysis of promoter activities in vivo and in vitro, and other studies indicate that the promoters of E. coli and cyanobacteria are found at similar positions relative to transcription initiation sites. Most cyanobacterial promoters display a conserved element at –10 from the transcription initiation site that conforms to the E. coli –10 promoter consensus. For the majority of genes, however, an element that conforms to the E. coli –35 promoter element is lacking. These data and other evidence suggest that many of the cyanobacterial genes thus far characterized are subject to regulation by activator proteins. A number of DNA binding proteins and cis regulatory regions have been identified that are implicated in the control of transcription initiation.

I. Introduction The cyanobacteria are a very ancient and diverse group of photosynthetic bacteria which have a wide distribution in freshwater, marine, and terrestrial environments. They have been called the creatures of ‘earth, wind, and fire’, a reference to their ability to inhabit harsh and varied environments. Given the tolerance of modern strains for nutrient deprivation, temperature and salinity extremes, desiccation, and variations in light level and quality, the longevity of cyanobacteria as a group is not surprising. As with other bacteria, the challenges of adverse environments are met in the short term by rapid adjustments in metabolic pathways. Though the genetic mechanisms governing these adjustments in cyanobacteria are largely unknown, there is considerable precedent in other bacterial systems for control at the level of transcription. Models for bacterial transcriptional control are founded on early studies of E. coli operons where repressor and activator proteins were found to play a role in the transduction of environmental signals to the transcriptional apparatus. In the longer term, cyanobacteria can respond to environmental challenges at the level of cell structure, differentiating a variety of morphologically and functionally distinct cell types. These adaptive processes, by contrast to simple regulation of a few metabolic pathways, involve global changes in gene expression. Again, other models for transcriptional regulation during

procaryotic differentiation likely provide insights into the genetic mechanisms controlling cyanobacterial development. Very little has been written in review of the studies of transcription in cyanobacteria, and this article is an attempt to summarize our knowledge to date. Since the information on cyanobacteria is reviewed in the context of our understanding of transcription in the model eubacterial system E. coli, a brief summary of the current understanding of certain aspects of transcription in this organism precedes, and provides a framework for the discussion of cyanobacteria. Although we are still far from a detailed understanding of transcription in cyanobacteria, it should be clear from the other articles in this series that researchers in the field rapidly are accumulating detailed information on a number of biological processes, and developing the tools required for sophisticated analyses of gene function and control. Thus, the years ahead show great promise for yielding insight into how cyanobacteria regulate gene expression.

II. Transcription in E. coli: Paradigms for Eubacteria Much of our understanding of transcription in eubacteria is derived from extensive studies of this process in the model organism E. coli. Transcription

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in E. coli is thus the paradigm to which transcription in all other eubacteria is compared and evaluated. To facilitate discussions of transcription in cyanobacteria, a brief summary of some of the features and principles of transcription in E. coli is presented in the following sections. The discussion of transcription in both E. coli and cyanobacteria is confined to the topics of the transcriptional apparatus and the regulation of transcription initiation.

A. RNA Polymerase Structure and Function Transcription of gene sequences in eubacteria is performed by a single type of DNA-dependent RNA polymerase. The RNA polymerase of E. coli consists of two components: a core (E), that catalyzes the polymerization of RNA, and an additional factor, the sigma subunit, that confers on the core specificity for subsets of DNA sequences. The core component is a multisubunit complex composed of three proteins designated and in the stoichiometry 2:1:1 (Burgess 1971). The core complex is catalytically competent, but binds DNA nonspecifically and initiates transcription inefficiently. Association of a single subunit with the core forms a holoenzyme which can recognize and initiate transcription from specific promoter sites on DNA (Burgess, 1971). The primary sigma factor of E. coli is denoted (superscript = molecular weight of the factor The E. coli holoenzyme, recognizes promoter sequences consisting of consensus hexamers centered approximately 10 and 35 bases upstream (–10 and –35) from the first transcribed base which is designated (+)1 (Hawley et al., 1983; Harley et al., 1987). Crosslinking studies with and promoter sequences (Chenchick et al., 1981, 1982) and analysis of sigma mutations (Gardella et al., 1989; Siegele et al., 1989) suggest that the sigma subunit recognizes and contacts the consensus hexamers of the promoter. This sequence-specific DNA binding of recently has been shown to require an allosteric interaction with the core of the RNA polymerase (Dombroski et al., 1992). In the initial step of transcription, binds to double-stranded DNA at the promoter site to form what is termed the ‘closed’ complex (Chamberlain, 1974). At this stage, the complex can either disassociate from the DNA or initiate transcription. The stability of the closed complex may be affected by other DNA binding proteins which activate or

615 repress transcription. Following stabilization of the complex, a region of the DNA helix is melted to form the ‘open’ complex. This process exposes the transcription start site on the template strand and allows the holoenzyme to catalyze the polymerization of the first few bases of the nascent mRNA (Chamberlain, 1974). The factor dissociates from the holoenzyme after transcription of the first eight to twelve bases while the core polymerase continues elongation of the message. Following dissociation, the factor may join other core enzymes and go through many cycles of initiation and disassociation (Burgess, 1971).

B. Alternate Sigma Factors Bacterial cells are thought to employ one principal sigma factor for the transcription of most genes during exponential growth. Features of the sigmacore interaction such as dissociation and cyclic reassociation suggested that a variety of factors could be utilized by the core enzyme (Burgess et al., 1969). These alternative factors could confer on the core the ability to initiate transcription from different sets of genes by modifying the promoter recognition specificity of the RNA polymerase. This hypothesis has been shown to be correct, and the utilization of alternative sigma factors for coordinate expression of genes during specialized conditions now is believed to be a general feature of eubacteria (Helmann and Chamberlain, 1988). Alternative sigma factors were first discovered in studies of phage infection and endospore development in Bacillus subtilis (reviewed in Doi et al., 1986), and temporal gene expression during these processes was proposed to be regulated through a cascade of alternative sigma factors (Losick et al., 1981). Indeed, the sequential activation of sets of genes during endospore formation in B. subtilis has been shown to require an intricate interplay of temporally regulated alternative sigma factors in both the mother cell and prespore (reviewed in Losick et al., 1992). Since the initial discoveries in Bacillus species, alternative sigma factors have been identified in a number of gram-negative and gram-positive eubacteria, and they have been shown to activate discrete sets of genes in response to a variety of conditions. Among these factors are those that control gene expression during bacteriophage infection, during motility and chemotaxis, in response to heat shock and other stresses, or at specific times during developmental

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616 programs (Helmann and Chamberlain, 1988). Another well studied class of alternative sigmas, the family, is able to regulate a number of different gene sets involved in a variety of physiological roles (Kustu et al., 1989).

C. Sigma Factor Structure The characterization and sequence analysis of sigma factors has led to their classification into two broad groups which differ in both their structure and regulation (Lonetto et al., 1992). One class of sigma factors is structurally similar to of E. coli and is designated the family. The second group has similarity to of E. coli and forms the family.

1. The

Family

Members of the family are homologous to the principal sigma factors of E. coli and B. subtilis. Regions of sequence similarity in alignment of family sequences generally fall into four domains, numbered 1 to 4 from amino to carboxyl terminus; these four domains are often further divided into subregions (Fig. 1; Helmann and Chamberlain, 1988; Lonetto et al., 1992). Analyses of mutations that alter sigma function together with secondary structure predictions of conserved motifs have provided information on the likely functions of sigma domains (reviewed in Helmann and Chamberlain, 1988; Lonetto et al., 1992). Regions 2 and 4 are the most highly conserved among all family members. Subregions 2.4 and 4.2 are thought to contact specific base pairs in the –10 and –35 regions, respectively, of cognate promoters

(Fig. 1). Other subdomains of region 2 are implicated in DNA melting. Regions 1 and 3 are less conserved and their functions are not well understood. Region 1 may play a role in structural integrity while region 3 may participate in core binding. Subdomains of these regions can be used to distinguish alternative and primary sigma factors. Alternative sigmas lack subregion 1.1, and in some cases subregion 3.2 (Fig. 1). This model of domain function suggests that there is a linear alignment of DNA binding domains and the conserved elements of cognate promoters (Lonetto et al., 1992). The –35 promoter element is in contact with the carboxy-terminal sigma domain (4.2) and the –10 promoter element is in contact with a domain near the N-terminus (2.4; Fig. 1). The family has been divided into three groups based on the role of the sigma factor and its similarity to other family members (Lonetto et al. 1992). The principal or primary sigma factors, which are essential for cell growth and display high sequence similarity, form Group 1. A number of family members are very similar in sequence to principal sigma factors but are distinguished by their dispensability during cellular growth. These sigmas, of which E. coli SigS and Streptomyces coelicolor HrdA, C and D are examples, are included in Group 2. The final Group, 3, is composed of alternative sigma factors, which direct the expression of specialized sets of genes in response to particular environmental conditions or cellular programs. The sequences of the sigmas in this group are the most diverged from that of the primary factors.

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2. The

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Family

Members of the family have been identified in a number of different bacteria (reviewed in Kustu et al., 1989). In contrast to alternative sigma factors of the family which control regulons expressed during a particular physiological response or cellular program, the homologs are able to direct the expression of genes involved in a wide variety of biological processes and physiological responses. These include, but are not limited to functions as diverse as nitrogen assimilation, nitrogen fixation, dicarboxylic acid transport, and flagellar synthesis (Kustu et al., 1989). Sigma factors of the family show little sequence similarity to proteins of the family. In addition, genes transcribed by differ from those transcribed by in both promoter structure and in the regulation of transcription initiation (see Section II D).

D. Structure and Function of Bacterial Promoters A promoter is a sequence of DNA recognized by the RNA polymerase and to which the RNA polymerase subsequently binds to initiate transcription. Promoters thus encode structural information that is interpreted by the transcription apparatus. To some degree, this information is carried by the base sequence of the promoter in the form of specific motifs with which the RNA polymerase directly interacts. These sites of interaction are held in an appropriate threedimensional conformation within the DNA helix. The function of promoters requires sequence conservation in some regions while other regions are free to vary as long as alternate sequences preserve the same preferred topology. Among eubacterial promoters, those of E. coli have been studied most extensively, and the general features of these promoters are conserved in many other eubacteria. E. coli promoters can be placed in two categories based on the type of factor or required for transcription.

1.

Promoters

The promoters recognized by can be placed in two different classes. For genes in the first class, the basic unit of control is the basal promoter which is limited to the core promoter elements. For these genes, the basal promoter alone specifies the

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frequency of transcription initiation. In the second class, the intrinsic activity of the basal promoter is modified by activation or repression. These regulated promoters consist of basal promoter elements upon which additional cis elements are superimposed .

a. Basal Promoters The most important features of the basal promoter are short, conserved sequence motifs which are apparent in alignments of even a few transcription initiation sites and 5' proximal sequences. A large survey of 263 E. coli promoters confirmed the prevalence of two consensus hexamer sequences upstream of transcription start sites at position +1: the sequence TTGACA centered at 35 bp upstream (–35), and the sequence TATAAT centered at 10 bp upstream (–10; Hawley and McClure, 1983; Harley and Reynolds, 1987). Although no base within either hexamer motif is invariant among all promoters of the database, the first three positions of the –35 element (TTGNNN) and three positions in the –10 element (TANNNT) are conserved in 80–90% of the promoters. The third element of the core, the spacer DNA between hexamers, is highly conserved in length; 92% of the promoters surveyed display spacer lengths of 17 ±1 bp. The distance between the –10 hexamer and the first transcribed base is also constrained. In the majority of promoters, transcription begins with a purine located 7 ± 1 bp downstream of the –10 hexamer (Hawley and McClure, 1983; Harley and Reynolds, 1987). The importance of the core promoter elements identified by sequence inspection has been demonstrated in a large number of functional studies. These studies, which include analysis of native promoters as well as mutant promoters with altered hexamer sequences and altered spacer lengths (reviewed in Gralla, 1990), led to the ‘consensus rule’. This rule states that the more similar promoters are to the consensus derived from comparison of promoter sequences, the higher the rate of transcription. A promoter is weakened by changes that lead away from the consensus –10 and –35 hexamer sequences; similarly, changes that alter the consensus spacer length weaken the promoter. Alterations in hexamer sequences are thought to affect the stability of the closed complex, while alterations in spacer length change the alignment of the hexamers along the helix and affect interactions with the RNA

618 polymerase. Sequences outside the core (within –45 to +20), have also been found to have some influence on transcription (Gralla, 1990). This expanded region coincides with the sequences contacted by RNA polymerase in the open complex (Siebenlist and Gilbert, 1980). As discussed earlier, alternative sigma factors are able to confer different promoter specificities on the core polymerase. The similarity in structure of with the alternative sigma factors of the family is mirrored in the similarity of promoter structures recognized by these sigma factors. Alternative factors of this class also recognize conserved promoter elements approximately centered at –10 and –35 from the transcription initiation site. Specificity is imparted, however, through the use of different consensus sequences at the two positions. The cognate promoters of individual alternative sigma factors generally differ from those of as well as the other alternative factors in the family in the sequences at both–10 and –35 (Helmann and Chamberlain, 1988). This is reflected in the fact that the sigma subdomains which recognize the consensus elements of the promoter (2.4 and 4.2) differ in sequence among alternative sigmas and the primary sigma factors (Lonetto et al., 1992).

b. Regulated Promoters Functionally important sequences in basal promoters are limited to the core elements and immediately adjacent regions that interact directly with RNA polymerase. However, many promoters are subject to more complex regulatory mechanisms involving interactions with trans-acting factors that are not part of the polymerase. These promoters are composed of both the core elements described above and additional regions that include binding sites for accessory factors. In such cases, the basic promoter region confers a basal transcriptional activity that can be modified by the binding of accessory factors (repressor or activator proteins). The principles which govern the location of binding sites relative to the core promoter and the functional nature of accessory protein-RNA polymerase interaction emerge from a survey of more than 100 regulated operons (Collado-Vides et al., 1991). The survey reveals that regulation can be elaborate. Many operons are subject to both activation and repression, and nearly half are considered ‘complex’ in that they are part of more than one

Stephanie E. Curtis and James A. Martin regulatory system or are transcribed from more than one promoter, each subject to repression or activation. Activators are accessory proteins that increase the rate of transcription initiation from basal promoters. Binding sites for activators of promoters are generally located in close proximity upstream from the –35 element of the basal promoter, and in more than two thirds of the operons surveyed the activator binding site overlaps the –40 position (ColladoVides et al., 1991). This configuration is thought to be important in allowing direct contact between the activator and RNA polymerase bound at the core elements. The function of the activator is negatively influenced by increasing distance from the basal promoter, and is also affected by the sequence of the core consensus hexamers (Gralla, 1991). These features reflect the role that the activator is thought to play in transcription initiation. Most promoters with activator binding sites show relatively weak conservation at the –35 core consensus hexamer and very low activities in the absence of bound activator. Thus, the stabilization of closed complex formation conferred by activators may substitute for well conserved–35 elements (Collado-Vides et al., 1991). Activation sites never overlap with the –10 hexamer elements, presumably because such a position would interfere with DNA melting which initiates in this region (Collado-Vides et al., 1991). Repression has a negative effect on transcription initiation, that unlike activation, can influence all stages of initiation including open and closed complex formation, and ‘promoter clearance’, the translocation of RNA polymerase during the initial stages of elongation. As with activation sites, one repressor site (operator) must be near the region of RNA polymerase binding. Operators are found in several positions relative to the RNA polymerase protected region: between the –10 and –35 elements, immediately upstream from the –35 element, just downstream from the –10 core element, or, rarely, at remote upstream or downstream sites (Collado-Vides et al., 1991). The first two sites are well positioned to disrupt closed complex formation since repressor binding would preclude polymerase binding at the same site. Operator sites downstream from the –10 element are positioned to disrupt later stages in initiation, such as promoter clearance or elongation. Analysis of the Collado-Vides et al (1991) database reveals that regulatory elements are often overlapping or present in multiple copies within the regulatory region. These arrangements facilitate regulatory

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mechanisms that are more complex than simple repression or activation. In many promoters, operator and activation sites overlap, and many of the remote operators are thought to act by preventing the binding of activators rather than direct inhibition of RNA polymerase. Multiple activator or operator sites also are common in the promoters surveyed. Operators for the same repressor are duplicated in approximately half of the promoters surveyed and these elements usually occur in a closely spaced arrangement. The extra operator is thought to enhance repressor binding through cooperative interactions between factors bound at adjacent sites (Collado-Vides et al., 1991). About a third of the promoters in the database that are subject to activation have multiple activator sites, either for the same or different activator proteins. As with duplicated operators, closely spaced activator sites for the same activator enhance regulation through cooperative interactions. Among the complex class of promoters are examples in which multiple binding sites for two different activators are interspersed. Binding of both activators leads to formation of a large multiprotein scaffold around which the regulatory DNA becomes wrapped. This serves to deliver the most distant regulatory elements toward the polymerase bound at the basal elements. The use of multiple binding proteins to bridge the distance between the polymerase and distal regulatory sites is reminiscent of mechanisms employed in the regulation of eucaryotic promoters (Gralla, 1991).

2.

Promoters

Operons transcribed by differ fundamentally from those transcribed by in both promoter structure and in the regulation of transcription initiation (reviewed in Kustu et al., 1989; ColladoVides et al., 1991; Gralla, 1991). While basal promoters include two conserved sequence elements, centered approximately at –12 and –24 from the site of transcription initiation, these motifs are distinct from those of promoters in both sequence and position. The processes of open and closed complex formation also differ in and transcription. can bind stably to a cognate promoter in a closed complex, but interaction with an activator protein as well as ATP hydrolysis is required for formation of an open complex and transcription initiation. Similar to the activation of promoters, the activator must contact the RNA

619 polymerase, however, the mechanism by which this is achieved differs in the two types of promoters. The activator sites for promoters are often multicopy and are enhancer-like, in that they are arranged at some distance from the polymerase binding site. Although the distance varies, the majority of activator sites are located about 110 bp upstream of the transcription initiation site (Collado-Vides et al., 1991). Contact between the activator and polymerase occurs through the looping out of intervening DNA between the activator site and basal promoter elements. The integration host factor (IHF) is often used as a coactivator, assisting in stabilization and activation (Kustu et al., 1989). As discussed earlier, members of the family direct the expression of genes involved in a wide variety of diverse biological processes. Transcription of different gene sets is controlled by the use of different activator proteins (Kustu et al., 1989). Regulation of activation is in turn achieved in part by modulation of the activator proteins under different physiological conditions. Modulation is mediated through a number of different mechanisms including phosphorylation and modification by other proteins (Kustu et al., 1989). Many features of transcription including the arrangement of regulatory elements and several aspects of transcription initiation are paralleled in the transcription of eucaryotic genes by RNA polymerase II (Gralla, 1991).

III. Transcription in Cyanobacteria A decade ago, a relatively small number of genes had been characterized from cyanobacteria, and virtually nothing was known about the regulation of gene expression in cyanobacteria. Our knowledge of the characteristics of cyanobacterial genes and their regulation has increased exponentially during the last ten years, facilitated by advances in molecular biology techniques, and the development of tools for genetic analysis in cyanobacteria (Haselkorn, 1991). Although many questions remain unanswered, a good deal of information on transcription in cyanobacteria has accumulated, particularly with regard to the transcriptional apparatus and transcription initiation. The information on these topics is discussed in the following sections, while aspects of transcription for which very little is known, such as transcription termination, are excluded.

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A. The Transcriptional Apparatus The subunit structure of the E. coli RNA polymerase has been found to be well conserved among eubacteria (Chamberlain, 1976; Zillig et al., 1985), and is generally thought to be universal. Early characterizations of cyanobacterial RNA polymerases (Herzfeld et al., 1971; Miller et al., 1978) were not in agreement about whether this rule extended to the cyanobacteria. The subunit composition of cyanobacterial RNA polymerases was resolved by studies of the enzyme from Anabaena sp. strain PCC 7120 (Schneider et al., 1987) that were subsequently extended to other cyanobacteria (Schneider et al. 1988b). These studies showed that although the basic features of the E. coli enzyme are preserved, cyanobacteria possess a RNA polymerase of unique structure.

1. RNA Polymerase Core Structure The unique aspect of cyanobacterial RNA polymerases is found in the composition of the core. This component of the cyanobacterial enzyme is very similar to that of E. coli with one notable exception. In addition to the and proteins, an extra polypeptide termed contributes to the core structure. The identified in Anabaena sp. strain PCC 7120, was shown to be a distinct holoenzyme subunit (Schneider et al. 1987), and a component of holoenzymes from fifteen other cyanobacteria representing three of the five major cyanobacterial subgroups (Schneider and Haselkorn, 1988b). Evidence that the subunit is serologically related to the subunit of E. coli led to the suggestion that the

Stephanie E. Curtis and James A. Martin domains of the subunit are split in cyanobacteria between the and subunits (Schneider and Haselkorn, 1988b). This hypothesis was borne out by the analysis of cyanobacterial RNA polymerase genes. The genes encoding the and subunits have been characterized from Nostoc commune UTEX 584 (Xie et al., 1989) and Anabaena sp. strain PCC 7120 (Bergsland et al., 1991), and in both cyanobacteria these genes are tightly linked in the order rpoB-rpoCl-rpoC2. It was apparent from the alignment of rpoCl and rpoC2 with the gene encoding the subunit of E. coli (rpoC), that the cyanobacterial and subunits correspond to the amino- and carboxy-terminal portions of E. coli respectively (Fig. 2). This arrangement is analogous to that in many chloroplast genomes in which the homolog of the E. coli subunit is encoded by two linked genes (rpoCl and rpoC2) for the plastid holoenzyme subunits designated and (Fig. 1; (Hu et al., 1991; Hudson et al., 1988; Shimada et al., 1990). Cyanobacteria and chloroplasts also share another organizational feature that differs from the E. coli arrangement. The rpoBC genes of E, coli are part of an operon with several ribosomal protein genes (rplKAJL-rpoBC), and the polymerase subunits are transcribed from the upstream ribosomal protein gene promoter (Downing et al., 1987). In contrast, the rpoBClC2 genes of cyanobacteria are not closely linked to the analogous ribosomal protein genes (Fig. 2; Sibold et al., 1990; Bergsland and Haselkorn, 1991). This is similar to the arrangement in plants, where the chloroplast rpoBC1C2 cluster is unlinked to the ribosomal protein genes which are located in the nuclear genome (Sibold and Subramanian, 1990).

Chapter 20

Transcription in Cyanobacteria

The rpoBC1C2 genes of chloroplasts are cotranscribed (Purton et al., 1988), however, it has been reported that the three genes do not form an operon in the cyanobacterium Nostoc commune UTEX 584 (Xie and Potts., 1990). In contrast to the rpoBC genes, the rpoA gene encoding the subunit of the holoenzyme core of chloroplasts and cyanobacteria displays a organization that resembles those of B. subtilis and E. coli. In E. coli the rpoA gene is included in an operon with ribosomal protein genes (rpmJ-rpsM-rpsK-rpsDrpoA-rplQ; Bedwell et al., 1985). A similar arrangement, rpmJ-rpsM-rpsK-rpoA-rplQ is found in B. subtilis (Boylan et al., 1989). Recent analysis of the rpoA gene locus from Synechococcus sp. strain PCC 7002 indicates an arrangement identical to that found in B. subtilis (L. Caslake and D. A. Bryant, unpublished). This same arrangement is also found in the cyanelle genome of Cyanophora paradoxa (V. L. Stirewalt and D. A. Bryant, personal communication). Interestingly, in chloroplasts the rpoA gene is linked to, and cotranscribed with, two ribosomal protein genes (rpmJ-rpsK-rpoA; Purton and Gray, 1988).

2. Sigma Factors The first cyanobacterial sigma factor described, SigA, was identified as a component of the major holoenzyme purified from vegetative cells of Anabaena sp. strain PCC 7120 that was able to confer promoter-specific transcriptional activity on the core enzyme in reconstitution experiments (Schneider et al., 1987). An oligonucleotide probe corresponding to partial amino acid sequence of the SigA subunit was used to isolate the corresponding gene (sigA; Brahamsha and Haselkorn, 1991). As in the case of rpoBC1C2, the relative organization of sigA differs from that of the analogous gene (rpoD) of E. coli. The sigA gene is monocistronic, while rpoD is included in an operon with genes for a DNA primase and a ribosomal protein (Burton et al., 1983). Alignment of the SigA amino acid sequence with that ofB. subtilis and E. coli revealed that the essential structure of these other principal factors is maintained in SigA. The four general regions of homology that characterize the protein family (Helmann and Chamberlain, 1988; Fig. 1) are all present in SigA. As with other members of the family, SigA domains 2 and 4, which are implicated in promoter contact, are the most similar to those of

621 and Regions 2 and 4 of SigA share 76% and 61% amino acid identity, respectively, with the corresponding regions of from E. coli (Brahamsha and Haselkorn, 1991). The pattern of sigA expression was examined in cells grown on nitrogen-replete and nitrogen-deplete medium. The transcript pattern is somewhat complex. Two sigA transcripts of 1.7 and 2.4 kb are detected in vegetative-cell RNA by Northern blot analysis, and five 5' transcript termini have been mapped, some of which vary in abundance when cells are grown in nitrogen-replete versus nitrogen-deplete conditions (Brahamsha and Haselkorn, 1991). Like many other eubacteria characterized, cyanobacteria also contain multiple species of sigma factors. In addition to sigA, two other sigma factor genes, sigB and sigC, from Anabaena sp. strain PCC 7120 have been characterized (Brahamsha and Haselkorn, 1992). The sigC gene was isolated by its homology to sigA, while sigB was isolated by homology to the rpoD oligonucleotide described by Tanaka et al. (1988). This probe corresponds to an amino acid sequence conserved in the principal sigma factors of E. coli and B. subtilis, but that is not conserved in alternative sigma species. The predicted protein products of SigB and SigC are very similar to each other and to SigA, and thus display features of principal sigma factors. Expression of sigB and sigC is not detected under nitrogen-replete growth conditions of Anabaena sp. strain PCC 7120. Transcripts for both genes appear transiently after nitrogen starvation (Brahamsha and Haselkorn, 1992), suggesting that their products might regulate the expression of genes involved in heterocyst development or nitrogen fixation. Generation of sigB and sigC single and double mutants by insertional inactivation, however, demonstrated that neither gene is required for vegetative growth, heterocyst differentiation or growth in the absence of combined nitrogen (Brahamsha and Haselkorn, 1992). Multiple sigma factor genes have also been identified by hybridization studies in three unicellular cyanobacteria, Synechococcus sp. strain PCC 7942 and Synechocystis sp. strain PCC 6803 (Tanaka et al., 1992b) and Synechococcus sp. strain PCC 7002 (L. Caslake and D. A. Bryant, personal communication). Four sigma genes have been characterized from Synechococcus sp. strain PCC 7942 (Tanaka et al., 1992a; Tanaka et al., 1992b); three sigma genes (of the estimated five-six total genes) from

622 Synechococcus sp. strain PCC 7002 have been cloned, sequenced, and overexpressed in E. coli (L. Caslake and D. A. Bryant, personal communication). All of the Synechococcus sp. sigma genes thus far characterized encode products with conserved domains characteristic of principal sigma factors of the class. In each of these cyanobacteria the gene with greatest similarity to sigA of Anabaena sp. strain PCC 7120 (rpoD1 in 7942 and sigA in 7002) is inferred to encode the sigma subunit of the major form of RNA polymerase. In Synechococcus sp. strain PCC 7002 the sigA gene is required for viability as judged by the inability to segregate interposonmutated alleles of this gene. Unlike the situation in Anabaena sp. strain PCC 7120, transcripts for sigA, sigB, and sigC are readily detected in Synechococcus sp. strain PCC 7002 cells growing in nitrogen-replete conditions. However, transcript levels for sigA and sigC decrease to barely detectable levels when cells are starved for combined nitrogen. Under these same conditions, transcript levels for sigB increase markedly (L. Caslake and D. Bryant, personal communication). Under nutrient-replete growth conditions, the growth rate of a sigB interposon mutant strain is identical to that of the wild type strain at high light intensity, although the growth rate of the mutant is somewhat slower than that of the wild type at low light intensity (L. Caslake and D. A. Bryant, personal communication). The roles of the multiple sigma factors in cyanobacteria remain to be elucidated. The ten sigma factors whose sequences have thus far been characterized all have features of principal sigmas and thus can be classified as family members. The apparent dispensability of SigB and SigC from Anabaena sp. strain PCC 7120 (Brahamsha and Haselkorn, 1992) would place these in Group 2 of the family. A pattern similar to that ofAnabaena sp. strain PCC 7120 is seen inStreptomyces coelicolor for which four sigma factor genes (hrdA-D) of the principal type have been characterized (Buttner et al., 1990; Tanaka et al., 1991). Disruption of the hrdB gene is lethal, suggesting that it encodes the principal sigma factor, while hrdC and hrdD mutants are viable and unaffected in gross morphology and differentiation (Buttner et al., 1990). Experiments to assess the effects of disruption of the various sigma genes of Synechococcus sp. strains PCC 7942 and 7002 are currently in progress (K. Tanaka, personal communication; L. Caslake and D. Bryant, personal communication). These studies should provide

Stephanie E. Curtis and James A. Martin information on the role of the multiple sigmas in these organisms and whether the general pattern observed in Anabaena sp. strain PCC 7120 and Streptomyces sp. is universal among cyanobacteria. By analogy with other eubacteria, sigma factors of the alternative type would be expected to play a role in adaptive processes, and in the temporal control of gene expression during development. Alternative sigma factors of the family have been shown to regulate gene expression during B. subtilis differentiation (Losick and Stragier, 1992), but such factors have not yet been shown to participate in heterocyst development, one of the best studied differentiation pathways in cyanobacteria. There is indirect evidence, however, that a cyanobacterial homolog of E. coli may exist in cyanobacteria. This is an alternative sigma factor of the family which controls the expression of heat shock genes (Grossman et al., 1984). Characterization of Synechococcus sp. strain PCC 7942 groESL (Webb et al., 1990), a heat-shockinduced operon, identified promoter sequences with similarity to those recognized by E. coli (Cowing et al., 1985; Fig. 5). The recA gene of Synechococcus sp. strain PCC 7002, which is heat-shock-inducible, also has sequence elements that resemble the promoter (Murphy et al., 1990; Fig. 4). In some bacteria, is responsible for the transcription of nitrogen fixation and nitrogen regulated genes (Kustu et al., 1989), and thus such genes in cyanobacteria are likely places to identify regulation. While conserved sequence motifs have been identified in the promoter regions of the small number of cyanobacterial nitrogen fixation operons for which transcription initiation sites have been mapped (Mulligan and Haselkorn, 1989), these conserved motifs do not conform to the promoter consensus sequences, and as yet there is no evidence that cyanobacterial nif genes are regulated by In addition, there is direct evidence that one of the best characterized genes in enteric bacteria, glnA, is not similarly regulated in cyanobacteria. Transcription assays performed in vitro with the Anabaena sp. strain PCC 7120 RNA polymerase showed that the glnA gene is transcribed by the holoenzyme bearing the principal sigma factor (Schneider et al., 1991). It should be noted that the glnA genes of several other bacteria also show a lack of dependence (Kustu et al., 1989). Although alternative sigma factors of either the or family have not been identified yet in cyanobacteria, hybridization experiments with sigA

Chapter 20

Transcription in Cyanobacteria

suggest that other sigma-like genes exist in the genomes of Anabaena sp. strain PCC 7120 (Brahamsha and Haselkorn, 1992)) and Synechococcus sp. strain PCC 7002 (L. Caslake and D. Bryant, personal communication). Thus, it is likely that additional sigma factors will be identified, and that some of these will be sigma factors of the alternative type.

B. Cyanobacterial Promoters The interaction of the RNA polymerase with the promoter is one of the initial steps in which transcription initiation is regulated. For most transcriptionally regulated genes, the basal promoter is a major determinant of the mechanisms required for regulation. The model of procaryotic gene regulation presented in previous sections suggests that basal promoters have an intrinsic activity that can be modified by accessory transcription factors or by the availability of RNA polymerases bearing cognate sigma factors. Thus, understanding the basis of this intrinsic activity can help to predict which of these mechanisms is operable for a given gene. A picture of the general features of Cyanobacterial promoters is emerging from several different types of studies on gene regulation. The information gained from sequence comparisons, and from transcription assays performed in vivo and in vitro is summarized in the following sections.

1. Sequence Comparisons Given the overall similarity in the structure of Cyanobacterial RNA polymerases to that of E. coli, Cyanobacterial basal promoter sequences are expected to lie within 50 bp upstream of transcription initiation sites. As in E. coli, comparison of such sequences should allow identification ofcore promoter elements. Transcripts now have been mapped for a number of genes (~50) from several different cyanobacteria using either primer extension and/or nuclease protection techniques (Sambrook et al., 1989). One caveat concerning such studies: The mapped 5' termini of RNA transcripts do not necessarily represent the 5' ends of primary transcripts. Primary transcripts may be processed or degraded, or the method of transcriptmapping mayproduce artifacts. Additional evidence that the transcript map site represents a transcription initiation site is necessarily shown by independent studies which demonstrate that the

623 transcript is a primary one (such as capping experiments) or demonstrate promoter function (in vivo or in vitro transcription assays). Figs. 3–6 show compilations of putative promoter regions from the four species of cyanobacteria for which the most information is available. With one exception, only sequences derived from mapping cellular transcripts during standard vegetative growth are included. The databases for several of the organisms are very small and even the overall database from all cyanobacteria is well below the size of that analyzed in E. coli (263 promoters; Harley and Reynolds, 1987). Despite this limitation, several features emerge from the promoter comparisons that are observed in >70% of the promoter sequences from all cyanobacteria, and thus are likely to be significant. These features include: 1) a conserved motif that conforms in both sequence and position to the –10 hexamer consensus of E. coli promoters, 2) transcription initiation 7 ±1 bp downstream from the conserved –10 motif and 3) transcription initiation with a purine. Less conserved but present in approximately half of the promoters is a sequence at the appropriate position that somewhat resembles the –35 consensus sequence ofE. coli promoters. The conservation of this motif is relatively weak compared to that observed in the E. coli database, where the first three positions of the hexamer are conserved in 80–90% of the promoter sequences. These three positions are conserved in less than 25% of the sequences in the Cyanobacterial database. This is in contrast to the Cyanobacterial –10 sequences in which the most highly conserved nucleotides of the E. coli hexamer (TANNNT) are conserved in > 70% of the Cyanobacterial sequences. Two promoter sequences from the cyanophage N-l (Fig. 1) are unique in that both the –10 and –35 elements closely approximate the canonical hexamers of E. coli promoters. For sequences in which the E. coli –35 motif is absent, no other conserved motif is apparent in all sequences at that position or elsewhere within 50 bp of the transcription initiation site. In some cases, however, operons which are coordinately expressed exhibit similar sequences in the –35 region. An example is the atp1 and atp2 operons of Anabaena sp. strain PCC 7120 which encode subunits of the ATP synthase; these subunits must be coordinately expressed for assembly of a functional complex. As seen in Fig. 3, the atp1 and atp2 operons have a sequence at –35 that conforms to AANTNT.

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Stephanie E. Curtis and James A. Martin

Chapter 20

Transcription in Cyanobacteria

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The promoter database for Anabaena sp. strain PCC 7120 genes is the largest, and as indicated in Fig. 3, a number of the sites to which the 5' termini of cellular transcripts have been mapped have been shown experimentally to be bona fide transcription initiation sites (see IIIB2,3). In an analysis of this database for sequences conserved outside the –10 and –35 regions, certain nucleotides were found to be present at specific positions at significantly greater than the expected frequency. A few ofthese coincide with positions and sequences that are also conserved in E. coli promoters. Whether these conserved residues are statistically or functionally significant awaits the outcome of analyses of a larger database, as well as functional studies on the relative importance of individual base pairs within the promoter. In summary, sequence comparisons suggest that cyanobacterial basal promoters share a core element with E. coli promoters that is positioned ten nucleotides from the transcription initiation site.

Stephanie E. Curtis and James A. Martin

While some promoters also have a sequence that resembles the –35 element of E. coli promoters, in the vast majority of cases this element is weakly conserved or absent.

2. In vitro Assays of Promoter Activity The ability to purify RNA polymerase makes possible in vitro assays of transcriptional activity. The advantage of in vitro transcription assays is that the contribution of individual components of the transcription machinery can be tested on specific DNA templates in a well defined system. For example, such assays were utilized in the identification of the sigma subunit of the Anabaena sp. strain PCC 7120 RNA polymerase holoenzyme (Schneider et al., 1987). In vitro transcription assays can also be applied to the measurement of transcriptional activity from specific DNA templates. In cyanobacteria, in vitro transcription assays have been used to verify

Chapter 20

Transcription in Cyanobacteria

transcription initiation sites initially identified by mapping the 5' termini of cellular transcripts, and to define the type of basal promoter sequences preferred by the RNA polymerase. The transcription of nine Anabaena sp. strain PCC 7120 genes expressed in vegetative cells (rbcL,petF, atp2, hupB,psbB,psbA, sigA, woxA), as well as two cyanophage N-1 promoters and a bacteriophage T4 promoter have been examined with in vitro transcription assays. These studies have been performed with purified RNA polymerase from Anabaena sp. strain PCC 7120 (woxA template; Borthakur, et al., 1990), crude extracts ofAnabaena sp. strain PCC 7120 active in transcription (sigA template; Brahamsha and Haselkorn, 1991) or both (rbcL, petF, atp2, hupB, psbB, psbA, N-1, T4 templates; Schneider et al., 1991). In all cases, the templates assayed were circular plasmid DNAs in which promoter regions were cloned upstream from a strong transcription terminator. Transcription from such templates resulted in transcripts whose lengths were compared to those expected based on the transcription initiation sites determined with cellular transcripts and the distance to the transcription

627

terminator. In all but a few exceptions, the transcription start points, identified by mapping studies with cellular transcripts were confirmed by the in vitro transcription assays (Borthakur et al., 1989; Brahamsha and Haselkorn, 1991; Schneider et al., 1991). This included genes for which multiple transcription start points had been identified (glnA, atp2, psbB , sigA). Qualitative comparisons of the in vitro activities of the rbcL,glnA,psbB,psbA, hupB,petF, atp2, N-1, and T4 promoters (Schneider et al., 1991) showed that the RNA polymerase is most active on templates that approximate the E. coli consensus promoter, such as those of T4 and cyanophage N-l (Fig. 3). The promoters for the other Anabaena sp. strain PCC 7120 genesfunctionrelativelyweakly inthese assays; these promoters share some but not all of the E. coli consensus features (Fig. 3). Although the activity of the rbcL andpsbA promoters was higher in the crude extracts than with purified polymerase, the activities were still much lower than those of the N-l and T4 promoters (Schneideret al., 1991). These data suggest that, as for the principal RNA polymerases of E. coli and B. subtilis, the consensus rule applies to

628 transcription in vitro by Anabaena sp. strain PCC 7120 RNA polymerase.

3. In vivo Assays of Promoter Activity The ability to introduce DNA into cells makes possible in vivo analyses of promoter activity. Such assays have the advantage that all of the transcription components that would normally function in gene expression are present in the particular cell type or under the particular growth condition in which the constructs are assayed. These components would include holoenzymes bearing minor sigma factors as well as trans-acting regulatory factors such as repressors and activators. Functional assays of promoter activity have been performed in vivo on a small collection of cyanobacterial genes. These studies make use of constructs in which isolated gene promoter regions are fused to promoterless reporter genes. The reporter gene fusions are introduced into cells and the ability of the cloned promoter to drive expression of the reporter gene in vivo is measured by quantitation of either the level of gene product, or the activity of the gene product in cell extracts. The type of reporter gene and the nature of the construct assayed differ in the cyanobacterial systems in which gene promoters have been studied. The regulation of Synechococcus sp. strains PCC 7942 and 7002 genes has been examined using chromosomally integrated gene (lacZ) fusions. The gene encoding chloramphenicol acetyl transferase (cat) carried on replicative plasmids has been used as a reporter in the study of genes from Anabaena sp. strain PCC 7120. Other ‘promoterprobe’ systems with similar features also have been developed (Ferino et al., 1989; Milkowshi et al., 1991; Scanlan et al., 1990) and should prove useful in promoter studies. The analysis of gene promoters in vivo has been important in the verification of transcription initiation sites, the identification of basal promoter elements, the assessment of promoter mutations, and the dissection of regulatory elements.

a. Verification of Transcription Initiation Sites and Identification of Minimal Basal Promoters The ability of cloned promoter fragments to drive the expression of reporter genes in vivo has been demonstrated for a number of cyanobacterial genes

Stephanie E. Curtis and James A. Martin including the rbcL, atp1, petF, psbAI operons of Anabaena sp. strain PCC 7120 (Martin et al., 1994a,b; J. Martin and S. E. Curtis, unpublished results) and the psbAI-III (Schaefer et al., 1989) and psbDII (Bustos et al., 1991) operons of Synechococcus sp. strain PCC 7942 and the cpcBACDEF operon of Synechococcus sp. strain PCC 7002 (Gasparich et al., 1987). In each of these cases, transcription ofthe reporter gene was shown to initiate from the same position to which the 5' termini of the cellular transcript had been shown to map, demonstrating that these positions represent transcription initiation sites (Figs. 3, 5). Many cyanobacterial genes appear to have multiple transcription initiation sites. For example, the atp2 (Curtis, 1987) and psbB (Lang et al., 1989) operons of Anabaena sp. strain PCC 7120 are each represented by two cellular transcripts that differ in length at their 5' termini. To determine whether the two transcripts from each gene result from processing or degradation events, or alternatively, derive from expression from separate promoters, constructs were designed in which the contribution of individual promoter regions could be tested. Assays of these constructs in vivo indicated that the two cellular transcripts are expressed from different promoters for both psbB (Lang, et al., 1991) and atp2 (Martin et al., 1994b). In the case of atp2, the two promoters have different activities by in vivo assays (Martin et al., 1994b). Greater activity is observed from the downstream promoter, atp2(I), mirroring the difference in abundance of the two native transcripts in cellular RNA (Curtis, 1987). This result is opposite to that obtained in the in vitro assays (Schneider et al., 1991), in which the upstream promoter, atp2(II), gives greater activity. Consistent with the overall results from in vitro transcription (see Section II B, 2), the atp2(II) promoter has a closer match to the E. coli consensus than does the atp2(I) promoter. The minimal region required to maintain basal transcription ofcyanobacterial genes has been studied with promoter deletion series. The constructs in these series contain promoter fragments with the same 3' end, but 5' ends with increasing truncations. Studies with the psbB(II) (Lang and Haselkorn, 1989), atp1 and atp2(II) (Martin et al., 1994b) promoters of Anabaena sp. strain PCC 7120 showed that maximal and faithful transcription is maintained with deletions that leave 70 to 100 bp upstream from the transcription initiation site. Severe promoter truncations have identified the likely boundaries of the basal pro-

Chapter 20

Transcription in Cyanobacteria

moter. A construct including the region –50 to +3 of the rbcL promoter of Anabaena sp. strain PCC 7120, gave a slightly higher level of reporter gene expression than constructs with longer 5' and 3' extensions, but transcription initiated at the appropriate site (Martin et al., 1994a). Similarly, deletion of the Synechococcus sp. strain PCC 7942 psbDII promoter to –42 had no effect on the level of transcription or the transcription initiation site. However, deletion of the promoter to – 25 abolished expression of the lacZ reporter gene (Bustos and Golden, 1991). All of the promoter deletion studies indicate that, as in E. coli, the basal promoter elements of cyanobacterial genes are within 50 bp of the transcription initiation site. The experiments with the psbDII promoter suggest that within the basal promoter, important element(s) are positioned in the region between –42 and –25.

b. Mutations in Basal Elements: The rbcL Promoter Assays of promoter mutations have been instrumental in establishing the importance of conserved sequence elements in E. coli promoters (reviewed in Gralla, 1990). Experiments with the rbcL gene of Anabaena sp. strain PCC 7120 have addressed the role of individual base pairs within one basal promoter element. The rbcL transcript for this gene initiates ~500 bp upstream from the coding region (Nierzwicki-Bauer et al., 1984). The ability of a range of rbcL promoter fragments to drive expression of the cat reporter gene was measured by in vivo assays (Martin et al, 1994a). It was established that transcription initiated accurately in a construct bearing the rbcL promoter fragment from –100 to +3, and that this construct gave the same level of cat expression as longer constructs that included the extended region from –1000 to +400. The rbcL promoter fragment from –100 to +3 was used as the basis for mutational studies. A single base substitution was introduced at each of the three most conserved sites within the promoter –10 element (TATAAT). These mutations had dramatic effects, with each abolishing expression of the cat reporter gene (Martin et al, 1994a). These most conserved bases of the –10 promoter element were presumed to be important for transcription by analogy with E. coli promoters, and because they are highly conserved among cyanobacterial promoter sequences. The rbcL mutational studies are the first direct evidence that

629

the –10 element plays an important role in transcription initiation in cyanobacteria.

c. Dissection of Regulatory Elements: The psbDII Promoter In vivo transcription assays have been used to identify cis elements involved in the regulation of basal transcription activity. In these studies, in vivo assays of promoter deletion series have been used to detect changes in reporter gene expression which are then correlated with the absence or presence of specific DNA segments within the region 5' to the gene. Studies with the psbA and psbD genes of Synechococcus sp. strain PCC 7942 suggest that cis elements within the untranslated leader region are involved in light regulation. The psbA and psbD genes encode the core components D1 and D2, respectively, of the Photosystem II complex. Synechococcus sp. strain PCC 7942 has two copies of the psbD gene (I-II) which encode identical products (Golden et al., 1988), and three copies of psbA (I-III) which encode two distinct products (Golden et al., 1986). The levels of psbAII, psbAIII and psbDII expression are low when cells are grown in low light, but increase at high light intensities; light induction of the protein products is controlled at the level of transcription (Bustos et al., 1992; Bustos et al., 1990; Kulkarni et al., 1992). DNA mobility-shift and footprinting assays with the psbDII promoter and soluble proteins from cells grown under high light intensity identified three protein binding sites between +11 and +84 ofthe untranslated leader region ofthe psbDII gene (Bustos and Golden, 1991). Gel mobility-shift assays with a DNA fragment upstream of the psbAII gene, and competition experiments with the psbAII and psbDII promoter fragments, suggest that the two genes bind the same or similar proteins. In vivo assays of psbDII promoter-lacZ fusions verified that the regions identified by mobility-shift and footprinting assays are involved in the expression of the psbDII gene. A deletion within the most 5' protected region of the footprint decreased the reporter gene activity to a small percentage of that of the undeleted control, but gene expression remained responsive to light. Deletion of the three protected regions completely abolished both gene expression and light induction (Bustos and Golden, 1991). These results suggest that the psbDII gene contains elements within the untranslated leader region that are required for light induction and

630 efficient expression, and that the psbAII gene may be similarly regulated. A more complete discussion of these results may be found in Chapter 23.

C. Regulation of Transcription Initiation By analogy with the eubacterial systems, the regulation of cyanobacterial genes in particular cell types or under particular growth conditions may be mediated by mechanisms of activation and repression by accessory factors. Alternatively, gene expression may be regulated through the use of RNA polymerases with different promoter specificities.

1. Accessory Factors As discussed in the previous section, DNA binding proteins have been identified that are correlated with the light regulated expression of the Synechococcus sp. strain PCC 7942 psbA and psbD genes. Factors that may play a role in light regulated gene expression during chromatic adaptation have also been characterized. Chromatic adaptation is a process by which certain cyanobacteria are able to adjust the composition of their light harvesting polypeptides, the phycobiliproteins, in response to changes in light quality (see Chapter 22). In this process, the synthesis of some of the phycobiliproteins varies according to the wavelength of incident light. In complementary chromatic adaptation, the phycocyanin (PC) and phycoerythrin (PE) phycobiliproteins are oppositely

Stephanie E. Curtis and James A. Martin regulated. In Calothrix sp. strain PCC 7601 (also called Fremyella diplosiphon), PC accumulates to high levels in red light, while the levels of PE are low. Conversely, in green light, the levels of PC are low and those of PE are high. These differences in expression are achieved primarily through transcriptional control of the operons which encode PE and its associated linker polypeptides, cpeBA and cpeCD, and the operon which encodes inducible phycocyanin 2, cpc2 (reviewed in Chapter 22 and Grossman et al., 1993). Factors which may participate in the green-light regulated expression of the cpeBA operon recently have been identified. Two proteins designated RcaA and RcaB, which are only detected in extracts from cells grown in green light, have been shown to interact with the cpeBA promoter in mobility-shift and footprinting assays (Sobczyk et al., 1993). Binding of RcaA is abolished when the protein is treated with alkaline phosphatase, suggesting that it is modulated by phosphorylation (Sobczyk et al., 1993). The RcaA protein footprints a region of the cpeBA promoter from –66 to –45, which contains a direct repeat separated by 4 bp, TTGTTA (Fig. 7). The binding of RcaA was abolished when mobility-shift experiments were performed with a DNA fragment that lacks the upstream repeat. The RcaB protein binding site is less clear, but it appears to map to a region between the RcaA binding site and the region bound by the RNA polymerase (Sobczyk et al., 1993).

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Binding of a protein (designated PepB) to the same region of the Calothrix sp. strain PCC 7601 cpeBA promoter as that bound by RcaA has been shown independently by another group using both in vivo and in vitro footprinting assays (Schmidt-Goff et al., 1993). In contrast to the RcaA protein however, the PepB protein was detected in extracts from cells grown in both red and green light. RcaA and PepB appear to be the same protein, but the issue of whether the protein is present in both red and green light regimes remains to be resolved. A possible explanation for these apparently contradictory results would be that the phosphorylated from of RcaA/ PepB binds DNA differently as observed for (NtrC) of E. coli (Weiss et al., 1992). The cpeBA promoter ofPseudanabaena sp. strain PCC 7409 was also shown to bind components of extracts from green-light grown Calothrix sp. strain PCC 7601 (Dubbs and Bryant, 1991). Although the specific binding sites were not characterized, it is likely that the RcaA and/or RcaB proteins are responsible for the binding activity. Consistent with this idea, comparison of the cpeBA promoters from several species of cyanobacteria that modulate PE synthesis in response to changes in light wavelength, reveals that the motif bound by RcaA is highly conserved (Fig. 7). This region is not conserved in the cpeBA promoter from Synechococcus DC2 (Sobczyk et al., 1993), a cyanobacterium which expresses PE constitutively. These data suggest that cyanobacteria that modulate PE synthesis in response to changes in light wavelength regulate cpeBA transcription through a common mechanism. The previously described DNA binding proteins are all implicated in aspects of light-regulated gene expression in cyanobacteria. Studies on the xisA gene of Anabaena sp. strain PCC 7120 identified other proteins that may play a role in cell-specific gene regulation. The xisA gene encodes a recombinase that mediates one rearrangement of nif genes in heterocysts (Lammers et al., 1986). Fusion of the xisA coding region to a heterologous promoter resulted in expression of xisA in vivo and ectopic DNA rearrangements in vegetative cells, but only if a region 5' to the xisA gene was deleted (Brusca et al., 1990). These data suggested that the deleted region is involved in repression of xisA in vegetative cells. Chastain et al. (1989) identified a factor (VF1) in vegetative cells of Anabaena so. strain PCC 7120 that bound in vitro to this xisA region. VF1 was later shown to bind the promoter region of nifH, another

631 heterocyst-specific gene (Ramasubramanian, Wei and Golden, personal communication). Although these data implicated VF 1 (now called bifA) as a repressor of gene expression in vegetative cells, the protein also was shown to bind the promoter regions of the rbcL and glnA genes, both of which are expressed in vegetative cells. A second DNA-binding protein designated BifB also was identified that binds the rbcL and xisA promoters (Ramasubramanian, Wei and Golden, personal communication). Thus, both BifA and BifB bind the promoters of genes expressed in vegetative and heterocyst cells. The roles of BifA and BifB in the regulation of Anabaena sp. strain PCC 7120 genes is as yet unknown, but it is possible that they have different roles in the regulation of different genes. Precedence for this is seen in E. coli with proteins that can act as both repressors and activators (Collado-Vides et al., 1991). The bifA gene has recently been cloned (Wei et al., 1993) and encodes a protein with strong similarity to NtcA, a global regulator involved in nitrogen control in Synechococcus sp. strain PCC 7942 that belongs to the Crp family ofbacterial regulators (Vega-Palas et al., 1992).

2. Promoter Switching? In addition to the regulatory mechanisms of repression and activation, gene expression can be controlled by the availability of holoenzymes bearing different sigma factors. By analogy with sporulation in B. subtilis (Losick and Stragier, 1992), alternative sigma factors might be expected to play a role in temporal and cell-specific regulation ofgene expression during heterocyst development. In the simplest model of this type of regulation, genes expressed at a particular time, or in a particular cell during development would be transcribed from promoters that are recognized by holoenzymes bearing cell-specific or development-specific sigma factors. Genes that are expressed throughout development would have multiple promoters, and exhibit ‘promoter switching’ during development ensuring continuous gene expression despite a changing RNA polymerase population. Such changes in promoter utilization have precedence in B. subtilis, where many operons have been shown to have multiple promoters with different holoenzyme specificities (Doi and Wang, 1986). The different promoters are often in close proximity and sometimes overlap (Doi and Wang, 1986).

632 Circumstantial evidence for promoter switching during heterocyst development comes from studies of the glnA and gnd genes encoding glutamine synthetase (GS) and 6-phosphogluconate dehydrogenase (6PGD), respectively, in Anabaena sp. strain PCC 7120. GS and 6PGD have roles in nitrogen assimilation and carbon metabolism in vegetative cells, but also perform essential functions in heterocysts. The ammonia produced by nitrogen fixation is converted to glutamine by GS in the heterocyst before transport to neighboring cells, while the oxidative pentose phosphate pathway (in which 6PGD participates) is thought to provide much of the reductant required for nitrogen fixation (reviewed in Haselkorn et al., 1992; see chapters 16 and 27). The activity of GS increases after nitrogen starvation (Orr and Haselkorn, 1982), and the activity of 6PGD is increased in heterocysts relative to vegetative cells (Winkenbach and Wolk, 1973). The glnA and gnd genes are each represented by vegetative-cell transcripts with multiple 5' termini: Four have been mapped for glnA (Tumer et al., 1983) and two for gnd (Curtis et al., 1994). Upon nitrogen starvation and the induction of heterocyst differentiation, a transcript with a unique 5' terminus is detected for each gene [glnA(I), Tumer et al., 1983 and gnd(II), Curtis et al., 1994]. The region that would serve as a promoter for the gnd II transcript overlaps the region that would serve as a promoter for one of the vegetative cell transcripts. Consistent with the idea that these new transcripts may be transcribed by a new form of the polymerase induced by nitrogen starvation, expression from the glnA(I) promoter was not detected with in vitro transcription assays using the RNA polymerase from vegetative cells grown on nitrogen-replete medium (Schneider et al., 1991). The DNA sequences upstream from the glnA(I) and gnd(II) transcription initiation sites are not strikingly similar, although both contain an E. coli consensus sequence at the –10 position. One possibility is that the transcripts induced by nitrogen starvation are expressed using sigma factors such as SigB or SigC whose expression is induced by nitrogen starvation (Brahamsha and Haselkorn, 1992). As these factors are similar in structure to the principal sigma factor, they are expected to recognize promoters with features such as the –10 element. While the existing data on the expression of the gnd and glnA genes support the idea that some genes may be regulated by promoter switching, direct evidence for such regulation remains to be

Stephanie E. Curtis and James A. Martin demonstrated. One of the central and yet unresolved questions in this regard is whether the nitrogen starvation-induced transcripts for the glnA and gnd genes are heterocyst-specific.

D. Conclusions and Future Directions The RNA polymerases of cyanobacteria are unique among those of the eubacteria in having two subunits and that correspond to the subunit of the E. coli enzyme. As the and subunits of cyanobacteria likely perform the same functions as the single subunit of E. coli, and there is correspondence between the remaining subunits of the enzyme, the RNA polymerases of cyanobacteria and E. coli are quite similar, despite a difference in subunit organization. Given this overall structural similarity, the general properties of transcription in cyanobacteria are expected to mirror those of E. coli. Transcription initiation in E. coli is controlled through interactions between the RNA polymerase and the promoter. The recognition and binding of the RNA polymerase to promoter sequences is mediated by the sigma subunit of the holoenzyme. Cyanobacterial sigma factors thus far characterized display the conserved features of proteins of the family, with the greatest conservation within regions of the factors that interact with promoter elements. This similarity in sigma structure leads to the prediction that the promoters of cyanobacteria and E. coli also are similar. Several independent lines of evidence support this idea: 1) Footprinting studies with the RNA polymerase of Calothrix sp. strain 7601 showed that the enzyme interacts with the cpeBA promoter from –45 to +15 (Sobczyk et al., 1993), similar to the region of promoters protected by the E. coli RNA polymerase. 2) Assays of promoter deletions in vivo suggest that, as in E. coli, basal promoter elements lie within 50 bp of the transcription initiation site. 3) Most of the cyanobacterial promoters examined display some features of the E. coli promoter consensus within the basal promoter region. In particular, a sequence is highly conserved at–10 that is very similar to the E. coli –10 promoter element. 4) The most conserved positions of the –10 sequence have been shown by mutageneis experiments with one gene to be essential for transcription of that gene. 5) E. coli genes clearly can be expresse in cyanobacteria as many E. coli genes serve as selectable markers in cyanobacteria. However, it has not been clear in these cases whether the E. coli

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promoter is recognized in cyanobacteria, or whether a fortuitously placed cyanobacterial promoter substitutes. 6) The converse is also true: There are many examples of cyanobactrial genes that are expressed from natve sequences in E. coli. Among these, only a few have been studied with regard to the specific sequences used to promote transcription in E. coli and cyanobacteria. Several genes have been shown to have identical transcription initiation sites in E. coli and cyanobacteria (Turner et al., 1985; Gasparich, 1989), although the apcF gene (Warner et al., 1994) is an exception. 7) Lastly, as judged by in vitro assays of transcriptional activity, the RNA polymerase of Anabaena sp. strain PCC 7120 prefers promoter sequenes that approximate the E. coli promoter consensus at positions –10 and –35 from the transcription initiation site. These data all indicate that the promoters of cyanobacteria and E. coli are similar in sequence and in their position relative to the transcription initiation site. Given this backgrond, it is somewhat paradoxical that promoters for the majority of characterized cyanbacterial genes expressed during vegetative growth lack good E. coli promoter sequences at both –10 and –35, including the promoters of many essential and highly expresse genes. This is reflected in transcription assays performed in vitro, where promoters from genes such as rbcL and psbAI give relatively weak activity comared to those that approximate the E. coli consensus such as the promoters of cyanophage N-l. It is noteworthy that the conditions defining the interaction of the polymerase with promoters may be qite different in vitro from those encountered in vivo. For exampe, the relatively high polymerase to template ratio of in vitro transcription systems makes open complex formation or prmoter clearance the rate limiting step for transcription initiation, rather than the formation of closed complexes. The same promoter can behave quite differently in vivo, where the polymerase concentration is typically much lower (Brunner et a., 1987). Even with this caveat in mind, it seems unlikely that the differences in gene expression observed in vitro ca be solely accounted for by conditions of the in vitro assays. The fact remains that the major RNA polymerase of Anabaena sp. strain PCC 7120 efficiently transcribes E. coli consensus promoter sequences, and that many genes lack such promoters. It is possible that promoters that conform weakly to the E. coli consensus simply are used less efficiently than those with consensus promoters. However, this

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seems ulikely given the high levels of expression of these genes and their central roles in essential cellular functions. The in vitro transcription assays performed to dte have used the major RNA polymerase from vegetative cells bearing the principal sigma factor. One possible explanation for the results from in vitro transcription assays is that at least some of the genes that lack E. coli promoter consensus sequences at – 35 can e inefficiently transcribed in vitro by the major RNA polymerase, but are expressed in vivo by holoenzymes with sigma factors other than the principal sigma factor. Thus, a number of different sigma factors might participate in gene expression during vegetative growth, with each recognizing different promoters. Given that most promoters characterized have the same type of conserved element at the –10 position, each ofthe sigma factors would be expected to recognize the –10 element and then a unique sequence at –35. The sequences at –35 in the cyanobacterial promoter database are quite diverse, and thus, such a scenario would require a large number of sigma factors. In support of this idea, multiple sigma factors have been identified in several cyanobacteria. All have properties of principal sigma factors, and thus would be expected to recognize promoters similar to the E. coli consensus. The nonprincipal sigma factors (SigB and SigC) thus characterized from Anabaena sp. strain PCC 7120 are not expressed during vegetative growth on nitrogen-replete medium, and inactivation of these sigma factors did not detectably affect vegetative growth. Thus, these factos cannot control the expression of essential genes in vegetative cells. Similar results have been obtained for Synechococcus sp. train PCC 7002, although in this case transcripts of the nonprincipal sigma genes are found in cells growing exponentially in replete medium. While there is circumstantial evidence that other yet uncharacterized sigma factors exist in cyanobacteria, there is no strong evidence that a large set of sigma factors are required for vegetative growth. A more likely explanation for the promoter paradox is that the majority of cyanobacterial genes require activation by accessory factorsfor maximal expression. Most of the E. coli genes that require activation display promoters with weak –35 elements for which activator proteins are thought to substitute. One of the striking features of cyanobacterial promoters thus characterized is, that although many display highly conserved –10 sequences, very few also have a well conserved –35 element. This feature

634 is consistent with the idea that such promoters may require accessory factors for transcription initiation. These factors would likely to be absent in assays with purified RNA polymerase, and thus genes that require them for maximal expression would give a relatively low level of activity in the transcription assays in vitro. In support of this idea, the activities of the psbA and rbcL promoters were higher in transcription assays performed in vitro with crude extracts than with purified polymerase (Schneider et al., 1991). Is there evidence for the activation of cyanobacterial genes? Analyses of E. coli promoters showed that the location of binding sites for accessory transcription factors is constrained by the requirement for direct interaction of these factors with RNA polymerase bound at the basal promoter. Activator binding sites are most often found in close proximity to the –35 region, allowing direct contact between the activator and polymerase bound at the basal promoter. The RcaA protein identified in Calothrix sp. strain PCC 7601 is a very good candidate for an activator of the classic type. This protein binds a region of the cpeBA promoter from –66 to –45 that is highly conserved among the promoter sequences of similarly regulated cpeBA genes from different cyanobacteria. The light regulated expression of the psbDII gene of Synechococcus sp. strain PCC 7942 has been shown to require cis elements in the untranslated leader region of the gene. Proteins identified that bind to ths region may serve as activators although the binding sites are an unusual location for activator binding sites in E. coli. Studies on the BifA and BifB proteins of Anabaena sp. strain PCC 7120 suggest that the regulation of cyanobacterial genes may be complex. These factors bind the promoters of both heterocyst cell-specific and vegetative cell-specific genes. The relative positions of binding sites vary among the genes, with locations characteristic of both repressor and activator binding sites in E. coli. BifA binds the rbcL promoter in two regions, one of which overlaps with a BifB binding region. These studies hint at the existence of cyanobacterial DNA-binding proteins with multiple roles in gene regulation. A possibility is that such binding proteins interact with other factors to regulate gene expression in different ways. This type of regulation has precedence in eucaryotic transcription, where gene-specific combinations of multple trans-acting factors interact to activate gene expression (Lewin, 1990). It should be clear from the preceding discussions that a great deal of information on transcription in

Stephanie E. Curtis and James A. Martin cyanobacteria has accumulated, but a number of questions remain unanswered. For example, what are the roles of the multiple sigma factors identified in cyanobacteria? This question can be answered in part by the analysis of isolated sigma factor genes and their products. It also appears likely that other sigma factor genes can be isolated by virtue of their homology to those in hand, although this strategy may not be successful with all genes. Do sigma factors of the alternative type, such as those that participate in adaptation to stress and developmental programs in other bacteria, exist in the cyanobacteria? A combination of biochemical and genetic approaches was used to identify alternative sigma factors in B. subtilis (Doi and Wang, 1986). If such factors exist in cyanobacteria, most likely it also will be necessary to eploy multiple approaches to their identification. A number of laboratories are actively using mutagenesis strategies to identify genes that regulate pathways such as chromatic adaptation and heterocyst and hormogonium development, for which alternative sigma factors might be expected to participate. These strategies should allow the identification of any sigma factor genes essential o these pathways. A complementary biochemical approach that has not been yet employed in the cyanobacteria, is the use of DNA templates with promoters not recognized by the major RNA polymerase (e.g., heat shock promoters) to probe for new RNA polymerase activities in vitro. What are the important elements of cyanobacterial basal promoters? The conserved features of E. coli basal promoters apparent from sequence comparisons were shown to be important by the analysis of promoters of varied sequence and the analysis of mutant promoters. Although some features of cyanobacterial promoters are emerging from sequence comparisons, the promoter database is still small and a much larger sample size is necessary to provide statistically significant information. Only small number of genes have been analyzed with regard to transcriptional activity by assays in vitro or in vivo. A large number of systematic studies on the relative activities of individual promoters carrying specific mutations, as well as studies of the relative activities of different promoters from the same organism need to be performed. Such studies should allow correlations to be made between variations in promoter sequence and transcriptional activity. Another strategy that should prove useful in the analysis of promoter sequences, is the comparison of

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sets of promoters that are coordinately regulated. Random insertions of reporter genes into the chromosome have been used to identify genes induced at the transcriptional level in response to certain environmental conditions (Wolk et al., 1991). Analysis of such sets of genes should allow the identification of common promoter elements involved in the coordinate control of gene expression. The mechanisms by which transcription initiation is regulated in cyanobacteria are largely unknown. Two of the obvious questions that derive from cyanobacterial promoter studies are whether some cyanobacterial genes require activation for maximal expression, and what roles the DNA binding proteins thus identified play in gene regulation. Although a few studies have been conducted that address these issues, it is clear that a much larger number of genes need to be analyzed in detail. The regulatory mechanisms controlling transcription initiation in cyanobacteria are likely to be diverse and complex, and the scope of mechanisms employed can only be discovered with a large database. Again, a combination of biochemical and genetic approaches likely will be necessary to address regulatory mechanisms. Promoter deletion studies and mobility-shift assays have been successfully used to identify cis elements important in regulation, and to identify proteins that bind these cis elements. More of these studies on a wider range of genes in different cyanobacteria need to be performed to develop an accurate picture of the types of regulation employed in cyanobacteria. Random mutagenesis approaches have identified regulatory genes involved in the chromatic adaptation and heterocyst development pathways. Some of these genes, such as rcaC of Calothrix sp. strain PCC 7601 (Chiang et al., 1992), hetR (Buikema and Haselkorn, 1991) and patA (Liang et al., 1992) of Anabaena sp. strain 7120 appear to act relatively early in the signal transduction pathways. However, the patB (Liang et al, 1993) encodes a product that may act as a transcriptional activator. Thus, such mutagenesis approaches will likely be very useful in the identification of factors that interact directly with promoter regions. In a similar approach, carefully designed strategies with specific promoter-reporter gene fusions in vivo could be used to identify mutations that abolish normal regulation of the promoter. The results from the characterization of the BifA and BifB DNA binding proteins of Anabaena sp. strain PCC 7120 are intriguing, and elucidation of their function should provide insight into the

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complexities of gene regulation in cyanobacteria. As noted by Gralla (1991), the distinctions between the regulation of procaryotic and eucaryotic transcription have begun to blur in recent years. Thus, it would behoove us to keep in mind mechanisms employed by both eucaryotes and procaryotes in evaluating cyanobacterial gene regulation. In conclusion, the cyanobacteria are an ancient group of bacteria that have adapted to a wide variety of ecological niches. Control at the level of gene transcription is likely to play a major role in both short and long term adaptations to environmental conditions. While many questions remain about transcription in cyanobacteria, the tools to address the role of transcription regulation in many interesting cyanobacterial systems are now available. Surely a review written in ten years time will provide answers to many of these questions and pose even more for us to resolve. Acknowledgments The authors thank the many colleagues who shared unpublished information. Special thanks to Don Bryant, Bob Haselkorn, and Martin Mulligan for informative discussions which helped solidify some of the ideas presented in this article. Work on transcription in the S.E.C. laboratory is supported by the National Science Foundation. References Alam J, Whitaker RA, Krogmann DW and Curtis SE (1986) Isolation and sequence of the gene for ferredoxin I from the cyanobacterium Anabaena sp. strain PCC 7120. J Bacteriol 168: 1265–1271 Anderson LK and Grossman AR (1990) Structure and lightregulated expression of phycoerythrin genes in wild-type and phycobilisome assembly mutants of Synechocystis sp. strain PCC 6701. J Bacteriol 172: 1297–1305 Bedwell D, Davis G, Gosink M, Post L, Nomura M, Kestler H, Zengel J and Lindahl L (1985) Nucteotide sequence of the alpha ribosomal protein operon of Escherichia coli. Nucl Acids Res 13: 3891–3903 Belknap WR and Haselkorn R (1987) Cloning and light regulation of expression of the phycocyanin operon of the cyanobacterium Anabaena. EMBO J 6: 871–884 Bergsland KJ and Haselkorn R (1991) Evolutionary relationships among eubacteria, cyanobacteria, and chloroplasts: Evidence from the rpoC1 gene of Anabaena sp. strain PCC 7120. J Bacteriol 173: 3446–3455 Borthakur D and Haselkorn R (1989) Nucleotide sequence of the

636 gene encoding the 33 kDa water oxidizing polypeptide in Anabaena sp. strain PCC 7120 and its expression in Escherichia coli. Plant Mol Biol 13: 427–439 Borthakur D, Basche M, Buikema WJ, Borthakur PB and Haselkorn R (1990) Expression, nucleotide sequence and mutational analysis of two open reading frames in the nif gene region of Anabaena sp. strain PCC 7120. Mol Gen Genet 221: 227–234 Boylan SA, Suh J-W, Thomas SM and Price CW (1989) Gene encoding the alpha core subunit of Bacillus subtilis RNA polymerase is cotranscribed with the genes for initiation factor 1 and ribosomal proteins B, S13, S11, and L17. J Bacteriol 171: 2553–2562 Brahamsha B and Haselkorn R (1991) Isolation and characterization of the gene encoding the principal sigma factor of the vegetativecell RNA polymerase fromthe cyanobacterium Anabaena sp. strain PCC 7120. J Bacteriol 173: 2442–2450 Brahamsha B and Haselkorn R (1992) Identification of multiple RNA polymerase sigma factor homologs in the cyanobacterium Anabaena sp. strain PCC 7120: Cloning, expression, and inactivation the sigB and sigC genes. J Bacteriol 174: 7273–7282 Brunner M and Bujard H (1987) Promoter recognition and promoter strength in the Escherichia coli system. EM BO J 6: 3139–3144 Brusca JS, Chastain CJ and Golden JW (1990) Expression ofthe Anabaena sp. strain PCC 7120 xisA gene from a heterologous promoter results in excision of the nifD element. J Bacteriol 172: 3925–3931 Buikema WJ and Haselkorn R (1991) Characterization of a gene controlling heterocyst development in the cyanobacterium Anabaena 7120. Genes Dev 5: 321–330 Burgess R (1971) RNA polymerase. Ann Rev Biochem 40: 771–735 Burgess R, Travers A, Dunn J and Bautz E (1969) Factor stimulating transcription by RNA polymerase. Nature 221: 43–44 Burton Z, Gross C, Watanabe K and Burgess R (1983) The operon that encodes the sigma subunit of RNA polymerase also encodes ribosomal protein S21 and DNA primase in E. coli K12. Cell 32: 335–349 Bustos SA and Golden SS (1991) Expression of the psbDII gene in Synechococcus sp. strain PCC 7942 requires sequences downstream of the transcription start site. J Bacteriol 173: 7525–7533 Bustos SA and Golden SS (1992) Light-regulated expression of the psbD gene family in Synechococcus sp. strain PCC 7942: Evidence for the role of duplicated psbD genes in cyanobacteria. Mol Gen Genet 232: 221–230 Bustos SA, Schaefer MR and Golden SS (1990) Different and rapid responses of four cyanobacterial psbA transcripts to changes in light intensity. J Bacteriol 172: 1998–2004 Buttner C, Chater K and Bibb M (1990) Cloning, disruption, and transcriptional analysis of three RNA polymerase sigma factor genes of Streptomyces coelicolor A3(2). J Bacteriol 172: 3367–3378 Chamberlain M (1974) The selectivity of transcription. Ann Rev Biochem 43: 721–767 Chamberlain M (1976) RNA polymerase – an overview. In: Losick R and Chamberlain M (eds) RNA Polymerase, pp 17– 68, Cold Spring Harbor Laboratory, Cold Spring Harbor, New

Stephanie E. Curtis and James A. Martin York Chiang GG, Schaefer, MR and Grossman AR (1992) Complementation of a red-light-indifferent cyanobacterial mutant. Proc Natl Acad Sci USA 89: 9415–9419. Chenchick A, Beabealashvilli R and Mirzabekov A (1981) Topography of interaction of Escherichia coli RNA polymerase subunits with the lacUV5 promoter. FEBS Lett 128: 46–50 Chenchick A, Beabealashvilli R, Mirzabekov A and Shik V (1982) Contact between subunits of Escherichia coli RNA polymerase and the nucleotides of the lacUV5 promoter. Molekuyarnaya Biologiya 16: 34–46 Collado-Vides J, Magasanik B and Gralla J (1991) Control site location and transcriptional regulation in Escherichia coli, Microbiol Rev 55: 371–394 Conley P, Lemaux P and Grossman A (1988) Molecular characterization and evolution of sequences encoding lightharvesting components in the chromatically adapting cyanobacterium Fremyella diplosiphon. J Mol Biol 199: 447–465 Cowing D, Bardwekk J, Craig E, Woolford C, Hendrix R and Gross C (1985) Consensus sequence for Escherichia coli heat shock gene promoters. Proc Natl Acad Sci USA 82: 2679–2683 Csiszar K, Houmard J, Damerval T and Tandeau de Marsac N (1987) Transcriptional analysis of the cyanobacterial gvpABC operon in differentiated cells: Occurrence of an antisense RNA complementary to three overlapping transcripts. Gene 60: 29–37 Curtis SE (1987) Genes encoding the beta and epsilon subunits of the proton-translocating ATPase from Anabaena sp. strain PCC 7120. J Bacteriol 169:80–86 Curtis SE and Haselkorn R (1984) Isolation, sequence and expression of two members of the 32 kd thylakoid membrane protein gene family from the cyanobacterium Anabaena 7120. Plant Mol Biol 3: 249–258 Curtis SE, Ligon PJB, Kim Y-H and Martin JA (1994) Expression of the gnd gene of Anabaena sp. strain PCC 7120 is upregulated during heterocyst development. Mol Microbiol, submitted Doi R and Wang L (1986) Multiple procaryotic ribonucleic acid polymerase sigma factors. Microbiol Rev 50: 227–243 Dombroski A, Walter W, Record J MT, Siegele D and Gross C (1992) Polypeptides containing highly conserved regions of transcription initiation factor exhibit specificity of binding to promoter DNA. Cell 70: 501–512 Downing W and Dennis P (1987) Transcription products from the rplKAJL-rpoBC gene cluster. J Mol Biol 194: 609–620 Dubbs J and Bryant D (1991) Molecular cloning and transcriptional analysis of the cpeBA operon of the cyanobacterium Pseudanabaena species PCC 7409. Mol Microbiol 5: 3073–3085 Federspiel NA and Grossman AR (1990) Characterization of the light-regulated operon encoding the phycoerythrin-associated linker proteins from the cyanobacterium Fremyella diplosiphon. J Bacteriol 172: 4072–4081 Ferino F and Chauvat F (1989) A promoter-probe vector-host system for the cyanobacterium, Synechocystis PCC 6803. Gene 84: 257–266 Fong S and Surzycki S (1992) Chloroplast RNA polymerase genes of Chlamydomonas reinhardtii exhibit an unusual structure and arrangement. Curr Genet 21: 485–497

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Gardella T, Moyle H and Susskind M (1989) A mutant Escherichia coli subunit of RNA polymerase with altered promoter specificity. J Mol Biol 206: 579–590 Gasparich G (1989) The effects ofvarious environmental stress conditions on gene expression in the cyanobacterium Synechococcus sp. PCC 7002. Ph.D. Thesis, The Pennsylvania State University Gasparich GE, Buzby JS, Bryant DA, Porter RD and Stevens, SE Jr (1987) The effects of light intensity and nitrogen starvation on the phycocyanin promoter in the cyanobacterium Synechococcus sp. PCC 7002. In: Biggins J, (ed) Progress in Photosynthesis Research, Vol. IV, pp 761–764, MartinusNijhoff, Dordrecht Golden SS and Stearns GW (1988) Nucleotide sequence and transcript analysis of three Photosystem II genes from the cyanobacterium Synechococcus sp. PCC 7942. Gene 67: 85– 96 Golden SS, Brusslan J and Haselkorn R (1986) Expression of a family of psbA genes encoding a Photosystem II polypeptide in the cyanobacterium Anacystis nidulans R2. EMBO J 5: 2789– 2798 Gralla J (1990) Promoter recognition and mRNA initiation by Escherichia coli Methods Enz 185: 37–54 Gralla J (1991)Transcriptional control – lessons from an E. coli promoter data base. Cell 66: 415–418 Grossman AD, Erickson J and Gross C (1984) The htpR gene product of E. coli is a sigma factor for heat-shock promoters. Cell 38:383–390 Grossman A, Lemaux P, Conley P, Bruns B and Anderson L (1988) Characterization of phycobiliprotein and linker polypeptide genes in Fremyella diplosiphon and their regulated expression during complementary adaptation. Photosynth Res 17: 23–56 Grossman A, Schaefer M, Chiang G and Collier J (1993) Environmental effects on the light-harvesting complex of cyanobacteria. J Bacteriol 175: 575–582 Harley C and Reynolds R (1987) Analysis of E. coli promoter sequences. Nucl Acids Res 15: 2343–2361 Haselkorn R (1991) Genetic systems in cyanobacteria. Methods Enz 204: 418–430. Haselkorn R and Buikema W (1992). Nitrogen fixation in cyanobacteria. In: Stacey G, Burris R, and Evans H (eds), Biological Nitrogen Fixation, pp 166–190. Chapman and Hall, New York Hawley D and McClure W (1983) Compilation and analysis of Escherichia coli promoter sequences. Nucleic Acids Res 1 1 : 2237–2255 Helmann JD and Chamberlain MJ (1988) Structure and function of bacterial sigma factors. Ann Rev Biochem 57: 839–872 Herzfeld F and Zillig W (1971) Subunit composition of DNAdependent RNA polymerase of Anacystis nidulans. Eur J Biochem 24: 242–248 Houmard J, Capuano V, Coursin T and Tandeau de Marsac N (1988) Genes encoding core components of the phycobilisome in the cyanobacterium Calothrix sp. strain PCC 7601: Occurrence of a multigene family. J Bacteriol 170: 5512–5521 Hu J, Troxler F and Bogorad L (1991) Maize chloroplast RNA polymerase: The 78-kilodalton polypeptide is encoded by the plastid rpoC1 gene. Nucl Acids Res 19: 3431–3434 Hudson G, Holton T, Whitfeld P and Bottomley W (1988) Spinach chloroplast rpoBC genes encode three subunits of the

637 chloroplast RNA polymerase. J Mol Biol 200: 639–654 Kulkarni RD, Schaefer MR and Golden SS (1992)Transcriptional and posttranscriptiona components of psbA response to high light intensity in Synechococcus sp. strain PCC 7942. J Bacteriol 174: 3775–81 Kustu S, Santero E, Keener J, Popham D and Weiss D (1989) Expression of (ntrA)-dependent genes is probably united by a common mechanism. Microbiol Rev 53: 367–376 Lammers PJ, Golden JW and Haselkorn R (1986) Identification and sequence ofa gene required for a developmentally regulated DNA excision in Anabaena. Cell 44: 905–911 Lang JD and Haselkorn R (1989) Isolation, sequence and trnscription of the gene encoding the Photosystem II chlorophyll-binding protein, CP-47, in the cyanobacterium Anabaena 7120. Plant Mol Biol 13: 441–457 Lang JD and Haselkorn R (1991) A vector for analysis of promoters in the cyanobacterium Anabaena sp. strain PCC 7120. J Bacteriol 173: 2729–31 Laudenbach D and Straus N (1988) Characterization of a cyanobacterial iron stress-induced gene similar to psbC. J Bacteriol 170: 5018–5026 Lewin, B (1990) Commitment and activation at PolII promoters: A tail of protein-protein interactions. Cell 61: 1161–1164 Liang JL, Scappino L and Haselkorn (1992) The patA gene product, which contains a region similar to CheY of Escherichia coli, controls heterocyst pattern formation in the cyanobacterium Anabaena 7120. Proc Natl Acad Sci USA 89: 5655–5659 Liang JL, Scappino L and Haselkorn R (1993) The patB gene product, required for growth of the cyanobacterium Anabaena sp. strain PCC 7120 undernitrogen-limitingconditions, contains ferredoxin and helix-turn-helix domains. J Bacteriol 175: 1697–1704 Lonetto M, Gribskov M and Gross C (1992) The family: Sequence conservation and evolutionary relationships. J Bacteriol 174: 3843–3849 Losick R and Pero J (1981) Cascades of sigma factors. Cell 25: 582–584 Losick R and Stragier P (1992) Crisscross regulation of celltype-specific gene expression during development in B. subtilis. Nature 355: 601–604 Luinenburg I and Coleman J (1992) Identification, characterization and sequence analysis of the gene encoding phosphoenolpyruvate carboxylase in Anabaena sp. PCC 7120. J Gen Microbiol 138: 685–691 Martin JA, Ligon PJB and Curtis SE (1994a) Analysis of the rbcL promoter of Anabaena sp. strain PCC 7120. J Bacteriol, submitted Martin JA, Ligon PJB and Curtis SE (1994b) Development of a system for promoter analysis in Anabaena sp. strain PCC 7120. J Biol Chem, submitted Milkowshi C and Quinones A (1991) Cloning ofpromoter-active DNA sequences from the cyanobacterium Synechocystis sp. PCC 6803 in an Escherichia coli host. Curr Microbiol 23: 333– 336 Miller S and Bogorad L (1978) Purification and characterization of RNA polymerase from Fremyella diplosiphon. Plant Physiol 62: 995–999 Mulligan ME and Haselkorn R (1989) Nitrogen fixation (nif) genes of the cyanobacterium Anabaena species strain PCC 7120. The nifB–fdxN-nifS-nifU operon. J Biol Chem 264:

638 19200–19207 Murphy R, Gasparich G, Bryant D and Porter R (1990) Nucleotide sequence and further characterization of the Synechococcus sp. strain PCC 7002 recA gene: Complementation of a cyanobacterial recA mutation by the Escherichia coli recA gene. J Bacteriol 172: 967–976 Nagaraja R (1986) Studies on the DNA-binding protein HU from the cyanobacterium Anabaena 7120. Ph.D. Thesis, University of Chicago Nierzwicki-Bauer S, Curtis S and Haselkorn R (1984) Cotranscription of genes encoding the small and large subunits of ribulose-1,5-bisphosphate carboxylase in the cyanobacterium Anabaena 7120. Proc Natl Acad Sci USA 81: 5961–5965 Orr J and Haselkorn R (1982) Regulation of glutamine synthetase activity and synthesis in free-living and symbiotic Anabaena spp. J Bacteriol 152: 626–635 Purton S and Gray J (1988) The plastid rpoA gene encoding a protein homologous to the bacterial RNA polymerase alpha subunit is expressed in pea chloroplasts. Mol Gen Genet 217: 77–84 Reith M, Laudenbach D and Straus N (1986) Isolation and nucleotide sequence analysis of the ferredoxin I gene from the cyanobacterium Anacystis nidulans R2.J Bacteriol 168: 1319– 1324 Rhiel E, Stirewalt VL, Gasparich GE and Bryant DA (1992) The psaC genes of Synechococcus sp. PCC 7002 and Cyanophora paradoxa: Cloning and sequence analysis. Gene 112: 123–128 Sambrook J, Fritsch E and Mantiatis T(1989). Molecular cloning: A laboratory manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. Scanlan D, Bloye S, Mann N, Hodgson DC and Carr NG (1990) Construction of lacZ promoter probe vectors for use in Synechococcus: Application to the identification of regulated promoters. Gene 90: 43–49 Schaefer MR and Golden SS (1989) Differential expression of members of a cyanobacterial psbA gene family in response to light. J Bacteriol 171: 3973–81 Schluchter WM, Zhao J and Bryant DA (1993) Isolation and characterization of the ndhF gene of Synechococcus sp. PCC 7002 and initial characterization of an interposon mutant. J Bacteriol 175: 3343–3352 Schmidt-Goff C and Federspiel N (1993) In vivo and in vitro footprinting of a light-regulated promoter in the cyanobacterium Fremyella diplosiphon. J Bacteriol 175: 1806–1813 Schneider GJ and Haselkorn R (1988a) Characterization of two early promoters of cyanophage N-l. Virology 167: 150–5 Schneider GJ and Haselkorn R (1988b) RNA polymerase subunit homology among cyanobacteria, other eubacteria, and archaebacteria. J Bacteriol 170: 4136–4140 Schneider GJ, Tumer NE, Richaud C, Borbely G and Haselkorn R (1987) Purification and characterization of RNA polymerase from the cyanobacterium Anabaena 7120. J Biol Chem 262: 14633–14639 Schneider GJ, Lang JD and Haselkorn R (1991) Promoter recognition by the RNA polymerase from vegetative cells of the cyanobacterium Anabaena 7120. Gene 105: 51–60 Shimada H, Fukuta M, Ishikawa M and Sugiura M (1990) Rice chloroplast RNA polymerase genes: The absence of an intron in rpoC1 and the presence of an extra sequence in rpoC2. Mol Gen Genet 221: 395–402 Sibold C and Subramanian A (1990) Cloning and characterization

Stephanie E. Curtis and James A. Martin of the genes for ribosomal proteins L10 and L12 from Synechocystis sp. PCC 6803: Comparison of gene clustering pattern and protein sequence homology between cyanobacteria and chloroplasts. Biochim Biophys Acta 1050: 61–68 Siebenlist U and Gilbert W( 1980) Contacts between Escherichia coli RNA polymerase and an early promoter of phage T7. Proc Natl Acad Sci USA 77: 122–126 Siegele D, Hu J, Walter W and Gross C (1989) Altered promoter recognition by mutant forms of the subunit of Escherichia coli RNA polymerase. J Mol Biol 206: 591–603 Sobczyk A, Schyns G, Tandeau de Marsac N and Houmard J (1993) Transduction of the light signal during complementary chromatic adaptation in the cyanobacterium Calothrix sp. PCC 7601: DNA-binding proteins and modulation by phosphorylation. EMBO J 12: 997–1004 Tanaka K, ShiinaTandTakahashi H (1991) Nucleotide sequence of genes hrdA, hrdC and hrdD from Streptomyces coelicolor A3(2) having similarity to rpoD genes. Mol Gen Genet 229: 334–340 Tanaka K, Masuda S and Takahashi H (1992a) The complete nucleotide sequence of the gene (rpoD1) encoding the principal sigma factor of the RNA polymerase from the cyanobacterium Synechococcus sp. strain PCC 7942. Biochim Biophys Acta 1132: 94–96 Tanaka K, Masuda S and Takahashi H (1992b) Multiple rpoDrelated genes of cyanobacteria. Biosci Biotech Biochem 56: 1113–1117 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 Tumer N, Robinson S and Haselkorn R (1983) Different promoters for the Anabaena glutamine synthetase gene during growth using molecular or fixed nitrogen. Nature 306: 337–342 Vega-Palas M, 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 Vrba JM and Curtis SE (1990) Characterization of a fourmember psbA gene family from the cyanobacterium Anabaena PCC 7120. Plant Mol Biol 14: 81–92 Warner L E, Stahel AW and Curtis SE (1994) Characterization of the apcF gene of Anabaena sp. strain 7120. J Bacteriol, submitted Webb R, Reddy K and Sherman L (1990) Regulation and sequence of the Synechococcus sp. strain PCC 7942 groESL operon, encoding a cyanobacterial chaperonin. J Bacteriol 172: 5079– 5088 Wei T-F, Ramasubramanian T, PuF and Golden J(1993) Anabaena bifA gene encoding a sequence-specific DNA-binding protein cloned by in vivo transcriptional interference selection. J Bacteriol 175: 4025–4035 Weiss V, Claverie-Martin F and Magasanik B (1992) Phosphorylation of nitrogen regulator I of Escherichia coli induces strong cooperative binding to DNA essential for activation of transcription. Proc Natl Acad Sci USA 89: 5088–5092 Winkenbach F and Wolk CP (1973) Activities of enzymes of the oxidative and the reductive pentose phosphate pathways in heterocyst of a blue-green alga. Plant Physiol 52: 480–483 Wolk CP, Cai Y and Panoff J-M (1991) Use of a transposon with

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luciferase as a reporter to identify environmentally responsive genes in a cyanobacterium. Proc Natl Acad Sci USA 88: 5355– 5359 Xie W and Potts M (1990) Gene cluster rpoBC1C2 in cyanobacteria does not constitute an operon. Arch Biochem Biophys 284: 22–25 Xie W-Q, Jager K and Potts M (1989) Cyanobacterial RNA polymerase genes rpoC1 and rpoC2 correspond to rpoC of Escherichia coli. J Bacteriol 171: 1967–1973

639 Zhao J, Snyder W, Muhlenhoff U, Rhiel E, Warren PV, Golbeck JH and Bryant DA (1993) Cloning and characterization of the psaE gene of the cyanobacterium Synechococcus sp. PCC 7002: Characterization of a psaE mutant and overproduction of the protein in Escherichia coli. Mol Microbiol 9: 1283–194. Zillig W, Schnabel R, Gropp F and Reiter W (1985)The evolution of the transcription apparatus. In: Schliefer, KH and Stackebrandt E (eds) Evolution of Prokaryotes, pp 45–72. Academic Press, London

Chapter 21 The Responses of Cyanobacteria to Environmental Conditions: Light and Nutrients Arthur R. Grossman, Michael R. Schaefer, Gisela G. Chiang and Jackie L. Collier The Carnegie Institution of Washington, Department of Plant Biology, 290 Panama Street, Stanford, CA 94305 Summary I. PBS Structure II. Chromatic Adaptation A. Historical Perspective B. Gene Expression C. Photobiology D. Regulatory Mutants E. Approaches for Isolating Regulatory Elements F. Model Describing the Regulation III. The Responses of Cyanobacteria to Nutrient Deficiency A. Nutrient-Specific Responses 1. Micronutrients 2. Macronutrients B. Intracellular Inclusions and Nutrient Reserves C. Pigment Changes and Altered Photosynthetic Activity D. Phycobilisome Degradation IV. Concluding Remarks Acknowledgments References

641 641 643 643 645 648 649 650 652 654 654 654 659 663 664 666 668 668 668

Summary Cyanobacteria are found in virtually all terrestrial niches and can be found in locations which exhibit widely fluctuating chemical and physical parameters including nutrient availability, light intensity, light wavelength, temperature, and water activity. Described throughout this volume are ways in which cyanobacteria respond to changes in their environment, and examples of the insights that molecular genetic analyses have provided into acclimation processes. This chapter will discuss the modification of the cyanobacterial light-harvesting apparatus in response to light quality and nutrient availability. Recent advances in understanding the regulation of nutrient acquisition systems during nutrient-limited growth will also be summarized. I. PBS Structure While all oxygen evolving organisms contain chlorophyll a in their photosynthetic reaction centers, the pigments ofthe antennae complexes that harvest light energy may vary markedly. In the procaryotic cyanobacteria and eucaryotic red algae phycobiliproteins are the major light-harvesting polypeptides D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 641–675. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

of the cell (see Chapter 6 for a more detailed description). Phycobiliproteins are a family of brilliantly pigmented, water soluble proteins that may constitute 50% of the soluble protein of the cyanobacterial cell. The major phycobiliproteins are phycoerythrin (PE, ), phycocyanin (PC, ) and allophycocyanin (AP, ~650 nm). These proteins comprise 85% of the

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macromolecular, light-harvesting complexes called phycobilisomes (PBS; Gantt, 1981; Glazer, 1985, 1987, 1989; Zuber, 1986; Bryant, 1991; also see Chapter 6) which are peripherally associated with the cytoplasmic side of the photosynthetic (thylakoid) membranes. In red algae and cyanobacteria the PBS appear in electron micrographs as rows of closely spaced granules on the stroma-exposed surface of the thylakoid membranes. The excitation energy absorbed by PE is transferred sequentially to PC, AP and then to the chlorophyll molecules associated with the reaction centers of photosynthesis. Each of the phycobiliproteins is composed of two different subunits termed and with molecular masses of 15 to 22 kDa. The chromophores, the light-absorbing molecules that are attached to the phycobiliproteins via thioether linkages, are linear tetrapyrroles. The numbers and types of chromophores associated with a particular phycobiliprotein subunit are usually invariant (with some exceptions; see Glazer, 1987, 1989), and the absorbances of these chromophores are strongly influenced by their conformation and interactions with amino acid residues of the protein moiety of the molecule. The phycobilin chromophores are very similar to the chromophore associated with the photoreceptor phytochrome, that is crucial for the development and differentiation of higher plants. Linker polypeptides, most of which are nonpigmented, are also integral to the PBS. These polypeptides range in molecular mass from 8 kDa to 120 kDa and may serve more than one function in the PBS. They help stabilize the PBS structure, determine the positions of specific phycobiliproteins in the complex, facilitate assembly of phycobiliproteincontaining substructures, modulate the absorption characteristics of the phycobiliproteins to promote unidirectional transfer of energy within the PBS and from the PBS to the chlorophylls of the photosynthetic reaction centers, and physically link the entire complex to the photosynthetic membranes (Glazer,

1985,1987,1989; Bryant, 1991; also see Chapter 6). A synthesis of the information generated over a score ofyears from electron microscopy, biochemical and biophysical studies, X-ray crystallography and molecular analyses of genes encoding PBS polypeptides has led to the development of a detailed model ofcyanobacterial PBS structure (Bryant et al., 1979; Gantt, 1981; Glazer et al., 1983; Glazer, 1987, 1989; Bryant, 1991). The structure ofa PBS from the filamentous cyanobacterium Calothrix sp. strain PCC 7601 (very similar to Fremyella diplosiphon; the two are used synonymously in this review) and the constituents that comprise each of the two PBS domains, the rods and the core, are diagramatically presented in Fig. 1. This structure is similar in its basic composition and structure to the PBS present in most cyanobacteria, although the composition of the Calothrix sp. strain PCC 7601 PBS is different in cells grown in red light (RL) and green light (GL). This phenomenon, called chromatic adaptation, is discussed in the next section. The Calothrix sp. strain PCC 7601 PBS are fanlike in appearance and contain a tricylindrical core structure and a set of six cylindrical rods that radiate from the core (see Glauser et al., 1992). Phycobiliproteins assemble into hexamers (double discs) that are stacked upon each other to form the cylinders that comprise both the rod and core substructures. The rods contain PE and PC hexamers that associate with specific linker (L) polypeptides. The core is composed primarily of AP subunits. A 35Å channel in the center of each hexamer is believed to be the site of linker polypeptide association. The core substructure directly contacts the thylakoid membranes and mediates the transfer of excitation energy from the PBS to the photosynthetic reaction centers. Essentially all genes encoding structural components of the PBS have been isolated and characterized from many different organisms (summarized in Tandeau de Marsac et al., 1990; Bryant, 1991; Grossman et al., 1993). This work has

Abbreviations: maximum; AP–allophycocyanin; –the and subunits of all ophycocyanin; –the and subunits of phycocyanin; –the and subunits of phycoerythrin; carbon; Cab polypeptides–chlorophyll a/b-binding polypeptides; FdB – Calothrix sp. strain PCC 7601 blue mutant; FdG – Calothrix sp. strain PCC 7601 green mutant; FdR – Calothrix sp. strain PCC 7601 red mutant; FRL – far red light; GL – green light; polypeptide in the core of the phycobilisome; polypeptide which serves as an interface between the core of the phycobilisome and the thylakoid membranes; of linker polypeptides in the phycobilisome rods; polypeptide that serves as the interface between the rod and core substructures of the phycobilisome; nblA – gene encoding small polypeptide involved in phycobilisome degradation; ORF – open reading frame; PBS – phycobilisome; PC – phycocyanin; phycocyanin: phycocyanin; PE – phycoerythrin; RL – red light; RcaA and RcaB – two proteins that bind to regions upstream of the genes encoding PE subunits and may be involved in regulating expression of these genes; rcaC– gene involved in complementing the FdR mutant; Rubisco – ribulose-1,5bisphosphate carboxylase/oxygenase; txlA – gene encoding protein that is required for normal growth and photosynthesis.

Chapter 21 Cyanobacterial Acclimation Processes

added enormously to our knowledge of PBS structure, the function of the individual components that comprise the complex, our ideas concerning the evolution of genes encoding PBS polypeptides, and our understanding of the ways in which the levels of the different phycobiliprotein components are modulated. Table 1 enumerates the different polypeptide constituents of the PBS, provides their gene designations and offers a brief description of the function of each (a more detailed discussion can be found in Chapter 6).

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Allen, 1984; Collier and Grossman, 1992). The degradation of the PBS under these conditions may allow for recycling of amino acids into proteins that aid in acclimation of the cell to nutrient-limited growth and/or may be important in preventing cellular damage via photo-oxidation. Finally, many cyanobacteria can alter the PBS composition in response to light quality (Bogorad, 1975; Tandeau de Marsac, 1983; Grossman, 1990). The next section will focus on this phenomenon.

A. Historical Perspective II. Chromatic Adaptation Many environmental parameters can significantly alter the composition or abundance of PBS. Low light intensities may stimulate the synthesis of PBS and cause an increase in size of the rod substructure (Öquist, 1974a,c; Lönneborg et al., 1985). Increased phycobiliprotein content is also observed in cyanobacteria that receive a preponderance of light wavelengths absorbed by PS I (i.e., chlorophyll; see Chapter 22). The increase in the PBS absorbance under such conditions may help balance the electron flow between the two photosystems (Melis et al., 1985; Manodori and Melis, 1986a,b). In contrast, macronutrient limitation results in extensive PBS breakdown during a response known as ‘chlorosis’ or bleaching (Foulds and Carr, 1977; Wood and Haselkorn, 1979, 1980; Yamanaka and Glazer, 1980;

Over a century ago it was observed that certain photosynthetic organisms can acclimate to their light environment by changing their cellular pigmentation (Engelmann, 1883a, b, 1884, 1902; Gaidukov, 1903 a, b, 1904, 1906). Based on their response to light quality (The term ‘light quality’ usually means a limited range of light wavelengths—e.g., red light, blue light, etc.—without regard to intensity.), cyanobacteria have been divided into three different groups (Tandeau de Marsac, 1977). Group I cyanobacteria can alter PBS size and number, along with photosystem stoichiometry, when grown in different light qualities, but do not dramatically alter the absorbance characteristics of their PBS through changes in phycobiliprotein composition. Group II cyanobacteria can alter the levels of PE in the PBS, and group III organisms can modulate both the PE and PC levels. Changes in pigmentation in

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photosynthetic organisms in response to light quality have been termed ‘chromatic adaptation.’ When the levels of PE alone, as in Group II organisms, or both the PE and PC pigments, as in the PBS of group III organisms, are modulated, the process is termed ‘complementary chromatic adaptation.’ Complementary chromatic adaptation has been most extensively examined in the cyanobacterium Calothrix sp. strain PCC 7601, although the phenomenon has been observed in a number of cyanobacteria (Tandeau de Marsac, 1977; Bryant and Cohen-Bazire, 1981; Bryant, 1981, 1982). This process is most dramatically demonstrated when the adapting organisms are compared after growth in RL and GL. In RL Calothrix sp. strain PCC 7601 accumulates high levels of the blue pigment PC and little of the red pigment PE. Conversely, in GL the organism has low levels of PC and high levels of PE. Since PC effectively absorbs RL while PE effectively absorbs GL, the change in character of the light-harvesting complex in response to the different wavelengths of light allows the cells to use the incident light efficiently. The events that trigger changes in phycobiliprotein gene expression during chromatic adaptation are probably initiated by the absorption of light by a

specific photoreceptor molecule(s). A number of different types of experiments have been performed to elucidate the nature of the photoreceptor. Early work concentrated on determining the action maxima for the light-triggered responses. The action maximum for the synthesis of PE for both Calothrix sp. strain PCC 7601 (Haury and Bogorad, 1977, Vogelman and Scheibe, 1978) and Tolypothrix sp. strain PCC 7101 (Tolypothrix tenuis; Fujita and Hattori, 1962; Diakoff and Scheibe, 1973) is between 540 nm and 550 nm, while the maximum for PC synthesis is between 650 and 660 nm. The accumulation of PC could also be promoted by shorter light wavelengths near 360 nm (Vogelman and Scheibe, 1978; Ohki et al., 1982). Similar observations have been made for the Type II chromatic adapter Synechocystis sp. strain PCC 6701 (Tandeau de Marsac et al., 1980). It is not known whether the short wavelength response is governed by the same regulatory elements as the RL/GL photoreversible response. It was first noted by Bryant and CohenBazire (1981; also see Bryant, 1981,1982) that more than one set of PC subunits was synthesized in chromatically adapting organisms of Type III grown in RL, while only a single set was present in cells

Chapter 21 Cyanobacterial Acclimation Processes grown in GL. These results implied that multiple PC genes were differentially controlled under the different conditions of illumination. This has been substantiated by more recent work (see below). The action spectrum for changes in the cellular phycobiliprotein content indicates that the photoreceptor involved in chromatic adaptation may to be a phycobiliprotein, like phytochrome, the photoreceptor in higher plants that controls many physiological and developmental processes (Schäfer and Briggs, 1986). Early attempts were made to isolate the photoreceptor involved in controlling chromatic adaptation, and it was suggested that AP was directly involved in photocontrol (Scheibe, 1972; Björn and Björn, 1980; Ohad et al., 1980). Cell extracts containing AP exhibited photoreversible absorption changes, as expected of a photoreceptor. However, photoreversibility of AP was only observed after partial denaturation or decomposition of AP and was not detectable within cells maintained under normal growth conditions (Ohki and Fujita, 1979a,b; 1981). Hence, the initial biochemical search for a photoreceptor was confounded by artifact. It was also likely that the photoreceptor would not be required at high levels and thus would be extremely difficult to identify amongst the sea of phycobiliproteins that constitute the structural components of the PBS. With the isolation of genes encoding both phycobiliproteins and linker polypeptides, the utilization of mutants that exhibit aberrant light responsiveness, and the development ofgene transfer technology, the processes controlling chromatic adaptation are now more amenable to detailed analysis.

B. Gene Expression Since almost all ofthe work referred to in this section concerns Calothrix sp. strain PCC 7601, a summary of the different genes encoding phycobiliproteins and linker polypeptides that have been isolated from this organism is presented in Table 2. The transcriptional characteristics of these genes have also been included. Nearly every gene listed in Table 2 encodes a structural component of the PBS. These genes have been characterized with respect to their sequences and the relative levels of their transcripts that accumulate in cells grown in RL and GL. Genes involved in regulating chromatic adaptation have only recently been isolated. One such gene, rcaC (Chiang et al., 1992a), has been included in Table 2.

645 Both our laboratory (Grossman et al., 1988; Grossman, 1990) and those of others (Tandeau de Marsac et al., 1988, 1990) have isolated and characterized genes encoding the PC, PE, and linker polypeptides from Calothrix sp. strain PCC 7601. The genome of this cyanobacterium contains three PC gene sets, and two of these are important to our discussion of chromatic adaptation. The mRNA from one accumulates constitutively while the mRNA from the second only accumulates in cells maintained in RL (Conley et al., 1985). The former gene set is cpcB 1A1 (the protein is or while the latter is cpcB2A2 (the protein is or ).The subscripts c and i denote constitutive and inducible, respectively. Similar results have been described for the Type III chromatic-adapter Pseudanabaena sp. strain PCC 7409 (Dubbs and Bryant, 1993), an organism that contains only two phycocyanin (cpc) operons. Transcripts from cpeBA, which encode the PE subunits, only accumulate to high levels in GL (Mazel et al., 1986, Oelmüller et al., 1988a,b; Dubbs and Bryant, 1991). The abundance of the mRNAs from cpcB1A1, cpcB2A2 and cpeBA reflects the polypeptide composition of the PBS under a given light condition. Furthermore, the half-lives of the mRNAs encoding and PE are not altered by exposure to different wavelengths of light (Oelmüller et al., I988a). These observations suggested that PBS biosynthesis in the different light qualities is to a great extent a consequence of differential transcription of the cpcB2A2 and cpeBA operons. Since specific linker polypeptides are associated with and PE, the expression of the genes encoding those polypeptides were also likely to be regulated appropriately in the different light qualities. The genes encoding linker polypeptides associated with are downstream of and cotranscribed with cpcB2A2 (Lomax et al., 1987). Overlapping transcripts are observed from this region of the genome. A transcript of 1.6 kb encodes the and subunits of while a transcript of 3.7 kb encodes both the and subunits of plus the three linker polypeptides required for assembly of onto the rod substructure (see Fig. 1). These linker polypeptides, and are encoded by the genes cpcH2, cpcI2 and cpcD2, respectively. The molecular masses of these proteins, indicated by the superscript to L, were calculated from knowledge of the gene sequences. The order of genes in the operon is cpcB2A2H2I2D2. There is the potential to form relatively stable, hairpin-loop structures at the 3'

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Chapter 21 Cyanobacterial Acclimation Processes ends of both the large and the small transcripts. The ratio of the 1.6- and 3.7-kb mRNAs from this operon, approximately 10:1, reflects the ratio of the subunits to the linker polypeptides in the PBS. The larger mRNA may be generated by transcriptional read-through approximately 10% of the time or may represent the primary transcript that is rapidly degraded to the more stable 1.6-kb transcript by a 3' exonuclease or a 5' exonuclease aided by endonuclease activity. The potential, large hairpin-loop structure at the 3' end of the 1.6-kb transcript (Conley et al., 1988) may serve as a transcription termination signal, a recognition site for RNA processing, or a structure that prevents further 3' exonuclease activity. This structure has a predicted of –31.7 kcal and is followed by a string of U residues that is characteristic of procaryotic rho-independent termination signals (Platt, 1986). The larger 3.7-kb transcript also terminates with an inverted repeat that has the potential to form a stable hairpin-loop structure with a predicted of –93 kcal While it is likely that the latter structure serves as a terminator since no longer transcripts have been detected from this operon, the role of the former in controlling the levels of the two transcripts and consequently the ratio of the PC subunits and linker polypeptides requires further investigation. The cpeBA genes, encoding the subunits of PE, have been sequenced from Calothrix sp. PCC 7601 (Mazel et al., 1986), Pseudanabaena sp. PCC 7409 (Dubbs and Bryant, 1991), and Synechocystis sp. strain PCC 6701 (Anderson and Grossman, 1990b). In Calothrix sp. strain PCC 7601, this gene set is transcribed as an approximately 1.5-kb mRNA (Mazel et al., 1986; Grossman et al., 1988) that is abundant in GL and present at low levels in RL. The cpeBA transcript from Synechocystis sp. strain PCC 6701 is 1.4-kb and is present at elevated levels in GL. Synechocystis sp. Strain PCC 6701 is a group II organism and while the level of the mRNA encoding PE is elevated in GL, light quality does not cause a change in the level of PC mRNA (Anderson and Grossman, 1990a). In Pseudanabaena sp. strain PCC 7409, the cpeBA operon was transcribed as a 1.4-kb transcript; this transcript was not usually detectable in total RNA isolated from cells grown in RL but was extremely abundant in cells grown in GL. Recently, Glauser et al. (1992) reported the presence of three linker polypeptides associated with PE hexamers of Calothrix sp. strain PCC 7601. The

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genes encoding these linker polypeptides have been characterized (Federspiel and Grossman, 1990, Federspiel and Scott, 1992). The protein sequences, as deduced from the nucleotide sequences of the genes, are similar to each other, and are also homologous to the PC linker polypeptides. In contrast to the situation for the linker polypeptide genes cpcH2I2D2 that are immediately downstream of cpcB2A2, these genes are not contiguous to cpeBA but are clustered into a separate operon. The PE linker genes are denoted cpeC, cpeD and cpeE, and transcripts from the operon containing these genes are 2.1 and 3.2 kb. The former transcript, which can accumulate to higher levels than the latter, encodes CpeC and CpeD (Federspiel and Grossman, 1990), while the latter encodes all three PE linker proteins (Federspiel and Scott, 1992). Located 3' of the translation stop codon of cpeD is a sequence that has a high potential to form a hairpin-loop structure (predicted This structure is followed by a string ofU residues that is characteristic of procaryotic rho-independent terminators (Platt, 1986). The occurrence of overlapping cpeCDE transcripts that accumulate to different steady state levels resembles the situation previously described for the operon encoding and the linker polypeptides, and may reflect a common mode of regulating the levels of different constituents of cyanobacterial macromolecular complexes. The 5' ends of transcripts from the cpeBA and cpeCDE operons of Calothrix sp. strain PCC 7601 have been mapped. The 1.4-kb mRNA encoding the PE and subunits begins 62–64 bases upstream of the translation initiation codon for the subunit while the cpeCDE transcripts have a leader of 187 bp. This latter sequence contains several small overlapping open reading frames (ORFs) of 14–21 amino acids that are associated with potential ribosome binding sites. Such short ORFs can be important for translational control in other bacteria, although the significance with respect to the cpeCDE transcript is not known. Similar ORFs are not present in the leader region of the cpeBA mRNA. The cpeCDE transcripts accumulate in parallel with cpeBA mRNA (Federspiel and Grossman, 1990). In an effort to delineate sequence motifs important for the light-wavelength-regulated expression of the phycobiliprotein genes, the regions upstream of the transcription start sites were examined. The constitutive PC gene set, cpcB1A1, may have both E.

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coli –10 (TATaat) and –35 (TTGaca) sequences. At position –10 to –15 is the sequence TATAGT while the sequence TTGACA is positioned –43 to –48 nucleotides upstream of the transcription start site. Sequences similar to the E. coli general transcription signals are associated with some of the regulated genes. For example, the cpcB2A2H2I2D2 operon has a TTGCAC at –29 to –35, the cpeBA operon has a TATGTT sequence at –8 to –13 and the cpeCDE operon has a TTGATG sequence at –35 to –40. It has not been established that these sequences play a functional role in the transcription of Calothrix sp. strain PCC 7601 genes. If they are important for the transcription of genes in this cyanobacterium, as in E. coli, they would likely serve as general transcription signals. Light-wavelength-responsive genes may contain additional cis-acting sequences upstream from these general motifs. Such sequences may be similar for similarly regulated genes. At this point too few similarly light-regulated genes have been identified and characterized, and only limited attempts have been made to identify sequences that confer specific regulatory characteristics to these genes. One candidate for a light-responsive regulatory element is located 83 bp upstream of the transcription start site of cpeBA. It is a 17 bp element TCCCCAGTCCCCAATCC that is also found, in the reverse complement, 195 bp upstream of the similarly-regulated cpeCDE operon. However, the function of this element in controlling GL-regulated transcription of cpeBA is doubtful since it resembles repetitive sequences that flank a number of genes in Calothrix sp. strain PCC 7601 (Mazel et al., 1990). The sequence CCCCA(A/G)T (the 17-bp element may represent contiguous repetitive elements) is repeated ten times in the region that precedes cpeBA. These sequences have been classified as short, tandemly repeated, repetitive sequences (STRR) and are scattered throughout the Calothrix sp. strain PCC 7601 genome (Mazel et al., 1990). A sequence of unknown significance GGATCAGG is repeated five times upstream of the cpeCDE gene cluster (Federspiel and Grossman, 1990). Ultimately, proving the role of a sequence motif in differential gene regulation will rely on defining the function of that motif in vivo. Such an analysis in turn, will require an appropriate genetic system. Recently, it has been shown that proteins bind to the promoter regions of both cpcB2A2 and cpeBA and possibly alter the transcriptional activity ofthese genes. Transcriptional control by such factors may

be modulated by light-driven phosphorylation events (see below). Similar factors may also control expression of cpeCDE.

C. Photobiology Action spectra for PE and PC synthesis in both Calothrix sp. strain PCC 7601 and Tolypothrix sp. strain PCC 7101 have been known for many years (Fujita and Hattori, 1962;Diakoff and Scheibe, 1973; Haury and Bogorad, 1977; Scheibe, 1972; Vogelman and Scheibe, 1978). Increased knowledge of the photobiology of chromatic adaptation provides some clues concerning the nature of the photoreceptor and the signal transduction pathway involved in the process. Photobiological characterizations of chromatic adaptation (Oelmüller et al., 1988a,b; 1989) suggest that: 1) Transcription from both cpcB2A2 and cpeBA can be triggered by a pulse of inductive light followed by darkness. For both gene sets the transcript population immediately increases following transfer to inductive light and reaches a maximum within 2 h at 25 °C. At elevated temperatures (32 °C) the response is more rapid. 2) The fluence required for altered transcription from the two gene sets is different. The response for both the increase and decrease in transcription from cpcB2A2 and cpeBA is saturated at a fluence of and 6 × respectively. These data imply that two distinct photoreceptors regulate expression from the two different phycobiliprotein gene sets or that there is a complex signal transduction chain between photoperception and the control of transcriptional activity that results in the different light responsiveness observed. 3) The kinetics of change in the rate of transcription from cpcB2A2 and cpeBA is different when cells are transferred from inductive to noninductive light. Transcription from cpcB2A2 declines rapidly upon transfer to noninductive light and is barely detectable after 2 h, while transcription from the cpeBA operon is still high even 10 h after the shift to noninductive light. Upon transfer to noninductive light, the level of cpeBA transcription decreases in concert with the generation time of the cells. It is reasonable to propose that a stable factor involved in positive regulation of the gene set is becoming progressively more dilute during cell growth. The rapid decline in transcriptional activity of cpcB2A2 upon shifting from inductive to noninductive light suggests that this gene set is

Chapter 21 Cyanobacterial Acclimation Processes controlled by a positive regulatory element that is unstable under noninductive light conditions or a negative regulatory element that is either synthesized de novo or activated under noninductive light conditions. We favor the idea of control of cpcB2A2 by either a negative regulatory element or both a positive and negative regulatory element. The alteration in the state of the photoreceptor by different wavelengths of light and the signal transduction pathway that communicates with the transcriptional machinery ofthe cell are not known. Many laboratories have attempted to isolate the photoreceptor by identifying a GL/RL photoreversible molecule, but have been unrewarded in these attempts as discussed above and as reviewed by Björn and Björn (1980). Although this photoreversible control system may involve a bilin-associated photoreceptor that resembles phytochrome, there are several important differences between the two photoresponsive systems. 1) The action maxima for the cyanobacterial photoreceptor are 540 nm (GL) and 640 nm (RL) while those for phytochrome are 660 nm (RL) and 730 nm (FRL). 2) Both forms of the cyanobacterial photoreceptor are biologically active in the sense that specific gene sets aretranscriptionally active at both action maxima. For phytochrome, only the RL-activated species promotes the biological response. 3) The cyanobacterial system exhibits no escape from photocontrol. Transcription from cpcB2A2 and cpeBA remains fully on or off immediately following the terminal irradiation and it is possible to reverse the effect fully with a pulse of complementary light, even after several hours of darkness following the terminal light pulse. In the phytochrome system, aperiod of darkness, following a terminal pulse of activating light, results in a diminution of photoreversibility. 4) Plants possess multiple genes encoding different forms of phytochrome. Some are tissue specific and may be developmentally regulated (Tomizawa et al., 1990). It is unknown whether there is only a single photoreceptor involved in chromatic adaptation and if that photoreceptor controls other cellular processes [It might play a role in the formation of hormogonia (Damerval et al., 1991; see Chapter 27)]. Despite extensive characterization of complementary chromatic adaptation at the level of gene expression, little is known about the photosensory and signal transduction components that govern this process. Photoregulation ofcellular processes is well documented for a number of higher plant systems,

649 however, the way in which the photoreceptor communicates with the transcriptional machinery of the cell is largely unknown. The photobiology of complementary chromatic adaptation in Calothrix sp. strain PCC 7601 suggests that the state of the photoreceptor is linked to the activity ofgene-specific transcriptional regulators. Since cyanobacteria are amenable to standard procaryotic genetic manipulation, they can be used as a relatively simple model to investigate photoresponsiveness at the molecular level. Especially beneficial for the analysis of this control process is the acquisition of mutants that display aberrant chromatic adaptation.

D. Regulatory Mutants Many Calothrix sp. strain PCC 7601 mutants that exhibit aberrations in chromatic adaptation have been isolated (Cobley and Miranda, 1983; Tandeau de Marsac, 1983; Beguin et al., 1985; Bruns et al., 1989; Mazel et al., 1991). These mutants may arise spontaneously and are easily identified by visual inspection. The exposure of wild-type cells to mutagenic agents or electric shock significantly increases the frequency at which pigment mutants appear. A genetic and molecular characterization of such mutants is extremely important in defining the detailed mechanism of chromatic adaptation. Cobley and Miranda (1983) were the first to report the characterization of pigment mutants after treatment of wild type cells with UV irradiation. Three classes of mutants were defined based on spectral examination of the cells. In green mutants the synthesis of PE did not occur in either GL or RL while PC synthesis was normal. In blue mutants both photoinduction of PE and photorepression of PC synthesis were impaired. In a black mutant PE was partially induced and PC partially repressed in RL. Hence, the black mutant responded to light quality in an opposite way to wild type cells. In 1983, a review article by Tandeau de Marsac (1983) described six classes of PBS regulatory mutants. These strains, isolated after exposure of wild-type cells to nitrosoguanidine, were postulated to represent six of eight possible mutant phenotypes in which PC and PE are abnormally regulated. At this point, a lack of detailed information about the different phenotypes does not allow for a critical evaluation of the model. Numerous mutants of Calothrix sp. strain PCC 7601 that exhibit abnormal chromatic adaptation have also been isolated and characterized in our

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laboratory. These mutants were generated during optimization of conditions for the transfer of DNA into this strain by electroporation. Three mutant classes (red, blue and green) were generated (Bruns et al., 1989) and characterized with respect to spectral analyses of whole cells and cell extracts, determination ofphycobiliprotein and linker polypeptide composition of intact PBS and the quantification of transcripts from cpeBA, cpcB1A1, cpcB2A2 and apcAB. These studies helped to distinguish regulatory mutants from mutants with lesions in the structural genes of the PBS or in PBS assembly processes. Cells of the red mutant class (designated FdR) have lost the ability to acclimate to RL. The FdR mutants have high levels of PE, normal levels of and no detectable under conditions of RL or GL. The levels of the individual phycobiliproteins are reflected in the levels of the mRNAs encoding those proteins. Hence, FdR mutants are locked in the GL regulatory mode and are indifferent to the presence ofRL. Blue mutants, designated FdB, exhibit normal regulation of cpeBA and elevated transcription from cpcB2A2 in both RL and GL. Our FdB mutants do not exhibit the impaired PE synthesis observed for the blue mutant class reported by Cobley and Miranda (1983). Cells of the green mutant class, designated FdG, have lost the ability to synthesize PE but regulate the levels of normally. The information gleaned from the characterization of these mutants has established certain general features of chromatic adaptation. Firstly, single lesions can alter the regulated transcription from both the cpcB2A2 and cpeBA operons, as exemplified in the phenotype of FdR strains—a phenotype frequently observed in mutant populations. This suggests the presence of at least one common element in the chain of events governing transcription of cpcB2A2 and cpeBA. Since expression of cpeBA and cpcB2A2 in FdR is locked in a mode in which the cells behave as if they are growing in GL, the putative activator and repressor that control expression of cpeBA and cpcB2A2, respectively, remain active even in RL. Secondly, there are distinct elements that control the expression of cpcB2A2 and cpeBA, as demonstrated in the phenotype of FdB. At this point, our conclusions concerning the control of cpcB2A2 by a repressor and the control of cpeBA by an activator are still tentative. Indeed, some results from both our laboratory (Oelmüller et al., 1989) and the laboratory of others (Houmard et al., personal communication) suggests that more than one regulatory element may

be involved in controlling the expression of each of the phycobiliprotein genes. In this respect the mutants described above are valuable since, in conjunction with a gene transfer system developed for Calothrix sp. strain PCC 7601, they will aid in the identification of regulatory elements integral to the phenomenon of chromatic adaptation. The generation and analysis of double mutants is also a powerful method for analyzing chromatic adaptation and the hierarchy of events involved in its regulation. When an FdR strain is used for the generation of double mutants, only the green mutant phenotype appears. Mutagenesis of the blue mutant FdB yields colonies of a purple phenotype (both PE and are expressed constitutively). No altered phenotype has thus farbeen obtained for a population of mutagenized green mutants. These results, which begin to establish a hierarchy of mutant phenotypes, suggest that the green mutant phenotype is dominant over the other pigment mutant phenotypes.

E. Approaches for Isolating Regulatory Elements The complementation of mutants of Calothrix sp. strain PCC 7601 exhibiting aberrant chromatic adaptation has recently been achieved. Unlike many gram-negative procaryotes and a number of unicellular cyanobacteria, Calothrix sp. strain PCC 7601 is not naturally competent for transformation with exogenous DNA. A gene transfersystem, initially established by Cobley and coworkers (Cobley et al., 1987), has been developed for the introduction of DNA into Calothrix sp. strain PCC 7601. This system employs a vector that can replicate both in E. coli and the cyanobacterium. This shuttle vector, designated pPL2.7, harbors a 2.7-kbp fragment from an endogenous Calothrix sp. strain PCC 7601 plasmid that contains all the necessary genetic elements for replication in the cyanobacterium. This plasmid also contains the ColE1 replication origin, the bom (basis of mobility) region which facilitates the conjugal transfer of the plasmid from an E. coli host to a recipient strain, and the gene encoding aminoglycoside 3'-phosphotransferase, that confers resistance to the antibiotic kanamycin. For complementation experiments a library of random Sau3A fragments of wild-type genomic DNA was ligated into pPL2.7 and the recombinant molecules introduced into Calothrix sp. strain PCC 7601 by either electroporation or conjugation. The plasmids are stably maintained once inside the cyanobacterial

Chapter 21 Cyanobacterial Acclimation Processes cells (Chiang et al., 1992b). Complementation of regulatory mutants also relies on a facile screen for choosing transformants that exhibit the wild-type phenotype. The dramatic pigmentationdifferences betweenwildtypeandPBS regulatory mutants provides a clear visual screen for complementation. For example an FdR mutant is reddish-brown when grown in RL whereas wild-type cells appear blue-green. After introduction of the genomic library into an FdR strain and growth ofthe transformed cells in RL, only the cells in which the lesion in the FdR mutant had been complemented would appear blue-green. They would be readily identified in a background of reddish-brown colonies containing cells that were transformed, and therefore resistant to kanamycin, but not complemented. Similarly, complemented FdG mutants should be identifiable as reddish-brown colonies among noncomplemented blue-green cells when grown in GL. Several genes encoding putative components of the signal transduction pathway have been isolated following complementation of members of the FdR and FdB mutant classes. The most thoroughly characterized gene, designated rcaC, was localized on a large plasmid that was rescued from a single complemented colony of the red mutant FdRl (Chiang et al., 1992b). Reintroduction of the complementing plasmid into the mutant strain resulted in a high frequency of complemented colonies. The selected plasmid could also confer the wild type phenotype to cells of a second red mutant, FdR2. Analysis of the complementing fragment revealed that the rcaC gene encodes a protein that has strong sequence identity with the Bacillus subtilis PhoP protein (Seki et al., 1987), a member of the superclass of regulatory proteins associated with bacterial two-component regulatory systems (for reviews, see Albright et al., 1989; Stock et al., 1990). Most response regulators have a conserved aminoterminal domain containing an aspartate residue that can be reversibly phosphorylated. The RcaC protein is unusual in that it contains conserved domains with potentially phosphorylated aspartate residues at both the amino- and carboxyl-termini. Southern-blot analysis of genomic DNA isolated from the two red mutants revealed gross, nonidentical structural aberrations at the rcaC locus, suggesting that the initial lesions were due to independent events involving mobile genetic elements. It had previously been reported that Calothrix sp. strain PCC 7601

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possesses multiple copies of two transposable elements (Mazel et al., 1991). These results imply that electroporation ofwild-type Calothrix sp. strain PCC 7601 cells triggered the movement of genetic elements, that in some cases caused a disruption in the rcaC locus. Genes whose products may be involved in chromatic adaptation were also isolated by rescuing plasmids thought to be involved in complementation of the blue mutant FdB1. Sequence analysis of the genomic DNA on the complementing plasmid revealed three large tandemly arranged open reading frames (ORFs). Two of the ORFs encode nearly identical proteins with strong identity to the E. coli protein (Miranda-Rios et al., 1987), a member of a superclass of sensory proteins associated with the aforementioned bacterial two-component regulatory systems. The third ORF encodes a protein with sequence similarity to rat myosin light-chain kinase (Roush et al., 1988), a member ofa eucaryotic class of serine/threonine kinases. A similar serine/ threonine kinase has been described in another procaryote, Myxococcus xanthus (Muñoz-Dorado et al., 1991). Although these genes were present on plasmids from independently isolated complemented strains, recomplementation of the mutant with the isolated plasmid was not achieved. Hence, the role of these kinases in the process of chromatic adaptation remains uncertain. While it is possible that complementation of FdB1 was the consequence of marker rescue (which is probably a very rare event in Calothrix sp. strain PCC 7601), the presence of the kinase genes on the recombinant pPL2.7 isolated from complemented strains of the blue mutant may be fortuitous. Further insight into the signal transduction mechanism governing chromatic adaptation can be achieved by studying the nature and activity of proteins that directly interact with the region of the cpeBA and cpcB2A2 genes upstream of the transcription initiation site. These interactions, that may be important for regulating gene activity, can be examined by extracting proteins from cells and incubating these proteins with specific DNA fragments. Theprotein-DNA complexeshave altered electrophoretic mobilities in a polyacrylamide gel matrix (gel retardation assay). The site specificity and binding activity of a transcriptional regulatory protein can be defined by gel retardation assays, and the putative regulatory gene encoding the binding protein may be isolated by screening a recombinant

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expression library with labeled target DNA (generally an oligomer of the specific region of the DNA that binds the protein). Gel retardation studies using DNA fragments from the vicinity ofthe transcription initiation site forcpeBA demonstrated the binding of specific proteins, only present in extracts from GLgrown cells (Dubbs and Bryant, 1991; N. Federspiel and A. Grossman, unpublished results). No binding to the same region was observed by proteins extracted from RL-grown cells. These data suggest that a trans-acting element is present or active only in cells cultured in GL and that it binds to and may facilitate transcription from the cpeBA promoter. Additional evidence for regulatory factors that interact with regions that precede the transcription initiation sites ofcpcB2A2 and cpeBA comes from recent studies by Sobczyk et al. (1993) and E. Casey and A. Grossman (unpublished results). Sobczyk et al. (1993) found that proteins from extracts of GL-grown cells were bound to DNA upstream of the transcription initiation site of cpeBA at two different locations. The binding proteins have been isolated, characterized and named RcaA and RcaB. One ofthe proteins loses its binding activity when it is treated with alkaline phosphatase (Sobczyk et al., 1993). Hence, two trans-acting proteins may interact with DNA sequences upstream of the transcription initiation site of cpeBA and alter transcription from that operon. These regulatory proteins may be controlled by phosphorylation events triggered by changes in the light environment. Proteins have also been shown to bind DNA sequences associated with cpcB2A2 (J. Houmard, personal communication; E. Casey and A. Grossman, unpublished results). One strong binding site, that is 30 nucleotides long and positioned approximately 120–150 nucleotides upstream of the cpcB2A2 transcription start site, has been observed for this operon. This region binds protein(s) from cells grown in both GL and RL. In-vivo analyses of chimeric genes in which regulated promoters are fused to reporter genes (GUS, luciferase) will help establish the significance of both cis-acting DNA sequences and trans-acting proteins in the regulated expression of phycobiliprotein genes. With the aid of a genetic system and the capabilities to evaluate promoter activity in vivo, coupled with in vitro, biochemical analyses of cis- and trans-acting elements associated with the light-regulated genes, the features of the cpcB2A2 and cpeBA promoter regions important for light-regulated transcription should be rapidly delineated.

F. Model Describing the Regulation A working model forthe RL/GL-regulated expression of genes during chromatic adaptation is developing. A reasonable assumption in generating a model is that the activity or abundance of transcriptional regulators is influenced by light quality. Before presenting the model, the features of chromatic adaptation crucial in developing this model will be summarized. Following the transfer of wild type cells from GL to RL, transcription from cpcB2A2 (the operon includes cpcB2A2H2I2D2) rapidly increases whereas transcription from the cpeBA and cpeCDE operons slowly decreases. When wild-type cells are transferred from RL to GL, transcription from cpcB2A2 rapidly decreases while transcription from cpeBA rapidly increases. A negative regulatory element may control transcription from the inducible PC operon while a positive regulator may control transcription from the PE operon. The initial model (Fig. 2), which simply provides a framework for examining light-regulated expression during chromatic adaptation, involves gene-specific regulation that is governed directly by trans-acting elements. In RL neither the represser of cpcB2A2 nor the activator of cpeBA transcription are functioning (represented as A and R which are not bound to cpeBA and cpcB2A2 in Fig. 2; this is only a pictorial representation and the nonfunctional transcription regulators may still bind DNA). This situation causes increased transcription from cpcB2A2 and low-level transcription from cpeBA. In GL the transcription regulators become active (represented as being bound to the DNA); this results in elevated transcription from cpeBA and repression of the cpcB2A2 operon. The work of Houmard and coworkers (Sobczyk et al., 1993; J. Houmard, personal communication), Chiang et al. (1992a), and in-vivo experiments in which proteins are phosphorylated under GL and RL (E. Casey and A. Grossman, unpublished results), suggests that a phosphorylation cascade is involved in controlling the activity of the regulatory elements. In Fig. 2 the phosphorylated elements are represented as functioning directly in the activation and suppression of transcription. The phosphorylation cascade contains the two elements X and Y (the number of elements in the cascade is arbitrary). Some ofthese elements might be histidine kinases, which are prominent in the signal transduction pathways in many procaryotes. The next question concerns how excitation of the

Chapter 21 Cyanobacterial Acclimation Processes

photoreceptor modulates the phosphorylation cascade. Based on the photobiology of chromatic adaptation and the limited studies of mutants that exhibit aberrant chromatic adaptation (Bruns et al., 1989; Oelmüller et al., 1988a,b; 1989), it is proposed that altered transcription from cpcB2A2 and cpeBA is governed by a single photoconvertible photoreceptor (this may be an oversimplification) and that in GL this photoreceptor is maintained in an inactive form Under these conditions Calothrix sp. strain PCC 7601 produces large amounts ofPE and minimal amounts of PC (both the repressor and activator are functioning). When the ratio of RL to GL becomes high, the photoreceptor is converted to an active form thatcan negatively regulate the phosphorylation cascade thereby preventing phosphorylation of the regulatory elements A and R. The transcription regulators would become inactive, and the cells would synthesize high levels of and low levels of PE. Additional support for this model comes from studies of PBS regulatory mutants. In the FdB mutants cpcB2A2 is expressed in both RL and GL while cpeBA is normally regulated. This phenotype could be the consequence of a lesion in a cpcB2A2-specific

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repressor, R. In the FdG mutants that have been characterized there is normal regulation of cpcB2A2 and no transcription from cpeBA in either RL or GL. This phenotype could be the consequence of inactivation of the positive regulator, A. Mutants of the FdR class exhibit constitutive expression of cpeBA and no expression of cpcB2A2 in either RL or GL. Such a phenotype could be due to a lesion in the photoreceptor (making it impossible to suppress the phosphorylation cascade) or an intermediate that links the RL signal perceived by the photoreceptor to inhibition of the phosphorylation cascade. Further support for the model comes from the determination of allowable phenotypes observed upon the creation of double mutants. For example, FdR mutants (constitutive functioning of the phosphorylation cascade) can be mutated to FdG (inactive phosphorylation cascade or the gene encoding the activator of cpeBA; A. Grossman, unpublished results). Secondary mutations in an FdB background could result in a new phenotype in which the organism becomes purple in both RL and GL. If the second lesion were in the photoreceptor or the mechanism by which the photoreceptor regulates the phosphorylation cascade,

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cpeBA would become constitutive, and the double mutant would appear purple in both RL and GL. As previously discussed, a gene, rcaC, that complements the FdR mutant has been isolated (Chiang et al., 1992a). In our model the FdR phenotype reflects a constitutively active state for regulators controlling cpcB2A2 and cpeBA; R is functioning as a repressor and A as an activator. Such a phenotype may be the consequence of a lesion in the photoreceptor or in components involved in the negative control mechanism immediately downstream of the photoreceptor. Hence, RcaC may be a transcriptional regulator necessary for expression of the photoreceptor gene or for genes encoding components of the negative signaling pathway. It is equally possible that RcaC is directly involved in negative signaling. Only through continued genetic studies will the details of the signaling pathway be firmly established. In summary, Fig. 2 presents a model proposing that the altered transcription of cpcB2A2 and cpeBA during chromatic adaptation is achieved by coordinated activation and deactivation of opposing, gene-specific transcriptional regulatory elements. These regulators are controlled by a phosphorylation cascade that can be inactivated in RL as a consequence of the accumulation of the form of the photoreceptor. Other environmental conditions such as light intensity and nutrient status ofthe cultures may also modulate the activity ofthis regulatory pathway. III. The Responses of Cyanobacteria to Nutrient Deficiency The myriad of responses exhibited by nutrientdeficient cyanobacteria have been previously reviewed (Healey, 1982; Allen, 1984; Bryant, 1986, 1991; Simon, 1987; Reithman et al., 1988). Studies of nutrient deficiency have been performed under a variety of conditions that can be grouped into two basic types. In some, cyanobacterial growth was limited by a nutrient provided at low levels, allowing growth (cell division) to continue under nutrientpoor conditions (nutrient-limitation), usually in a chemo- or turbidostat. In other experiments a nutrient was completely eliminated from the medium and the cyanobacteria were unable to continue growth (nutrient-deprivation or depletion). The terms ‘limited’ and ‘deprived’ (or ‘depleted’) will be used in these specific senses here, and the term ‘deficient’

will be used in a more general sense to encompass any condition under which growth is restricted by the availability of a nutrient. Some ofthe responses ofcyanobacteria to nutrient deficiency are specific and may be triggered by deficiency ofone particular nutrient, while others are general and occur during deficiency for any of a number of different nutrients. The former category includes increased synthesis of specific transport systems and enzymes that transform inaccessible forms of a nutrient into those that the cell can use. The latter category includes changes in cellular morphology, intracellular nutrient reserves, and in the activity of numerous physiological processes, including photosynthesis. Recent progress in identifying genes involved in the specific nutrientuptake processes will first be discussed. The more general responses of changes in the abundance of inclusion bodies and alterations in the photosynthetic apparatus will then be explored. Synechococcus sp. strain PCC 7942, Synechococcus sp. strain PCC 6301 (Anacystis nidulans), and Synechococcus leopoliensis are very similar strains, and where appropriate they will be grouped as Synechococcus sp. in the discussion. A number of Anabaena sp. strains are also treated as a group.

A. Nutrient-Specific Responses 1. Micronutrients Specific responses of cyanobacteria to deficiency of a given nutrient often aid the cell in elevating intracellular concentrations of that nutrient. In addition, when the limitingnutrient is amicronutrient such as Fe or Cu, cellular demand for it may be reduced. To reduce the need for Fe (which is absolutely essential for viability) during periods of low Fe availability, many but not all cyanobacteria (Entsch et al., 1983) replace the Fe-containing protein ferredoxin, a component of the photosynthetic electron transport chain, with flavodoxin, an electron carrier that does not contain Fe (Bryant, 1986; Laudenbach et al., 1988; Filiat et al., 1991; Bottin and Lagoutte, 1992; see Chapters 12 and 25). This replacement causes no significant loss of photosynthetic function (Sandmann et al., 1990). Accumulation of flavodoxin under low-Fe conditions is regulated at the level of transcription (Laudenbach et al., 1988; Fillat et al., 1991), while the abundance of ferredoxin mRNA is unaffected by the Fe status of

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Chapter 21 Cyanobacterial Acclimation Processes the cell (Laudenbach et al., 1988; Van der Plas et al., 1988). The importance ofthis acclimation process in fresh water environments is supported by the finding of natural cyanobacterial populations containing only flavodoxin (Ho et al., 1979). Potential limitation by Fe in a marine environment has been inferred from studies in which field-collected samples of the cyanobacterium Trichodesmium sp. exhibited increased C- and N-fixation in response to Fesupplementation (Rueter, 1988). During Fe-deficiency cyanobacteria may also exhibit an increased capacity to take up Fe, which results from the synthesis of Fe-scavenging siderophores (Kerry et al., 1988; Hutchins et al., 1991; Mahasneh, 1991). In cells limited for Fe, new cytoplasmic andoutermembraneproteinsaccumulate (Scanlan et al., 1989). Some ofthese proteins may be constituents of an energy-dependent ferric-siderophore transport system (Lammers and Sanders-Loehr, 1982). Recently, a gene encoding a protein thought to be involved in Fe uptake or storage has been characterized (Reddy et al., 1988). A more detailed description of the acquisition and utilization of iron by cyanobacteria is found in Chapter 25. When limited for Cu, some cyanobacteria are able to replace the Cu-containing photosynthetic electron carrier plastocyanin with cytochrome (Ho and Krogmann, 1984; Bryant, 1986; Sandmann, 1986; Briggs et al., 1990). Expression of both plastocyanin and cytochrome is regulated at the transcriptional level (Van der Plas et al., 1989, Zhang et al., 1992), although post-transcriptional regulation my also occur (Briggs et al., 1990, Zhang et al., 1992). Some natural populations of cyanobacteria contain no detectable plastocyanin (Ho et al., 1979). The absence of plastocyanin may be due either to Cu-deficiency in the environment or the presence of cyanobacteria that cannot synthesize plastocyanin. It is interesting to note that Synechococcus sp. strain PCC 7942, which cannot synthesize plastocyanin, remains viable when the cytochrome gene is inactivated (Laudenbach et al., 1990). This implies that an alternate, as yet uncharacterized, electron carrier may substitute for cytochrome in the photosynthetic electron transport chain, or that electrons can be transported directly from the cytochrome complex to PS I. A similar observation was made for Rhodobacter capsulatus. When the electron carrier cytochrome was inactived, the organism was still able to perform photosynthesis. This was shown to be due to the presence of a novel, membrane-

659 associated c-type cytochrome (Jenny and Daldal, 1993). It is important to note that some natural populations of cyanobacteria that are lacking in ferredoxin and plastocyanin may also be lacking in cytochrome (Ho et al., 1979). Hence, habitats in which cyanobacteria grow may impose both Fe and Cu limitations on the organisms and alternative electron carriers and mechanisms for reducing the cellular demand for Fe and Cu may be of considerable ecological importance.

2. Macronutrients Many cyanobacteria can actively take up inorganic carbon both as carbon dioxide and bicarbonate, which allows them to concentrate within the cell up to 1000-fold over external levels (Pierce and Omata, 1988; Miller et al., 1990; Kaplan et al., 1991). This process requires ATP, the majority of which is probably generated by cyclic photophosphorylation (Pierce and Omata, 1988; Kaplan et al., 1991; Ogawa, 1992). The uptake of carbon dioxide may be constitutive while that of bicarbonate increases when levels limit growth (Badger and Price, 1990, Espie et al., 1991); this increased uptake capacity is reflected in a decreased of photosynthesis for (Turpin et al., 1984). The external concentration that causes cyanobacteria to increase their bicarbonate transport capabilities in culture is similar to the level present in freshwater (Miller et al., 1984), suggesting that elevated uptake is important in the natural environment. Interestingly, marine cyanobacteria examined to date may not possess an inducible mechanism (Karagouni et al., 1990). This finding may reflect the greater availability of bicarbonate in marine as compared to freshwater environments. Recent progress in understanding and identifying systems involved in the uptake and metabolism of are discussed in detail elsewhere in this volume (see Chapter 15). We will only briefly discuss nitrogen (N) acquisition and metabolism since it has been reviewed recently (Guerrero and Lara, 1987), and is described in detail in Chapters 16 and 27. Cyanobacteria can use a variety of sources of N, including ammonium, nitrate, and nitrite. Most can also use organic compounds such as urea and many are able to fix dinitrogen. Motile, marine cyanobacteria are chemotactic toward specific N-sources (Wiley and

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Waterbury, 1989). Ammonium is the preferred source of N, and its inclusion in growth medium often represses the synthesis of systems required for the uptake and assimilation of other N-containing compounds (Kashyap and Singh, 1985; Guerrero and Lara, 1987; Prasad and Kashyap, 1990; Singh, 1990). In some cyanobacteria, the presence of the proper substrate may also be required for the expression of a specific transport system. Recently, nitrate uptake in Synechococcus sp. strain PCC 7942 was characterized, and the for the substrate was reported to be (Rodriguez et al., 1992). The same carrier may also transport nitrite with a slightly lower affinity. Genes encoding the nitrate transporter and nitrate- and nitrite-reductases have been identified in Synechococcus sp. (Omata, 1991; Luque et al., 1992). These genes are clustered in an operon, and are not transcribed in ammonium-containing medium, but are transcribed when nitrate is the sole N-source (Suzuki et al., 1992). In N-fixing cyanobacteria, both nitrate and nitrite can repress the development of heterocysts and Nfixation in the presence or absence of dinitrogen (Guerrero and Lara, 1987). The expression of nitrogenase itself is transcriptionally controlled by the availability of other N-sources (Martin-Nieto et al., 1991). The assimilation of most N sources proceeds through the enzyme glutamine synthetase, which catalyzes the addition of ammonium to glutamate to form glutamine, although alternative pathways also exist (Wagner, et al. 1993). Cells grown on nitrate have higher levels of glutamine synthetase than those grown on ammonium (Edmond et al., 1979; Merida et al., 1990, 1991). Increases in the levels ofglutamine synthetase transcripts (Wagner et al., 1993) and activity (Paone and Stevens, 1981; Merida et al., 1991) have also been observed in cells deprived of N, although this is not always the case (Edmond et al., 1979). The molecular details of the regulation of Nassimilation in cyanobacteria are not well understood. Glutamine synthetase activity is regulated (reversibly) in the short term by ammonium (Merida et al., 1991), although cyanobacterial glutamine synthetase does not appear to be covalently modified in the same manner as its enterobacterial homologue, suggesting that cyanobacteria use a different mode of regulation. However, a component of the enterobacterial N-control system has been identified in cyanobacteria (Tsinoremas et al., 1991), and is modified in response to both photosynthetic activity

and ammonium availability. Furthermore, a locus involved in N-regulation has been cloned from Synechococcus sp. strain PCC 7942 (Vega-Palas et al., 1990), and recently the protein encoded by this locus was shown to be NtcA, a global nitrogen regulator belonging to the Crp family of transcriptional regulators (Vega-Palas et al., 1992; see Chapter 16). This recent work suggests new and exciting areas of exploration. Protein turnover, which is an important and poorly understood aspect of the cyanobacterial response to nutrient deficiency, has been examined most thoroughly in N-deficient cells. A large proportion of cellular protein is degraded in Anabaena sp. during the differentiation of heterocysts after transfer from medium providing a source of fixed N to medium lacking N (Ownby et al., 1979; Thiel, 1990). Both phycobiliproteins and other cellular proteins are degraded in N-deprived Synechococcus sp. (Boussiba et al., 1984). In Spirulina sp., a considerable fraction of cellular PC may be degraded before the growth rate is affected by decreasing N availability (Boussiba and Richmond, 1980). Similar observations have been made for PE in a marine cyanobacterium (Wyman et al., 1985). The material released as a consequence of protein degradation may provide substrates for de novo synthesis of polypeptides required for acclimation to lower N availability. Proteolysis may be energy dependent, and relies upon new protein synthesis following the onset of N deficiency. Both general proteases and those that specifically degrade PC have been detected (Foulds and Carr, 1977; Boussiba and Richmond, 1980; Wood and Haselkorn, 1979, 1980; Yamanaka and Glazer, 1980; Elmorjani and Herdman, 1987). The gene encoding a general protease has been cloned from Anabaena sp. (Lockau et al., 1988; Maldener et al., 1991). It appears to be of the serine type, but shows no significant similarity to other known serine proteases. This protease is believed to be identical to the protease that was reported to be induced during N-deficiency, as heterocyst differentiation began in Anabaena sp. transferred from medium containing nitrate to medium without a source of fixed N( Wood and Haselkorn, 1979,1980). The activity of this protease has now been shown to be identical in nitrate-grown and N-fixing Anabaena sp. (Lockau et al., 1988, Maldener et al., 1991). A transient increase in the activity of the protease may occur during the mass differentiation of heterocysts after transfer from growth on fixed N to N-fixing

Chapter 21 Cyanobacterial Acclimation Processes conditions, but is not maintained during steady-state growth of N-fixing cultures. Furthermore, while inactivation of this gene eliminated the dependent protease activity, there was no effect on heterocyst differentiation (Maldener et al., 1991). When cyanobacteria are deficient in P, they synthesize periplasmic or extracellular phosphatases and exhibit an increased capacity to take up phosphate. The review by Healey (1982) summarizes the work on P-nutrition up to about 1980. Phosphate uptake may involve more than one transport system in cyanobacteria. For example, Synechococcus sp. may have three phosphate transporters. The regulation of these transporters has not been examined in detail, but in Synechococcus sp. P-limitation results in a 50fold increase in the for P-transport, while the remains unchanged (Grillo and Gibson, 1979). Although most cyanobacteria appear unable to take up organic P-containing molecules, the production of multiple extracellular or periplasmic phosphatases under P-deficient conditions (Doonan and Jensen, 1980; Healey, 1982; Marco and Orus, 1988a, b; Whitton et al., 1990; Islam and Whitton, 1992b) allows the cells to scavenge phosphate by hydrolyzing it from a wide variety of organic P-sources. Some cyanobacteria are also able to use hydroxyapatite as a source of both and P (Cameron and Julian, 1988). Progress is being made in identifying components of cyanobacterial P-acquisition systems. The iphP gene for a phosphatase from Nostoc commune has been cloned (Xie et al., 1989) and overproduced (Potts et al., 1993), and appears to be a periplasmic protein with a mass of 74 kDa. The alkaline phosphatase that accumulates in P-deficient Synechococcus sp. strain PCC 7942 is a periplasmic protein with a molecular mass of approximately 150 kDa (Block and Grossman, 1988). This alkaline phosphatase (PhoA) appears to be transcriptionally controlled, and mRNA encoding PhoA only accumulates in P-deficient cells. Characterization of PhoA has demonstrated that it is unlike any other described phosphatase (Ray et al., 1991). The first one third of the protein has homology to the subunit of the coupling factors (ATPases) of chloroplasts and 5' nucleotidases ofbacteria. A region of about 50 amino acids shows strong homology to the UshA protein, a UDP sugar hydrolase of E. coli that has 5' nucleotidase activity. The last half of the protein has sequences homologous to P-loops, which are motifs found in kinases that are involved in

661 binding nucleotide triphosphates via the terminal phosphate ester. The presence of multiple P-loops in the alkaline phosphatase might allow for the binding ofphosphate groups attached to a variety ofdifferent compounds, thereby expanding the substrate specificity of the enzyme. This would be a desirable trait for an enzyme utilizing diverse phosphorylated compounds in its natural environment. Since P is most often the limiting nutrient in freshwater ecosystems (Hecky and Kilham, 1988), the phosphatases of cyanobacteria probably serve a vital role in natural cyanobacterial populations. Synechococcus sp. are able to use a number of organic and inorganic S compounds, including sulfate, thiosulfate, cysteine, cystine, reduced glutathione, and thiocyanate (Schmidt et al., 1982; Lawry and Jensen, 1986; Laudenbach and Grossman, 1991). The utilization of sulfate is currently the most thoroughly understood. Sulfate uptake is an active, light-dependent process that varies with both temperature and pH (Utkilen et al., 1976; Jeanjean and Broda, 1977). Metabolic poisons such as DCCD and CCCP, sulfate analogues such as selenate and chromate, and the S-compounds thiosulfate and sulfite all inhibit the transport of sulfate into the cell. The capacity for sulfate uptake increases in cells deprived of S. Utkilen and coworkers (Utkilen et al., 1976) reported a of and a of 0.7 pmol for sulfate transport at 42 °C in Sstarved Synechococcus sp. Periplasmic components were implicated in the transport process since osmotic shock treatment was inhibitory. Once sulfate is transported into Synechococcus sp. cells it is reduced via the PAPS sulfotransferase pathway (Schmidt and Christen, 1978). Other cyanobacteria can reduce sulfate via the APS sulfotransferase pathway (Tsang and Schiff, 1975). Initial physiological studies with Synechococcus sp. strain PCC 7942 (Green and Grossman, 1988) demonstrated that while the for sulfate transport increased between 10- and 20-fold during Sdeprivation, the was approximately in both S-sufficient and S-deficient medium. This suggested that S deficiency caused elevated accumulation of a single sulfate transport system. A region of the Salmonella typhimurium genone that was genetically defined to contain genes important for growth of the bacterium on sulfate was used to isolate an analogous region of the cyanobacterial genome (Green et al., 1989; Laudenbach et al., 1991; Laudenbach and Grossman, 1991). A map of this region is shown in

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Fig. 3. Transcripts from many of the genes in this region are either undetectable or were detected at very low levels in cells grown in nutrient-replete medium. A marked increase in the accumulation of the mRNA transcribed from these genes is apparent when the cells are deprived of S. The first gene from this region to be characterized encoded a nucleotide binding protein (gene designation cysA) of a periplasmic permease system. Periplasmic transport systems generally contain four components (Ames, 1986). One of the components, localized to the periplasmic space, is a polypeptide that binds to the substrate (Sbp, substratebindingprotein). Thisprotein has two globular domains that are separated by a flexible hinge. When the substrate binds to the globular domains there is a conformational change in the protein and bending ofthe hinge which results in trapping of the substrate molecule. The protein ligand complex then interacts with two hydrophobic proteins (CysT and CysW) that are thought to span the cytoplasmic membrane and form a pore. This interaction causes the release of the substrate which traverses the pore and is brought into the cell against a concentration gradient via the hydrolysis of ATP. Complete sequence characterization ofthe region of the cyanobacterial genome identified with the Salmonella typhimurium DNA probe revealed the presence of genes encoding all of the components of a periplasmic transport system (cysA, cysT, cysWand sbpA). To prove that the system was involved in the transport of sulfate, each of these genes was inactivated by interposon mutagenesis. The results of these experiments confirmed that the sulfate permease genes had been isolated, although some findings suggested that not all of the components of the system were absolutely required for the transport ofsulfate. Ifthe genes encoding either ofthe integral membrane components or the nucleotide binding protein were inactivated, the cells could no longer

transport sulfate and required an alternate S source for growth. In contrast, if the gene encoding the periplasmic sulfate binding protein (sbpA) was inactivated, the cells could still grow if sulfate were supplied as the sole S source, although they did not exhibit the dramatic increase in transcript accumulation from the permease genes exhibited by wild type cells deprived of S. These results suggested either that the sulfate binding protein was not absolutely required for growth on sulfate, or that a second gene encoding a sulfate binding protein was present in another location on the cyanobacterial genome. Southern hybridizations at low stringency were not successful in detecting another gene encoding the sulfate binding protein, supporting the former hypothesis. The situation in the sbpA mutant may be similar to that in E. coli. Although there have been extensive searches for E. coli mutants unable to grow when sulfate was supplied as the sole S source, none harbored lesions in the gene encoding the binding protein [the gene encoding the binding protein was fortuitously isolated (Hellinga and Evans, 1985)]. It is thought that the hydrophobic membrane components of the transport system do contain binding sites forthe substrate which are only exposed when these proteins interact with the substrate binding protein. However, they may also become exposed by mutagenesis, as has been suggested in studies of the maltose permease system (Shuman, 1982). In the case of the sulfate permease, the binding sites of the integral membrane proteins may be exposed, and therefore the binding protein is not absolutely required for sulfate transport (Laudenbach and Grossman, 1991). Alternatively, a second binding protein, encoded by a gene that doesn’t readily hybridize with sbpA, may substitute forsbpA in the permease system. The sulfate permease may also be utilized for the uptake of thiosulfate. Thiosulfate transport can be augmented by the binding of thiosulfate to a specific

Chapter 21 Cyanobacterial Acclimation Processes substrate binding protein designated CysP in E. coli (Hryniewicz et al., 1990). Based on sequence homology to the E. coli gene, the cysP gene in Synechococcus sp. strain PCC 7942 is immediately downstream of cysW. The cyanobacterial cysP gene is 5' to two genes (cysU and cys V) that have homology to cysT and cysW. These may be part of a transport system specific for the uptake of thiosulfate. Mutational analysis ofthis region is being performed to confirm the proposed gene functions. There does not appear to be a cysA-like gene associated with this system. However, preliminary results suggest that the CysA constituent of the sulfate permease may also function with this thiosulfate enhancer system. Another gene depicted on the map, located in the middle of the sulfate permease operon, has been designated cysR. The cysR gene product resembles DNA-binding transcriptional regulators such as those encoded by fixK and crp (Laudenbach and Grossman, 1991). When the cysR gene is interrupted, thiocyanate can no longer serve as a sole S source, although other S sources can support growth of the cysR- strain. Recent evidence suggests that the genes controlled by CysR and involved in the transport and utilization of thiocyanate are located on the large plasmid of Synechococcus sp. strain PCC 7942 (D. Laudenbach, personal communication). These results suggest that during S-limited growth of Synechococcus sp. strain PCC 7942, systems involved in the acquisition and utilization of diverse S-containing compounds are activated. A number of these compounds may be found in the natural habitats of this cyanobacterium. Finally, positioned downstream of the putative thiosulfate transport genes is a gene designated rhdA, which encodes a protein that exhibits some similarity to the enzyme rhodanese. Rhodanese is a thiosulfatesulfur transferase that can cleave a sulfane bond and transfer the thiol group to a thiophilic acceptor molecule (Laudenbach et al., 1991). This protein is located in the periplasmic space and becomes very abundant in S-deficient Synechococcus sp. strain PCC 7942. It is still uncertain whether this protein displays rhodanese activity or just functions in binding certain S-containing molecules. Hence, the protein may have a catalytic role in the acclimation process, or simply deliver specific S-containing compounds from the environment to cytoplasmic membranelocalized permeases. This latter function would also prevent leakage of specific S-containing molecules from the cell. Further studies on the function of the rhdA gene product should help elucidate the

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physiological processes that are important for optimizing survival ofthe cyanobacterium during Sdeficiency.

B. Intracellular Inclusions and Nutrient Reserves In addition to the biochemical changes described above, nutrient-deficient cyanobacteria exhibit alterations in the abundance and types of intracellular inclusion bodies, many of which serve as nutrient reserves (Allen, 1984; Simon, 1987). These changes are summarized in Table 3 (A, B, C, D, and E). The N-status of N-fixing cyanobacteria requires special considerations. Transfer of cyanobacteria capable of N-fixation from growth on a source of fixed-N to dinitrogen results in a transient N-deficiency which is relieved once N-fixation begins. In some cyanobacteria N-fixation occurs in specialized cells called heterocysts which protect the nitrogenase from inactivation by oxygen (see Chapter 27). Heterocystous N-fixing cyanobacteria (Anabaena sp., Calothrix parietina, and Mastigocladus laminosus in Table 3E) can be deprived of N by the removal of both fixed-N and dinitrogen; nonheterocystous Nfixing cyanobacteria (Plectonema boryanum inTable 3E) can be deprived of N by the removal of fixed-N and dinitrogen or by the removal of fixed-N in the presence of oxygen. Glycogen, which is a storage form of fixed-C, is generally present at low levels in exponentially growing cells. The cellular content of glycogen increases in Fe-, S-, and P-deficient cyanobacteria. Glycogen also accumulates in N-deficient cells (not in heterocysts), but does not accumulate in C-deficient cells. In general, the pattern ofglycogen accumulation matches its role as a fixed-C storage compound. Carboxysomes, present in all cyanobacteria during exponential growth, consist mainly of Rubisco and are the sites of fixation, although they have also been considered to be N- or C-storage granules. The carboxysome content of C-limited Synechococcus sp. generally increases, while that of Fe-deficient Synechococcus sp. decreases. In contrast, Fedeficiency in the marine cyanobacterium Synechococcus sp. strain PCC 7002 causes little alteration in carboxysome levels. Carboxysome levels remain relatively constant in S- or P-deficient Synechococcus sp. and in N-deficient, non-N fixing cyanobacteria. However, carboxysome content declines in N-fixing cyanobacteria that are N-deprived, and in heterocysts (which do not fix photosynthetically).

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Polyphosphate granules serve cyanobacteria (like many other organisms) as intracellular P-reserves. Polyphosphate is generally present in exponentially growing cells, and may accumulate to high levels in N- or S-deficient cells (except in heterocysts). Polyphosphate may be degraded in Fe-and C-deficient cells, as well as in P-deficient cells. Some cyanobacteria produce cyanophycin, a nonribosomally synthesized copolymer of aspartic acid and arginine (Allen, 1984). This compound serves as both a N- and C-reserve, and is present only at low levels in exponentially growing cells. Cyanophycin may accumulate transiently as N is exhausted from the medium (Mackerras et al., 1990a), but is eventually degraded in both C- and N-deprived cells. In contrast, cyanophycin accumulates in P- and Sdeficient cells, and may also reappear in the vegetative cells of heterocystous cyanobacteria once N-fixation has begun. PBS, the major light-harvesting complexes of the cell, are sometimes considered to be nutrient reserves, mainly for N. PBS are known to be degraded in Nand S-deprived cells, and may also be degraded under other conditions. There is evidence that PBS content can decline during N-deficiency in some cyanobacteria without affecting their growth rate (Boussiba and Richmond, 1980; Wyman et al., 1985). In general, the intracellular reserves for a specific nutrient decline when that nutrient is limiting growth, but increase when it is in excess, or when levels of another nutrient limit growth. Hence, both the types and numbers ofcellular inclusion bodies may provide a gauge for evaluating the nutrient status of the cell. The regulation ofstorage body accumulation has not been investigated thoroughly, but evidence suggests that enzyme activities for both synthesis and degradation are always present. For example, the activity of cyanophycin synthetase (polyphosphate synthetase) is detectable in extracts of N (P) deficient cyanobaeteria, even though the level ofcyanophycin (polyphosphate) is low (Grillo and Gibson, 1979, Mackerras et al., 1990a). The persistence of the activities involved in the synthesis of these storage bodies may explain the over-accumulation ofreserve material that occurs when N (P) is resupplied to N (P) deficient cells. This ‘overplus’ phenomenon results in an initial, massive accumulation of cyanophycin or polyphosphate granules, respectively, which are subsequently degraded as the cells return to nutrient-replete exponential growth. This massive accumulation exceeds immediate cellular demand

and occurs despite the presence of cyanophycinase (polyphosphatase) activity. It may allow cyanobacteria to take advantage of pulses of available nutrients that are common in patchy, nutrient-poor environments. There is some evidence that the transient accumulation of cyanophycin which precedes N-deprivation may be due to an increase in cyanophycin synthetase activity, although this does not necessarily occur during ‘overplus’ accumulation (Mackerras et al., 1990a). Many open questions, including the role of these reserve compounds in the normal, daily cycle of biosynthetic activities (Mackerras et al., 1990b), remain to be investigated.

C. Pigment Changes and Altered Photosynthetic Activity Cyanobacterial cells deficient for any of a number of different nutrients exhibit a decline in cellular pigmentation known as chlorosis, or bleaching (see Tables 3A, B, C, D, and E). In some cases chlorosis involves the degradation of pigments, while in other cases there is no active degradation, but the levels of the different pigments per cell (or by weight) decline because cell division (or the synthesis of other cell components, such as storage compounds) continues after pigment synthesis stops. Both types ofbleaching are observed, and the distinction may be important for understanding the modulation of photosynthetic activity and function ofthe photosynthetic apparatus during various stress conditions. The decline of pigmentation in batch cultures on a volume basis after the initiation of nutrient deficiency indicates that degradation of the pigments must be occurring; other situations may be more difficult to interpret. The cellular levels of both chlorophyll and phycobiliproteins decline during Fe-limitation (Table 3A). However, while the ratio of PC (the most frequently measured phycobiliprotein) to chlorophyll in Fe-limited Synechococcus sp. declines, it remains steady or increases in other cyanobacteria. Differences among experiments may reflect a diversity of responses to Fe-deficiency, or simply different experimental conditions and severities of deficiency. Specific changes of pigment-protein complexes may also be a consequence of Fe-limited growth. In Synechococcus sp. strain PCC 7942, Fe-limitation causes the appearance of a new chlorophyll-protein complex associated with a shift in the chlorophyll absorbance maximum (Reithman et al., 1988). Although the genes required for these changes have

Chapter 21 Cyanobacterial Acclimation Processes not been positively identified, a gene (isiA) with homology to psbC has been sequenced and is part of an operon upregulated under conditions of low Fe availability (Laudenbach and Straus, 1988). The accumulation of this new chlorophyll-protein complex is in part responsible for a number of other changes within the photosynthetic apparatus of Fedeficient cells, including changes in fluorescence emission spectra and an increase in the ratio of chlorophyll to PS I (Öquist, 1974b; Guikema and Sherman, 1983; Hardie et al., 1983a; Sandmann, 1985). The new pigment protein complex may represent a stored form of chlorophyll that can be rapidly converted into a photochemically active form once Fe deficiency is relieved. The ratios of PC to chlorophyll, of chlorophyll to carotenoid and of PS II to PS I all decline in Cdeficient cells (Table 3B), although the extent to which these changes occur depends on the precise experimental conditions. The altered pigment and photosystem ratios in C-limited cells may reflect the increased need for ATP generated via cyclic electron flow around PS I to drive active accumulation (Manodori and Melis, 1984; Pierce and Omata, 1988; Kaplan et al., 1991; Ogawa, 1992). C-limited Synechococcus sp. accumulates a 42 kDa ‘carotenoidbinding protein’, once thought to be involved in uptake. The abundance of this polypeptide also increases with higher levels of illumination (Reddy et al., 1989; Omata et al., 1990). Interestingly, this protein has a high degree of identity with a protein encoded by a gene that has been identified as part of the nitrate transporter (Omata et al., 1990). P-deficient cells also generally exhibit a decline in cellular PC and chlorophyll contents, along with declines in PC to chlorophyll and chlorophyll to carotenoid ratios (Table 3C). In some cases there is also a decline in total culture PC and chlorophyll content. In N- or S-deprived cultures, PC contents can decrease to virtually undetectable levels, while chlorophyll contents are often much less affected; the resulting decline in PC to chlorophyll ratio can be dramatic, however, (see Table 3D, E). Similar changes are observed in non-N-fixing strains depleted of N, and in N-fixing strains during the transient Ndeficiency that precedes the development of efficient N-fixation. Oxygen evolution and PS II activities often decline in parallel with the loss of PC, so that nutrient-deprived cells with little PC also have little detectable PS II activity. Elimination of PS II function during nutrient-deprivation may be advantageous

665 since, in the absence of anabolic processes, the formation of excited chlorophyll molecules that are unable to transfer energy to the reaction centers might generate toxic oxygen species. In this regard, it is interesting to note that both the chlorophyll and carotene levels in N-, S-, or P-deprived cells decrease relative to the levels of xanthophylls, and in particular relative to zeaxanthin (Gombos and Vigh, 1986; Fresnedo et al., 1991; Grossman et al., 1992). Maintenance of high zeaxanthin levels may be beneficial to nutrient-limited cells since zeaxanthin can quench oxygen radicals generated by the photochemically nonproductive excitation of chlorophyll molecules (Demmig-Adams, 1990). The photosynthetic apparatus remaining in heterocysts may be similar to that found in S-deprived, and in N-deprived, non-N-fixing, cyanobacteria. Heterocysts do not have functional PSII generated by PS II would inactivate nitrogenase), but they maintain an active PS I for the production ofATP via cyclic electron flow. The phycobiliprotein content of heterocysts is low but variable. The phycobiliproteins that remain after the differentiation of a heterocyst, consisting almost entirely of PC, are not assembled into PBS but can still transfer harvested light energy to PS I (Peterson et al., 1981; Tyagi et al., 1981; Yamanaka and Glazer, 1983). The generation of ATP by cyclic electron flow in heterocysts would help support the energetically expensive process of Nfixation. Similarly, cells deprived of S for 24 hours maintain PS I activity and the production ofATP (S. Herbert, J. Collier, D. Fork, and A. Grossman, unpublished results) via cyclic photophosphorylation. The continued production of ATP in this manner would provide energy to drive metabolic processes that remain essential in nutrient deficient cells. This energy would also be available for transporting the limiting nutrient into the cell once it does become available. The levels of mRNAs encoding the phycobiliproteins have also been reported to decline during nutrient-deficient growth, de Lorimier and Bryant (de Lorimier et al., 1984) suggested that the level of cpcBA mRNA decreases in N-starved Synechococcus sp. strain PCC 7002. Additional studies have shown that 3-5 h after the initiation of N-deprivation, transcripts from the cpcBACDEF operon are essentially undetectable (Bryant, 1991; Zhou et al., 1992). Data obtained by Gasparich et al. (1987), using a cpcB-lacZ fusion suggests that N-depletion results in a marked decrease in transcription of the

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cpcBA operon. In Synechococcus sp. strain PCC 7942 the levels of mRNA encoding both PC and AP decline rapidly during either N- or S-deprivation and less rapidly (and not to as low a level) during Pdeprivation. However, all of these mRNA species remain at 5-10% ofthe level found in nutrient replete cells (Collier and Grossman, 1992), even 48 h after cells are transferred to medium lacking N or S. The change in the steady state levels of phycobiliprotein mRNAs may be a consequence of both altered rates of transcription and mRNA turnover. The low levels ofPC and AP mRNA that remain in nutrient-deprived cells may not be translated (T. Allen, J. Collier and A. Grossman, unpublished results), suggesting that in addition to transcriptional regulation, phycobiliprotein synthesis in nutrient deprived cells is controlled at the post-transcriptional level. When the N-fixing cyanobacterium Anabaena variabilis is deprived of fixed-N and begins to differentiate heterocysts, the initial bleaching response includes a decrease in the level of mRNA encoding PC and AP (Johnson et al., 1988, Wealand et al., 1989). This decrease is temporary, and normal levels of both cpcBA and apcAB mRNAs are reestablished in vegetative cells (Belknap and Haselkorn, 1987), but not in heterocysts, after N-fixation begins.

D. Phycobilisome Degradation The loss of PC in N-deprived Synechococcus sp. strain PCC 6301, as determined spectrophotometrically, correlates with a loss of PC apoprotein, as determined immunologically (Lau et al., 1977). Furthermore, the synthesis of new PC is depressed during growth in N-deficient medium and is restored upon addition of N to the medium. The net loss of PC that occurs in N- and S-deprived cells is due to a rapid and nearly complete degradation of the PBS (Yamanaka and Glazer, 1980; Collier and Grossman, 1992). It should be emphasized that a decline in PC (or PBS) content per cell or per gram of tissue does not necessarily mean that the PBS has been degraded. For example, P-depleted Synechococcus sp. cells have a low PBS content, but very little, if any, PBS degradation occurs (Collier and Grossman, 1992). As discussed above, degradation of the PBS (as well as other cellular proteins) during N-deficiency could provide amino acids for the synthesis of proteins or other cellular constituents important for the acclimation process. The use of phycobiliproteins as amino acid storage molecules may be especially

important for marine cyanobacteria (Wyman et al., 1985; but see Yeh et al., 1986), since nitrogen may frequently be limiting in marine environments. PBS are a poor source of S-containing amino acids, but are nevertheless degraded in S-deprived cells. In fact, Calothrix sp. strain PCC 7601 has a cpc operon encoding proteins that are devoid of all but the essential S-containing amino acids and that are only expressed when the cells are maintained on low-S medium (Mazel and Marlière, 1989). PBS destruction during nutrient-deficiency may also help protect the cells from phototoxicity (discussed above), although other mechanisms that dramatically reduce the transfer of energy from the PBS to chlorophyll also exist (see below). The degradation of the Synechococcus sp. PBS during N- and S-deprivation is virtually identical and is an ordered process (Fig. 4). Degradation begins at the periphery of the complex with the elimination of the terminal PC hexamer of the rod substructure and its associated 30 kDa linker polypeptide (Yamanaka and Glazer, 1980; Collier and Grossman, 1992). This is followed by degradation of the next PC hexamer and its associated 33 kDa linker polypeptide and then the loss of some entire rods. The degradation of PBS polypeptides results in a decrease in the PBS size (sedimentation coefficient) and a reduction in the ratio of PC to AP. A potentially similar ‘trimming’ of PBS in N-deprived Synechocystis sp. strain PCC 6308 has also been observed (Duke et al., 1989). The smaller PBS still functions in harvesting light energy. Continued nutrient deprivation results in the complete degradation of the remaining PBS structure. Hence, cellular phycobiliprotein content is controlled at the level of both PBS size and number. Upon adding the limiting nutrient back to deprived cultures, new PBS are rapidly synthesized to normal levels before growth resumes. The existence of a protease involved in PBS degradation, that is synthesized de novo in response to N-deprivation, has been suggested in a number of studies withAnabaena, Spirulina, and Synechococcus species (Boussiba and Richmond, 1980, Elmorjani and Herdman, 1987, Foulds and Carr, 1977, Wood and Haselkorn, 1979, Wood and Haselkorn, 1980, YamanakaandGlazer, 1980).Toexaminethisprocess from a different perspective, Synechococcus sp. strain PCC 7942 was mutated and screened for organisms unable to degrade their PBS (Collier and Grossman, 1994). Such colonies appeared blue-green when allowed to grow on agar substantially free of S, while

Chapter 21 Cyanobacterial Acclimation Processes

wild-type colonies bleached. Some mutants obtained did not degrade their PBS during either S- or Ndeprivation, although no new PBS were synthesized during the stress treatment. Surprisingly, these cells grew at a similar rate and exhibited a similar susceptibility to high light during nutrient deficiency as wild-type cells. Hence, even though the cells had a considerable number of PBS when N- or S-deprived, they appeared to be no more photosensitive than normally acclimating cells. Additional experiments suggested that the PBS present in the mutant organisms deprived of N or S were not able to efficiently transfer harvested light energy to the reaction center of PS II; this uncoupling of the light harvesting complex from the primary photochemical reactions of PS II may explain the lack of photosensitivity in these strains and be an important photosynthetic control mechanism in wild-type cells. To define the lesion responsible for the ‘nonbleaching’ phenotype, mutant strains were complemented back to the bleaching, wild-type phenotype and the DNA responsible for complementation was characterized. A 2.0-kbp fragment of genomic DNA was shown to be effective in complementation (Collier and Grossman, 1994). Three genes, designated orf134, nblA and txlA were located on this fragment. orf134 is constitutively expressed and does not appear to be involved in the bleaching process. The txlA gene product resembles both a thioredoxin and a protein disulfide isomerase. The role of txlA in the response of cells to N- and S-limitations is still uncertain, although kinetic experiments and insertional inactivation of txlA suggests that it is required for normal photosynthetic efficiency. The nblA gene contains an open reading frame of

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59 amino acids with an initiator methionine preceded by a ribosome binding site. A small transcript covering this gene accumulates at high levels in cells deprived of N or S. Larger transcripts initiated at the beginning of nblA can extend through txlA (which is encoded on the opposite strand). Very low levels of these transcripts can be detected in cells maintained in complete medium, and low levels are present in Pdeprived cells. Insertional inactivation of nblA results in a nonbleaching phenotype, while inactivation of orf l 34 and txlA does not prevent bleaching during nutrient deficiency. The strain initially isolated as a nonbleaching mutant contained a single base change in nblA that altered a serine codon to a phenylalanine codon (Collier and Grossman, 1994). When the nblA gene is placed on a multicopy plasmid and transformed into Synechococcus sp. strain PCC 7942, the cells bleach to some extent even in nutrientreplete medium and break down their PBS much more quickly than wild type cells during nutrient deprivation. The cells also degrade the PBS during P-deprivation if nblA is placed under the control of the derepressible phoA promoter (Collier and Grossman, 1994). Hence, any condition that favors increased expression ofnblA triggers PBS degradtion. No homology was observed between the protein encoded by nblA and any sequence in the GenBank databases. A number of interesting ideas can be gleaned from these studies. Firstly, since multiple copies of nblA can induce bleaching to some extent when cells are maintained in complete medium, and since P-deprivation can cause PBS breakdown in a strain in which nblA is placed under the control ofthe alkaline phosphatase promoter, it is likely to be the only gene whose activity needs to be directly increased

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(NblA itself may be involved in the activation of other genes) during S- or N-deprivation to provoke bleaching. Secondly, the small size of this protein and the lack of similarity to any known protease suggest that it is not a protease itself. There are a number of possible functions that this polypeptide may have that would explain why it is absolutely required for bleaching. 1) NblA may function to activate a protease, such as the one studied by Wood and Haselkorn (1980), that degrades the PBS. 2) NblA may trigger PBS degradation by interacting with the constituents of the complex and altering their susceptibility to proteolysis. Such interactions may involve covalent attachment, similar to the binding ofubiquitin to proteins in eucaryotes, which marks them for degradation (Hershko, 1988). Alternatively, this small peptide may disrupt hydrophobic and/or ionic interactions among various constituents of the PBS, rendering them susceptible to degradation. 3) Finally, NblA may be involved in activating other genes that are directly responsible for causing PBS degradation. Three other cyanobacteria that have been tested contain sequences that hybridize to both nblA and txlA at low stringency. The similar pattern of PBS degradation observed in Synechocystis sp. strain PCC 6308 and Synechococcus sp. strain PCC 7942 suggests that nblA homologues are responsible for PBS degradation in other cyanobacteria. PBS degradation during the development of a heterocyst may also use a similar mechanism. In any case, defining the events necessary for PBS degradation may be important for a general understanding of turnover of macromolecular complexes in procaryotic organisms.

IV. Concluding Remarks The size, structure and number of PBS in cyanobacteria are exquisitely sensitive to growth conditions. The responses of certain cyanobacteria to light quality may provide a broader understanding of photoperception in photosynthetic microbes, and may also yield information on proteins ancestral to the ubiquitous photoreceptor in higher plants, phytochrome. Studies concerning environmentallyregulated degradation ofthe PBS have already begun to provide insights into processes involved in the targeting of macromolecular complexes for degradation and the machinery that implements this

degradation. They may also lead to a greater understanding ofthe overall control ofenergy transfer within the photosynthetic apparatus. We have tried to raise many questions concerning the ways in which photosynthetic microbes sense and respond to their environment and view it as a very fertile field for future studies.

Acknowledgments The authors are grateful to Jean Houmard for sharing unpublished data and Kirk Apt, Elena Casey, and John Davies for helpful discussions. Jane Edwards was invaluable in typing and formatting of the manuscript and Glenn Ford was always available to aid us with computer assisted analysis of the phycobilisome polypeptides and genes. We are also very grateful to The National Institutes of Health, The National Science Foundation and The Carnegie Institution of Washington, agencies that have supported our work for the last ten years. This is a CIW publication No. 1162.

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Chapter 21 Cyanobacterial Acclimation Processes plastocyanin from the cyanobacterium Anabaena sp. PCC 7937: Isolation, sequence and regulation. Mol Microbiol 3: 275–284 Vasil’eva VE and Levitin MG (1974) Effect of carbon dioxide starvation on certain blue-green algae. Fiziologiya Rastenii 21: 1207–1211 Vega-Palas MA, Madueño F, Herrero A and Flores E (1990) Identification and cloning of a regulatory gene for nitrogen assimilation in the cyanobacterium Synechococcus sp. Strain PCC 7942. J Bacteriol 172: 643–647 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 Vogelman TC and Scheibe J (1978) Action spectrum for chromatic adaptation in the blue-green alga Fremyella diplosiphon. Planta 143:233–239 Wagner SJ, Thomas SP, Kaufman RI, Nixon BT and Stevens SE Jr (1993) The glnA gene of the cyanobacterium Agmenellum quadruplicatum PR-6 is nonessential for ammonium assimilation. J Bacteriol 175: 604–612 Wanner G, Henkelmann G, Schmidt A and Köst H-P (1986) Nitrogen and sulfur starvation of the cyanobacterium Synechococcus 6301. An ultrastructural, morphometrical, and biochemical comparison. Z Naturforsch 41 c: 741–750 Wealand JL, Myers J A and Hirschberg R (1989) Changes in gene expression during nitrogen starvation in Anabaena variabilis ATCC 29413. J Bacteriol 171: 1309–1313 Whitton BA, Potts M, Simon JW and Grainger SLJ (1990) Phosphatase activity of the blue-green alga (cyanobacterium) Nostoc commune UTEX 584. Phycologia 29: 139–145 Willey JM and Waterbury JB (1989) Chemotaxis toward nitrogenous compounds by swimming strains of marine Synechococcus spp. Appl Environ Microbiol 55: 1888–1894 Wood NB and Haselkorn R (1979) Proteinase activity during

675 heterocyst differentiation in nitrogen-fixing cyanobacteria. In: Cohen GN and Holzer H (ed) Limited Proteolysis in Microorganisms, pp 159–166. US DHEW Publication No. (NIH) 79-1591, Bethesda, MD Wood NB and Haselkorn R (1980) Control of phycobiliprotein proteolysis and heterocyst differentiation in Anabaena. J Bacteriol 141: 1375–1385 Wood P, Peat A and Whitton BA (1986) Influence of phosphorus status on fine structure of the cyanobacterium (blue-green alga) Calothrix parietina. Cytobios 47: 89–99 Wyman M, Gregory RPF and Carr NG (1985) Novel role for phycoerythrin in a marine cyanobacterium, Synechococcus strain DC2. Science 230: 818–820 Xie W-Q, Whitton BA, Simon JW, Jäger K., Reed D and Potts M (1989) Nostoc commune UTEX 584 gene expressing indole phosphate hydrolase activity in Escherichia coli. J Bacteriol 171:708–713 Yamanaka G and Glazer AN (1980) Dynamic aspects of phycobilisome structure. Phycobilisome turnover during nitrogen starvation in Synechococcus sp. Arch Microbiol 124: 39–47 Yamanaka G and Glazer AN (1983) Phycobiliproteins in Anabaena 7119 heterocysts. In: Papageorgiou GC and Packer L (ed) Photosynthetic Prokaryotes: Cell Differentiation and Function, pp 69–90. Elsevier Science Publishers, Amsterdam Yeh SW, Ong LJ and Glazer AN (1986) Role of phycoerythrin in marine picoplankton Synechococcus spp. Science 234: 1422– 1423 Zhang L, McSpadden B, Pakrasi HB and Whitmarsh J (1992) Copper-mediated regulation of cytochrome and plastocyanin in the cyanobacterium Synechocystis 6803. J Biol Chem 267: 19054–19059 Zuber H (1986) Structure of light harvesting antenna complexes of photosynthetic bacteria, cyanobacteria and red algae. Trends Biochem Sci 11:414–419

Chapter 22 Short-term and Long-term Adaptation of the Photosynthetic Apparatus: Homeostatic Properties of Thylakoids Yoshihiko Fujita, Akio Murakami, Katsunori Aizawa Department of Cell Biology, National Institute for Basic Biology, Okazaki, Aichi 444, Japan

Kaori Ohki School of Marine Sciences and Technology, Tokai University, Shimizu, Shizuoka 424, Japan Summary I. Introduction II. Short-term Adaptation: The State Transition A. Historical Background B. Possible Mechanism in PBS-Chl a System III. Long-Term Adaptation: Regulation of PS I:PS II Stoichiometry A. Historical Background B. PS I:PS II Change C. Possible Mechanism IV. Relationship Between Short-Term and Long-Term Adaptation Acknowledgments

677 678 679 679 681 683 683 684 686 689 690

Summary Light-energy conversion in thylakoids is accomplished by cooperative interactions between two photoreactions. The balance between these two photoreactions determines the efficiency of energy conversion. The efficiency ofenergy conversion is maintained at a high level by at least two regulatory mechanisms: short-term adaptation and long-term adaptation. Short-term adaptation, i.e. the state transition, is a regulatory mechanism that controls the distribution of excitation energy transfer from the light-harvesting antenna complexes, the phycobilisomes (PBS), to the two photosystems. The transfer of energy trapped by PBS to the Chl a of Photosystem I (PS I) or Photosystem II (PS II) is regulated either at the transfer point from the PBS to the two photosystems or at a transfer point between the Chl a of PS II and PS I. Energy transfer from the PBS to PS I increases, and to PS II decreases, when most of PS II centers are closed, and the opposite occurs upon a shift to conditions under which most PS I centers are closed. This regulation occurs in response to the state ofbalance between the two photoreactions through monitoring of the redox status of electron transport between the two photosystems or through monitoring the electrochemical potential ofthe membrane around the two photosystem complexes. This regulatory process is called ‘short-term adaptation’ because this regulation can occur rather rapidly (usually completed within several minutes or less). ‘Long-term adaptation’ refers to a regulated change in the Stoichiometry of PS I and PS II in the thylakoids. The PS I:PS II ratio becomes greater—values of 2.0 to 3.0 are typical—when cells are grown under conditions where most PS II centers are closed (e.g., green-rich light). The ratio becomes small, approximately 1.0 , upon a shift to conditions where most of PS I centers are closed (e.g., growth in red-rich light). The PS I:PS II ratio is also regulated in response to the redox state of electron transport between two photosystems. Regulation of PS I synthesis appears to be the general pattern in D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 677–692. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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cyanobacteria. Synthesis of PS I complexes apparently is controlled at the assembly level but not at the level of apoprotein synthesis. This regulation seems to occur by controlling Chl a synthesis or transport to the site for PS I assembly. The regulation of PS I synthesis is as rapid as state transition. However, the PS I:PS II ratio changes more slowly over a period of hours or days, and hence this process is referred to as ‘long-term adaptation.’ These short-term and long-term adaptation regulatory responses operate together to produce fineand coarse-tuning adjustments, respectively, and to balance the activities ofthe two photoreactions in response to changing photosynthetic environments. The phenomena associated with these responses as well as possible mechanisms for their regulation are discussed. I. Introduction Cyanobacteria obtain their energy for growth and development primarily from oxygenic photosynthesis. Although some strains are able to support growth in the dark by heterotrophic respiration (see Chapter 13), most cyanobacteria are obligate photoautotrophs. Cyanobacteria have had a very long life-history on Earth. The oldest fossils suggest their occurrence on the Earth about 3.5 Gyr ago (Schopf and Walter, 1982), and the development of oxygenic photosynthesis by cyanobacteria is generally believed to have caused the oxygenation of the Earth’s atmosphere. The secret to their long lifehistory may be attributed to their growth mode. Oxygenic photosynthesis is ideally suited to the Earth’s environment. This type of photosynthesis requires light as the energy source, as theelectron donor for electrochemical reactions, and as the carbon source—all of which are abundantly present in most of the surface environments of the Earth. However, is a very stable molecule, and its oxidation requires a large amount of energy. During oxygenic photosynthesis, is photochemically oxidized into molecular and protons with the concomitant production of a weak reductant (plastoquinol) by PS II (see Chapter 8). However, fixation of requires a strong reductant, such as NADPH, and large amounts of ATP. PS I can reoxidize the weak reductant generated by PS II and produces a strong reductant, reduced ferredoxin, which can subsequently be used during noncyclic electron transport to reduce NADP+ or which can return Abbreviations: CCCP – carbonylcyanide m-chlorophenylhydrazone; Chlide a – ohlorophyllide a; Cyt – cytochrome; DBMIB – 2,5-dibromo-3-methyl-6-isopropyI-p-benzoquinone; ETS – electron transport system; HQNO – 2-n-heptyl-4hydroxyquinoline N-oxide; LHCII – light-harvesting chlorophyll a/b protein complex; PBP – phycobiliprotein; PBS – phycobilisome; PQ – plastoquinone; MV – methylviologen; Pchlide – protochlorophyllide.

electrons via the cytochrome complex to during cyclic electron transport to produce additional ATP for fixation or other reactions (see Chapters 9 and 10). High-efficiency light energy conversion during oxygenic photosynthesis thus requires a careful balancing ofthe reactions catalyzed by these two photosystems. In natural environments the light regime for photosynthesis of cyanobacteria is not necessarily optimal for balancing the two photochemical reactions. A typical example ofa biased environment is the light regime in the ocean. In the open ocean only light in the blue-green wavelength region (maximal wavelength around 480 nm) is available for photosynthesis, while in the coastal seas only wavelengths in the green region ofthe spectrum (560 nm) are available. These occur because ofthe removal of light at shorter wavelengths by dissolved and suspended organic materials and at longer wavelengths by absorption by molecules themselves (Jerlov, 1976). Another example is the light environment within a deep forest; a thick layer of tree leaves becomes an optical filter that decreases the quantity and modifies the wavelengths (quality) of light that penetrate to the Earth’s surface. In cyanobacteria the main light-harvesting antennae for photosynthesis are the PBS, which are primarily composed of phycobiliproteins (PBP) and exist on the surface of thylakoids as supramolecular structures (see Chapter 7). The major pigments in the PBS are phycoerythrin (absorption maximum ~570 nm) or phycocyanin (absorption maximum ~630 nm), and these proteins provide the primary input of light energy for photosynthesis in these organisms. However, the PBS have been known to act preferentially as a light-harvesting antennae for PS II, although some of the energy trapped by the PBS is also transferred to PS I as will be explained later. Besides the PBS, Chl a molecules associated with each photosystem complex act as light-harvesting pigments for each photochemical reaction. The

Chapter 22 Short-Term and Long-Term Adaptation number of Chl a molecules in a PS I complex is greater than that in PS II complexes: 100–150 Chl a molecules are present in a PS I complex while 35–60 Chl a molecules, about a halfthe number found in PS I, are present in PS II (Myers et al., 1980; Fujita and Murakami, 1987). Thus, light of wavelengths absorbed by Chl a preferentially excites PS I. Moreover, since the Chl a molecules in PS I complexes are present in an environment producing an absorption band at longer wavelength than those molecules associated with PS II (see Chapters 8 and 10), light in the near far-red wavelength region (i.e., longer than 680 nm) preferentially excites PS I. These characteristics of the light-harvesting antennae in the cyanobacterial thylakoid system exaggerate the imbalance between the twophotochemical reactions. For autotrophic growth of these organisms, cells must obtain inorganic nutrients, such as nitrate, sulfate, and phosphate (see Chapter 21), in addition to a source ofcarbon or see Chapters 14 and 15) from their environment. When ionic nutrients are present at low concentration, such as in the open ocean, they must be actively concentrated within cells by a pumping system. Such pumping systems require extra energy beyond that minimally required for photosynthetic carbon fixation. For example, the energy dependencies for growth on two different carbon sources, and ion, are greatly different. Cells must also consume extra energy to provide protection from physicochemical environments such as those of high osmotic pressure. These energy requirements may be met by changing the proportion of non-cyclic versus cyclic electron transport in thylakoids, and these changes can lead to further imbalances in the two photochemical reactions. Environments for cell growth are not fixed but often fluctuate. Thus, the photosynthetic thylakoid system must respond to variations that cause an imbalance of the two photochemical reactions. Two types of adjustments to variations in photosynthetic environments have been recognized. The first mechanism provides an adjustment of energy distribution from the PBS to the two photosystem complexes and is called ‘state transition.’ Since this process occurs on the time scale of seconds to minutes, it is referred to as ‘short-term adaptation.’ The second mechanism adjusts the stoichiometry between PS I and PS II. Although this regulatory response also occurs quickly, the full expression of the response depends upon the synthesis of new proteins and

679 thylakoids. This slower change, with a time scale of hours or days, is referred to as long-term adaptation (see Anderson, 1986). This chapter provides a short review of the study of these regulatory processes in cyanobacteria and will attempt to explain their mechanisms. Since there are many publications concerning short-term adaptation, only some salient points will be covered here. The reader should consult other more detailed reviews for additional references (see Williams and Allen, 1987; Biggins and Bruce, 1989; Bennett, 1991; Allen, 1992).

II. Short-term Adaptation: The State Transition

A. Historical Background In cyanobacteria, the action spectrum for the quantum yield of evolution shows that the maximum yield is always found in the wavelength region where PBP absorption occurs. Quantum yield values in this wavelength range may be as high as the theoretical one derived from the model of two light reactions (Emerson and Lewis, 1942; Myers, 1963). This indicates that the light energy absorbed by PBP is transferred not only to PS II but is also transferred, either directly or indirectly, to PS I. A constantly high quantum yield also suggests that such transfer is regulated so as to balance the excitations arriving at the two photosystems. Murata (1969) first noted the occurrence of this type of regulation during studies with the red alga Porphyridium cruentum. He found that the fluorescence intensity emitted in vivo from the Chl a of PS I and PS II at 77 K is not constant but depended upon the conditions under which the cells had been kept prior to freezing. When cells had been preilluminated for a sufficient time period with light absorbed by PBP, the emission from PS II Chl a, as well as that from PBS, became less intense while the emission from PS I Chl a increased. This suggested that energy transfer from PBS to PS II decreased and to PS I increased, and cells exhibiting such energy distribution are said to be in state 2. Preillumination ofcells with light absorbed by Chl a caused a reversed effect leading to an increase in the emission from PS II Chl a and a decrease in the emission from the Chl a of PS I. Cells exhibiting this pattern of emission are said to be in state 1; dark incubation of cells led to a state close to state 2. These results indicate that the distribution of light-energy absorbed by PBS to

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the two photosystems is regulated in response to the light regime. Preferential excitation of PS II causes an increase in the transfer to PS I and decrease in that to PS II, and preferential PS I excitation causes the opposite effects. This regulation of energy distribution between two photosystems has been called the ‘state transition.’ Observations of this regulatory phenomenon are usually made at liquid-nitrogen temperature, since at this temperature protein complexes cannot move within the thylakoids. Murata (1969) interpreted his observations as follows: (1) PBS-absorbed light energy is first transferred to PS II and then from PS II to PS I by spillover (see Myers, 1963); (2) regulation occurs in the process of spillover from PS II to PS I complexes. Murata (1970) also examined this phenomenon with the red alga Porphyra yezoensis and confirmed the occurrence of this regulation by determining the emission from PS II Chl a at room temperature. At about the same time, Bonaventura and Myers (1969) found the same type of regulation in the green alga Chlorella pyrenoidosa. They determined photosystem activity not only by fluorescence emission but also by evolution measurements. A correlation between the shift to state 2 and the phosphorylation state of LHC II was found in the chloroplasts of vascular plants (see Barber, 1983; Bennett, 1991; Allen, 1992). Dephosphorylation of LHC II causes an increase in the transfer ofenergy to PS II. These findings have been interpreted as follows: (1) energy transfer from LHC II to the two photosystems is regulated by phosphorylation of LHC II; (2) non-phosphorylated LHC II transfers energy exclusively to PS II, but LHC II can transfer energy to PS I once phosphorylated; (3) phosphorylation of LHC II causes the electrochemical nature of the protein surface to become more negatively charged; and (4) phosphorylation of LHC II causes repulsive forces which lead to destacking of granal membranes, where most PS II centers are located; LHC II moves laterally to the non-stacked stromal thylakoid membranes, where most PS I centers are found, and transfer energy to PS I. Indeed, the time course of phosphorylation correlates well with the transition to state 2. The protein kinase for LHC II phosphorylation is one of the kinases located in thylakoid membranes, and its activity depends upon redox potential. Redox titration of the activity indicates that the midpoint potential is very close to that of with n = 2 in Nernst Equation (Horton et al., 1981).

Correlation of redox characteristics between PQ and activation of the kinase suggests that the redox state of the PQ pool is monitored for regulation of kinase activity. Among kinases in thylakoids, a species with a molecular mass of 64–65 kDa may be the enzyme for LHC II phosphorylation (Coughlan and Hind, 1986; Gal et al., 1990). The enzyme can phosphorylate LHC II, and it is activated by when it forms aggregates with Cyt complex, suggesting that the activity is redox-dependent. Although the redox state of PQ is the most probable target to be monitored for regulation of LHC II phosphorylation, the idea that the state of the Cyt complex is also associated with regulation of phosphorylation has also been proposed by experiments with the inhibitors of Cyt oxidation (Gal et al., 1988; see also Bennett, 1991; Allen, 1992) and mutants lacking the Cyt complex (Lamaire et al., 1987; Gal et al., 1987; Bennett et al., 1988; Coughlan, 1988). These studies suggest that Cyt complex reduction is required for activation of protein phosphorylation. Dephosphorylation seems to occur by a constitutive phosphatase independent ofredox-regulation. Based on the findings described above, the model shown schematically in Fig. 1 is currently accepted for regulation in plants and green algae. When PS II is preferentially excited and turns over faster than PS I, PQ and/or the Cyt complex become more reduced and activate the kinase for LHC II phosphorylation. Phosphorylated LHC II moves laterally to unstacked membranes and acts as light-harvesting antenna for PS I. Energy transfer to PS II decreases while transfer to PS I increases. Preferential excitation of PS I causes an opposite set ofchanges. The kinase is inactivated due to low level of reduced PQ and Cyt complex, and phosphorylated LHC II is dephosphorylated by the constitutive phosphatase. This causes a return movement of LHC II to the stacked granal membranes. In summary, the state transition in the chloroplasts of vascular plants and green algae seems to occur as a regulation ofthe energy transfer at the step from the light-harvesting antenna to the two photosystems but not between the two photosystems. However, molecular details ofthe mechanism, such as how the protein kinase is activated and deactivated, remain unknown. An important element of the proposed mechanism is the lateral heterogeneity of the two photosystems in chloroplast thylakoids: PS II complexes occur mostly in the grana stacks while PS I complexes are found in the non-stacked stromal

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thylakoids. The PBS-covered thylakoids of red algae and cyanobacteria are not stacked, however, and thus some difference or modification may occur in the mechanism for state transition in these organisms.

B. Possible Mechanism in PBS-Chl a System The mechanism for the state transition in PBS-Chl a systems is presently controversial. However, the phenomena in cyanobacteria and red algae can be summarized as follows. (1) Light-energy absorbed by PBP is preferentially transferred to PS II, and energy absorbed by Chl a is mainly transferred to PS I. (2) Energy transfer from PBP to PS I-associated Chl a occurs and increases under the light conditions for PBS-absorption (state 2); however, the increase in PS I excitation is not so great as the decrease in PS II excitation. (3) Such energy transfer to PS I is insignificant or zero when cells are adapted to conditions where most light is absorbed by Chl a (state 1). (4) The functioning of the light-harvesting pigment system of cells grown in the dark is similar to that found in cells illuminated with PBS-absorbed light. Thus, these phenomena suggest that regulation of energy transfer occurs at the level of energy transfer from PBS to the two photosystems or between the two photosystem complexes. Since the efficiency of energy transfer depends upon the distance and orientation between pigment molecules, geometric changes must occur among the PBS, PS I and PS II protein complexes. Figure 2 schematically shows three possible models for the state transition in PBS-containing organisms.

In model I modification ofthe structure occurs at the site for the connection of the PBS with photosystem complexes in thylakoid membranes; the core part of PBS including anchor-polypeptide (ApcE) and allophycocyanin B (ApcD) are primary candidates (see Chapter 7 and below). According to this model, a modification occurring when cells are illuminated with PBS-absorbed light must cause a repulsion, probably electrostatic in nature, between the PBS and PS II complexes but allows a higher affinity between the PBS and PS I complexes. The opposite changes would occur under Chl a-absorbed light. This is the same in principle as the widely accepted mechanism for the state transition in green plants. If the regulation occurs at the level of photosystem complexes, as Murata (1969) first assumed (model II), the modification must occur in the structure(s) of the chlorophyll-protein complexes themselves. The surface structure of either the PS I or PS II complex, or both, would be modified to become repulsive or attractive to the counterpart complex. In these two models, compensatory changes in energy transfer to the respective photosystems should occur; however, this is inconsistent with the observation that changes in energy transfer to PS II

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are always greater than those in the transfer to PS I (Salehian and Bruce, 1992). This discrepancy can be explained by the detachment model (model III). According to this model, some PBS become detached from PS II without association with PS I during illumination with PBS-absorbed light (shiftto state 2). None of models or combination of models can adequately explain the experimental observations so far reported. Models I and II can be distinguished by differences in changes in effective absorption crosssections of the photosystems in cells exposed to either PBS- or Chl a-absorbed light. For model I, the effective absorption cross-section of Chl a in the two photosystem complexes should not be altered, but variation should only occur in the cross-section due to the PBS. A recent determination with P700 photooxidation as the index for the PS I reaction was consistent with this prediction (Mullineaux, 1992). However, fluorescence from PS I after emission at 77 K was found to occur upon excitation of either PBS or Chl a, indicating the occurrence ofchanges in the effective cross-section of Chl a associated with the state transition for Porphyridium cruentum (Murata, 1969, Ley and Butler, 1980) and for Synechococcus sp. strain PCC 6301 (Salehian and Bruce, 1992). Further, changes in effective cross-section were also observed with a PBS-less mutant of Synechococcus sp. strain PCC 7002 (Bruce et al., 1989). These findings favor spillover regulation, model II. Allen and his colleagues (Allen et al., 1985; Sanders and Allen, 1987; Sanders et al., 1989) found with Synechococcus sp. strain PCC 6301 that the lightinduced shift to state 2 causes phosphorylation of 15- and 18.5-kDa proteins in vivo and in vitro. Dephosphorylation occurred under the light-induced shift to state 1 (Sanders et al., 1989). The 18.5-kDa protein was identified as a PBS-associated component (Sanders and Allen 1987; Sanders et al., 1989). This phosphorylation was postulated to cause mobilization of PBS on the thylakoid surface like LHC II phosphorylation in the system ofgreen plants. Based on this protein phosphorylation, Allen and Holmes (1986) proposed a model for the state transition in which PBS-absorbed light causes phosphorylation of a PBS component; the phosphorylated PBS moves laterally on thylakoid surface to PS I, and the energytransfer pattern becomes that typical of state 2. However, Biggins et al. (1984a) and Kirschner and Senger (1986) failed to observe light-induced protein phosphorylation with Porphyridium cruentum. They observed phosphorylation of many proteins, but all

protein phosphorylations were light-independent. Furthermore, the conditions for light-induced phosphorylation described by Allen et al., (1985) were reported to be inadequate for inducing the state transition (Biggins and Bruce, 1989). Katoh and Gantt (1979) have reported the isolation of vesicles in which PS II is associated with PBS; in such vesicles, the PBS transfers energy to PS II complexes as efficiently as the PBS do in vivo. However, the isolation of complexes between PS I and PBS have not been reported. Further examination of this problem is required. Besides fluorescence at 77 K and reactions specific to the respective photosystems, photoacoustic measurements have recently been adopted for analysis of the state transition (Canaani, 1986; Malkin et al., 1990; Bruce and Salehian, 1992). Photoacoustic determination ofenergy storage using a 2-msec pulse revealed that energy storage during the msec timescale remains unaltered under either state 1 or 2 (Bruce and Salehian, 1992) and indicating that the detachment model (model III) seems not to occur during the state transition. Bruce and Salehian (1992) stated in their recent paper that ‘none of the three proposed mechanisms for the state transition in cyanobacteria (mobile antenna, spillover and PBS detachment) can fully explain the data’. Details of research on the state transition can be found in the reviews written by Biggins and Bruce (1989) (from the school in favor of spillover model) and Williams and Allen (1987) and Allen (1992) (from the school in favor of mobile antenna model). Much of the controversy surrounding this subject seems to arise from the quality of the indices adopted for the determination of the state transition, and a more quantitatively accurate index is probably required. Determination of the rise and decay kinetics of fluorescence may be a property that satisfies such a demand (Mullineaux and Holzwarth, 1991; Mullineaux et al., 1990). As explained later in this chapter, the photosystem stoichiometry in cells is variable, and a consideration of such changes in photosystem ratio in cells is necessary in order to interpret the observed properties ofcells grown under different conditions. The state transition in green plants probably occurs in response to the redox state of PQ and/or the Cyt complex. Is this also true for organisms that contain PBS and Chl a? Satoh and Fork (1983) and Biggins et al. (1984b) suggested that cyclic electron flow is related to the state transition in red algae and

Chapter 22 Short-Term and Long-Term Adaptation cyanobacteria. In these organisms, the addition of CCCP as well as DBMIB inhibited the shift to state 1 and caused a shift to state 2 in cells already in state 1. They assumed that an electrochemical gradient around the photosystem complexes is induced by cyclic electronflowand causes changes in the affinity of the PS complexes that leads to the state transition. On the other hand, Mullineaux and Allen (1990) proposed that the state transition occurs in response to the redox status of PQ and/or Cyt complex as in green plants. Their prediction was obtained from the effects of added reagents that affect the intersystem electron transport chain. In the former studies (Satoh and Fork, 1983; Biggins et al., 1984b), DBMIB and MV were found to suppress the state transition from state 2 to state 1 as well as cyclic electron flow,and in the latter studies (Mullineaux and Allen, 1990), the effects of various conditions for reducing and oxidizing the intermediate electron transport components, such as PQ, between two photosystems were examined. The photosynthetic and respiratory electron transport chains of cyanobacteria are still not completely understood, however (see Chapters 9, 10, and 13). Additional studies are required to identify the signal(s) responsible for the state transition in cyanobacteria. Molecular genetics provides a promising approach to a more detailed understanding of the state transition phenomenon in cyanobacteria. A large number of well-defined mutations affecting components of PBS, PS I, PS II, and ETS components have been created in recent years (see Chapters 7–10, 12, and 13 for details), and some of these mutations affect the state transition. For example, a null mutation in the psaL gene of Synechococcus sp. strain PCC 7002 (encoding a subunit of PS I; see Chapter 10), causes the transition from state 2 to state 1 to occur more rapidly than in the wild-type strain (W. M. Schluchter, J. Zhao, and D. A. Bryant, personal communication). Mutations affecting cyclic electron transport (e.g., Zhao et al., 1993), the NADH dehydrogenase (e.g., Schluchter et al., 1993), or both Yu et al., 1993), also affect the kinetics of state transition changes, probably through the effects of these mutations on the redox behavior of the components of the ETS between the two photosystems (J. Zhao and D. A. Bryant, personal communication). The detailed characterization ofthese mutants may provide new insights into the signaling mechanism for the state transition. The most interesting mutations affecting state

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transitions identified thus farare mutations affecting core components of the PBS. Zhao et al. (1992) found that a mutation in the apcD gene, encoding the allophycocyanin B subunit; of Synechococcus sp. PCC 7002 cannot perform the state transition and that cells of the mutant appear to remain in state 1 under all light regimes. Although the apcD mutant grew as rapidly as the wild-type strain under highintensity white light, the doubling time ofthe mutant was ~35% greater than the wild type when cells were grown in green light (PBS light). Zhao et al. (1992) also studied a mutation of the apcF gene, encoding the allophycocyanin subunit and found that it had very low fluorescence emission at 695 nm from cells adapted to state 2 but essentially normal emission from cells adapted to state 1. Zhao et al. (1992) proposed a model in which the subunit of the PBS plays a critical role in distributing light energy between the two photosystems. Under state 2 conditions, this subunit was proposed to direct light energy to PS I but during state 1 conditions, light energy is redirected towards PS II (also see Chapter 7). Although details of the molecular mechanism are still largely unknown, the regulated distribution of energy between the two photosystems, i.e. the state transition, maintains a balanced state ofthe two light reactions so that highly efficient photosynthesis can occur. However, the dynamic range for adjustment by the state transition is rather narrow. When the state transition cannot compensate for an imbalance in energy distribution to the two photosystems, a long-term adaptation occurs in which the PS I:PS II ratio changes. III. Long-Term Adaptation: Regulation of PS I: PS II Stoichiometry

A. Historical Background Changes in the Stoichiometry of thylakoid components, such as changes in the ratio of the two photosystems, were first observed as a change of quantum yield of the light absorbed by Chl a. Yocum and Blinks (1958), in studies with Porphyra nereocystis, and Brody and Emerson (1959), in studies with Porphyridium cruentum, independently found that the quantum yield of light absorbed by Chl a is variable and dependent upon the light regime for cell growth. When cells are grown under PBS-absorbed light, the quantum yield for light absorbed by Chl a

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is lowest and shows steep red- and blue-drops (Emerson and Rabinowitch, 1960). However, the quantum yields for light absorbed by Chl a become higher when cells are grown under Chl a-absorbed light, although the quantum yield for PBS-absorbed light decreases significantly under these conditions. Thus, the phenomenon can be summarized as follows: (1) Cells grown in light absorbed by PBS produce a pigment system showing a low quantum yield for the light absorbed by Chl a but a higher quantum yield for the light absorbed by PBS. (2) Cells grown in Chl a-absorbed light have a pigment system characterized by low-quantum yield for PBS-absorbed light but higher quantum yield for Chl a-absorbed light. This phenomenon appears to be an adaptation of the pigment system to chromatic light and thus was called ‘chromatic adaptation’ at that time (not to be confused with complementary chromatic adaptation; see Chapter 21). Indeed, the absorption spectra of cells grown under the two types of chromatic light are different, indicating that the pigment compositions of such cells are different. Cells exhibiting a higher quantum yield for PBS-absorbed light actually contain smaller amounts of PBP relative to Chl a; on the other hand, Chl a abundance becomes lower relative to PBP in cells showing a higher quantum yield for Chl a-absorbed light. The occurrence of this phenomenon in cyanobacteria was confirmed by Myers and his colleagues (Myers, 1963; Jones and Myers, 1964). Since the variation of pigment composition appears to be opposite to the changes in quantum yield, this phenomenon was mysterious for a long time. However, the development of knowledge concerning the molecular properties of thylakoid components, especially the PS I and PS II complexes, brought about new approaches for the analysis of this phenomenon. Myers et al. (1980) showed that changes in the PS I:PS II stoichiometry occur accompanying this phenomenon; the PS I:PS II ratio becomes greater in cells grown under PBS-absorbing light, but decreases under Chl a-absorbed light. At about the same time, Kawamura et al. (1979) reported that the PS I:PS II ratio in cyanobacteria and red algae is greater than 1.0 when cells are grown under weak white light, but decreases to 1.0 or less when cells are grown under strong light. These observations revealed that (1) the stoichiometry of thylakoid components, such as PS I and PS II, is variable; and (2) the stoichiometry ofthese components is adjusted in response to photosynthetic conditions such as light regime. If one re-examines the original observations in

view of the adaptive variation of PS I:PS II stoichiometry, one can explain those findings as follows. The PS I:PS II ratio is highest in cells grown under PBS-absorbed light and decreases when cells are grown under Chl a-absorbed light. Since Chl a is only contained in the two photosystem complexes, and since the number of Chl a molecules in PS I complexes is approximately two-fold greater than that found in PS II complexes, the quantum yield for Chl a -absorbed light increases when the PS I:PS II ratio decreases under Chl a-absorbed light. The stoichiometry between PBS and PS II is constant in most cyanobacteria (Ohki et al., 1987), and thus a decrease in PS I abundance relative to PS II causes higher PBS abundance relative to Chl a. Thus, the phenomenon observed by Yocum and Blinks (1958) and Brody and Emerson (1959) can be recognized as the manifestation of regulation of PS I:PS II stoichiometry in repines to light regime.

B. PS I:PS II Change Table 1 summarizes changes observed in PS I:PS II ratio in various cyanobacteria grown under different conditions. All strains tested so far clearly show changes in PS I: PS II ratio as a function light quality and quantity. Although the values for the PS I:PS II ratio clearly fluctuate among the organisms tested, the pattern that the ratio is greater in cells grown under weak, PBS-absorbed light than that in cells grown under weak, Chl a-absorbed light is common among all organisms. Since PBS-absorbed light excites mainly PS II (henceforth referred to as PS II light) and Chl a-absorbed light preferentially excites PS I (henceforth called PS I light), the pattern tells one that when excitation of PS II exceeds that of PS I, the PS I:PS II ratio is adjusted to become greater, and the ratio is adjusted to be smaller when PS I is preferentially excited. Indeed, changes in the PS I:PS II ratio improve the efficiency of photosynthesis under the respective light regimes (Murakami and Fujita, 1988; Melis et al., 1989). The chromatic lights used in these experiments were so weak that the size of the PBS remained constant at the maximum size in cells of either type. Since the stoichiometry between PBS and PS II is fixed in most of cyanobacteria (see Ohki et al., 1987), the amount of PBS relative to Chl a becomes higher in cells grown under PS I light than that in cells grown under PS II light. This variation is the same as that observed in the earlier studies mentioned above. Changes in PS I:PS II ratio are induced not only by chromatic

Chapter 22 Short-Term and Long-Term Adaptation

light but also by intensity differences for white light. Since PBP absorb a greater fraction of visible light, a weak white light, such as fluorescent light, produces a PS I:PS II ratio similar to that produced by PBSabsorbed light (i.e., the PS I:PS II ratio becomes higher). However, when cells are illuminated by high-intensity white light (at levels sufficiently high to produce saturation of photosynthesis), the PS I:PS II ratio becomes smaller and is similar to that observed in cells grown under Chl a-absorbed light. When cells are illuminated with a white light of very high intensity, the abundance of the two photosystems is markedly reduced; however, the PS I:PS II ratio is still maintained at a low value—similar to that found under a light intensity sufficient to produce saturation of photosynthesis (Yokoyama et al., 1991). Changes in the PS I:PS II ratio are not only produced by differences in the light regime but are also caused by variations in other photosynthetic conditions such as differences in the source of inorganic carbon versus (Eley, 1971; Manodori and Melis, 1984). The PS I:PS II ratio is

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greater when cells are grown on than when they are grown on Heterotrophic growth in the dark causes a greater PS I:PS II ratio, similar to growth under weak white light (Fujita et al., 1987). The question then arises as to which abundance is variable: PS I, PS II, or both. The abundance of PS II seems to be more variable than that of PS I when the abundance is expressed on the basis Chl a, as usually adopted for various photosynthetic reactions. However, both PS I and PS II complexes contain Chl a, and the number of Chl a molecules in a PS I complex is ~ 100–150 Chl a; this is approximately two-fold greater than the number of Chl a in one PS II complex (35–60 Chl a) (Myers et al., 1980; Fujita and Murakami, 1987). Thus, the variation of PS II abundance is greater when the abundance is expressed on the basis of Chl a. However, Chl a is also a variable component in this case, and thus it is not suitable to express changes in the abundance of photosystem complexes on the basis of this cell component. Since one wishes to know the concentration of thylakoid components per functional unit,

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the basis must be the ‘unit thylakoid’ in an ideal sense. For convenience, however, cell or particle (chloroplast) units can be used instead of thylakoid units, when thylakoid concentration does not vary significantly during the experimental incubation, Table 2 shows the variation of thylakoid components in the unicellular strain Synechocystis sp. strain PCC 6714 expressed as the number of complexes per cell. The occurrence of variation in PS I abundance is clear. The abundances of neither the PS II nor the Cyt complexes are significantly changed. This pattern indicates that the abundance of PS I is regulated in repines to light regime. This was confirmed by immunological determination of component proteins (Aizawa et al., 1992). However, Cunnigham et al. (1990) found that variation occurs not only in the abundance of PS I but also in the abundance of PS II in the red alga Porphyridium cruentum. Variation of PS II abundance has also been confirmed with the red alga Porphyra yezoensis (S. Abe, personal communication). This difference will be attributed to a difference in the association of PS II with PBS as discussed below. The pattern that the abundance of PSI complexes, but not PS II complexes, is regulated in response to photosynthetic conditions is common in organisms that contain typical hemidiscoidal PBS; in such cases, one PBS is associated with one PS II complex (Ohki et al., 1987). The next question is whether synthesis or degradation of PS I causes the variations in its abundance. The effect of chloramphenicol on PS I variation suggested that synthesis or assembly of PS I is accelerated under PS II light or weak white light and is suppressed under PS I light or strong white light (Fujita et al., 1988; Murakami and Fujita, 1991b). More direct examination using pulse-labeling and pulse-chase experiments has confirmed the above assumption (Aizawa and Fujita, 1992). The rate of PSI synthesis is two-fold faster under PS II light than that under PS I light, and changes in the rates of synthesis of PS II components (PsbA and PsbC) were insignificant. The rates of degradation of either PS I or PS II are very slow and remain constant. Changes in the rates of PS I synthesis occur rapidly. Upon a shift of light regime, acceleration or deceleration of PS I synthesis occurs with a of 2 to 4 min. The changes in the PS I:PS II ratio are produced by regulating the rate of synthesis of the PS I complex in response to photosynthetic conditions to bring about balanced rates of electron transport by the two photosystems. As shown in Table 3 for a

vascular plant (pea), the stoichiometry of PS II, Cyt and ATP synthase complexes remain constant while PS I abundance decreases in plants grown under high light intensities (Leong and Anderson, 1986). Thus, regulatory variations in PS I content also seem to occur for the thylakoid system of vascular plant chloroplasts. The ratio of PS I to PS II (PS I:PS II), rather than the ratio PS II:PS I, is proposed as the index for monitoring the regulation of photosystem stoichiometry, since PS I synthesis appears to be regulated in all organism so far examined. Because this regulation is achieved by de novo synthesis of PS I complexes, regulation occurs only during the development of thylakoids coupled with cell proliferation. Thus, this regulatory phenomenon is easily observable in algal systems. In vascular plants, such changes can only be seen in developing seedlings but not in adult plants.

C. Possible Mechanism Questions arise as to what is the signal for regulation of the PS I:PS II ratio, how is the signal transduced and how is the synthesis of the PS I (and also PS II) complexes controlled. However, at present the detailed mechanism for the regulation of the PS I:PS II stoichiometry remains unknown. Nonetheless, evidence suggesting possible answers to these questions will be described. As noted above, changes in the PS I:PS II ratio occur not only in response to light regime—changes in light quality and quantity—but also to changes in cellular energy requirement such as an increased requirement for ATP when serves as the carbon source for fixation. Melis et al. (1985) first proposed that the ATP:NADPH ratio in cytosol, which has been determined by a counterbalance

Chapter 22 Short-Term and Long-Term Adaptation

between consumption and production, is the signal for regulation of PS I:PS II stoichiometry. Although Fujita et al. (1985) assumed a similar model for the signal when they observed PS I:PS II ratio changes in numerous organisms, they subsequently found that the PS I:PS II ratio change corresponds well with the ratio of open versus closed PS I and PS II reaction centers. The PS I:PS II ratio becomes greater when cells are placed under conditions where electron carriers acting between two photosystems, PQ and the Cyt complex, remain in a highly reduced state (see Fujita et al., 1987). The light regime is not the sole effective environmental factor controlling the PS I:PS II ratio. Indeed, the PS I:PS II ratio becomes high even under dark heterotrophic conditions where all ETS components remain in a highly reduced state because the rate-limitation in respiration occurs at Cyt oxidase (see Fujita et al., 1987; Adhikary et al., 1990). In Synechocystis sp. strain PCC 6714, respiration depends on thylakoid ETS (K. Ohki and Y. Fujita, unpublished results). Based on correlation between state ofthylakoid ETS and the change in PS I:PS II ratio, it was proposed that regulation ofthe PS I:PS II ratio, i. e., regulation ofPS I synthesis, occurs in response to the redox state of ETS components acting between two photosystems, PQ and/or Cyt complex (Fujita et al., 1987; Murakami and Fujita, 1991a). According to their model, the redox state of such component(s) is monitored in some manner that is probably similar to that for short-term adaptation, i.e., the state transition. However, the PS I:PS II ratio becomes smaller under strong white light while PQ must remain in a highly reduced state due to the rate-limitation of photosynthesis oxidation) under such conditions (Murakami and Fujita, 1991b). In an examination of flash-induced oxidation-reduction of Cyt f in vivo, Murakami and Fujita (199la) found that the change in PS I:PS II ratio greater values, caused by an acceleration of PS I synthesis, correlates well with Cyt oxidation in

687 the Q-cycle (Murakami and Fujita, 1991a). Furthermore, Murakami and Fujita found that HQNO at a concentration suppressing Cyt oxidation at the decelerates PS I synthesis that is accelerated by PS II light (Fujita et al., 1992). Although the approaches for examining the nature of the controlling signal have remained at a physiological level and were made by using rather indirect methods, a model involving Cyt oxidation as the signal for the regulation mechanism appears to be most plausible. HQNO-sensitive Cyt oxidation may be involved in the signal-monitoring mechanism for the regulation; however, definitive proof and additional details of the signaling process remain unknown. It is possible that the same signal-monitoring system acts in the state-transition process. If this is correct, then adjustment of the thylakoid system is achieved by a short-term adaptation, the state transition, and a long-term adaptation, changes in the PS I:PS II ratio, by a common signal-monitoring system. The next question to arise concerns how the signal transduction process occurs—how a redox measurement is used to control the rate of synthesis of the PS I complex. At present, no evidence is available to suggest a mechanism for this process. More knowledge of the molecular mechanism of signalmonitoring systems or of the control of component reaction(s) in PS I synthesis are necessary before a mechanism for the signal transduction process can be formulated. The PS I complex is a very large and complicated protein complex (see Chapter 10). Its de novo assembly in the thylakoid membrane requires both the synthesis of proteins and of prosthetic groups such as Chl a, quinones, and Fe-S centers. Thus, the control of PS I synthesis can be achieved not only by regulation of apoprotein synthesis but also by regulation of synthesis and/or supply of prosthetic groups to the sites for complex assembly. Glick et al. (1986) reported that the abundance of the mRNA for the psaAB genes, that encode the two largest apoproteins of the PS I complex, is higher in pea seedlings grown in PS II light but is lower in seedlings grown under PS I light. Since the mRNA levels seemed to correlate with the rate and extent of PS I synthesis, these workers assumed that transcription of the psaAB genes is regulated during the change in PS I:PS II ratio. However, the psaAB transcripts turn over rather rapidly (Aizawa and Fujita, 1992), and their levels do not necessarily correspond with changes in transcription rate.

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Yoshihiko Fujita, Akio Murakami, Katsunori Aizawa and Kaori Ohki

Klein et al. (1988a, b) and Eichacker et al. (1990) have found that synthesis of the PsaA and PsaB polypeptides in etiolated barley seedlings is regulated by the Chl a supply for their assembly. When Chl a is not supplied (e.g., during growth in the dark), peptide elongation on the thylakoid-bound polysomes is stopped (Klein et al., 1988a) but can be re-initiated by Chl a supply due to illumination or exogenous addition of Chlide a and geranylgeranyl pyrophosphate, the precursor of phytol (Eichacker et al., 1990). Thus, synthesis of the PsaA and PsaB polypeptides in barley can be controlled not at the transcriptional level but at the translational level—at peptide elongation on polysomes. A similar mechanism may be imagined for regulation of PS I synthesis during the change in PS I:PS II ratio in cyanobacteria. In the green plants, Chl a supply to complex assembly occurs hierarchically among the three complexes: PS II, PS I and LHC II complexes, with the affinity being higher in this order (Fujita et al., 1989). If the same hierarchical supply of Chl a to assembly of the two photosystem complexes occurs in cyanobacteria, then the regulation ofChla synthesis could control the PS I:PS II stoichiometry. Indeed, Fujita et al. (1990) found that PS I formation is selectively suppressed by the inhibitors of Chl a synthesis, even when cells were grown under PS II light, so that the PS I:PS II ratio becomes smaller as occurs in cells grown in PS I light. It is noteworthy that Myers et al. (1982) showed that Anacystis nidulans TX20 accumulates the Chl a precursor, Pchlide a, when grown under PS I light and that accumulated Pchlide a disappears when cells are shifted from PS I light conditions to PS II light conditions. Pchlide a accumulation occurs in all strains tested at present. This finding suggests that PS I light suppresses Chl a synthesis, and that PS II light releases such suppression. Chromatic light appears to control Chl a synthesis at the terminal step ofsynthetic pathway (see Chapter 17) or during transport to the site(s) for assembly ofthe photosystem complexes. The time-course of Pchlide a disappearance upon the shift to PS II light conditions correlates well with that of the increase in the rate of PS I synthesis determined by pulse-labeling upon shift to PS II light (Y. Fujita and K. Aizawa, unpublished results). Regulation of Chl a synthesis, observed as Pchlide a accumulation under various conditions, correlates well with the regulation of PS I synthesis. If PS I synthesis is regulated by Chl a supply, synthesis of the PS I complex may be regulated

at the level of its assembly; the flux of Chl a supply to PS I assembly would be regulated by control of Chl a synthesis itself and/or the regulation could occur in the affinity for supply of Chl a to the assembly of the two respective photosystem complexes. Thus, a decrease in Chl a synthesis would cause a selective regulation of PS I synthesis as Fujita et al. (1990) found in the effect of artificial inhibitors. PS II light would release Chl a synthesis from the suppression that occurs under PS I light on one hand and would increase the affinity for Chl a supply to PS I assembly on the other. However, this suggested mechanism is speculative and is not supported by direct experimental evidence at present. Evaluation of this hypothesis must await for further developments in our knowledge ofthe terminal steps of Chl a synthesis. Figure 3 shows schematically a working model for regulation of PS I: PS II stoichiometry. When turnover of PS II exceeds PS I, or when an extra requirement of ATP occurs, electron flux through Cyt of the Cyt complex increases. The net signal reaction, Cyt oxidation, would then occur and would activate reductively (?) a factor in the signal transduction system. The activated factor would release the Chl a supply for PS I assembly from suppression and would stimulate PS I assembly. In the case of Porphyridium cruentum, acceleration of Chl a supply to PS I assembly would cause a decrease in Chl a supply to PS II assembly so that suppression of PS II assembly would simultaneously occur. This variation in PS II assembly is possible in red algae such as Porphyridium cruentum and Porphyra yezoensis because multiple PS II complexes are associated with one PBS. In such a case, the stoichiometry between PS II and PBS could be rather flexible. The abundance of Cyt oxidase, another terminus of the thylakoid ETS (for details, see Chapters 9 and 13), seems to be regulated in parallel with PS I (Adhikary et al., 1990). Synthesis of Cyt oxidase might be regulated at a different step(s) from PS I synthesis but by a common signal coming from the ETS. If this notion is proven correct, a pattern that the abundance of a terminal ETS component is regulated so as to balance the electron influx to, and the efflux from, the ETS by monitoring the redox status of intermediate ETS components, such as the Cyt complex, may be a common character of not only photosynthetic but also respiratory ETS. Complementary chromatic adaptation, a process that produces changes in the molecular species of the

Chapter 22 Short-Term and Long-Term Adaptation

peripheral rods of PBS in response to light wavelength, and sun-shade adaptation, a process that produces changes in the size (and sometimes composition) ofthe peripheral rods ofPBS depending on irradiance intensity, are also types of long-term adaptation ofthe light-harvesting antenna system in cyanobacteria. These topics will not be considered in this chapter. Complementary chromatic adaptation has been discussed in detail elsewhere in this volume (see Chapters 7 and 21); information concerning the mechanism ofsun-shade adaptation is presently rather limited (however, also see Chapters 7 and 21).

IV. Relationship Between Short-Term and Long-Term Adaptation Both short-term adaptation, the state transition, and long-term adaptation, changes in the PS I:PS II ratio, are homeostatic capabilities of the thylakoid system designed to maximize light-energy conversion. Conditions for growth, such as the light regime and nutrient availability, are not always ideal for the photosynthetic process, and the efficiency of lightenergy conversion must therefore be lowered. Changes in light quality and quantity cause an imbalance between the two photoreactions: an increase in the closed state of either photosystem lowers the efficiency of light-energy conversion. When cells must grow under nutritionally poor conditions, extraATP is required by pumping systems for the incorporation and concentration ofnutritional ions within cells. Under these conditions, the proportion of cyclic electron transport relative to

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non-cyclic electron transport increases, and an imbalance between the two photosystems occurs. However, such imbalance is monitored by the redox state of PQ and/or Cyt complex (most probably Cyt and a short-term adaptation, the state transition, and a long-term adaptation, a change in the PS I:PS II ratio, probably occur simultaneously. The state transition adjusts the balance between the two photoreactions very quickly but within a rather limited dynamic range. As cell growth continues and thylakoids develop, long-term adaptation also occurs. An adjustment in the rate of PS I synthesis occurs as rapidly as for the state transition. However, the effect of such an adjustment appears slowly and depends on the rate of thylakoid development and cell proliferation. Although the rate ofchange is slow, the range of the PS I:PS II ratio change can produce a large adjustment in the photoreactions. Both adjustments, acting respectively as fine and coarse tuning mechanisms to changes in photosynthetic environments, serve to maintain the optimal functioning of thylakoid photoreaction systems (see Fig. 4). The overall efficiency of light-energy conversion in photosynthesis is very high. Such a high efficiency is attributed not only to the elaborate molecular architecture and mechanisms of the antenna proteins, reaction centers and electron transport molecules, but also to an ability for homeostatic regulation of system function during both short-term and longterm adaptation. The purpose ofthis article has been to describe the phenomena surrounding the state transition, whichrefers tothe short-termredistribution of energy to the photosystems, as well as the changes

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in PS I:PS II ratio that lead to long-term optimization of the photosynthetic apparatus. As explained above, studies of both types of regulation have thus far remained predominantly at the descriptive and physiological levels. However, detailed knowledge of thylakoid components is accumulating rapidly due to advances brought about by the combination of molecular genetics, biochemistry, and biophysics. Based upon more detailed understanding of the component protein complexes, the molecular details of these regulatory processes should be elucidated in the near future.

Acknowledgments The studies described in this article from the laboratory of the authors were supported in part by Grants-in-Aid for Scientific Research from the Ministry of Education, Sciences and Culture, Japan.

References Adhikary SP, Murakami A, Ohki K and Fujita Y (1990) Photoregulation of respiratory activity in the cyanophyte Synechocystis PCC 6714: the possibility of the simultaneous regulation of the amount of PSI complex and the activity of respiratory terminal oxidase in thylakoids. Plant Cell Physiol 31: 527–532 Aizawa K and Fujita Y (1992) Regulation of PSI formation induced by light quality observed with Synechocystis PCC 6714. In: Murata N (ed) Research in Photosynthesis, Vol IV, pp 329–332. Kluwer, Dordrecht

Aizawa K, Shimizu T, Hiyama T, Satoh K, Nakamura Y and Fujita Y (1992) Changes in composition of membrane proteins accompanying the regulation of PSI/PSII stoichiometry observed with Synechocystis PCC 6803. Photosynthesis Res 32: 131–138 Allen JF (1992) Protein phosphorylation in regulation of photosynthesis. Biochim Biophys Acta 1098: 275–335 Allen JF and Holmes NG (1986) A general model for regulation of photosynthetic unit function by protein phosphorylation. FEBS Lett 202: 175–181 Allen JF, Sanders CE and Holmes NG (1985) Correlation of membrane protein phosphorylation with excitation energy distribution in the cyanobacterium Synechococcus 6301. FEBS Lett 193: 271–275 Anderson JM (1986) Photoregulation of the composition, function, and structure of thylakoid membranes. Annu Rev Plant Physiol 37: 93–136 Barber J (1983) Membrane conformational changes due to phosphorylation and the control of energy transfer in photosynthesis. Photobiochem Photobiophys 5: 181–190 Bennett J (1991) Protein phosphorylation in green plant chloroplasts. Annu Rev Plant Physiol Plant Mol Biol 42: 281– 311 Bennett J, Shaw EK and Michel H (1988) Cytochrome complex is required for phosphorylation of light-harvesting chlorophyll a/b complex II in chloroplast photosynthetic membranes. Eur J Biochem 171: 95–100 Biggins J and Bruce D (1989) Regulation of excitation energy transfer in organisms containing phycobilins. Photosynthesis Res 20: 1–34 Biggins J, Campbell CL and Bruce D (1984a) Mechanism of the light state transition in photosynthesis. II. Analysis of phosphorylated polypeptides in the red alga, Porphyridium cruentum. Biochim Biophys Acta 767: 138–144 Biggins J, Campbell CL, Creswell LL and Wood EA (1984b) Mechanism of the light state transition in Porphyridium cruentum. In: Sybesma C (ed) Advances in Photosynthesis Research, Vol III, pp 303–306. Martinus Nijhoff/Dr W. Junk Publishers, The Hague

Chapter 22 Short-Term and Long-Term Adaptation Bonaventura C and Myers J (1969) Fluorescence and oxygen evolution from Chlorella pyrenoidosa. Biochim Biophys Acta 189: 366–383 Brody M and Emerson R (1959) The quantum yield of photosynthesis in Porphyridium cruentum, and the role of chlorophyll a in the photosynthesis of red algae. J Gen Physiol 43: 251–264 Bruce D, Brimble S and Bryant DA (1989) State transition in a phycobilisome-less mutant of the cyanobacterium Synechococcus sp. PCC 7002. Biochim Biophys Acta 974: 66–73 Bruce D and Salehian O (1992) Laser-induced optoacoustic calorimetry of cyanobacteria. The efficiency of primary photosyntheticprocesses in state 1 andstate 2. Biochim Biophys Acta 1100: 242–250 Canaani O (1986) Photoacoustic detection of oxygen evolution and state 1–state 2 transitions in cyanobacteria. Biochim Biophys Acta 852: 74–80 Coughlan SJ (1988) Chloroplast thylakoid protein phosphorylation is influenced by mutations in the cytochrome bf complex. Biochim Biophys Acta 933: 413–422 Coughlan SJ and Hind J (1986) Purification and characterization of a membrane-bound protein kinase from spinach thylakoids. J Biol Chem 261: 11378–11385 Cunnigham FX Jr, Dennenberg RJ Jursinic PA and Gantt E (1990) Growth under red light enhances Photosystem II relative to Photosystem I and phycobilisomes in the red alga Porphyridium cruentum. Plant Physiol 93: 888–895 Eichacker LA, Soll J, Lauterbach P, Rüdiger W, Klein RR and Mullet JE (1990) In vitro synthesis of chlorophyll a in the dark triggers accumulation of chlorophyll a apoproteins in barley etioplasts. J Biol Chem 265: 13566–1357 Eley JH (1971) Effect of carbon dioxide concentration on pigmentation in the blue-green alga Anacystis nidulans. Plant Cell Physiol 12: 311–316 Emerson R and Lewis CM (1942) The photosynthetic efficiency of phycocyanin in Chroococcus, and the problem of carotenoid participation in photosynthesis. J Gen Physiol 25: 579–595 Emerson R and Rabinowitch E (1960) Red drop and role of auxiliary pigments in photosynthesis. Plant Physiol 35: 477– 485 Fujita Y and Murakami A (1987) Regulation of electron transport composition in cyanobacterial photosynthetic system: Stoichiometry among Photosystem I and II complexes and their light-harvesting antennae and cytochrome complex. Plant Cell Physiol 28: 1547–1553 Fujita Y, Ohki K and Murakami A (1985) Chromatic regulation of photosystem composition in the photosynthetic system of red and blue-green algae. Plant Cell Physiol 26: 1541–1548 Fujita Y, Murakami A and Ohki K (1987) Regulation of photosystem composition in the cyanobacterial photosynthetic system: the regulation occurs in response to the redox state of the electron pool located between the two photosystems. Plant Cell Physiol 28: 283–292 Fujita Y, Murakami A, Ohki Kand Hagiwara N (1988) Regulation of photosystem composition in cyanobacterial photosynthetic system: Evidence indicating that Photosystem I formation is controlled in response to the electron transport state. Plant Cell Physiol 29: 557–564 Fujita Y, Iwama Y, Ohki K, Murakami A and Hagiwara N (1989) Regulation of the size of light-harvesting antennae in response to light intensity in the green alga Chlorella pyrenoidosa. Plant

691 Cell Physiol 30: 1029–1037 Fujita Y, Murakami A and Ohki K (1990) Regulation of the Stoichiometry of thylakoid components in the photosynthetic system of cyanophytes: Model experiments showing that control of the synthesis or supply of Chl a can change the stoichiometric relationship between the two photosystems. Plant Cell Physiol 31: 145–153 Fujita Y, Murakami A and Aizawa K (1992) Light acclimation of thylakoid system in cyanophytes: Regulation of PSI formation in response to light regime. In: Murata N (ed) Research in Photosynthesis, Vol IV, pp 301–308. Kluwer, Dordrecht Gal A, Shahak Y, Schuster G and Ohad I (1987) Specific loss of LHCII phosphorylation in the Lemna mutant 1073 lacking the cytochrome complex. FEBS Lett 221: 205–210 Gal A, Schuster G, Frid D, Canaani O, Schwieger HG and Ohad (1988) Role of cytochrome complex in the redox controlled activity of Acetabularia thylakoid protein kinase. J Biol Chem 263: 7785–7791 Gal A, Hauska G, Herrmann R and Ohad I (1990) Interaction between light-harvesting chlorophyll-a/b protein (LHC II) kinase and cytochrome complex. In vitro control of kinase activity. J Biol Chem 265: 19742–19749 Glick RE, McCauley SW, Gruissem W and Melis A (1986) Light quality regulates expression ofchloroplast genes and assembly of photosynthetic membrane complexes. Proc Natl Acad Sci USA 83: 4287–4291 Horton P, Allen JF, Black MT and Bennett J (1981) Regulation of phosphorylation of chloroplast membrane polypeptides by the redox state of plastoquinone. FEBS Lett 125: 193–196 Jerlov NG (1976) Marine Optics. Elsevier Scientific, Amsterdam Jones LW and Myers J (1964) Enhancement in the blue-green alga, Anacystis nidulans. Plant Physiol 39: 938–946 Katoh T and Gantt E (1979) Photosynthetic vesicles with bound phycobilisomes from Anabaena variabilis. Biochim Biophys Acta 546: 383–393 Kawamura M, Mimuro M and Fujita Y (1979) Quantitative relationship between two reaction centers in the photosynthetic system of blue-green algae. Plant Cell Physiol 20: 697–705 Kirschner J and Senger H (1986) Thylakoid protein phosphorylation in the red algae Porphyridium cruentum. In: Akoyunoglou G and Senger H (eds) Regulation of Chloroplast Differentiation, pp 339–344. Alan R Liss Inc, New York Klein RR, Gamble PE and Mullet JE (1988a) Light-dependent accumulation of radiolabeled plastid-encoded chlorophyll aapoproteins requires chlorophyll a. I. Analysis of chlorophyll deficient mutants and phytochrome involvement. Plant Physiol 88: 1246–1256 Klein RR, Mason HS and Mullet JE (1988b) Light-regulated translation of chloroplast proteins. I. Transcripts of psaApsaB, psbA and rbcL are associated with polysomes in darkgrown and illuminated barley seedlings. J Cell Biol 106: 289–301 Lemaire C, Girard-Bascou J and Wollman FA (1987) Characterization of the complex subunits and studies on the LHC-kinase in Chlamydomonas reinhardtii using mutant strains altered in the complex. In: Biggins J (ed) Progress in Photosynthesis Research, Vol IV, pp 655–658. Martinus Nijhoff, Dordrecht Leong TY and Anderson JM (1986) Light-quality and irradiance adaptation of the composition and function of pea-thylakoid membranes. Biochim Biophys Acta 850: 57–63

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Ley AC and Butler WL (1980) Energy distribution in the photochemical apparatus of Porphyridium cruentum in state I and state II. Biochim Biophys Acta 592: 349–363 Malkin S, Herbert SK and Fork DC (1990) Light distribution, transfer and utilization in the marine red alga Porphyra perforata from photoacoustic energy-storage measurements. Biochim Biophys Acta 1016: 177–189 Manodori A and Melis A (1984) photochemical apparatus organization in Anacystis nidulans (Cyanophyceae). Effect of concentration during cell growth. Plant Physiol 74: 67–71 Manodori A and Melis A (1986) Cyanobacterial acclimation to Photosystem I or Photosystem II light. Plant Physiol 82: 185– 189 Melis A, Manodori A, Glick RE, Ghirardi ML, McCauley SW and Neale PJ (1985) The mechanism of photosynthetic membrane adaptation to environmental stress conditions: A hypothesis on the role of electron-transport capacity and of ATP/NADPH pool in the regulation of thylakoid membrane organization and function. Physiol Veg 23: 757–765 Melis A, Mullineaux CW and Allen JF (1989) Acclimation of the photosynthetic apparatus to Photosystem I or Photosystem II light: Evidence from quantum yield measurements and fluorescence spectroscopy of cyanobacterial cells. Z Naturforsch 44c: 109–118 Mullineaux CW (1992) Excitation energy transfer from phycobilisomesto Photosystem I in a cyanobacterium. Biochim Biophys Acta 1100: 285–292 Mullineaux CW and Allen JF (1990) State l–state 2 transitions in the cyanobacterium Synechococcus 6301 are controlled by the redox state of electron carriers between Photosystem I and II. Photosynthesis Res 23: 297–311 Mullineaux CW and Holzwarth AR (1991) Kinetics of excitation energy transfer in the cyanobacterial phycobilisomePhotosystem II complex. Biochim Biophys Acta 1098: 68–78 Mullineaux CW, Bittersmann E, Allen JF and Holzwarth AR (1990) Picosecond time-resolved fluorescence emission spectra indicate decreased energy transfer from the phycobilisome to Photosystem II in light-state 2 in the cyanobacterium Synechococcus 6301. Biochim Biophys Acta 1015: 231–242 Murakami A and Fujita Y (1988) Steady state of photosynthesis in cyanobacterial photosynthetic systems before and after regulation of electron transport composition: Overall rate of photosynthesis and PSI/PSII composition. Plant Cell Physiol 29: 305–311 Murakami A and Fujita Y (1991 a) Steady state of photosynthetic electron transport in cells of the cyanophyte Synechocystis PCC 6714 having different stoichiometry between PSI and PSII: Analysis of flash-induced oxidationreduction of cytochrome f and P700 under steady state of photosynthesis. Plant Cell Physiol 32: 213–222 Murakami A and Fujita Y (1991b) Regulation of photosystem stoichiometry in the photosynthetic system of the cyanophyte Synechocystis PCC 6714 in response to light-intensity. Plant Cell Physiol 32: 223–230 Murata N (1969) Control of excitation transfer in photosynthesis. I. Light-induced change of chlorophyll a fluorescence in Porphyridium cruentum. Biochim Biophys Acta 172: 242–251 Murata N (1970) Control of excitation transfer in photosynthesis IV. Kinetics of chlorophyll a fluorescence in Porphyra

yezoensis. Biochim Biophys Acta 205: 379–389 Myers J (1963) Enhancement. In: Kok B and Jagendorf AT (ed) Photosynthetic Mechanisms of Green Plants, pp 301–317. National Academy of Sciences—National Research Council, Washington, DC Myers J, Graham J-R and Wang RT (1980) Light harvesting in Anacystis nidulans studied in pigment mutants. Plant Physiol 66: 1144–1149 Myers J, Graham J-R and Wang RT (1982) Protochlorophyll(ide) in a blue-green alga. Plant Physiol 69: 549–550 Ohki K, Okabe Y, Murakami A and Fujita Y (1987) A comparative study of quantitative relationship between phycobiliproteins and Photosystem II in cyanobacteria and red algae. Plant Cell Physiol 28: 1219–1226 Salehian O and Bruce D (1992) Distribution of excitation energy in photosynthesis: quantification of fluorescence yields from intact cyanobacteria. J Lumin 51: 91–98 Sanders CE and Allen JF (1987) The 18.5 kD phosphoprotein of the cyanobacterium Synechococcus 6301: A component of the phycobilisome. In: Biggins J (ed) Progress in Photosynthesis Research, Vol II, pp 761–764. Martinus Nijhoff, Dordrecht Sanders CE, Melis A and Allen JF (1989) In vivo phosphorylation of proteins in the cyanobacterium Synechococcus 6301 after chromatic acclimation to Photosystem I or Photosystem II light. Biochim Biophys Acta 976: 168–172 Satoh K and Fork DC (1983) A new mechanism for adaptation to changes in light intensity and quality in the red alga, Porphyra perforata. I. Relation to state 1–state 2 transitions. Biochim Biophys Acta 722: 190–196 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 Publishers, Oxford Schluchter WM, Zhao J and Bryant (1993) Isolation and characterization of the ndhF gene of Synechococcus sp. PCC 7002 and initial characterization of an interposon mutant. J Bacteriol 175: 3343–3352 Williams WP and Allen JF (1987) State 1/state 2 changes in higher plants and algae. Photosynthesis Res 13: 19–45 Yocum CS and Blinks LR (1958) Light-induced efficiency and pigment alterations in red algae. J Gen Physiol 41: 1113–1117 Yokoyama E, Murakami A, Sakurai H and Fujita Y (1991) Effect of supra-high irradiation on the photosynthetic system of the cyanophyte Synechocystis PCC 6714. Plant Cell Physiol 32: 827–834 Yu L, Zhao J, Mühlenhoff U, Bryant DA, and Golbeck JH (1993) PsaE is required for cyclic electron flow around Photosystem I in the cyanobacterium Synechococcus sp. PCC 7002. Plant Physiol 103: 171–180 Zhao J, Zhou J and Bryant DA (1992) Energy transfer processes in phycobilisomes as deduced from analyses of mutants of Synechococcus sp. PCC 7002. In: Murata N (ed) Research in Photosynthesis, Vol 1, pp 25–32. Kluwer, Dordrecht Zhao J, Snyder WB, Muhlenhoff U, Rhiel E, Warren PV, Golbeck JH and Bryant DA (1993) Cloning and characterization of the psaE gene of the cyanobacterium Synechococcus sp. PCC 7002: characterization of a psaE mutant and overproduction of the protein in Escherichia coli. Mol Microbiol 9: 183–194

Chapter 23 Light-Responsive Gene Expression and the Biochemistry of the Photosystem II Reaction Center Susan S. Golden Department of Biology, Texas A&M University, College Station, TX 77843-3258, USA

Summary I. PS II: Agent and Target of Environmental Variation II. PS II Genes of Cyanobacteria A. psbA Genes 1. psbA Multigene Families 2. psbA Gene Structure 3. Mutations in psbA that Cause Herbicide Resistance B. psbB C. psbC and psbD Genes D. Genes Encoding Small PS II Polypeptides and the Manganese Stabilizing Protein III. Response of psbA Genes to Changes in Light Intensity A. Experimental Challenges in Studying Light-Mediated Responses B. Dissecting the High-Light Response of Synechococcus sp. strain PCC 7942 psbA Genes C. Anatomy of a Light-Responsive Promoter D. Long-Term Adaptation to High Light IV. Response of psbD Genes to Changes in Light Intensity A. Regulatory elements of the psbDII gene 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 A. Photoinhibition, D1 Metabolism, and psbA Expression in Wild-Type Synechococcus sp B. The Role of the Two Forms of D1 in Synechococcus sp. strain PCC 7942 VIII. Cyanobacteria as Models for Studying Photoinhibition Mechanisms in vivo IX. Future directions Acknowledgments References

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Summary Characterization of Photosystem II genes from strains of cyanobacteria that can be genetically manipulated provides a new avenue for studying PS II structure, function, and regulation. Much has been learned about the functions of specific PS II components by examining the phenotypes of strains that carry null, defective, or otherwise altered alleles of PS II genes. Molecular approaches have expanded the analysis of PS II regulation to probe the mechanisms that control synthesis of components of the complex. The best-studied examples are thepsbA andpsbD gene families, which encode the D1 and D2 proteins ofthe PS II reaction center, respectively. The presence of multiple genes for these proteins is a novel property of cyanobacteria which suggests a mechanism to regulate PS II synthesis through the differential regulation of the members of the gene families. D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 693–714. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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In Synechococcus sp. strain PCC 7942 the three psbA genes encode two different forms of the D1 protein, and two psbD genes encode identical D2 proteins. Each gene family has members that exhibit light-responsive regulation, in which transcription is induced by exposure ofcells to a higher light intensity and the abundance of both psbA and psbD messages is thereby increased. Light-responsive expression of the psbAII and psbAIII genes is driven by specific cis elements which lie downstream of the promoters in the regions that correspond to the untranslated leaders of their mRNAs. A post-transcriptional mechanism also functions in which a lightinduced factor accelerates the degradation of the psbAI and psbAIII messages during growth at high light intensity. The result is a shift in the psbA mRNA population from the low-light situation, in which psbAI messages predominate, to a high-light population favoring psbAII and psbAIII messages. The difference in D1 proteins encoded by psbAI versus psbAII and psbAIII suggests a qualitative alteration of PS II during transition to a new light environment. I. PS II: Agent and Target of Environmental Variation The PS II complex of cyanobacteria holds an infamous role in history as the instrument of oxidative conversion of the earth’s atmosphere (Schopf and Walter, 1982). Over the millennia, PS II has changed the global environment and has itselfbeen influenced by environmental variations as cyanobacteria and plants colonized a range of habitats. In a given habitat, temporary and cyclic fluctuations of light, temperature, and availability of nutrients and water influence PS II function; some variations are potentially hazardous if the complex is not able to modify function, synthesis, and degradation of its components. The potential for free-radical formation during primary photochemistry necessitates mechanisms to control the absorption oflight and movement of electrons through the primary and secondary reactions. Cyanobacteria and plant chloroplasts share this challenge, but it is likely that they have developed some different strategies for regulation at the genetic level, given the differences in genome and cellular structure. Major advances in genetic transformation of both groups of oxygenic photosynthetic organisms promise new insights into the mechanisms that regulate the photosynthetic apparatus in response to environmental signals. The focus ofthis chapter is the regulation of genes encoding cyanobacterial PS II reaction center proteins by changes in the light environment (For a discussion of PS II structure and function, the reader should see Chapter 8). Light is an environmental variable that is Abbreviations: ATCC – American Type Culture Collection; MSP – Manganese Stabilizing Protein; PCC – Pasteur Culture Collection; PPFD – Photosynthetic Photon Flux Density; UTEX – University of Texas Culture Collection

clearly central to the life ofcyanobacteria: Sufficient light to run the photosynthetic reactions is a necessity, but excess photic energy can result in photoinhibition. Aside from its role as the ultimate energy source in cyanobacteria, light also serves as a signal that triggers control steps in cellular processes. Specific light wavelengths have been shown to affect phycobilisome composition in strains that undergo complementary chromatic adaptation (Bogorad, 1975; Tandeau de Marsac, 1983; Grossman, 1990; see Chapter 21), the ability ofSynechocystis sp. strain PCC 6803 to grow heterotrophically (Anderson and McIntosh, 1991), and differentiation of hormogonia in Calothrix sp. strain PCC 7601 (Damerval et al., 1991). Several recent reports have demonstrated that cyanobacteria regulate some cellular processes by a circadian pacemaker, a mechanism previously thought to function only in eucaryotes (Mitsui et al., 1986; Huang et al., 1990; Chen et al., 1991; Kondo et al., 1993). Light/dark cycles are major pacesetting inputs of circadian clocks, adding an additional role to the list ofinfluences photons can exert on cyanobacteria. Variations in the intensity of white light have been shown to affect the expression ofPS II genes and the synthesis and degradation of their products. It is this environmental stimulus that is the central theme of the following review. II. PS II Genes of Cyanobacteria PS II genes have been characterized from a number of cyanobacteria, including the four species that are most amenable to genetic manipulation: PCC strains 6803 (Synechocystis sp.), 7002 (Synechococcus sp.), 7120 (Anabaena sp.), and 7942 (Synechococcus sp.) (See Chapter 19). Gene transfer procedures are available in other species as well, including Calothrix

Chapter 23 Light Responsive Regulation of PS II Genes sp. strain PCC 7601 and the facultative heterotroph Anabaena sp. strain ATCC 29413, a useful organism for mutagenesis of PS I genes (See Chapter 19). Table 1 documents the sequences of cyanobacterial PS II genes psbA-psbO which have been deposited in the GenBank or EMBL databases or otherwise have been published. These data highlight one feature of cyanobacterial PS II genes that distinguishes them from their chloroplast homologs: The genes that encode the reaction center proteins D1 and D2 are present as multigene families in all cyanobacteria that have been examined.

A. psbA Genes 1. psbA Multigene Families The psbA gene encodes the D1 reaction center protein of PS II. The known complement of cyanobacterial psbA genes ranges from two in Prochlorothrix hollandica (Morden and Golden, 1989) to four in Anabaena sp. strain PCC 7120 (Vrba and Curtis, 1989). The entire gene family has been sequenced from two other strains, Synechococcus sp. strain PCC 7942 (Golden et al., 1986) and Synechocystis sp. strain PCC 6803 (Osiewacz and McIntosh, 1987; Ravnikar et al., 1989; Metz et al., 1990), each of which has three psbA genes. In all cases except P. hollandica, the multiple genes encode two distinct forms of the D1 protein. One of the psbA genes in Synechocystis sp. strain PCC 6803 is sufficiently divergent from the other two to be distinguished by the difference in its hybridization signal on Southern blots with some psbA probes (Jansson et al., 1987); this has also been observed for the psbA family in Synechocystis sp. strain PCC 6714 (Ajlani et al., 1989a). The divergent psbA gene of Synechocystis sp. strain PCC 6803 has been reported to be inactive; however, the strain from which the genes were sequenced is not a wild-type strain, but a mutant selected for its glucose tolerance and improved growth under photoheterotrophic conditions (Williams, 1988). It is possible that a mutation which led to this phenotype might affect expression of PS II genes. The functional significance of multiple psbA and psbD genes and the potential to encode alternate forms of D1 in cyanobacteria has not been established. However, experiments like those described in this chapter demonstrate differential regulation of the genes under different environmental conditions and suggest that the coordinated expression of the

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gene families is important to the overall fitness ofthe cell.

2. psbA Gene Structure The psbA genes of phycobilisome-containing cyanobacteria are distinct from their chloroplast and prochlorophyte counterparts in that they encode a seven-amino-acid domain near the carboxy terminus of the 360-residue D1 pre-protein which is absent in the 345- to 353-residue polypeptide from chlorophyll b-containing organisms (Curtis and Haselkorn, 1984). The significance of this domain, which is not present in the mature protein (Nixon et al., 1992), is unclear. However, the correlation between its presence and the absence of chlorophyll b as an accessory pigment is striking: a 360-codon sequence is present in phycobilisome-containing cyanobacteria (Table 1), the red alga Cyanidium caldarium (Maid et al., 1990), the xanthophyte Bumilleriopsis filiformis (Scherer et al., 1991) and the flagellate Cyanophora paradoxa (Janssen et al., 1989), but not in Chlamydomonas reinhardtii (Erickson et al., 1984), higher plants (Zurawski et al., 1982), or the prochlorophyte P. hollandica (Morden and Golden, 1989). This distinction could represent a specific phylogenetic relationship, or a functional constraint of the polypeptide that pairs a specific D1 carboxyterminal structure with stacked or non-stacking thylakoids. The body of data refuting a specific close relationship between P. hollandica and chloroplasts (Turner et al., 1989; Morden and Golden, 1991; Scherer et al., 1991; Golden et al., 1992; Swift and Palenik, 1992), and placing the prochlorophytes within the cyanobacterial radiation (Turner et al., 1989; Morden and Golden, 1991), directs attention to the latter suggestion. The position of this domain within a cleaved peptide is consistent with the notion that the region may be important in targeting. Nixon et al. (1992) have shown that a mutant D1 protein, synthesized in its mature form (without the carboxyterminal extension), is functional in the thylakoids of Synechocystis sp. strain PCC 6803; however, the efficiency of targeting was not addressed by their study.

3. Mutations in psbA that Cause Herbicide Resistance Alleles of psbA that cause resistance to the herbicides

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Chapter 23 Light Responsive Regulation of PS II Genes

atrazine, DCMU, and/or ioxynil, have been selected in Synechocystis sp. strain PCC 6714 (Ajlani et al., 1989a,b; Creuzet et al., 1990), Synechococcus sp. strain PCC 7002 (Gingrich et al., 1988), and Synechococcus sp. strain PCC 7942 (Golden and Haselkorn, 1985; Hirschberg et al., 1987). In each species, a single member of the psbA gene family is responsible for the atrazine- or DCMU-resistant phenotype; however, Brusslan and Haselkorn (Brusslan and Haselkorn, 1989) demonstrated that it is possible to propagate mutants of Synechococcus sp. strain PCC 7942 on DCMU when the herbicideresistance allele is in any one of the three loci. Pecker et al. (1987) obtained different results, reporting that a merodiploid of Synechococcus sp. strain PCC 7942, which carries both herbicide-resistance and wildtype alleles of psbAI, and wild-type alleles of psbAII and psbAIII, is phenotypically herbicide-sensitive. The conflicting outcomes of these two studies are likely explained by differences in the relative contributions of the three Synechococcus sp. strain PCC 7942 psbA genes to the D1 pool during growth under different light intensities, as described later in this chapter. Variations in the ratio of herbicidesensitive and -resistant reaction centers as a function

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of light intensity were reported recently in mutants of Synechocystis sp. strain PCC 6803 that carry different types of psbA alleles from Synechocystis sp. strain PCC 6714 in the active psbA loci (Bouyoub et al., 1993). Additional herbicide-resistance mutations have been engineered in the psbA genes of Synechococcus sp. strain PCC 7942 (Horvitz et al., 1989; Ohad and Hirschberg, 1990) and Synechocystis sp. strain PCC 6803 (Ohad and Hirschberg, 1992) which have provided important information regarding the structure and function of the D1 polypeptide in vivo. The latter study confirmed the validity of the structural analogy between the D1 polypeptide and the L subunit of the purple bacterial reaction center in the region of the site (Trebst, 1987). In addition, some of the eight unique mutations (Ohad and Hirschberg, 1992) revealed a distinction between the binding sites of herbicides and the quinone.

B. psbB The psbB gene encoding the CP47 chlorophyll abinding polypeptide of PS II has been sequenced from Synechocystis sp. strain PCC 6803, Anabaena

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sp. strain PCC 7120, and Synechococcus sp. strain PCC 7942, as well as from P. hollandica (Table 1). Alignment of deduced amino acid sequences from these genes and from those of chloroplast psbB genes revealed a unique eight-amino-acid insertion in the P. hollandica CP47 sequence in the aminoterminal portion of the polypeptide (Kulkarni et al., 1993). Two transcript 5' ends were reported for the genes from all of these species except Synechocystis sp. strain PCC 6803, for which the message has not been mapped. No functional data are available regarding psbB promoters which may give rise to these transcripts.

C. psbC and psbD Genes The psbD genes encoding the D2 reaction center protein of PS II have been sequenced from Synechocystis sp. strain PCC 6803 and Synechococcus sp. strains PCC 7002 and 7942 (Table 1). Each contains two psbD genes: One overlaps the psbC gene, which encodes the CP43 chlorophyll abinding protein, as is observed in higher plant chloroplasts (Holschuh et al., 1984); a second is not closely linked to psbDC. Southern and partial nucleotide sequence analyses suggest that this number and arrangement of psbD genes is present in P. hollandica as well (M. S. Nalty, T. Mor, L. Cooksey, I. Ohad, and S. S. Golden, unpublished). A single psbC gene has been reported thus far for each strain, and no PS II genes other than psbA and psbD are known to be present as multigene families in cyanobacteria. Both psbD genes were shown to be functional in Synechococcus sp. strain PCC 7942 (Golden et al., 1989a). However, the monocistronic psbDII gene is expressed at very low levels under typical growth light intensities and is induced at higher light intensities (Bustos and Golden, 1992). The monocistronic psbDII genes of Synechocystis sp. strain PCC 6803 and Synechococcus sp. strain PCC 7002 produce messages that are present at steady-state levels comparable to those of the psbDC operon under standard growth conditions (Gingrich et al., 1990; Vermaas and Yu, 1990). In Synechococcus sp. strains PCC 7002 and 7942, psbC is transcribed only as a dicistronic message that also carries the open reading frame of the psbDI gene (Golden and Stearns, 1988; Gingrich et al., 1990). However, a psbC probe recognizes at least three

Susan S. Golden messages for Synechocystis sp. strain PCC 6803, including one which appears to be a monocistronic psbC transcript (Vermaas and Yu, 1990).

D. Genes Encoding Small PS II Polypeptides and the Manganese Stabilizing Protein A few sequences are available for cyanobacterial psbE, psbF, psbH, psbI, psbJ, psbK, psbL, and psbN genes (Table 1). The genes encoding cytochrome psbE and psbF, have been sequenced from Synechocystis sp. strain PCC 6803 (Pakrasi et al., 1988). They are cotranscribed as part of an operon which also includes thepsbL andpsbJ genes (Lind et al., 1993). Although the function of cytochrome is still not known, inactivation of psbEF blocks photosynthetic electron transport (Pakrasi et al., 1988) and destabilizes the PS II reaction center (Pakrasi et al., 1990). Either psbJ (Lind et al., 1993) or psbK (Ikeuchi et al., 1991) can be inactivated in Synechocystis sp. strain PCC 6803 without abolishing photoautotrophic growth; however, mutants lacking function of either gene have a decreased number of PS II reaction centers and a slower photoautotrophic growth rate. The psbH gene has been sequenced from P. hollandica and Synechocystis sp. strain PCC 6803 (Table 1). In chloroplasts psbH is part of an operon that includes psbB, petB, and petD (Westhoff et al., 1986), but this organization is not found in either of the cyanobacterial examples (Mayes and Barber, 1991; Greer and Golden, 1992). The role of the PsbH protein in PS II was investigated by inactivation of psbH in the chromosome of Synechocystis sp. strain PCC 6803 (Mayes et al., 1993). Mayes et al. concluded from the phenotype of their mutants that PsbH is not strictly required for PS II activity, but that it functions to optimize electron flow between and The psbN gene is upstream of psbH and divergently transcribed in Synechocystis sp. strain PCC 6803, as is the case in chloroplast (Mayes et al., 1993) and cyanelle (see Chapter 4) genomes. A mutant in which psbN was deleted along with psbH showed no phenotype other than that of a psbH null mutant, suggesting that PsbN function is not essential for photoautotrophic growth. The psbI gene was identified in Synechococcus sp. strain PCC 6301 by an oligonucleotide probe that was designed from the N-terminal amino-acid sequence of the purified PsbI polypeptide from

Chapter 23 Light Responsive Regulation of PS II Genes Synechococcus vulcanus (Chen et al., 1990). This gene was not useful as a probe to identify psbI from Synechocystis sp. strain PCC 6803 (W Vermaas, personal communication). However, the Synechocystis sp. strain PCC 6803 psbI gene has been cloned recently following purification and N-terminal amino-acid sequence analysis of the PsbI protein from that organism (M. Ikeuchi, personal communication). The gene that encodes the manganese stabilizing protein (MSP) associated with water oxidation was originally identified in the cyanobacterium Synechococcus sp. strain PCC 7942 and it was named woxA. Its locus designation has since been changed to psbO in keeping with PS II gene nomenclature (Hallick, 1989). Inactivation of the gene in Synechocystis sp. strain PCC 6803 and Synechococcus sp. strain PCC 7942 has demonstrated that the protein is not essential for water oxidation or photoautotrophic growth (Bockholt et al., 1991; Burnap and Sherman, 1991; Mayes et al., 1991; Philbrick et al., 1991), although the mutant strains are more susceptible than the wild type to photoinhibition (Mayes et al., 1991; Philbrick et al., 1991). Inactivation of this gene in Synechococcus sp. strain PCC 7942 increased the L-amino acid oxidase activity of a flavoprotein that copurifies with oxygen-evolving PS II particles (Gau et al., 1989), and which is suppressed by the presence of and (Bockholt et al., 1991). This finding lends support to the provocative hypothesis put forward by that research group that the MSP helps to stabilize and associated with the L-amino acid oxidase flavoprotein, which they propose to be an integral component of the water oxidizing enzyme (Bockholt et al., 1991).

III. Response of psbA Genes to Changes in Light Intensity Several groups have reported changes in cyanobacterial psbA gene expression as a function of light intensity. In 1988 Lönneborg et al. (1988) reported that the level of psbA message is higher in Synechococcus sp. strain PCC 6301 cells grown at than in cells grown at Mohamed and Jansson (1989) studied the influence of light on variations in levels of transcripts from a number of photosynthesis-related genes, including psbA, in the glucose-tolerant strain of Synechocystis

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sp. strain PCC 6803 (Williams, 1988). They reported that the psbA transcript pool (potentially contributed by all members ofthe gene family) requires light in order to accumulate, and that the level of psbA messages is higher from cells grown at 1500 than from those grown at (Mohamed and Jansson, 1989). The same pattern was seen in cells in which only psbA-2 or psbA-3 is active, suggesting that these two genes respond similarly to light; transcripts from the psbA-1 gene are not detectable in this strain (Mohamed and Jansson, 1989). Bouyoub et al. (1993) foundevidence for differential regulation of the Synechocystis sp. strain PCC 6803 genes by measuring the level of herbicide resistance in transgenic strains that carry a DCMU-resistance allele of the Synechocystis sp. strain PCC 6714 psbAI gene at the Synechocystis sp. strain PCC 6803 psbA-2 locus and a wild-type allele at the psbA-3 locus (psbAI from Synechocystis sp. strain PCC 6714 is highly conserved with the psbA2 and psbA-3 loci of Synechocystis sp. strain PCC 6803). They found that a higher proportion of reaction centers are DCMU sensitive following incubation at high light, suggesting relatively higher expression at high light from the psbA-3 locus than from psbA-2. Schaefer and Golden (1989) determined that light affects the individual psbA genes differently in Synechococcus sp. strain PCC 7942: Expression of psbAI is inversely related to light intensity, whereas that of psbAII and psbAIII is directly proportional to light intensity. Exploring the mechanism of lightresponsive changes inpsbA gene expression required the development of experimental growth apparatus and protocols that control the effective light intensity perceived by a growing culture.

A. Experimental Challenges in Studying LightMediated Responses The close relationship between light intensity and growth rate in cyanobacteria, and the efficiency of the cyanobacterial light-harvesting antenna, pose special problems for studying light as an environmental signal. The amount of light able to penetrate a culture that has a cross-section of several centimeters is determined as much by cell density as by the intensity of light at the surface of the culture vessel (Schaefer and Golden, 1989a; see Fig. 1). This complicates experiments designed to assay responses to high light: Cell density increases more rapidly

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when light intensity is increased and quickly reduces the effective light intensity penetrating the culture. In order to observe changes in gene expression in response to light intensity, it is necessary to establish conditions that limit the effect ofchanging cell density. This can be done by working at low cell density or minimal culture cross-section, and completing the experiment before the growth reaches a stage that limits light penetration. This strategy was used in many of the experiments described below for analyzing light-responsive PS II gene expression in Synechococcus sp. strain PCC 7942 (Bustos et al., 1990; Bustos and Golden, 1991; Kulkarni et al., 1992). When sustained exposure to high light is important, the cell density must be controlled over the course of the experiment. Some researchers have achieved this by manual periodic dilution of the culture (Krupa et al., 1990). An automated alternative is to use a turbidostat culture device that maintains constant cell density (Bustos and Golden, 1992). Many laboratories use turbidostat continuous culture vessels which are based on the original design of Myers and Clark (1944). As described in their paper, the turbidostat provides a source of experimental material of high uniformity and a means of limiting internal variables so that a specific environmental variable can be studied. The basic design is of concentric glass tubes which comprise

Susan S. Golden

an annular culture vessel surrounded by a water jacket. Cell density control is provided by two photosensors; one is placed inside the culture annulus where it is screened by the culture, and another outside the apparatus where it senses light emitted directly from the source. A change in the light reading between the two electrodes triggers a switch that activates a mechanism to add fresh culture medium to the apparatus. Essential information for building the glass components ofa turbidostat can be found in the descriptions provided in the following references: Myers and Clark, 1944; Bustos and Golden, 1992; and Wünschmann and Brand, 1992. The electronic plans for a Mach IIb turbidostat controller are available from Robert Nagy, Department of Botany, University ofTexas, Austin TX 78713. A key feature of the original design was a culture cross-section of only 6–7 mm, which allows very good light penetration, but seriously limits the capacity of the vessel. Other designs increase the width ofthe annulus (Bustos and Golden, 1992), but at high cell density, the light intensity at the interior of the apparatus can be significantly lower than at the surface. Appropriate compromises of volume and density should be based on the needs of a particular experiment and the likely effect of a light gradient on the outcome.

Chapter 23 Light Responsive Regulation of PS II Genes

B. Dissecting the High-Light Response of Synechococcus sp. strain PCC 7942 psbA Genes The initial observation that the three Synechococcus sp. strain PCC 7942 psbA genes respond differently to changes in light intensity was based on measurements of activity driven by translational psbA-lacZ fusions genes which were recombined into the chromosome (Schaefer and Golden, 1989a). Reporter strains growing in 8-liter carboys were sampled over a time course during which the light penetrating the culture decreased with increasing cell density (Fig. 1). These experiments showed that activity from apsbAII-lacZ or psbAIII-lacZ strain is highest soon after inoculation of the culture, when light intensity penetrating the culture is above and drops sharply as light availability decreases. Conversely, activity from psbAI-lacZ increases during the course of the experiment. Additional experiments showed that the changes inpsbA-lacZ expression correlate with light intensity rather than cell density (Schaefer and Golden, 1989a). The same pattern of response is observed when wild-type cells are shifted from a photosynthetic photon flux density (PPFD) of to higher or lower light intensities and mRNA levels of the three psbA genes are assayed (Bustos et al., 1990). In this experimental protocol cell density is the same in all samples, the surface area to culture cross-section ratio is high, and the samples are harvested within 15–30 min of exposure to the new light intensities. The wild-type psbA messages respond rapidly to changes in incident light intensity as predicted by the reporter gene activities: The psbAI message level increases when cells are shifted to a lower PPFD and decreases at higher light intensities. Conversely, psbAII and psbAIII message levels increase sharply when cells are shifted to higher light intensities and decrease at a lower intensity. The magnitude ofthe increase at high light is proportional to the light intensity, with a greater response observed in cells shifted to than to The agreement between these two types of experiments suggests a transcriptional component for the increase ofpsbAII and psbAIII expression at high light: The reporter genes could reflect changes in transcription or translation, but their transcripts do not appear to have the same stabilities as the native

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messages (Schaefer and Golden, 1989a); the steadystate message levels should reflect changes in transcription and transcript stability. Thus only a transcriptional response should produce comparable effects in the two analyses. The transcription inhibitor rifampicin blocks the response, providing additional evidence for transcriptional induction of psbAII and psbAIII at high light (Kulkarni et al., 1992). Inhibitor addition also revealed a post-transcriptional component, as the rapid disappearance of the psbAI message is slowed when rifampicin is present prior to the increase in light intensity. The accelerated degradation of the psbAI transcript at high light appears to require translation as well as transcription, since addition of chloramphenicol in the absence of rifampicin also slows disappearance of the message. These results suggest that a factor is quickly synthesized when cells are shifted to high light, and that it is necessary for the rapid disappearance ofthe psbAI transcript (Kulkarni et al., 1992). As this hypothesis would predict, exposure to high PPFD for 10 min before the addition of rifampicin is sufficient to trigger accelerated transcript degradation. This experiment, in which transcription was blocked after a brief period of exposure to high light, revealed that the psbAIII message is also susceptible to posttranscriptional control through decreased half-life. The high-level transcriptional induction ofpsbAIII usually masks the accelerated degradation when messages are analyzed by northern blots of RNA from uninhibited cells, because an increase in light intensity always results in a higher level of psbAIII transcripts. When rifampicin is added before the shift to high light intensity, transcriptional induction is blocked, but so is transcription of the presumptive high-light-induced degradation factor. A delay in rifampicin addition allows the induction ofpsbAIII and the presumptive factor; a decreased half-life for the psbAIII message at high light intensity can then be observed after rifampicin treatment. The half-life of the psbAII message is unaffected by the time of addition ofrifampicin, arguing against an artifactual effect caused by the use of inhibitors. Addition of chloramphenicol prior to the light-shift causes a higher-than-normal increase in psbAIII message level, which would be predicted for blocking synthesis of the putative degradation factor, but not induction of the transcript. The identification of a post-transcriptional component in regulation of psbAIII explains an earlier

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paradox: lacZ translational fusions indicate that psbAIII is expressed at a higher level than psbAII at high light intensity (Schaefer and Golden, 1989a), whereas assays of the mRNA population at high PPFD show that the psbAII message level is much higher than that of psbAIII (Brusslan and Haselkorn, 1989; Kulkarni and Golden, 1994). The difference in stabilities of the psbAII and psbAIII messages would account for different observations depending on whether expression is assayed by a reporter which is not a good indicator of post-transcriptional control, or by steady-state message levels which reflect the combined effects of transcription and transcript degradation. The transcriptional and post-transcriptional responses to high light intensity are separable, as protein synthesis appears to be necessary for accelerated transcript degradation, but not for transcriptional induction (Kulkarni et al., 1992). Whether both responses are stimulated by the same sensory pathway has not been determined. It is likely that a similar adaptive response to high light occurs in Synechocystis sp. strain PCC 6803. Mohamed and Jansson (1991) reported that inhibition of photosynthetic electron transport affects degradation but not synthesis ofpsbA transcripts. This is in agreement with the necessity for protein synthesis to produce a specific transcript degradation factor. Electron transport inhibitors block the increase in galactosidase from psbAII-lacZ and psbAHI-lacZ reporter genes at high light intensity, even though transcriptional induction of psbAII and psbAIII is evident by northern analysis (M. R. Schaefer and S. S. Golden, unpublished). We interpret this as an inability to translate the reporter enzyme when photosynthetic electron transport is blocked.

C. Anatomy of a Light-Responsive Promoter The light-responsive induction of psbAII and psbAIII transcription in Synechococcus sp. strain PCC 7942 is driven by specific regulatory elements upstream of their open reading frames (Li and Golden, 1993). These regions were identified by designing apromoter assay vector that fuses specific putative regulatory elements to a promoterless lacZ gene. The vector targets the transcriptional fusion to a region of the Synechococcus sp. chromosome, termed a neutral site, into which insertions are tolerated without detectable phenotypic effect (Bustos and Golden, 1992). Homologous recombination between the

Susan S. Golden neutral site segments on the vector and the native locus on the chromosome transfers a spectinomycin/ streptomycin-resistance cassette (a modified Omega cassette; Bustos and Golden, 1992) and the transcriptional fusion to the chromosome. Vector sequences, based on the Escherichia coli plasmid pBR328, are lost upon recombination. Fig. 2 depicts the promoter assay vector pAM990. Derivatives of this and other neutral-site vectors that can be transferred to the cyanobacterium through conjugation from E. coli are now available (A. L. Kutach, N. Tsinoremas, and S. S. Golden, unpublished). Approximately 1 kb of DNA from the upstream region of each of the psbA genes, including 60 bp of open reading frame, confers patterns of lightresponsive expression on lacZ that match expression of the native psbA genes: psbAII-lacZ and psbAIIIlacZ expression increases upon a shift to high light intensity, and a small but reproducible decrease in galactosidase activity is detected from psbAI-lacZ after the shift (Li and Golden, 1993). Successively smaller fragments ofthepsbAII andpsbAIII upstream regions were generated by the polymerase chain reaction, and their ability to drive light-responsive lacZ expression was tested. These experiments revealed the following control regions (Fig. 3). A basal promoter element is spaced equivalently to E. coli promoters, having a left end of –38 or –39 relative to the transcription start site, and a right end of –1 in the case of psbAIII or +12 for psbAII; a fragment having a shorter right end for the psbAII promoter shows no activity. Expression driven by each of these basal promoters is unaffected by an increase in light intensity. Light-responsive expression is acquired by the extension of the right ends of the promoter elements to include the transcribed, untranslated leader regions of the genes. The untranslated leader regions are also able to confer a small but reproducible level of lightresponsive expression on a heterologous promoter, indicating that they represent specific light-responsive elements. Inclusion ofadditional upstream sequences beyond the basal promoters of either psbAII or psbAIII has the unexpected effect ofdecreasing overall expression from the lacZ reporter fusions (Li and Golden, 1993). However, the light-responsive expression pattern of each construct is unaffected by these sequences. These data suggest that in addition to a basal promoter and downstream light-responsive elements, the psbAII

Chapter 23 Light Responsive Regulation of PS II Genes

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andpsbAIIIcontrol regions include upstreamnegative elements (Fig. 3). Whether these regions repress heterologous promoters has not been tested.

D. Long-Term Adaptation to High Light The transcriptional and post-transcriptional responses described for thepsbA genes occur within minutes of exposure of cells to a higher light intensity (Bustos et al., 1990; Kulkarni et al., 1992). The result is that the composition of the psbA mRNA population changes dramatically, with the ratio of psbAI:psbAII:psbAIII transcripts shifting from about 16:3:1 at to about 1:16:3 within 30 min of a transfer to (Kulkarni and Golden, 1994; Fig. 4), However, when cells are maintained at high light intensity, the message population shifts again, so that after 6 h, the ratio of psbAI:psbAII:psbAIII transcripts is about 7:11:2. This is accounted for by a rebound in psbAI message level 2–3 h after exposure to high light intensity, which plateaus at a level approximately 2 times its value before the light shift. The overall result is that the total psbA message pool is 3- to 4-fold higher

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after prolonged exposure to high light than when cells are maintained at 100 Fig. 5 summarizes in cartoon form the short- and long-term responses of the Synechococcus sp. strain PCC 7942 psbA genes to exposure to high light intensity. IV. Response of psbD Genes to Changes in Light Intensity A member ofthepsbD gene family ofSynechococcus sp. strain PCC 7942 also exhibits light-responsive expression. Translational gene fusions between the lacZ gene and psbDI, psbC, or psbDII, when recombined into a neutral site of the chromosome, show that only psbDII responds significantly to an

Susan S. Golden increase in light intensity (Bustos and Golden, 1992). Northern analysis also indicates a rapid increase in psbDII message, but not the psbDI/C message, after a shift to high light.

A. Regulatory elements of the psbDII gene The Synechococcus sp. strain PCC 7942 psbDII gene has a potential promoter region which matches exactly the canonical E. coli ‘–10’ element in both spacing and nucleotide sequence. However, no sequence corresponding to a ‘–35’ element is evident (Golden and Stearns, 1988; see Chapter 20 for additional discussion of promoters). Exonuclease digestion was performed to remove sequences upstream of the –42 position of the psbDII-lacZ translational fusion. The level of

Chapter 23 Light Responsive Regulation of PS II Genes produced by this construct and the response to high light intensity are comparable to that of the original reporter fusion, which carries approximately 400 bp of DNA upstream of the transcription start site (Bustos and Golden, 1991). This indicates that all of the information necessary for transcription, and for lightresponsive expression, is within or downstream of a promoter element comparable to the position of an E. coli promoter. The transcribed but untranslated leader of the psbDII transcript is approximately 100 nucleotides long, which is approximately twice the length of the corresponding upstream leaders of the psbAII, psbAIII, and psbDI transcripts (Golden et al., 1986; Golden and Stearns, 1988). Deletions within this region provide evidence for its involvement inpsbDII expression and in binding trans-acting factors (Bustos and Golden, 1991). Electrophoretic mobility-shift assays show binding of proteins to a fragment that extends from approximately 100 bp upstream of the transcription start site to approximately 50 bp inside the psbDII open reading frame. Deletions that remove one, two, or three regions of apparent protein-DNA interaction affect the number of complexes formed in mobility-shift assay and dramatically decrease expression of psbDII-lacZ reporter genes. However, recent identification ofnegative elements immediately upstream of the psbAII and psbAIII promoters suggests that at least a portion of the decrease in psbDII-lacZ expression in the deletion mutants may result from additional bases that are present upstream of the promoter in these constructs relative to the undeleted control. Even deletions which have a dramatic effect on overall expression of psbDII do not completely abolish light-responsive expression (Bustos and Golden, 1991). Therefore an element specifically responsible for conferring the light response was not defined by the study.

V. Functional Significance of LightResponsive Regulation The psbA gene family potentially offers a means of altering the composition of PS II qualitatively, since the three genes encode two different forms ofthe D1 protein (Golden et al., 1986; Schaefer and Golden, 1989b). The composition of the message pool soon after the shift to high light intensity is highly biased in favor oftranscripts frompsbAII andpsbAIII which encode Form II of D1 (Bustos et al., 1990; Kulkarni

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and Golden, 1993). This shift in message population is striking, and may define a transition period during which the cells adapt to high light intensity before psbAI expression again increases. Even after longterm adaptation to high light, Form II messages predominate (Kulkarni and Golden, 1993). The lightresponsive expression of psbAII and psbAIII is important for growth at high light intensity: A mutant that is deficient in both genes (R2S2C3) shows growth impairment at high light, although it is able to recover after prolonged exposure (Kulkarni and Golden, unpublished results). A major question still unresolved is whether the difference in primary sequence between Form I and Form II is a significant variable, or whether the benefit of light-responsive regulation is a matter of quantitative variation in the production of psbA message. R2S2C3, the mutant lacking Form II transcripts, has a very low level of psbA message for the first few hours after the shift to high light while the only remaining transcript, that frompsbAI, is down-regulated (Kulkarni and Golden, unpublished results). This period oflowpsbA steadystate message level, regardless of the form of D1 being produced, may account for impaired growth. The D2 amino acid sequences predicted by the psbDI and psbDII genes are identical, and thus there is no potential for a qualitative change in the PS II reaction center as a result of light-responsive expression of psbDII. Induction of this gene at high light boosts the overall level of messages encoding the D2 protein, although the monocistronic, lightinduced transcript still does not reach the level of the psbDI/C transcript (Bustos and Golden, 1992). Transfer of Synechococcus sp. strain PCC 7942 to high light results in an increase in the D2 level as detected by Western analysis of thylakoid proteins. However, a mutant in which the psbDII gene is inactivated shows a decrease in D2 level over the same time course. An experiment in whichthepsbDIIinactivated mutant was co-cultured with wild-type cells showed that the mutant grows as well as the wild type at standard growth light intensity of but decreases in the population when the light intensity is increased to (Bustos and Golden, 1992). These results suggest that induction of the psbDII gene at high light intensity helps the cell maintain the necessary level of D2 synthesis under high-light conditions. If increased expression of psbD at high light is beneficial, is there an advantage in activating a second psbD gene, rather than regulating expression of the

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psbDI/C operon? One possible explanation is that the need for D2 production at high light may exceed the need for CP43, and the structure of the psbDI/C operon may lock in the ratio of synthesis of the two polypeptides. The dicistronic operon structure also occurs in higher plant chloroplasts, where there is no additional monocistronic psbD gene. However, the chloroplast operon gives rise to a complex set of transcripts during light-induced development, and not all are dicistronic (Gamble et al., 1988). Synechocystis sp. strain PCC 6803 also shows several transcripts which contain psbC sequences, including one which is likely to be monocistronic (Vermaas and Yu, 1990). Levels of both psbD transcripts from this strain appear to increase at high light intensity, but no difference between the response of the two is detectable (Mohamed and Jansson, 1989). VI. Light Quality and psbA Expression Light-responsive expression of the Synechococcus sp. strain PCC 7942 psbA and psbD genes poses questions regarding the nature of the environmental signal, the primary sensor, and elements ofthe signal transduction pathway. The nature of the signal is not straightforward, as the shift to high light intensity used as an environmental cue in most of these experiments may inadvertently alter another parameter that is the actual signal. Increased white light intensity is likely to alter the redox state of the cell through changes in PS II photochemical activity, and it may cause a slight change in the spectrum of incident light. Addition ofelectron transport inhibitors prior to the shift to high light intensity showed that the induction of psbAII and psbAIII transcripts is not tied to photosynthetic activity (Tsinoremas et al., 1994). A comparison of the spectral breakdown of the white light used for low-light and high-light exposures did not reveal any significant differences. However, additional experiments demonstrated that light wavelength is important. When cells incubated under of white light are placed inside a blue filter which blocks penetration of other wavelengths, the cells respond as though they are at high light intensity, even though the actual light intensity is only about 10% of the white-light fluence (Tsinoremas et al., 1994). No other wavelength of light induces this response. Red light has little or no effect on psbA

Susan S. Golden message levels itself, but red-light incubation after a pulse of blue light cancels or attenuates the bluelight induction ofpsbAII and psbAIII. It remains to be determined whether this response is equivalent to the high-light response, or represents a second pathway by which light information is transduced to the PS II genes. The identities ofthe sensor and other components of the signal transduction pathway are still unknown, although proteins have been shown to bind upstream of the three psbA genes and psbDII in Synechococcus sp. strain PCC 7942 (Bustos and Golden, 1991; Mueller, 1991). VII. Light-Regulated Gene Expression and the Biochemistry of PS II Proteins The changes in expression of the psbA and psbD genes at high light intensity ultimately increase the message pools encoding the D1 and D2 polypeptides (Bustos and Golden, 1992; Kulkarni and Golden, 1993). This response is in keeping with evidence from chloroplasts that these two polypeptides have accelerated turnover at high light, which would demand high rates of synthesis to maintain PS II function (Mattoo et al., 1984; Virgin et al., 1988). Light-dependent turnover of D1 has been confirmed in cyanobacteria (Goloubinoff et al., 1988 Ohad et al., 1990; Koenig, 1992). The regulation of D1 synthesis and turnover should be considered in the context of the phenomenon of high-light-induced inhibition of photosynthetic capacity, which is generally termed photoinhibition. Photoinhibition has been studied extensively in higher plants and algae (reviewed in Andersson and Styring, 1991). Although the precise source ofdamage is still debated, most studies support a mechanism whereby the D1 polypeptide is specifically damaged, cleaved, and removed from the PS II complex; recovery requires synthesis of new D1 polypeptides in excess of the rate of damage (Ohad et al., 1984; Ohad et al., 1985; Schuster et al., 1988).

A. Photoinhibition, D1 Metabolism, and psbA Expression in Wild-Type Synechococcus sp. The body of data obtained for photoinhibition in Synechococcus sp. strain PCC 6301 and its very close relative, Synechococcus sp. strain PCC 7942 (Golden et al., 1989b), provide a biochemical context in which to interpret the data on regulation of the

Chapter 23 Light Responsive Regulation of PS II Genes psbA genes by light. Samuelsson et al. (1985) found that Synechococcus sp. strain PCC 6301, adapted to a PPFD of rapidly decreases its evolution rate following exposure to 250, 500 or and that inhibition is directly proportional to light intensity. Recovery in dim light appears to require protein synthesis but not transcription, based on the effects ofthe inhibitors chloramphenicol and rifampicin. Additional experiments showed that cells adapted to growth at higher light intensities are more resistant to photoinhibition than those grown at lower intensities, primarily because ofa higher capacity for these cells to recover from the inhibition (Samuelsson et al., 1987). Synechococcus sp. strain PCC 7942 shows the same properties (Krupa et al., 1990). Wünschmann and Brand (1992) determined that high-light-induced decline in yield of photosynthetic evolution displays time-dependentfirst-orderkinetics; likewise, recovery of cells at lower light, restoring full photosynthetic activity, is a time-dependent firstorder process. They compared actual photoinhibition ofa population ofcells with an equation that predicts photosynthetic activity when photoinhibition and recovery occur simultaneously. The precision of fit ofexperimental data to the theoretical curve indicates that net photoinhibition is the sum of rates of simultaneous photoinhibition and recovery, and strongly supports the protein turnover model for photoinhibition. The high-light conditions used to assess lightresponsive gene expression in Synechococcus sp. strain PCC 7942 are favorable for growth of the organism: Parameters are usually an increase in PPFD from to in a turbidostat, or in 2-cm diametertesttubes submerged in a 30 °C waterbath, with provision of 1% inair. Wild-type cells grow better under these conditions (doubling time of about 5 h) than at lower light intensity (10–11 h at Krupa et al. (1990) reportphotoinhibition forwild-type Synechococcus sp. strain PCC 7942 cells shifted from to for 90 min, as manifested by a decrease in evolution to about 35% of the control rate. However, in these experiments, recovery was prevented by addition of streptomycin. Uninhibited cells decrease to approximately 60% of control rates with the same light treatment (Krupa et al., 1990). The difference between the degree of photoinhibition in these two experiments indicates that some recovery occurs even during the photo-

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inhibitory treatment. Wünschmann and Brand (1992) report that Synechococcus sp. strain PCC 6301 cells maintained continuously in photoinhibitory light for more than 1 hour gradually recover to some extent; their photoinhibitory PPFD treatments were approximately 700 and Given these data, the observation that wild-type Synechococcus sp. strain PCC 7942 cells grow well during highlight-shift growth may not be contradictory. Cells pre-adaptedto should be more resistant to photoinhibition after a shift to than those grown at and thus may suffer little decline in evolution rate, and any reduction may be transient. A brief period of reduction in photosynthetic capacity, though observable by evolution or fluorescence measurements, would not be detected by monitoring turbidostatically-controlled growth. A key outcome, however, is overall superior growth of the cells at the higher light intensity. The period of time used for photoinhibition experiments (90 min) corresponds well to the period during which wild-type Synechococcus sp. strain PCC 7942 cells show a dramatic shift in the psbA mRNA population in favor of psbAII and psbAIII transcripts. If left at high light, however, the psbAI message level begins to rise after 2–3 h, until it is more abundant than before the increase in PPFD (Kulkarni and Golden, 1994). The first few hours after exposure to high light may represent an important transition period in which the ability to alter expression from the three genes adapts the cell to high-light growth. Mutants that are unable to activate the light-induced psbA or psbD genes show impaired growth that is consistent with transient photoinhibition and recovery (Bustos and Golden, 1992; Kulkarni and Golden, unpublished results).

B. The Role of the Two Forms of D1 in Synechococcus sp. strain PCC 7942 Genes that potentially encode two forms of D1 have been identified in the three phycobilisome-containing cyanobacteria in which the entire gene family has been sequenced (Table 1), and DNA hybridization data suggest that this will also be the case in Synechocystis sp. strain PCC 6714 (Ajlani et al., 1989a). Changes in PPFD have an effect on psbA expression in Synechococcus sp. strain PCC 7942 that are detectable at the level of D1 protein incorporated into thylakoid membranes: the ratio of Form I to Form II changes as a function of light

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intensity (Schaefer and Golden, 1989b; Kulkarni and Golden, 1994). These observations suggest that the difference in primary sequence between the two forms may confer somewhat different biochemical properties such that the qualitative change in PS II reaction center composition is beneficial under some conditions. Experiments thus far do not clearly distinguish between the effects of qualitative and quantitative changes in D1 synthesis directed by the changes in gene expression. A strain of Synechococcus sp. strain PCC 7942, which lacks function ofthepsbAII andpsbAIII genes, R2S2C3 (Golden et al., 1986), responds similarly to wild type following photoinhibitory treatment, whereas a strain which lacks psbAI function, R2K1 (Golden et al., 1986), is more resistant (Krupa et al., 1990). Initial experiments suggested that this resistance is due to a higher capacity for recovery (i.e., re-synthesis of D1) in R2K1; however, at lower photoinhibitory PPFD, some evidence for streptomycin-resistant, and thus intrinsic, resistance was observed (Krupa et al., 1991). The levels ofmessages from the psbAII and psbAIII genes are very low in wild-type cells cultured but they are elevated in strain R2K1, which lacks the psbAI message, even under these conditions (Kulkarni and Golden, unpublished results). Thus R2K1 has Form II of D1 incorporated into its thylakoids prior to photoinhibitory treatment. If Form II is a high-lightadaptive form, this may be significant. The higher recovery rate of R2K1 may also be due to the unusually high levels of the psbAII message prior to photoinhibition; it is very stable, and is not susceptible to the active degradation that affects levels of the psbAI and psbAIII transcripts at high light intensity (Kulkarni et al., 1992). Mutant strain R2S2C3 grows as well as R2K1 and wild-type Synechococcus sp. strain PCC 7942 when it is incubated in mixed culture with those strains at However, its representation in a mixed population drops upon exposure to 500 (Kulkarni and Golden, unpublished results). Interestingly, it is not lost from the population even after many generations, but appears to recover and continue to maintain its minority fraction of the cell population. It is tempting to assume that the disadvantage strain R2S2C3 suffers at high light is a consequence of the lack of Form II; however, this strain is quantitatively impaired in psbA message pool during the putative critical transition period following exposure to high PPFD. The lone psbA

Susan S. Golden gene in this strain, psbAI, produces a message that is susceptible to accelerated degradation at high light (Kulkarni et al., 1992), and its message level rebounds only after 2–3 h of continuous exposure to high light (Kulkarni and Golden, 1994). Consequently, total psbA message level is low in strain R2S2C3 soon after a shift to high light, and this factor alone may impair photosynthetic function (increase photoinhibitory effects) during the transition period until message levels normalize. Krupa et al. (1991) report poor survival of R2K1 and R2S2C3 grown in continuous culture at or above , and imply that all strains, including the wild type, show symptoms of cell degradation after 120 h of continuous growth even at 50 . Kulkarni and Golden (1994) have not observed difficulty in continuous culture of any of these strains at 125 or Parameters of growth conditions which could account for the difference in strain viability between labs are not obvious. Fig. 6 presents a model for the Synechococcus sp. strain PCC 7942 PS II complex in which the turnover of D1 and D2 is accelerated at high light, and newly synthesized proteins expressed from the high-lightinduced psbA and psbD genes are incorporated into the reaction center. The model is based on the biochemical and genetic data described above. Messenger RNA levels suggest that the D1 composition of the PS II reaction center should be qualitatively different when compared under low light, soon after exposure to high light, and after several hours’ adaptation to high light. Specific elements of the model are now being tested by measurement of PS II composition and protein turnover under these conditions (Kulkarni and Golden, 1994). VIII. Cyanobacteria as Models for Studying Photoinhibition Mechanisms in vivo The ability to transform strains Synechocystis sp. strain PCC 6803 and Synechococcus sp. strain PCC 7942 allows the engineering of isogenic strains that carry single site-directed mutations or loss offunction in particular genes, and many of these mutants show alterations in sensitivity to photoinhibition. van der Bolt and Vermaas (1992) examined four mutants that carry amino-acid substitutions in the D2 polypeptide and that show greatly increased rates of photo-

Chapter 23 Light Responsive Regulation of PS II Genes

inactivation (defined as loss ofPS II electron transport at high light intensity, as opposed to D1 degradation). They found rapid photoinactivation in two mutants that appear to have defects at or near the watersplitting complex and conclude that increased susceptibility to damage is likely caused by the lack ofsufficient donation ofelectrons to highly oxidizing species such as and This rinding is consistent with experiments in which the MSP is absent through inactivation of the psbO gene. Mayes et al. (1991) and Philbrick et al. (1991) report that genetically MSP-depleted mutants of Synechocystis sp. strain

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PCC 6803 show enhanced sensitivity to photoinhibition. Removal of the MSP from Synechocystis sp. strain PCC 6803 thylakoids by washing with Tris also results in light-induced photoinactivation as measured by electron transport from diphenyl carbizide to 2,6-dichlorophenolindophenol (van der Bolt and Vermaas, 1992). Herbicide-resistant mutants of Synechococcus sp. strain PCC 7942 and Synechocystis sp. strain PCC 6714 indicate that perturbations near the binding pocket affect susceptibility to photoinhibition and stability of the D1 protein. Amino-acid substitutions

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at positions 264 and/or 255 of the PCC 7942 D1 protein confer various degrees of resistance to a number ofherbicides including atrazine and DCMU (Koenig, 1990; Ohad et al., 1990) and cause accelerated turnover of D1 (Ohad et al., 1990). These mutants exhibit destabilization of as measured by in vivo thermoluminescence emission, suggesting that destabilization of the semireduced quinone facilitates light-induceddamage in D1 whichsignals its degradation (Ohad et al., 1990). Kirilovsky et al. (1989) observed high-light sensitivity in an atrazineresistant mutant of Synechocystis sp. strain PCC 6714 that carries two mutations in the pocket region ofthe D1 protein. Degradation ofD1 is slower in thylakoids isolated from this mutant than in those from wild-type cells. Modifications other than alteration of the D1 and D2 polypeptides can affect sensitivity of cyanobacterial cells to photoinhibition; many of these can now be addressed genetically. The identification of genes that encode specific fatty acid desaturase enzymes allows the analysis of photoinhibition in cells that have altered membrane lipids. A strain of Synechocystis sp. strain PCC 6803 designated Fad6/ desA::Kmr contains only monounsaturated lipids due to two mutations, one of which is a recombinational inactivation of the desA gene (Gombos et al., 1992). This strain is highly susceptible to lowtemperaturephotoinhibition whereas theparent strain, Fad6, which carries a single mutation and produces di-unsaturated lipids, is unaffected relative to wild type. Thus, tri-unsaturated membrane lipids, produced in wild type but not in Fad6, are not important for resistance to photoinhibition, but di-unsaturated membrane lipids are. The loss of a major superoxide dismutase activity was tested for its affect on photoinhibition in Synechococcus sp. strain PCC 7942, based the role this enzyme plays in alleviating oxygen-radical toxicity (Herbert et al., 1992). The sodB gene, encoding iron superoxide dismutase, was inactivated by an insertion mutation. Although the mutant cells show an increased in sensitivity to damage by active oxygen, photoinhibition of PS II by exposure to high light is not affected. The regulation of PS II genes by light intensity suggests a protective mechanism to avoid photoinhibition, or to repair photoinhibitory damage once it occurs. Although experiments to measure changes in gene activity have used high light as a trigger, the possibility exists that this stimulus actually induces a generalized stress response, which includes the actual

Susan S. Golden cue for changes in gene regulation. However, stimulation ofthe heat shock response, which rapidly induces the groESL operon in Synechococcus sp. strain PCC 7942 (Webb et al., 1990), does not induce the changes in psbA expression that characterize the high light response (Kulkarni and Golden, 1994). Conversely, a shift to high light does not trigger the changes in groESL message that are observed during heat shock. IX. Future directions A combination ofbiochemical, biophysical, genetic, molecular, and physiological approaches greatly improves the chances of thoroughly understanding PS II dynamics: optimization and protection of function, mechanism of damage, repair, and energetics. Concerted efforts to normalize growth parameters between laboratories that assess different aspects of PS II function should greatly improve the applicability ofinformation derived from one type of analysis to another. A big advantage for understanding PS II in cyanobacteria is the focus of many researchers, including those whose work is purely biophysical or physiological, on a few strains that are amenable to genetic manipulation. Perhaps the most important insight that is needed to reach our goal of understanding is to keep the physiology of the organisms in mind, and to interpret data in the context of a vital organism that is constantly adapting to signals from the world around it. The mechanism by which those signals are relayed to the PS II genes is an intriguing secret that the cyanobacteria may be persuaded to reveal in the coming years. Acknowledgments I am grateful to all the members of my laboratory who provided advice and criticism ofthe manuscript. Resham Kulkarni, Rixin Li, and Nikos Tsinoremas deserve special thanks for agreeing to share their data prior to publication, as do Wendy Button and Alan Kutach for their drafts of two of the figures. Many of my colleagues whose work is cited in this chapter provided reprints, suggestions, and unpublished information; I thank them all. Research on the light-responsive PS II genes of Synechococcus sp. strain PCC 7942 was supported by grants from the American Cancer Society (JFRA-

Chapter 23 Light Responsive Regulation of PS II Genes 224), the National Institutes of Health (GM 37040) and the National Science Foundation (DMB8958089).

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rhythm in amino acid uptake by Synechococcus RF-1. Plant Physiol 97: 55–59 Chisholm D and Williams JGK (1988) Nucleotide sequence of psbC, the gene encoding the CP-43 chlorophyll a-binding protein of photosystem II, in the cyanobacterium Synechocystis 6803. Plant Mol Biol 10: 293–301 Creuzet S, Ajlani G, Vernotte C and Astier C (1990) A new ioxynil-resistant mutant in Synechocystis PCC 6714: Hypothesis on the interaction of ioxynil with the D1 protein. Z Naturforsch 45 c: 436–440 Curtis SE and Haselkorn R (1984) Isolation, sequence and expression of two members of the 32 kd thylakoid membrane protein gene family from the cyanobacterium Anabaena 7120. Plant Moi Biol 3: 249–258 Damerval T, Guglielmi G, Houmard J and Tandeau de Marsac N (1991) Hormogonium differentiation in the cyanobacterium Calothrix: A photoregulated developmental process. Plant Cell 3: 191–201 Dzelzkalns VA and Bogorad L (1988) Molecular analysis of a mutant defective in photosynthetic oxygen evolution and isolation of a complementing clone by a novel screening procedure. EMBO J 7: 333–338 Erickson JM, Rahire M and Rochaix J-D (1984) Chlamydomonas reinhardii gene for the 32 000 mol. wt. protein ofphotosystem II contains four large introns and is located entirely within the chloroplast inverted repeat. EMBO J 3: 2753–2762 Fukuda M, Meng BY, Hayashida N and Sugiura M (1989) Nucleotide sequence of the psbK gene of the cyanobacterium, Anacystis nidulans 6301. Nucl Acids Res 17: 7521 Gamble PE, Sexton TB and Mullet JE (1988) Light-dependent changes in psbD and psbC transcripts of barley chloroplasts: Accumulation of two transcripts maintains psbD and psbC translation capability in mature chloroplasts. EMBO J 7:1289– 1297 Gau AE, Wälzlein G, Gärtner S, Kuhlmann M, Specht S and Pistorius E (1989) Immunological identification of polypeptides in photosystem II complexes from the cyanobacterium Anacystis nidulans. Z, Naturforsch 44 c: 971–975 Gingrich JC, Buzby JS, Stirewalt VL and Bryant DA (1988) Genetic analysis of two new mutations resulting in herbicide resistance in the cyanobacterium Synechococcus sp. PCC 7002. Photosynth Res 16: 83–99 Gingrich JC, Gasparich GE, Sauer K and Bryant DA (1990) Nucleotide sequence and expression of the two genes encoding D2 protein and the single gene encoding the CP43 protein of Photosystem II in the cyanobacterium Synechococcus sp. PCC 7002. Photosynth Res 24: 137–150 Golden SS and Haselkorn R (1985) Mutation to herbicide resistance maps within the psbA gene of Anacystis nidulans R2. Science 229: 1104–1107 Golden SS and Stearns GW (1988) Nucleotide sequence and transcript analysis of three photosystem II genes from the cyanobacterium Synechococcus sp. PCC7942. Gene 67: 85– 96 Golden SS, Brusslan J and Haselkorn R (1986) Expression of a family of psbA genes encoding a photosystem II polypeptide in the cyanobacterium Anacystis nidulans R2. EMBO J 5: 2789– 2798 Golden SS, Cho DC and Natty MS (1989a) Two functional psbD genes in the cyanobacterium Synechococcus sp. strain PCC 7942. J Bacteriol 171: 4707–4713

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Susan S. Golden resistant mutant of Synechocystis 6714. Plant Mol Biol 13: 355–363 Koenig F (1990) Anacystis mutants with different tolerances to DCMU-type herbicides show differences in architecture and dynamics of the photosynthetic apparatus, depending on site and mode of the amino acid exchanges in the D1 protein. Z Naturforsch 45 c: 446–451 Koenig F (1992) Development of shade-type appearance – light intensity adaptation and regulation of the D1 protein in Synechococcus. In: Argyroudi-Akoyunoglou JH. (ed) Regulation of chloroplast biogenesis, pp 545–550. Plenum Press, New York. Kondo T, Strayer CA, Kulkarni RD, Taylor W, Ishiura M, Golden SS and Johnson CH (1993) Circadian rhythms in prokaryotes: Luciferase as a reporter of circadian gene expression in cyanobacteria. Proc Natl Acad Sci USA 90: 5672–5676 Krupa Z, Öquist G and Gustafsson P (1990) Photoinhibition and recovery of photosynthesis in psbA gene-inactivated strains of cyanobacterium Anacystis nidulans. Plant Physiol 93: 1–6 Krupa Z, Öquist G and Gustafsson P (1991) Photoinhibition of photosynthesis and growth responses at different light levels in psbA gene mutants of the cyanobacterium Synechococcus. Physiol Plant 82: 1–8 Kulkarni RD and Golden SS (1994) Adaptation to high light intensity in Synechococcus sp. strain PCC 7942: Regulation of three psbA genes and two forms of the D1 protein. J Bacteriol 176: 959–965 Kulkarni RD, Schaefer MR and Golden SS (1992) Transcriptional and posttranscriptional components of psbA response to high light intensity in Synechococcus sp. strain PCC 7942. J Bacteriol 174: 3775–3781 Kulkarni RD, Mueller UW and Golden SS (1993) Nucleotide sequence of psbB from Synechococcus sp. strain PCC 7942. Biochim Biophys Acta 1173: 329–332 Kuwabara T, Reddy KJ and Sherman LA (1987) Nucleotide sequence of the gene from the cyanobacterium Anacystis nidulans encoding the Mn-stabilizing protein involved in photosystem II water oxidation. Proc Natl Acad, Sci USA 84: 8230–8234 Lang JD and Haselkorn R (1989) Isolation, sequence and transcription of the gene encoding the photosystem II chlorophyll-binding protein, CP-47, in the cyanobacterium Anabaena 7120. Plant Mol Biology 13: 441–456 Li R and Golden SS( 1993) Enhancer activity of light-responsive regulatory elements in the untranslated leader regions of cyanobacterial psbA genes. Proc Natl Acad Sci USA 90: 11678–11682 Lind LK, Shukla VK, Nyhus KJ and Pakrasi HB (1993) Genetic and immunological analyses of the cyanobacterium Synechocystis sp. PCC 6803 show that the protein encoded by the psbJ gene regulates the number of photosystem II centers in thylakoid membranes. J Biol Chem 268: 1575–1579 Lönneborg A, Kalla SR, Samuelsson G, Gustafsson P and Öquist G (1988) Light-regulated expression of the psbA transcript in the cyanobacterium Anacystis nidulans. FEES Lett 240: 110– 114 Maid U, Valentin K and Zetsche K (1990) The psbA-gene from a red alga resembles those from cyanobacteria and cyanelles. Curr Genet 17:255–259

Chapter 23 Light Responsive Regulation of PS II Genes Mattoo AK, Hoffman-Falk H, Marder JB and Edelman M (1984) Regulation of protein metabolism: Coupling of photosy nthetic electron transport to in vivo degradation of the rapidly metabolized 32-kilodalton protein of the chloroplast membranes. Proc Natl Acad Sci USA 81: 1380–1384 Mayes SR and Barber J (1990) Nucleotide sequence of the psbH gene of the cyanobacterium Synechocystis 6803. Nucl Acids Research 18: 194 Mayes SR and Barber J (1991) Primary structure of the psbNpsbH–petC-petA gene cluster of the cyanobacterium Synechocystis PCC 6803. Plant Mol Biol 17: 289–293 Mayes SR, Cook KM, Self SJ, Zhang Z and Barber J (1991) Deletion of the gene encoding the Photosystem II 33 kDa protein from Synechocystis sp. PCC 6803 does not inactivate water-splitting but increases vulnerability to photoinhibition. Biochim Biophys Acta 1060: 1–12 Mayes SR, Dubbs JM, Vass I, Hideg É, Nagy L and Barber J (1993) Further characterization of the psbH locus of Synechocystis sp. PCC 6803: Inactivation of pxbH impairs QA to QB electron transport in photosystem 2. Metz J, Nixon PJ and Diner B (1990) Nucleolide sequence of the psbA3 gene from the cyanobacterium Synechocystis PCC 6803. Nucl Acids Res 18: 6715–6715 Mitsui A, KumazawaS, Takahashi A, Ikemoto H, Cao S and Arai T (1986) Strategy by which nitrogen-fixing unicellular cyanobacteria grow photoautotrophically. Nature 323:720–722 Mohamed A and Jansson C (1989) Influence of light on accumulation of photosynthesis-specific transcripts in the cyanobacterium Synechocystis 6803. Plant Mol Biol 13: 693–700 Mohamed A and Jansson C (1991) Photosynthetic electron transport controls degradation but not production of psbA transcripts in the cyanobacterium Synechocystis 6803. Plant Mol Biol 16:891–897 Morden CW and Golden SS (1989) psbA genes indicate common ancestry of prochlorophytes and chloroplasts. Nature (London) 337: 382–385 Morden CW and Golden SS (1991) Sequence analysis and phylogenetic reconstruction of the genes encoding the large and small subunits of ribulose-1,5-bisphosphatc carboxylase/ oxygenase from the chlorophyll b-containing prokaryote Prochlorothrix hollandica. J Mol Evol 32: 379–395 Mueller UW (1991) Identification of protein binding sites in the promoter regions of a light-responsive gene family in a cyanobacterium. M.Sc. thesis, Texas A&M University. Mulligan B, Schultes N, Chen L and Bogorad L (1984) Nucleotide sequence of a multiple-copy gene for the B protein of photosystem II of a cyanobacterium. Proc Natl Acad Sci USA 81:2693–2697 Myers J and Clark LB (1944) Culture conditions and the development of the photosynthetic mechanism II. An apparatus for the continuous culture of Chlorella. J Gen Physiol 28:103– 112 Nixon PJ, Trost JT and Diner BA (1992) Role of the carboxy terminus of polypeptide D1 in the assembly of a functional water-oxidizing manganese cluster in photosystem 11 of the cyanobacterium Synechocystis sp. PCC 6803: Assembly requires a free carboxyl group at C-terminal position 344. Biochemistry 31: 10859–10871 Ohad N and Hirschberg J (1990) A similar structure of the herbicide binding site in photosystem II of plants and

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cyanobacteria is demonstrated by site specific mutagenesis of the psbA gene. Photosynth Res 23: 73–79 Ohad N and Hirschberg J (1992) Mutations in the D1 subunit of photosystem II distinguish between quinone and herbicide binding sites. Plant Cell 4: 273–282 Ohad I, Kyle DJ and Arntzen CJ (1984) Membrane protein damage and repair: Removal and replacement of inactivated 32-kilodalton polypeptides in chloroplast membranes. J Cell Biol 99:481–485 Ohad I, Kyle DJ and Hirschberg J (1985) Light-dependent degradation of the in isolated pea thy lakoids. EMBO J4: 1655–1659 Ohad N, Amir-Shapira D, Koike H, Inoue Y, Ohad I and Hirschberg J (1990) Amino acid substitutions in the D1 protein of photosystem II affect stabilization and accelerate turnover of D1. Z Naturforsch 45 C: 402–408 Osiewacz HD and McIntosh L (1987) Nucleotide sequence of a member of the psbA multigene family from the unicellular cyanobacterium Synechocystis 6803. Nucl Acids Res 15: 10585–10585 Pakrasi HB, Williams JG 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 I I . EMBO J 7: 325–332 Pakrasi HB, Nyhus KJ and Granok H (1990) Targeted deletion mutagenesis of the beta subunit of Cytochrome protein destabilizes the reaction center of photosystem II. Z Naturforsch, C, Biosci 45: 423–429 Pecker I, Ohad N and Hirschberg J (1987) The chloroplastencoded type of herbicide resistance is a recessive trait in cyanobacteria. In: Biggins J. (ed) Progress in Photosynth Res, Vol III, pp 811–814. Martinus Nijhoff Publishers, Dordrecht. Philbrick JB and Zilinskas BA (1988) Cloning, nucleotide sequence and mutational analysis of the gene encoding the Photosystem II manganese-stabilizing polypeptide of Synechocystis 6803. Mol Gen Genet 212: 418–425 Philbrick JB, Diner BA and Zilinskas BA (1991) Construction and characterization of cyanobacterial mutants lacking the manganese-stabilizing polypeptide of photosystem II. J Biol Chem 266: 13370–13376 Ravnikar PD, Debus R, Sevrinck J, Saetaert P and McIntosh L (1989) Nucleotide sequence of a second psbA gene from the unicellular cyanobacterium Synechocystis 6803. Nucl Acids Res 17: 3991 Samuelsson G, Lönneborg A, Rosenqvist E, Gustafsson P and Öquist G (1985) Photoinhibition and reactivation of photosynthesis in the cyanobacterium Anacystis nidulans. Plant Physiol 79: 992–995 Samuelsson G, Lönneborg A, Gustafsson P and Öquist G (1987) The susceptibility of photosynthesis to photoinhibition and the capacity of recovery in high and low light grown cyanobacteria, Anacystis nidulans. Plant Physiol 83: 438–441 Schaefer MR and Golden SS (1989a) Differential expression of members of a cyanobacterial psbA gene family in response to light. J Bacteriol 171: 3973–3981 Schaefer M R and Golden SS (1989b) Light availability influences the ratio of two forms of D1 in cyanobacterial thylakoids. J Biol Chem 264: 7412–7417 Scherer S, Herrmann G, Hirschberg J and Böger P (1991) Evidence for multiple xenogenous origins of plastids: Comparison of

714 psbA-genes with a xanthophyte sequence. Curr Genet 19: 503–507 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. Schuster G, Timberg R and Chad I (1988) Turnover of thylakoid photosystem II proteins during photoinhibition of Chlamydomonas reinhardtii. Eur J Biochem 177: 403–410 Swift H and Palenik B (1992) Prochlorophyte evolution and the origin of chloroplasts: Morphological and molecular evidence. In: Lewin RA. (ed) Origins of Plastids: Symbiogenesis, Prochlorophytes, and the Origins of Chloroplasts, pp 123–139. Chapman & Hall, New York. Tandeau de Marsac N (1983) Phycobilisomesandcomplementary adaptation in cyanobacteria. Bull Inst Pasteur 81: 201–254 Trebst A (1987) The three-dimensional structure of the herbicide binding niche on the reaction center polypeptides of photosystem II. Z Naturforsch 42c: 742–750 Tsinoremas NF, Schaefer MR and Golden SS (1994) Blue and red light reversibly control psbA expression in the cyanobacterium Synechococcus sp. strain PCC 7942. J Biol Chem, in press Turner S, Burger-Wiersma T, Giovannoni SJ, Mur LR and Pace NR (1989) The relationship of a prochlorophyte Prochlorothrix hollandica to green chloroplasts. Nature (London) 337:380–382 van der Bolt F 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 Vermaas WFJ and Yu J (1990) Transcript levels and synthesis of photosystem II components in cyanobacterial mutants with inactivated photosystem II genes. Plant Cell 2: 315–322 Vermaas WF, Williams JG and Arntzen CJ (1987) Sequencing and modification of psbB, the gene encoding the CP-47 protein of photosystem II, in the cyanobacterium Synechocystis 6803.

Susan S. Golden Plant Mol Biol 8: 317–326 Virgin I, Styring S and Andersson B (1988) Photosystem II disorganization and manganese release after photoinhibition of isolated spinach thylakoid membranes. FEBS Lett 233: 408–412 Vrba JM and Curtis SE (1989) Characterization of a fourmember psbA gene family from the cyanobacterium Anabaena PCC 7120. Plant Mol Biol 14: 81–92 Webb R, Reddy KJ and Sherman LA (1990) Regulation and sequence of the Synechococcus sp. strain PCC 7942 groESL operon, encoding a cyanobacterial chaperonin. J Bacteriol 172: 5079–5088 WesthoffP, 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 Williams JGK. (1988) Construction of specific mutations in photosystem II photosynthetic reaction center by genetic engineering methods in Synechocystis 6803. Meth Enzymol 167:766–778 Williams JGK and Chisholm DA (1987) Nucleotide sequences of both psbD genes from the cyanobacterium Synechocystis 6803. In: Biggins J. (ed) Progress in Photosynth Res, Vol. IV, pp 809–812. Martinus Nijhoff, Dordrecht Wünschmann G and Brand JJ (1992) Rapid turnover of a component required for photosynthesis explains temperature dependence and kinetics of photoinhibition in a cyanobacterium, Synechococcus 6301. Planta 186: 426–433 Zhang ZH, Mayes SR and Barber J (1990) Nucleotide sequence of the psbK gene of the cyanobacterium Synechocystis 6803. Nucl Acids Res 18: 1284 Zurawski G, Bohnert HJ, Whitfeld PR and Bottomley W (1982) Nucleotide sequence of the gene for the Mr 32 000 thylakoid membrane protein from Spinacea oleracea and Nicotiana debneyi predicts a totally conserved primary translation product of Mr38,950. Proc Natl Acad Sci USA 79: 7699–7703

Chapter 24 Thioredoxins in Cyanobacteria: Structure and Redox Regulation of Enzyme Activity Florence K. Gleason Department of Plant Biology, University of Minnesota, St. Paul, MN 55108, USA Summary I. Introduction A. General Occurrence of Thioredoxins B. Thioredoxin in Photosynthetic Organisms 1. Thioredoxin in Higher Plants 2. Thioredoxins in Cyanobacteria 3. Thioredoxin Genes in Cyanobacteria II. Structure of Cyanobacterial Thioredoxins III. Reduction of Thioredoxins A. NADPH-Dependent Reductase B. Ferredoxin-DependentReductase IV. Functions of Thioredoxins in Cyanobacteria A.Thioredoxin as a Protein Disulfide Reductase 1. Relation to Chloroplast Thioredoxin-m 2. Modulation of C-3 (Calvin) Cycle Enzymes a. Fructose-1,6-Bisphosphatase b. Phosphoribulokinase 3. Glucose Catabolism-Modulation of Glucose-6-Phosphate Dehydrogenase 4. Thioredoxin and the Enzymes of Nitrogen Metabolism a. NADP-Dependent Isocitrate Dehydrogenase b. Glutamine Synthetase B. Thioredoxin as a Reducing Agent in Cyanobacteria 1. 3'-Phosphoadenosine-5'-Phosphosulfate Reductase 2. Ribonucleotide Reduction C. What is the Essential Function of Thioredoxin in Cyanobacteria? References

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Summary Thioredoxin is a small disulfide-containing redox protein that reduces disulfide bonds in other proteins. It can act both as a modulator ofenzyme activity by reducing structurally important disulfide bonds in a target protein and also as a reducing agent. Unlike other eubacteria, the cyanobacteria have two distinct thioredoxins with approximately 39% amino acid identity. One of the thioredoxins (T1) is similar to bacterial thioredoxins both in structure and general redox activity. The other protein (T2) is relatively unstable and seems to be unique to cyanobacteria. Several enzymes in cyanobacteria are regulated by a disulfide redox mechanism that can be effected by either T1 or T2, including enzymes for fixation, carbon catabolism and nitrogen metabolism. In addition, cyanobacterial thioredoxins can function as reducing agents in the phosphoadenosine phosphosulfate reductase and ribonucleotide reductase reactions. Thioredoxins, in turn, are reduced by the products of PS I, thus providing a biochemical link between light reactions and regulation of metabolism. An attempt to inactivate the T1 gene in Synechococcus sp. strain PCC 7942 did not succeed in producing thioredoxin-minus mutants which implies that T1 performs some essential function in photosynthetic organisms that cannot be D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 715–729. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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efficiently substituted by T2 or other redox systems. Although its role in regulating carbon metabolism seems to be most crucial for photoautotrophic growth, definitive evidence for the in vivo functions ofthioredoxin in cyanobacteria is still lacking.

I. Introduction

A. General Occurrence of Thioredoxins

The cyanobacteria, as do other microorganisms, regulate enzyme activity in response to changing environmental conditions. For photosynthetic organisms, light is the most potent effector of metabolic activity. Light can exert effects both at the gene level, by initiating gene transcription and translation, and at the protein level, by modulating enzyme activity. The effects of light are often not direct. A signal is perceived by a light-sensitive receptor and relayed by a variety of biochemical mechanisms to the appropriate targets. Light-initiated effects on metabolic activity in photosynthetic organisms are mediated by a number of different intracellular changes. For example, shifts in the pH and ion concentration of the cytoplasm due to photosynthetic electron transport affect the activity of ribulose 1,5-bisphosphate carboxylase/oxygenase, the first enzyme in fixation (Portis, 1992; see Chapter 14). Light-initiated protein phosphorylation can redistribute electron flow in the photosynthetic electron transport system (Allen et al., 1985) or alter the activity ofenzymes such as phosphoenolpyruvate carboxylase (Jiao et al., 1990). Light obviously affects the redox state of a photosynthetic cell or organelle, leading to increased ratios of reduced ferredoxin and NADPH relative to the oxidized forms. This change in redox state can be relayed to other enzymes and processes by reduction ofa small disulfide-containing protein, thioredoxin. This review will focus on the structure and activity of thioredoxin in the cyanobacteria and its similarity to proteins and functions in other organisms, in particular, higher plants.

Thioredoxin was first described as a reducing agent for sulfate metabolism in yeast (Wilson et al., 1961). The best characterized protein is from Escherichia coli in which thioredoxin was initially found to function as a reducing agent in ribonucleotide reduction (Laurent et al., 1964). It has since been reported to occur in most living organisms and has been assigned a number of probable rolls in metabolism, mostly based on in vitro activities. These can be divided into three categories. As initially reported, thioredoxin can serve as a reducing agent, i.e., channeling reducing equivalents from a reduced cofactor such as NADPH to some ongoing enzymatic process. These would include reactions such as ribonucleotide reduction (Scheme 1), sulfate reduction and methionine sulfoxide reduction (for a review, see Gleason and Holmgren, 1988). Secondly, thioredoxin is an effective protein disulfide reductase and can modulate the activity of other enzymes in which it reduces structural disulfide bonds. The reduction of a target enzyme can lead to enhanced activity as in the case of chloroplast fructose-1,6-bisphosphatase (Scheme 2). Several examples are described for the redox regulation of enzymes in photosynthetic organisms (see Buchanan, 1991, for a review). This protein disulfide reductase activity of thioredoxin may also facilitate protein folding (Pigiet and Schuster, 1986). Finally, thioredoxin is found as a structural component of some multi-protein complexes. The bestcharacterized example is the bacteriophage T7 DNA polymerase. E. coli thioredoxin and the viral gene 5 protein form a 1:1 complex which then constitutes a

Abbreviations: ATCC – American Type Culture Collection; CCAP – Cambridge Collection of Algae and Protozoa; Coupling Factor 1 ATPase; Coenzyme – 5'-adenosylcobalamin; FAD – flavin adenine dinucleotide; FBPase – fructose-1,6bisphosphatase; FdTR – ferredoxin-dependent thioredoxin reductase; G6PDH – glucose-6-phosphate dehydrogenase; GS – glutamine synthetase; GuHCl – guanidine hydrochloride; NADP-IDH – NADP-dependent isocitrate dehydrogenase; NADP-MDH – NADPdependent malate dehydrogenase; PAPS – 3'-phosphoadenosine-5'–phosphosulfate; PCC – Pasteur Culture Collection; PRK – phosphoribulokinase; RNRase – ribonucleotide reductase; Tl – cyanobacterial thioredoxin type 1; T2 – cyanobacterial thioredoxin type 2; Tf– chloroplast thioredoxin type f; Tm – chloroplast thioredoxin type m; UTEX – University of Texas at Austin Culture Collection.

Chapter 24 Thioredoxins in Cyanobacteria

highly processive DNA polymerase (Tabor et al., 1987). Although the reduced form of thioredoxin is required for complex formation, site-directed mutagenesis has demonstrated that no redox function is involved (Huber et al., 1986). This complexed form of thioredoxin is found in commercially available preparations of T7 DNA polymerase. Thioredoxin is also required for the replication of coliphages f1 and M13. Despite the list of vital functions for thioredoxin, mutants of E. coli that lack thioredoxin are viable. Strains such as E. coli BH2012 have extended lag periods, but normal growth rates during the exponential phase. The only phenotype which distinguishes them from the wild type is their inability to reduce methionine sulfoxide and to support the growth of bacteriophage such as T7, f1, and M13 (Lim et al., 1985a; Lim et al., 1985b; Russel and Model, 1985). Further characterization of these mutants revealed that the role of thioredoxin in reactions such as ribonucleotide reduction is effectively substituted by a related protein called glutaredoxin. Unlike thioredoxin, the redox-active disulfide in glutaredoxin is reduced by glutathione. Although present in bacterial cells in much smaller amounts than thioredoxin, it is a more efficient reducing agent for ribonucleotide reductase, and this may be one of its primary roles in vivo (Holmgren, 1985). Glutaredoxin has a tertiary structure similar to that of thioredoxin (Sodano et al., 1991) but in its primary structure and activity it is more closely related to a number of glutathione-dependent redox

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proteins called thiol transferases which have been described in mammalian tissues (Yang and Wells, 1991).

B. Thioredoxin in Photosynthetic Organisms 1. Thioredoxin in Higher Plants Most organisms contain only one type of thioredoxin with the exception of Corynebacterium nephridii (McFarlan et al., 1989) and yeast (Muller, 1991) in which two proteins are found with very similar amino acid sequences. This is in contrast to the situation in higher plant tissues where multiple thioredoxins have been reported (Jacquot et al., 1978; Bestermann, 1983). A combination of protein chemistry (Kamo et al., 1989; Maeda et al., 1986) and gene isolation (Lepiniec et al., 1992; Wedel et al., 1992) has clearly demonstrated that there are two distinct thioredoxins associated with higher plant chloroplasts. These are encoded by nuclear genes and the proteins are transported into the chloroplast after synthesis in the cytoplasm. Some ofthe amino-terminal variants may result from cleavage ofthe signal sequence at different sites. One of the chloroplast thioredoxins from spinach has 46% amino acid identity to the E. coli protein (see Table 1) and is especially efficient in reducing the chloroplast -dependent malate dehydrogenase (NADP-MDH). It has been designated thioredoxin-m (Tm). The other chloroplast thioredoxin bears only 30% amino acid identity to the E. coli protein (Table 1) and was first described as an

718 activator of the chloroplast fructose- 1,6-bisphosphatase (FBPase); thus, it is designated thioredoxin-f (Tf). These thioredoxins have been shown to reduce a number ofadditional proteins in plants. For example, Tm will stimulate the coupling factor 1-ATPase and deactivate chloroplast glucose-6phosphate dehydrogenase (G6PDH). Tf, at relatively higher concentrations, will also reduce the NADPMDH and In addition, this thioredoxin can reduce other enzymes in the C-3 cycle, including, sedoheptulose-1,7-bisphosphatase, NADP-glyceraldehyde-3-phosphate dehydrogenase and phosphoribulokinase (PRK; Crawford et al., 1988).

2. Thioredoxins in Cyanobacteria Initial studies with the cyanobacterium Anabaena sp. strain PCC 7119 (Nostoc muscorum) suggested that homologs of Tm and Tf were also present in this organism. This conclusion was based on cross reaction of partially purified fractions with the spinach chloroplast enzymes (Yee et al., 1981). An ‘m-type’ thioredoxin was purified to homogeneity from Anabaena sp. strain PCC 7119. Based on its activity as a general protein disulfide reductase and its ability to serve as a reducing agent for ribonucleotide reductase, it was assumed to be similar to the protein from E. coli (Gleason and Holmgren, 1981). Subsequent characterization of a second, ‘f-type’ thioredoxin from Anabaena sp. strain PCC 7119 by Whittaker and Gleason (1984) lead to some skepticism as to whether this protein was actually a thioredoxin. Unlike most thioredoxins which have masses of approximately 12 kDa, this protein is twice as large (~25.5 kDa). It can activate spinach FBPase but is not effective with the Anabaena sp. strain PCC 7119 enzyme nor can it reduce the disulfide bonds in insulin, a general property of all known thioredoxins (Holmgren, 1979). It can not act as a reducing agent for ribonucleotide reductase. The presence of only a single thioredoxin fraction in cyanobacteria was further suggested by the purification ofthe protein from Anabaena cylindrica strain CCAP 1403/2a. This thioredoxin has protein disulfide reductase activity characteristic of the ‘mtype.’ The amino-terminal amino acid sequence showed a high degree of similarity to that of E. coli thioredoxin. In addition, antibodies to the cyanobacterial thioredoxin reacted with only one protein in extracts of Anabaena cylindrica, ruling out the presence of a second immunologically-related

Florence K. Gleason thioredoxin (Ip et al., 1984). However, subsequent immunological studies did suggest the presence of a minor amount of another thioredoxin in Anabaena cylindrica (Darling et al., 1986).

3. Thioredoxin Genes in Cyanobacteria Initial attempts to isolate the gene for Anabaena sp, strain PCC 7119 thioredoxin also led to the conclusion that this cyanobacterium had only one thioredoxin gene. The entire E. coli thioredoxin gene was used as a probe of Anabaena sp. strain PCC 7119 genomic DNA and only one hybridizing DNA fragment was found on Southern blots. Subsequent cloning and sequencing of the gene showed that it encoded a protein that was identical to the one which had been previously characterized in Anabaena sp. strain PCC 7119 (Lim et al., 1986). The gene was inserted into a pUC plasmid vector and used to transform an E. coli strain which lacks thioredoxin (BH2012). Relatively large amounts of cyanobacterial thioredoxin could be produced in this strain. The Anabaena sp. strain PCC 7119 thioredoxin, with 48% amino acid identity to the E. coli protein, restored the wild-type phenotype in strain BH2012 except for T7 phage production indicating that the heterologous protein, when present at high concentration, could effectively replace all redox functions in the bacterium. A single gene for thioredoxin from the unicellular cyanobacterium Synechococcus sp. strain PCC 7942 was also isolated and characterized. The gene sequence coded for a protein with 47% amino acid identity to E. coli thioredoxin and 85% identity to the Anabaena sp. strain PCC 7119 protein (see Table 1). Attempts to replace the gene in Synechococcus sp. strain PCC 7942 with a defective thioredoxin construct interrupted with a kanamycin-resistance cassette yielded no transformants lacking thioredoxin. The small number of transformed cells that were obtained were found to have added the defective gene to the chromosome either upstream or downstream from the gene encoding thioredoxin (Muller and Buchanan, 1989). Thus it appears, that cyanobacteria, unlike E. coli, may require a functional thioredoxin gene and lack an alternate redox system that can substitute for the protein. Using a nucleotide probe constructed to recognize the active site coding region of the Anabaena sp. strain PCC 7119 thioredoxin gene, Alam and coworkers (1989) detected two putative thioredoxin genes in Anabaena sp. strain PCC 7120 genomic

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720 DNA. Sequencing of one of these genes showed that it encoded an unusual thioredoxin with 43% amino acid identity to the E. coli protein and only 39% identity to the Anabaena sp. strain PCC 7119 thioredoxin. Although this unusual thioredoxin could be produced in E. coli BH2012, it did not restore the wild-type phenotype. The purified protein had protein disulfide reductase activity and also served as a reducing agent for Anabaena sp. ribonucleotide reductase. It cross-reacted weakly with polyclonal antibodies to Anabaena sp. strain PCC 7119 thioredoxin but could not be detected in extracts of Anabaena sp. strain PCC 7120 on immunoblots. Subsequent production of antibodies specific to the cloned Anabaena sp. strain PCC 7120 thioredoxin clearly demonstrated that the protein did occur in Anabaena sp. strain PCC 7120 and also in strain PCC 7119. Cross-reacting protein could also be found in Synechococcus sp. strain PCC 7942 and Synechocystis sp. strain PCC 6803, but not in eucaryotic algae such as Euglena gracilis strain UTEX 753 or spinach extracts. The immunological evidence indicates that the cyanobacteria possess two thioredoxin fractions: one that is readily isolated and that is similar to the m-type thioredoxins of E. coli and spinach; and a second thioredoxin that is unique to cyanobacteria and that is produced in relatively small quantities. Additional confirmation was obtained by separation and isolation of the two thioredoxin fractions from extracts of Anabaena sp. strain PCC 7120; the more abundant protein, T1, that has extensive homology to other thioredoxins, and T2, the unusual fraction, can be separated on an ionexchange column (Gleason, 1992). Cyanobacteria thus seem to be similar to the chloroplasts of higher plants in having two distinct but related thioredoxins: one of which is highly conserved throughout evolution, and a second, more divergent protein that is perhaps unique to this class of photosynthetic organisms. Although the data are limited, this conclusion is supported by recent isolation and sequencing of two thioredoxins from Chlamydomonas reinhardtii. Thioredoxin-2 from this organism is an abundant protein with extensive sequence identity to spinach Tm and Anabaena sp. strain PCC 7119 T1 (Decottignies et al., 1990). In contrast, C. reinhardtii thioredoxin-1 is present in relatively small amounts and although it resembles other thioredoxins, it is definitely unique to this chlorophyte (Decottignies et al., 1991). The primary

Florence K. Gleason structures ofsimilar and divergent thioredoxins from photosynthetic organisms are grouped together in Table 1. Unlike the situation in most heterotrophs, it appears that the proper functioning of an oxygenevolving photosynthetic cell requires two different thioredoxins. The results of the gene replacement experiments of Muller and Buchanan (1989) also suggest that the unusual thioredoxin cannot substitute for the more abundant fraction as does glutaredoxin in the corresponding E. coli mutants. The T2 of Anabaena sp. strain PCC 7119 does have some ofthe properties of a glutaredoxin in that it can be reduced by glutathione. The in vitro reaction requires relatively high concentrations of reduced glutathione (5 mM) and its physiological significance is unknown (Gleason, 1992). It is possible that the unusual cyanobacterial thioredoxin has a somewhat less negative redox potential than T1 making it more easily reduced by glutathione. II. Structure of Cyanobacterial Thioredoxins As seen in Table 1, all the thioredoxins from photosynthetic organisms have the same active site sequence as that of E. coli, Trp-Cys-Gly-Pro-Cys. In addition to the disulfide, structurally important residues are conserved such as Pro40 and Pro76 (numbering from the E. coli sequence). As yet, there are no three-dimensional structures of the plant thioredoxins. However, given the high degree of primary structural conservation between Anabaena sp. strain PCC 7119 T1 and E. coli thioredoxins, a tertiary structure has been predicted for the cyanobacterial protein. The X-ray structural coordinates for E. coli thioredoxin were used together with molecular modeling techniques in which the residues were replaced by those present in Anabaena sp. strain PCC 7119 T1 (Eklund et al., 1991). The model shows that the E. coli thioredoxin tertiary structure is conserved in the cyanobacterial T1. As seen in Fig. 1, the protein consists ofa core of comprised of five antiparallel flanked by four Most of the non-conservative residue changes between the E. coli and Anabaena sp. thioredoxins occur on the exterior of the protein and would have little effect on the overall threedimensional structure. Only three residue changes occur in the active site region which may explain why Anabaena sp. strain PCC 7119 T1 can replace E.

Chapter 24 Thioredoxins in Cyanobacteria

coli thioredoxin in many reactions and in vivo. These amino acid substitutions include a glutamic acid at position 30 substituted with proline and the lysine at position 36 substituted with arginine as shown in Fig. 1, and glycine 74 substituted by serine which would be positioned immediately below the disulfide bridge. These same substitutions at positions 30 and 74 are also seen in the related thioredoxins from spinach, Synechococcus sp. strain PCC 7942, and Chlamydomonas reinhardtii. The substitution of the larger arginine at position 36 for a lysine residue occurs on the exterior of the protein and should have little effect on the structure of the active site. Most thioredoxins have a positively charged residue at this position that was believed to stabilize a thiolate anion on Cys32 in the reduced protein (Kallis and

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Holmgren, 1980). However, mutation of Lys36 to a glutamic acid in E. coli thioredoxin showed that this residue has little effect on redox potential and functions mainly to optimize interaction with thioredoxin reductase and probably other proteins as well (Navarro et al., 1991). One can speculate that the arginine in the cyanobacterial thioredoxin serves a similar function in optimizing protein-protein interactions. It has also been proposed that the Gly74 in E. coli thioredoxin is essential for interaction with the T7 gene 5 protein which would explain why the cyanobacterial protein with a serine substitution will not support T7 replication (Eklund et al., 1991). Similar attempts to model the unusual Anabaena sp. strain PCC 7120 T2 have not been especially successful. Although this protein probably has the

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typical thioredoxin tertiary structure shown in Fig. 1, several additional minor changes in the backbone and positions of the side chains must be made to yield a chemically reasonable model structure. This indicates that T2 has additional tertiary structural modifications which cannot be readily predicted (F. K. Gleason, unpublished results). This proposal is supported by a recent study of the stability of thioredoxins. Figure 2 shows a denaturation curve of oxidized T2 in the presence ofvarying concentrations of guanidine hydrochloride (GuHCl). The midpoint for denaturation of T2 is at 0.91 M GuHCl, as compared to a midpoint of 2.0 M and 2.4 M GuHCl obtained for Anabaena sp. strain PCC 7119 T1 and E. coli thioredoxin, respectively (R. K. Harris and F. K. Gleason, unpublished results). In general, thioredoxins are unusually stable proteins due to the large percentage of residues involved in secondary structures. The significantly lower stability of Anabaena sp. strain PCC 7120 T2 indicates that it has less highly organized secondary elements. It also accounts for the low yields ifa heat-treatment is used during purification. Ill. Reduction of Thioredoxins

A. NADPH-Dependent Reductase In E. coli, thioredoxin is reduced by NADPH in a reaction catalyzed by the flavoprotein, thioredoxin reductase. This enzyme is composed oftwo identical subunits (approximately 30 kDa each), each containing an active-site disulfide and a bound FAD (Thelander, 1968). A similar enzyme has been purified from mammalian tissues (Luthman and Holmgren, 1982) and the photosynthetic bacterium, Rhodobacter sphaeroides (Clement-Metral, 1987). Although an NADPH-dependent thioredoxin reductase has been detected in a number of plant tissues such as roots and seeds (Suske et al., 1979; Johnson et al., 1987), it was not found in photosynthetic tissues. However, careful fractionation of spinach cytosol has revealed the presence ofa flavoprotein reductase which seems to be specific for the cytosolic forms of thioredoxin (Florencio et al., 1988). NADPH-dependent reductase has also been detected in extracts of Anabaena sp., but the enzyme has not been purified (Gleason, 1986).

B. Ferredoxin-Dependent Reductase Chloroplast and cyanobacterial thioredoxins can be reduced by a ferredoxin-dependent system. The ferredoxin-thioredoxin reductase (FdTR) has been purified from Anabaena sp. PCC 7119 (Nostoc muscorum). The cyanobacterial FdTR is a [4Fe-4S] protein with a mass of 28 kDa. It is composed oftwo dissimilar subunits with masses of 14 and 7 kDa. The 14 kDa subunit is immunologically similar to the large subunit found in the enzyme isolated from higher plants, while the smaller subunit seems to be unique to each class of photosynthetic organism (Droux et al., 1987). The activity is dependent on light-reduced ferredoxin, and given the complexity of the assay system, little is known about specificity and mechanism of action. Presumably both types of thioredoxin are reduced equally well. This reaction is unusual in that ferredoxin, a one electron redox protein, transfers electrons one at a time to thioredoxin, a two electron (and two proton) carrier

Chapter 24 Thioredoxins in Cyanobacteria mediated by an Fe-S protein. Oxidized ferredoxin will form a relatively tight complex with the reductase (Hirasawa et al., 1988). The gene for the small, variable subunit of the FdTR has been isolated and characterized from Synechococcus sp. (Szekeres et al., 1991). The single-copy open reading frame encodes a protein of 73 amino acids which has no cysteine residues. Therefore, the [4Fe-4S] center and redox-active disulfide must both occur on the larger, more conserved subunit. The small subunit contains the sequence, Asn-Gly-Lys-Pro, which is the same as the ferredoxin binding site in spinach ferredoxinoxidoreductase. This suggests a mechanism in which reduced ferredoxin binds to the variable subunit, as suggested by the work of Hirasawa and coworkers (1988), and transfers its electron to the [4Fe-4S] center in the large subunit. After two electrons have been transferred, a redox-active disulfide on this subunit is reduced and can then transfer electrons and protons to thioredoxin. The FdTR provides a more direct link between lightgenerated reducing equivalents and reduction of thioredoxin than does an NADPH-dependent reduction. Reduced thioredoxin, as a protein disulfide reductase, can modulate the activity of a variety of enzymes which are indirectly regulated by light.

IV. Functions of Thioredoxins in Cyanobacteria

A. Thioredoxin as a Protein Disulfide Reductase

1. Relation to Chloroplast Thioredoxin-m Chloroplast Tm was originally isolated as the activator ofthe NADP-dependent malate dehydrogenase. This enzyme is important in C-3 plants for shuttling NADPH equivalents across the chloroplast membranes. In C-4 plants it is required in the reduction of oxaloacetate to malate in the initial fixation of into C-4 dicarboxylic acids. Thioredoxin-m has been shown to reduce a disulfide bond between Cys10 and Cys15 in the maize enzyme (Decottignies et al., 1988). However, neither of the MDH functions is important in cyanobacteria where there is no transmembrane NADPH shuttle and C-4 acids are produced mainly by the of phosphoenolpyruvate (Owttrim and Colman, 1988). MDH activity in cyanobacteria is very low and not

723 susceptible to redox activation (for example, see Ip et al., 1984). Although cyanobacterial T1 is structurally homologous to chloroplast Tm, it must have some other major function in these organisms.

2. Modulation of C-3 (Calvin) Cycle Enzymes

a. Fructose-1,6-Bisphosphatase Chloroplast FBPase is specifically activated by Tf. Tm isalsoeffective butatmuchhigherconcentrations. The sequence of the phosphatase from spinach chloroplast has been determined. It exhibits 40% amino acid identity to gluconeogenic FBPases. However, the chloroplast enzyme has a 16-residue insert that is not present in FBPases from heterotrophic organisms. This insert contains two cysteine residues separated by four amino acids. Molecular modeling suggests that these would form a disulfide bridge on the surface of the enzyme and present an obvious site for reduction by thioredoxin (Marcus and Harrsch, 1990). There is no comparable structural information on cyanobacterial FBPases. Ip and coworkers (1984) showed that FBPase in crude extracts of Anabaena cylindrica could be activated by a dithiothreitol (DTT)-reduced thioredoxin comparable to the T1type. However, T2 from Anabaena sp. strain PCC 7120 is much more active in reducing spinach chloroplast FBPase than is T1 (Gleason, 1992). Whittaker and Gleason (1984) also showed that partially purified Anabaena sp. FBPase can be fully activated at pH 8 by elevated, but physiological, concentrations of Resolution of the lightregulation of this enzyme in cyanobacteria will require additional study of the structure and activity of the FBPase.

b. Phosphoribulokinase Spinach chloroplast phosphoribulokinase contains two cysteine residues at positions 16 and 55. Reduction of a disulfide bond between these residues by Tf results in a fully activated enzyme. Cysl6 is positioned near the active site and plays a role in binding ATP (Porter et al., 1988). PRK has been isolated from the cyanobacterium, Chlorogloeopsis fritschii strain CCAP 1411/16. The enzyme is composed of similar subunits with a mass of40 kDa as is the spinach PRK. The C. fritschii enzyme

724 requires for activity and could be activated in extracts by adding DTT or reduced glutathione (Marsden and Codd, 1984). Further information on cyanobacterial PRK was obtained from the characterization of light-sensitive mutants of Synechocystis sp. strain PCC 6803 (Su and Bogorad, 1991). Mutant cells will grow photosynthetically in dim light (300–500 lux), but not in bright light (3000–5000 lux). The mutation occurs in an open-reading frame which encodes a 39 kDa polypeptide with extensive homology to spinach PRK. The amino acid sequence obtained from the gene shows approximately 60% identity to PRK from Chlamydomonas reinhardtii and higher plants but not to the enzyme found in photosynthetic bacteria (see Chapter 14). Cysteine residues at positions 19 and 41 correspond to those found in the spinach enzyme which are the site of thioredoxin redox activation. However, the bright-light sensitive mutation occurs in the codon for Ser222 which is altered to a phenylalanine. The mutant enzyme has a higher forATP, suggesting that the ATP binding site in cyanobacterial PRK may not be the same as that of the spinach enzyme. Alternatively, one can assume that significant changes occur in the tertiary structure of the enzyme after reduction. Additional information on the structure of PRK will be needed to fully describe the role of thioredoxin in the modulation of its activity.

3. Glucose Catabolism-Modulation of Glucose6-Phosphate Dehydrogenase The major pathway for glucose catabolism in cyanobacteria is via the hexose monophosphate shunt (pentose-phosphate pathway). It is also a major source of reductant (NADPH) in nitrogen-fixing species. Deactivation of the G6PDH has been reported to result from reduction of the enzyme by the m-type thioredoxin. The G6PDH has been purified from Anabaena variabilis strain ATCC 29413. It has a mass of 250 kDa and is probably a tetramer (Cossar et al., 1984). DTT-reduced thioredoxin inhibits activity but not in the presence of substrate, indicating that both compounds bind to the same or closely related sites. The gene for G6PDH has been cloned from Synechococcus sp. strain PCC 7942 (Scanlan et al., 1992). The open reading frame codes for a protein of 524 amino acids with 41% amino acid sequence identity to the protein from E. coli (Scanlan et al.,

Florence K. Gleason 1992). However, the Synechococcus sp. strain PCC 7942 G6PDH has two additional cysteine residues at positions 191 and 454 that may form the regulatory disulfide. Cysl91 is located just before the active site, and indicates a juxtaposition of the substrate and effector sites. This result agrees with results obtained with the Anabaena variabilis strain ATCC 29413 enzyme. The cloning of the gene makes possible overproduction of the protein and a more thorough study of the interaction with effectors.

4. Thioredoxin and the Enzymes of Nitrogen Metabolism a. NADP-Dependent Isocitrate Dehydrogenase Thioredoxin has been implicated in the regulation of two enzymes in cyanobacteria which function in nitrogen metabolism: isocitrate dehydrogenase and glutamine synthetase. NADP-dependent isocitrate dehydrogenase (IDH) may be the source ofreducing equivalents, (NADPH) in heterocysts that are required for the generation of reduced ferredoxin and also for production of 2-oxoglutarate for glutamine synthesis from (Bothe and Neuer, 1988). Early work by Papen and coworkers (1983) showed that the activity of IDH was higher in heterocysts than in vegetative cells of Anabaena sp. strain PCC 7119 (Nostoc muscorum) and Anabaena cylindrica. In crude extracts, the enzyme exhibited hysteretic kinetics and could be activated, presumably to a classical Michaelis-Menton type of activity, by addition of DTT and partially purified thioredoxin from A. cylindrica or by addition of spinach Tm (Papen et al., 1983). However, immunocytochemical localization of thioredoxin in A. cylindrica and A. variabilis showed that heterocysts contained far less thioredoxin than vegetative cells (Cossar et al., 1985). Recent purification of NADP-IDH from the non-nitrogen fixing Synechocystis sp. strain PCC 6803 showed no effect on enzyme activity by added DTT, Tm from spinach, or reduced glutathione (Munro-Pastor and Florencio, 1992). These same workers reported a 3–4 fold increase in IDH activity in Anabaena sp. strain PCC 7120 when the organism was grown without a source of fixed nitrogen. It seems likely that there may be two different forms of IDH, one that is constitutive such as that described in Synechocystis sp. strain PCC 6803 which is not subject to redox control, and a second form that is

Chapter 24 Thioredoxins in Cyanobacteria specific to heterocysts and may be redox regulated. This regulation may be one of the functions of cyanobacterial T2, which was not detected by the immunological studies cited above.

b. Glutamine Synthetase Glutamine synthetase (GS) from Anabaena cylindrica can be activated by cyanobacterial T1 or spinach Tm. The results of GS reduction are obvious at pH 7, but at pH 8, the enzyme is fully activated by high but physiological concentrations of (Papen and Bothe, 1984). This is similar to results obtained with the FBPase ofAnabaena sp. strain PCC 7120. GS in the unicellular Synechococcus sp. strain PCC 6301 also responds to activation by light. Although this suggests redox control ofthe enzyme, no reactivation of dark-inactivated enzyme was found with added DTT or spinach Tm (Marques, et al., 1992). Clarification of a role for thioredoxin, if any, in this system will require further biochemical characterization of GS.

B. Thioredoxin as a Reducing Agent in Cyanobacteria 1.3'-Phosphoadenosine-5'-Phosphosulfate Reductase Cyanobacteria, like other bacteria and plants, take up sulfur as and reduce it to sulfide which is then incorporated into cysteine. Several reports suggested that cyanobacteria are similar to higher plants in that sulfate is carried as adenylyl sulfate (APS) and

725

reduced to sulfite and sulfide while bound to a carrier, perhaps thioredoxin (for a review, see Anderson, 1990). In contrast, enterobacteria and yeast produce PAPS from sulfate and 3'-phosphoadenosine-5'phosphate which is subsequently reduced to free sulfite by a reaction which uses thioredoxin as a reducing agent (see Kredich et al., 1987). However, using the cysH gene from E. coli, which codes for PAPS reductase, as a probe, Niehaus and coworkers detected an homologous gene in Synechococcus sp. strain PCC 7942 (Niehaus et al., 1992). Comparison of the derived amino acid sequence from the cyanobacterial gene with PAPS reductase from E. coli showed 58% amino acid identity including a conserved cysteine residue near the carboxylterminal. Moreover, the cyanobacterial gene will substitute for that from E. coli in cysH mutants. Southern blot analysis by Schwenn’s group also showed gene sequences similar to the cysH gene in Synechocystis sp. and the photosynthetic bacterium, Rhodospirillum rubrum (Gisselmann et al., 1992). It thus appears that photosynthetic organisms use the same mechanism of sulfate reduction as previously established in E. coli and yeast. Sulfate is bound to the 5'-phosphate ofADP and reduced to sulfite. The reaction is catalyzed by PAPS reductase dimer and thioredoxin serves as the reducing agent (Krone et al., 1991). By analogy to the E. coli system, cyanobacterial T1 probably functions in this reaction although T2 may also substitute. In mutants of E. coli that lack both thioredoxin and glutaredoxin, cysteine must be added to the medium for cells to survive. Either protein will serve as reducing agent in the PAPS

Florence K. Gleason

726 reductase system but this is an essential function of these proteins in E. coli. Ribonucleotide reduction, on the other hand, is not inhibited in double mutants indicating an additional reducing system can be utilized for this reaction (Russel et al., 1990). Given the homology between E. coli and cyanobacterial thioredoxins and the glutathione reduction of T2, it seems likely that a similar mechanism operates in cyanobacteria at least with respect to sulfate metabolism.

2. Ribonucleotide Reduction Thioredoxin-1 from Anabaena sp. strain PCC 7119 was isolated on the basis of insulin disulfide reductase activity. It was also shown to be an effective reducing agent for both E. coli and Anabaena sp. strain PCC 7119 ribonucleotide reductases (RNRase). The reductases differ considerably between the two organisms. E. coli RNRase is composed of two different subunits—one containing iron and a tyrosyl free radical—while the other subunit contains substrate, effector, and redox binding sites. The RNRase from Anabaena sp. strain PCC 7119 has only one polypeptide chain with the iron protein functionally replaced by adenosylcobalamin (Coenzyme Gleason and Frick, 1980). This latter form of the enzyme is more common in procaryotes, particularly the cyanobacteria (Gleason and Wood, 1976). There is no information as yet on the structure of cyanobacterial RNRases. Analysis of the E. coli enzyme and the Coenzyme reductase from Lactobacillus leichmannii, show that either thioredoxin or glutaredoxin can reduce a disulfide on the surface of the substrate-binding polypeptide (Åberg et al., 1989; Lin et al., 1987). There is then an intramolecular relay of reducing equivalents to a disulfide at the active site. In E. coli, glutaredoxin is a more efficient reducing agent than thioredoxin. In Anabaena sp. strain PCC 7120 both T1 and T2 serve this function, but T1 has approximately 10-fold greater catalytic efficiency (Alam et al., 1989). This suggests that T1 is the usual reducing agent in this reaction in vivo. No glutaredoxin has been described in cyanobacteria, but T2 may be the equivalent.

C. What is the Essential Function of Thioredoxin in Cyanobacteria? Unlike the situation in E. coli, the results of Muller and Buchanan (1989) have shown that mutants of the

obligate photoautotroph, Synechococcus sp. strain PCC 7942 are non-viable if they lack thioredoxin 1. These workers have proposed that this is due to an essential regulatory role of thioredoxin in light/dark modulation of carbon metabolism—particularly the G6PDH reaction. However, like E. coli, cyanobacteria may have a back up system in thioredoxin 2. It has similar activity to T1 in vitro, i.e., it serves as a reducing agent and protein disulfide reductase. It may not be able to replace T1 in vivo, perhaps due to its altered specificity which is as yet unknown or its instability which suggests a high turnover and contributes to its low concentration in cells. The role of thioredoxin in carbon metabolism might be better explored in a heterotrophic cyanobacterium such as Synechocystis sp. strain 6803 or in conditionallylethal thioredoxin mutants of cyanobacteria. Alternatively, given the ease of transformation of some cyanobacteria, thioredoxin could be replaced by a protein with specific residues altered by sitedirected mutagenesis to produce a less-efficient reducing agent. Given that two dissimilar thioredoxins seem to occur in all photosynthetic organisms from cyanobacteria to chloroplasts of angiosperms, it seems quite likely that this protein has some essential and unique role in the photosynthetic life style. Alternative essential functions may now be postulated—e.g., an essential role in sulfate reduction. Future studies should reveal the role of these important and widespread redox proteins. References Åberg A, Hahne S, Karlsson M, Lursson Å, Örmo M, Ahgren A and Sjöberg B-M (1989) Evidence for two different classes of redox-active cysteines in ribonucleotide reductase from Escherichia coli. J Biol Chem 264: 12249–12252 Alam J, Curtis S, Gleason FK, Gerami-Nejad M and Fuchs JA (1989) Isolation, sequence and expression in Escherichia coli of an unusual thioredoxin gene from the cyanobacterium Anabaena sp. strain PCC 7120. J Bacteriol 171: 162–171 Allen JF, Sanders CE and Holmes NG (1985) Correlation of membrane protein phosphoryhition with excitation energy distribution in the cyanobacterium Synechococcus 6301. FEBS Lett 193: 271–275 Anderson JW (1990) Sulfur metabolism in plants. In: Stumpf PK and Conn, EE (eds) The Biochemistry of Plants, Vol 16, pp 327–381. Academic Press, San Diego Bestermann A, Vogt F and Follmann H (1983) Plant seeds contain several thioredoxins of regular size. Eur J Biochem 131: 339–344 Bothe H and Neuer G (1988) Electron donation to nitrogenase in heterocysts. Meth Enzymol 167 496–501

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Florence K. Gleason Owttrim GW and Colman, B (1988) Phosphoenol pyruvate carboxylase mediated carbon flow in a cyanobacterium. Biochem Cell Biol 66: 93–99 Pace CN (1980) Determination and analysis of urea and guanidine hydrochloride denaturation curves. Meth Enzymol 131: 266– 280 Papen H and Bothe H (1984) The activation of glutamine synthetase from the cyanobacterium Anabaena cylindrica by thioredoxin. FEMS Microbiol Lett 23: 41–46 Papen H, Neuer G Refaian M and Bothe H (1983) The isocitrate dehydrogenase from cyanobacteria. Arch Microbiol 134:73–79 Pigiet VP and Schuster BJ (1986) Thioredoxin-catalyzed refolding of disulfide-containing proteins. Proc Natl Acad Sci USA 83: 7643–7647 Porter MA, Stringer CD and Hartman FC (1988) Characterization of the regulatory thioredoxin site of phosphoribulokinase. J Biol Chem 263: 123–129 Portis AR (1992) Regulation of ribulose 1,5-bisphosphate carboxylase/oxygenase activity. Annu Rev Plant Physiol Mol Biol 43: 415–437 Russel M and Model P (1985) Thioredoxin is required for filamentous phage assembly. Proc Natl Acad Sci. USA 82: 29–33 Russel M, Model P and Holmgren A (1990) Thioredoxin or glutaredoxin in Escherichia coli is essential for sulfate reduction but not for deoxynucleotide synthesis. J Bacteriol 172: 1923– 1929 Scanlan DJ, Newman J, Sebaihia M, Mann NH and Carr, NG (1992) Cloning and sequence analysis ofa glucose-6-phosphate dehydrogenase gene from the cyanobacterium Synechococcus PCC 7942. Plant Mol Biol 19: 877–880 Sodano P, Xia TH, Bushweller JH, Björnberg O, Holmgren A, Billeter M and Wuthrich K (1991) Sequence-specific H-1 NMR assignments and determination of the three-dimensional structure of reduced Escherichia coli glutaredoxin. J Mol Biol 221: 1311–1324 Su X and Bogorad L (1991) A residue substitution in phosphoribulokinase of Synechocystis PCC 6803 render the mutant light sensitive. J Biol Chem 266: 23698–23705 Suske G, Wagner W and Follmann H (1979) N ADPH-dependent thioredoxin reductase and a new thioredoxin from wheat. Z Naturforsch 34c: 214–221 Szekeres M, Droux M and Buchanan BB (1991) The ferredoxinthioredoxin reductase variable subunit gene from Anacystis nidulans. J Bacteriol 173: 1821–1823 Tabor S, Huber HE and Richardson CC (1987) Escherichia coli thioredoxin confers processivity on the DNA polymerase

activity of the gene 5 protein of bacteriophage T7. J Biol Chem 262: 16212–16223 Thelander L (1968) Studies on thioredoxin reductase from Escherichia coli B. The relation of structure and function. Eur J Biochem 4:407–422 Wedel N, Clausmeyer S, Herrmann RG, Gardet-Salvi L and Schurmann P (1992) Nucleotide sequence of cDNAs encoding the entire precursorpolypeptide of thioredoxin m from spinach chloroplasts. Plant Mol Biol 18: 527–533 Whittaker MM and Gleason FK (1984) Isolation and characterization of thioredoxin f from the filamentous cyanobacterium, Anabaena sp. 7119. J Biol Chem 259: 14088–14093

Chapter 24 Thioredoxins in Cyanobacteria Wilson LG, Asahi T and Bandurski RS (1961) Yeast sulfatereducing systems. T. Reduction of sulfate to sulfite. J Biol Chem 236: 1822–1829 Yang Y and Wells WW (1991) Identification and characterization of the functional amino acids at the active center of pig liver thiol transferase by site-directed mutagenesis. J Biol Chem

729 266: 12759–12765 Yee BC, DeLaTorre A, Crawford NA, Lara C, Carlson DE, and Buchanan BB (1981) The ferredoxin/thioredoxin system of enzyme regulation in a cyanobacterium. Arch Microbiol 130: 14–18

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Chapter 25 Iron Deprivation: Physiology and Gene Regulation Neil A. Straus Department of Botany, University of Toronto, Toronto, Ontario, Canada M5S 3B2

Summary I. Introduction II. Iron in Photosynthetic Electron Transport III. Responses to Iron Deprivation A. Retrenchment 1. Ultrastructural Symptoms a. Iron in the Synthesis of Phycobiliproteins 2. Physiological Response B. Compensation 1. Ferredoxin and Flavodoxin a. Iron-Dependent Appearance of Ferredoxin and Flavodoxin b. Gene Regulation 2. An Iron-Stress-Induced, Chlorophyll-Binding Protein a. The isiA Gene C. Acquisition 1. Siderophores: Iron-Specific Chelators 2. Iron Transport in E. coli 3. Iron Transport in Cyanobacteria a. Siderophore Production and Nitrogen Fixation IV. The Control of Gene Expression by Iron A. The Fur Represser of E. coli B. Iron-Regulated Genes in Cyanobacteria Acknowledgments References

731 732 732 734 734 734 734 735 735 735 736 737 738 738 740 740 740 741 743 743 743 745 745 747

Summary Iron is an essential redox component of most living organisms. It is particularly important to photoautotrophs, such as cyanobacteria, in which it is a crucial part ofthe protein complexes involved in photosynthetic electron transport. However, in oxic ecosystems, the low solubility of above neutral pH limits the biological availability of iron to aquatic microorganisms, that must double their iron content with every round of cell division. As a result, cyanobacteria and other microorganisms have evolved a number of responses to cope with frequently occurring conditions of iron deficiency. These responses include a system of genetic regulation that alters the profile of soluble electron carriers in advance of extreme iron deprivation, that induces the synthesis of an efficient iron-scavenging system and that may also differentially control the loss of iron-requiring physiological structures which become expendable during extreme iron deprivation.

D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 731–750. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

732 I. Introduction With the exception of some lactobacilli, iron is an essential requirement of all organisms (Archibald, 1983). As an essential component of heme and iron sulfur centers in a variety of proteins from cytochromestoferredoxins, itformsversatile electron transfer agents whose redox potentials span a range of about 1000 mV These proteins are important components ofbasic physiological processes such as photosynthesis, respiration, nitrogen assimilation, dinitrogen fixation, chromophore biosynthesis and deoxyribonucleotide synthesis. From superficial considerations, one would not expect iron acquisition to be a problem for most organisms, since iron is the fourth most abundant element by weight in the Earth’s crust. However, in aqueous oxic environments isquicklyoxidized to At physiological pHs, forms insoluble hydroxides, severely reducing the biological availability of iron. For example, at pH 7 the free concentration is only about ions per ml which is substantially lower than the to ions required by a single bacterial cell (Braun et al., 1990). This limitation in iron availability results in a major environmental challenge for microbial organisms which must double the amount of iron ions they assimilate with every round ofcell division. Because of this, efficient iron acquisition is essential to microbial survival. The selective pressure of limited iron availability has led to sophisticated systems of iron acquisition involving iron-binding molecules called siderophores and a vigorous competition between organisms for environmental iron. This competition can turn particularly nasty for eucaryotes since some virulent bacteria produce toxins during iron deprivation in a less than subtle effort to acquire iron from a stingy host (Braun, 1985; Crosa, 1989). Different aspects of iron transport have been reviewed very frequently over the years; because of the large numbers, an arbitrary selection is referenced in this article. Reviews on bacterial systems for iron acquisition are particularly noteworthy because of the incredible advances that have been made in unraveling the regulation of gene expression in Abbreviations: ATCC – American Type Culture Collection; CP – chlorophyll protein; EDDA – ethylene diamine di-ohydroxyphenyl acetic acid; IU – Indiana University Culture Collection; PCC–Pasteur Culture Collection; UTEX– University of Texas Culture Collection

Neil A. Straus response to iron stress. Many reviews have focused on the detection, chemistry and function ofbacterial siderophores, iron-specific biological ligands, (for examples see: Neilands, 1981a, 1981b, 1984; Hider, 1984; Braun and Winkelmann, 1987; Matzanke, 1991). Other reviews cover the molecular biology and molecular genetic aspects of siderophore production and iron transport systems (for examples see: Neilands, 1982; Bagg and Neilands, 1987; Braun et al., 1987; Crosa, 1989; Braun et al., 1990; Braun and Hanke, 1991). Since iron chelation and uptake by cyanobacteria has been reviewed by Boyer et al. in 1987, this review will focus more on the physiological implications of iron deprivation, particularly those affecting photosynthesis, and on what has been learned about cyanobacteria undergoing iron stress through molecular biology and molecular genetics. II. Iron in Photosynthetic Electron Transport Since most cyanobacteria are obligate photoautotrophs, the photosynthetic electron transport pathway performs the most important energytransducing reactions in the cell. Indeed, it is this nearly complete dependence on light energy that distinguishes the biochemical strategy of cyanobacteria from most other bacteria. Although a large part of this review focuses on the effects of iron deficiency on photosynthetic electron transport, it should be noted that this approach also covers a significant part of the respiratory pathway, since much ofthe respiratory electron transport pathway is a subset of the photosynthetic pathway (located between NADH dehydrogenase and cytochrome oxidase (Peschek and Schmetterer, 1982; Alpes et al., 1984; see Chapter 13). Iron plays a major role in the photosynthetic electron transport chain in cyanobacteria. The components of the photosynthetic pathway of cyanobacteria have been reviewed in depth by Bryant (1986) and are individually reviewed in different chapters of this book. Although iron is not a component of the light-harvesting phycobilisome complex, it plays an important role in the synthesis of the phycobilin chromophores of their constituent phycobiliproteins (see section III, A.1a). Likewise, iron is not a component of the oxygen-evolving system. However, iron is a major constituent of all the remaining protein complexes ofthe photosynthetic pathway from PS II to ferredoxin, the terminal

Chapter 25 Iron Stress, fur Repressor, Gene Regulation electron acceptor on the reducing side of PS I. The positions of these iron ions will be briefly overviewed because iron deprivation has a marked effect on the photosynthetic apparatus and because a number of iron-regulated genes are part of the photosynthetic pathway. The organization and function ofthe components of PS II and its associated light-harvesting complexes have been recently reviewed in detail for both eucaryotes and cyanobacteria (Vermaas and Ikeuchi, 1991; see Chapter 8). The major protein components of PS II include: the reaction center proteins D1 and D2, the chlorophyll-binding proteins CP47 and CP43 and cytochrome Primary charge separation has been observed in D1-D2-cytochrome complexes isolated from chloroplasts and cyanobacteria (Nanba and Satoh, 1987; Danelius et al., 1987; Takahashi et al., 1987; Gounaris et al., 1989), indicating that CP47 and CP43 are probably antenna proteins responsible for the transfer of excitation energy to P680. CP47 appears to be more closely associated with the reaction center than CP43 since CP43 is more readily dissociated from the complex (Yamaguchi et al., 1988; Ghanotakis et al. 1989). However, both CP43 and CP47 appear to be required for stable PS II assembly in the cell, since psbB and psbC mutants of Synechocystis sp. strain PCC 6803 that lack either CP47 or CP43 are not able to grow photoautotrophically (Vermaas et al., 1988; Yu and Vermaas, 1990; Rögner et al., 1991). Cytochrome is thought to form a heterodimer of two polypeptides encoded by the psbE and psbF genes. Each polypeptide contains a single histidine that participates in the binding of the heme of cytochrome Cytochrome appears to play an essential role in photosynthesis since deletion mutations of the genes encoding cytochrome in Synechocystis sp. strain PCC 6803 result in a loss of PS II activity (Pakrasi et al., 1988, 1989). However, there is some question about the number of cytochromes in the core complex. Although some results indicate two cytochromes per P680, others favor only one (Miyazaki et al., 1989). PS II also contains a nonheme iron between theprimary electron accepting quinone that is bound to D2, and the plastoquinone that binds to D1 and dissociates upon complete reduction. The function of the nonheme iron is not known and no redox turnover has been observed under physiological conditions. However, the presence of the nonheme iron is essential since its extraction or replacement with other ions

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results in a loss of PS II activity (Vermaas and Ikeuchi, 1991). Electrons passed to plastoquinone from PS II are subsequently passed to the cytochrome complex. This complex is shared by both the photosynthetic and respiratory pathways in cyanobacteria. The structure and function of this complex is reviewed by Kallas in Chapter 9; the cyanobacterial complex and the related complexes in otherphotosynthetic bacteria, chloroplasts and mitochondria, have been reviewed previously (Hauska et al., 1983; Hauska, 1985). The complex functions as an electrogenic proton translocator; it passes electrons from plastoquinone to plastocyanin or cytochrome and translocates protons across the thylakoid membrane in the process thus contributing to the protonmotive force thatdrives ATP synthesis (Hurt et al., 1982). It consists of four major subunits: cytochrome which contains two hemes; cytochrome f which is a c-type cytochrome with one heme; a Rieske Fe-S protein that has a 2Fe2S center; and a 17 kDa polypeptide frequently called subunit IV. Finally PS I catalyzes the photo-oxidation of plastocyanin/cytochrome and the photoreduction of ferredoxin. The structure and function of this complex is reviewed by Golbeck in Chapter 10 and has been recently reviewed by Golbeck and Bryant (1991), Chitnis and Nelson (1991), and Bryant(1992). PS I contains 11–12 protein subunits and three 4Fe4S centers t and that participate in electron transfer within the complex. Thus, a minimum of 21–23 iron atoms are needed for a functional photosynthetic apparatus in cyanobacteria under normal nutrient conditions. PS II requires two or three irons depending on the number ofcytochromes that actually occur in the PS II complex. The cytochrome complex has five iron atoms, PS I contains twelve iron atoms per P700, and the terminal electron acceptor ferredoxin has two irons in its single 2Fe-2S center. The need for the 23rd iron depends on whether the cyanobacterium uses plastocyanin or cytochrome to shuttle electrons between the cytochrome complex and PS I; this issue is covered in more detail by Morand et al. in Chapter 12. Since the complexes and soluble carriers do not exist in 1:1 molar ratios, a physiologically meaningful minimal ‘photosynthetic unit’ would probably require an even greater number of iron atoms.

734 III. Responses to Iron Deprivation Studies on the effects of iron deprivation reveal a variety ofresponses that range from the reduction or loss of structures or molecules that are important to the cell under non-stress conditions to the synthesis of new molecules. For the sake of discussion, these responses will be divided into three different categories: retrenchment, compensation and acquisition. Retrenchment is characterized by symptoms of iron stress including alterations and reduction in cellular structures and physiological activity. Many of these symptoms have been studied in cells undergoing extreme iron deprivation. As a result, this response is often characterized by large decreases in the amount of iron-containing molecules and molecules whose synthesis is directly or indirectly dependent on iron. Compensation is characterized by the production of new proteins or increased amounts of proteins that are capable of performing the function of proteins that are present in reduced amounts during iron stress. Acquisition is a response by which the cell greatly enhances its ability to scavenge iron from the environment by producing special iron-chelating molecules called siderophores.

A. Retrenchment 1. Ultrastructural Symptoms There have been three detailed studies on the ultrastructural symptoms of iron deprivation. Calothrix parietina cultures grown under iron stress showed an increase in heterocyst frequency, the formation of colorless multicellular hairs and the development offalse branching (Douglas et al., 1986). Vegetative cells in the basal part of the trichome developed small intrathylakoidal vacuoles and showed several forms of membrane degradation. Vegetative cells exhibited no change in the carboxysome content, a decrease in the content of cyanophycean granules, and a large increase in polyglucoside granules. Surprisingly, although the chlorophyll content on a dry weight basis decreased, the yield as estimated by dry weight showed very little change from control cultures (Douglas et al., 1986). The structures of control and iron-starved cells of Synechococcus sp. strain PCC 7942 were compared, and changes during temporal recovery from iron deprivation were followed by Sherman and Sherman

Neil A. Straus (1983). Although iron-stressed cells doubled at approximately the same rate as control cells, they were only one-half to two-thirds as long. Electron micrographs showed decreases in the quantities of membranes, phycobilisomes and carboxysomes, and large increases in glycogen storage granules in ironstressed cells compared to cells grown in iron-replete medium. If iron was returned to the medium, the number of carboxysomes increased within a sixhour period by which time there was evidence of phycobilisome assembly (Sherman and Sherman, 1983). Ultrastructural studies on iron deficiency in Synechococcus sp. strain PCC 7002 documented the appearance of iron-stress symptoms over a 200-hour period. Polysaccharide granules began to accumulate at the onset of iron deficiency; this was followed by ribosome degradation and later, by the degradation ofmembranes. However, iron starvation appeared to have no effect on carboxysomes (Hardie et al., 1983b).

a. Iron in the Synthesis of Phycobiliproteins The studies by Sherman and Sherman (1983) and Hardie et al. (1983) noted that iron-stress symptoms, such as the loss of phycobilisomes and the accumulation of glycogen, were also symptoms of nitrogen starvation which they suggested might result from iron deprivation. However, at this point it is not clear that the above-described symptoms result from a physiologically induced nitrogen-stress condition resulting from iron deprivation. Evidence on the effect ofiron stress on nitrogen assimilatory enzymes appears to be contradictory. Peschek (1979) reported that iron deficiency in Synechococcus sp. strain PCC 6301 resulted in substantial reductions in the enzymatic activities of both nitrate reductase and nitrite reductase; however, in studies on Synechococcus sp. strain PCC 7002, Hardie et al. (1983a) reported initial increases in the activity of these enzymes followed by a steady decrease. In addition, symptoms such as phycobilisome loss may be more directly affected by heme availability for the synthesis of the phycobiliprotein chromophores than by other indirect factors. In support of the notion that ironstressed cultures were nitrogen-limited, Gasparich (1989) found that periodic addition of nitrogen to iron-stressed cultures of Synechococcus sp. strain PCC 7002 prevented most of the phycobiliprotein and chlorophyll losses due to long-term iron limitation.

Chapter 25 Iron Stress, fur Repressor, Gene Regulation The biosynthesis of phycobilins is reviewed in detail by Beale in Chapter 17. Phycocyanobilin and phycoerythrobilin aresynthesized by theferredoxindependent reduction of biliverdin, which is formed by the oxidative ring-opening activity of heme oxygenase on protoheme (Beale and Cornejo, 1983, 1984, 199 l; Rhie and Beale, 1992). Therefore, heme oxygenase is the first enzyme in a heme-dependent pathway dedicated to the production of phycobiliprotein chromophores. There are now a number of lines of evidence that indicate heme availability, or the production of the chromophores themselves, may play a role in the regulation of phycobiliprotein biosynthesis. Troxler et al. (1989) showed that heme and heme precursors were capable of inducing the appearance of allophycocyanin and phycocyanin mRNAs and apoproteins, which were normally absent in dark grown cells of red alga Cyanidium caldarium. Recently, cpcE and cpcF have been identified as the genes responsible for the covalent attachment of phycocyanobilin to the phycocyanin a subunit apoprotein (Fairchild et al., 1992). Mutants, in which either cpcE, cpcF, or both genes were insertionally inactivated, failed to accumulate normal levels of phycocyanin (Zhou et al., 1992). If lack of chromophore attachment results in reduced levels of phycocyanin, it would appear reasonable then that reduced chromophore synthesis during iron stress could also result in a reduction of phycobiliproteins. Since heme oxygenase is the first enzyme in a heme- and ferredoxin-dependent pathway for chromophore synthesis, it is tempting to assume that the gene for heme oxygenase might be regulated by iron availability. Such a regulatory step would preserve heme for cytochrome synthesis and reduce the demand for ferredoxin. This model will be easily tested once the gene for heme oxygenase is isolated.

2. Physiological Response Iron deficiency has been shown to induce a number of alterations in the photosynthetic physiology of cyanobacteria. Studies on iron-stressed Synechococcus sp. strain PCC 6301 and Synechococcus cedrorum (Synechococcus sp. strain PCC 6908) revealed large increases in initial and maximal fluorescence at room temperature and dramatic alterations in the low temperature fluorescence profile (Öquist, 1974b; Guikema and Sherman, 1983;

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Guikema, 1985). Absorption studies on whole cells and phycocyanin-free lamellae of Synechococcus sp. strain PCC 6301 showed an iron-deficiency induced blue shift of 5–6 nanometers in the main red chlorophyll absorption band (Öquist, 1971, 1974a; Guikema and Sherman, 1983). Öquist attributed this shift to a different distribution ofin vivo chlorophyll a forms in PS I. However, recently a similar blue shift was reported for isolated PS II complexes from a mutant of Synechocystis sp. strain PCC 6803 that lacks CP43 (Rögner et al., 1991). Since chlorophyll binding proteins of PS II appear to have a characteristic iron-stress response (Section III, B 2), it seems reasonable that the cause of this spectral shift may well come from alterations to PS II. In any case the question merits reexamination. Physiological observations in Synechococcus sp. strains PCC 7002, PCC 6301, PCC 6908, and PCC 7942, and Synechocystis sp. strain PCC 6714 that parallel the ultrastructural observations for cyanobacterial cells under iron stress include reduced contents of phycocyanin and chlorophyll (Hardie et al., 1983a; Guikema and Sherman, 1983; Sandmann, 1985). Additionally, studies on Synechocystis sp. strain PCC 6714 showed iron-stress-induced decreases in all normal components of photosynthetic electron transport that were examined, including cytochrome of PS II, cytochromes b and/of the cytochrome complex and all the Fe-S centers (especially those of PS I); in these studies, EPR spectroscopy permitted the identification of all the Fe-S centers involved in photosynthetic electron transport (Sandmann and Malkin, 1983; Sandmann, 1985). Recent studies on Anabaena sp. strain PCC 6309, a cyanobacterium that is unable to synthesize flavodoxin, examined the dry-weight content of chlorophyll a, phycocyanin, ferredoxin I and ferredoxin II in cells grown at five different iron concentrations from to The report showed parallel decreases in chlorophyll a and phycocyanin content. There were also substantial decreases in both ferredoxins; ferredoxin I decreased more rapidly than ferredoxin II (Pardo et al., 1990).

B. Compensation 1. Ferredoxin and Flavodoxin Numerous studies have reported substantial decreases in the content of ferredoxin at ‘intermediate levels’

736 of iron stress. However, because many of these studies also reported a reciprocal production of flavodoxin, that can replace ferredoxin in many redox reactions, thedecline inferredoxin andthereciprocalproduction of flavodoxin have been viewed as compensatory responses. By substituting for ferredoxin, flavodoxin functionally relieves the cell of the iron demand required by the iron-sulfur centers offerredoxin. The assumption that this is a programmed compensatory response seems reasonable because all ofthe reported transitions occurred at levels of iron above those that cause the more drastic symptoms of iron starvation.

a. Iron-Dependent Appearance of Ferredoxin and Flavodoxin The appearance of flavodoxin (phytoflavin) in response to increasing iron stress was first observed in Synechococcus sp. strain PCC 6301 (Smillie, 1965a, 1965b). This and subsequent work demonstrated that flavodoxin, which was isolated from iron-stressed cells, was able to replace ferredoxin in the photoreduction of NADP (Entsch and Smillie, 1972). Subsequent work, mostly in non-photosynthetic bacteria indicated that the production of flavodoxin in response to iron stress was quite a general bacterial response and that flavodoxin was capable of substituting for ferredoxin in a range of reactions (Yoch and Valentine, 1972; Mayhew and Ludwig, 1975; Tollin and Edmondson, 1980). The structures and functions of flavodoxin and ferredoxin are discussed in more detail by Morand, Ho and Krogmann in Chapter 12. The first careful documentation of the reciprocal accumulation of flavodoxin and decrease in the levels of ferredoxin was reported by Hutber et al. in 1977. They systematically studied the dry weight concentrations of ferredoxin I, ferredoxin II and flavodoxin in Nostoc sp. strain MAC (PCC 8009) that was grown in media with different iron concentrations. They found that the levels of both ferredoxins I and II remained fairly constant as the concentration of iron in the culture medium was decreased to about Below this iron level the concentration of both ferredoxins decreased rapidly; most of the decrease occurred in a fairly narrow window of iron concentration—between and The content of flavodoxin, on the other hand, was negligible under iron-replete conditions and only began increasing around flavodxin content began to level off around

Neil A. Straus In Synechocystis sp. strain PCC 6714, the decrease in ferredoxin preceded all other observed iron stress effects and occurred at moderate iron deficiency, as opposed to extreme iron deficiency at (Sandmann and Malkin, 1983). In addition, the decrease in ferredoxin was paralleled by an increase in flavodoxin concentration that achieved a concentration level more than twice that offerredoxin under iron-replete conditions. Unlike the previous study which measured dry-weight concentrations, this study measured the concentration relative to the chlorophyll content, which also decreases under iron stress. Since most ofthe changes in the concentrations of ferredoxin and flavodoxin occur at intermediate iron-stress levels and since the more severe changes in glycogen accumulation, cell size and chlorophyll reduction occur at much lower iron concentrations, the major discrepancy between these two methods would appear as differences in end-point concentrations. More recently, Sandmann et al. (1990) studied the iron-dependent formation of ferredoxin and flavodoxin in three different Anabaena sp. strains: ATCC 29413, ATCC 29211 (PCC 6309) and ATCC 29151 (PCC7119). When Anabaena sp. ATCC 29413 was exposed to decreasing concentrations of iron, there were large decreases in the amount offerredoxin and corresponding increases in the amount of flavodoxin. Most of the reciprocal change between ferredoxin and flavodoxin occurred between and On the other hand Anabaena sp. strain PCC 6309, which is incapable of synthesizing flavodoxin (Pardo et al., 1990), appeared to loose most of its ferredoxin at lower iron concentrations, between and In Anabaena sp. strain ATCC 29413, a comparison of oxygen evolution and nitrogen fixation midway through the transition from ferredoxin to flavodoxin indicated that photosynthetic activity was about the same but nitrogen fixation decreased to less that halfthe control rate. Since ferredoxin and flavodoxin exhibited the same values as electron donors to nitrogenase (Fillat et al., 1988; Sandmann et al., 1990), the authors concluded that iron concentration may affect the concentration of nitrogenase. Measurements of flavodoxin content in the heterocysts of Anabaena sp. strain ATCC 29151 (PCC 7119) indicated that flavodoxin was constitutively produced in the heterocysts of this species; the authors suggested that this may be a more general observation for heterocystous cyanobacteria.

Chapter 25 Iron Stress, fur Repressor, Gene Regulation

b. Gene Regulation The gene for ferredoxin I has been cloned from Synechococcus sp. strain PCC 7942 (Reith et al., 1986; Van der Plas et al., 1986). Southern-blot hybridizations indicated that the genome contains only one copy of this gene. Transcriptional analysis using Northern-blot hybridizations indicated that the gene was transcribed into a monocistronic message of about 450 nucleotides; the open reading frame coding for ferredoxin contains 297 nucleotide pairs (Reith et al., 1986). The first molecular genetic study on the comparative regulation ofgene expression for ferredoxin and flavodoxin was done using Synechococcus sp. strain PCC 7942 (Laudenbach et al., 1988). The gene for flavodoxin (isiB or petI) was cloned and Southern-blot hybridizations under reduced stringency indicated only one copy of the gene per Synechococcus sp. strain PCC 7942 genome. Transcript analysis indicated that the gene for flavodoxin was the second open reading frame (513 nucleotide pairs) on an iron-stress-induced (isi) mRNA that was about 1900 nucleotides long. Transcript analysis showed that the operon containing flavodoxin actually produced two messages and that a shorter message of about 1100 nucleotide was about 7–9 times more abundant than the message containing flavodoxin. The transcriptional patterns of the genes for ferredoxin and flavodoxin were compared at different iron concentrations and at different time intervals after the addition of iron to iron-deficient media. The RNA/DNA hybridization patterns indicated that ferredoxin is constitutively transcribed but that the transcription of flavodoxin is tightly controlled by iron concentration. No mRNA for flavodoxin could be detected in iron-replete conditions. The messages transcribed by the flavodoxin operon rapidly disappeared after the addition of iron to the iron-deficient medium; all of the messages were degraded within one hour of the addition of iron. Studies on Synechococcus sp. strain PCC 7942 by Van der Plas et al. (1988) confirmed that the concentration offerredoxin was drastically decreased in iron-stressed cells; immunoblot analyses indicated a 30-fold reduction in the amount of ferredoxin in cells grown under iron stress compared to those grown in iron-replete medium. This study noted only a moderate decrease in the abundance of iron transcripts under iron stress. The combined results of both regulatory studies indicate that, in Synecho-

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coccus sp. strain PCC 7942, the expression of flavodoxin is tightly regulated at the transcriptional level while the appearance of ferredoxin is regulated post-transcriptionally. Recent results indicate that the post-transcriptional regulation offerredoxin may be controlled by mRNA stability (Bovy et al., 1993). More recently the organization and expression of the genes for ferredoxin and flavodoxin have been studied in the unicellular marine cyanobacterium, Synechococcus sp. strain PCC 7002 (Leonhardt and Straus, 1992). In this organism the isiB gene for flavodoxin is organized in the same way as was shown for Synechococcus sp. strain PCC 7942; it is the second open reading frame in a dicistronic operon that also codes for a putative PS II chlorophyllbinding protein (Laudenbach and Straus, 1988). The transcriptional regulation of this operon, the production of two mRNAs, the sizes of the two transcripts, and the relative abundance ofthe different messages closely parallels the situation observed in Synechococcus sp. strain PCC 7942. DNA sequencing, Southern-blot hybridization and transcript analysis indicated that, as in Synechococcus sp. strain PCC 7942, the petFI gene for ferredoxin I is encoded in a single monocistronic operon that is constitutively expressed in different iron regimes. Continuing work indicates that the chromosomal organization of the isiAB operon is similar in Synechocystis sp. strain PCC 6803 (Ferreira and Straus, 1994). Therefore, the pattern of organization and expression that was originally observed for the flavodoxin gene of Synechococcus sp. strain PCC 7942 may be quite widely shared by unicellular cyanobacteria. The organization ofthe gene for flavodoxin and its transcriptional pattern appear to be quite different in Anabaena sp. strain PCC 7119. Northern-blot hybridization patterns for RNAs from cells grown in media with and without iron indicatedthat flavodoxin was tightly regulated at the transcriptional level. However, the size of the resultant message, 1250 nucleotides, was much smaller than the dicistronic messages produced by Synechococcus sp. strain PCC 7942 and Synechococcus sp. strain PCC 7002. This indicates that the gene encoding flavodoxin is probably transcribed into a monocistronic message (Fillat et al., 1991). In Anabaena sp. strain PCC 7120, the gene for flavodoxin is the first open reading frame of an iron-regulated transcript (K. G. Loenhardt andN. A. Straus, unpublished data). Since heterocysts of Anabaena sp. strain PCC 7119 appear to produce flavodoxin constitutively (Sandmann et al., 1990), it

738 appears that filamentous, nitrogen-fixing strains of cyanobacteria are quite different from unicellular organisms in the arrangement and expression of the gene for flavodoxin.

2. An Iron-Stress-Induced, Chlorophyll-Binding Protein Between 1984 and 1988 Sherman and colleagues conducted a series of studies to define the effect of iron stress on the profile of polypeptides in the photosynthetic membranes of Synechococcus sp. strain PCC 7942. Their studies not only documented the decrease in a large number of membrane polypeptides (Guikema and Sherman, 1984) but also revealed a new iron-stress-induced, PS II-associated chlorophyll-protein complex denoted CPVI-4 (Pakrasi et al., 1985). The complex did not contain PS II core proteins but contained a 34 kDa chlorophyll-binding polypeptide whose synthesis was induced by iron deprivation (Riethman and Sherman, 1988a,b). It has been suggested that complex CPVI4 may function as an auxiliary light-harvesting complex that compensates for the loss of phycobilisomes during iron stress and may also function as a chlorophyll reservoir that contributes to the assembly of reaction center complexes in the early stages of recovery from iron stress (Riethman and Sherman, 1988b).

a. The isiA Gene DNA sequences ofthe isiAB operons from Synechococcus sp. strains PCC 7942 and PCC 7002 indicate that the first open reading frame coded for a putative polypeptide of 342 amino acids. Homology searches indicated that the deduced amino acid sequence of this polypeptide showed a great deal of similarity to CP43 (Laudenbach and Straus, 1988; Leonhardt and Straus, 1992). Recently, the product of isiA was identified as the iron-stress-induced, 34 kDa protein of the PS II chlorophyll-protein complex CPVI-4. The CPVI-4 complex was shown to be absent under iron-stress conditions in a mutant strain of Synechococcus sp. PCC 7942 in which the isiA gene had been insertionally inactivated (Burnap et al., 1993). Many features of the deduced IsiA protein sequence indicate that it may have functions similar to those of CP43. Hydropathy plots show that all the hydrophobic regions of CP43 have been conserved in the IsiA

Neil A. Straus protein, indicating that both probably assume similar membrane spanning structures. In addition, the conservation of histidine residues in the potential membrane spanning regions suggest chlorophyll ligand capabilities similar to CP43 (Laudenbach and Straus, 1989; Bricker, 1990; Vermaas and Ikeuchi, 1991). Figure 1 compares the amino acid sequences for the IsiA polypeptides with the amino acid sequences of CP43 from Synechococcus sp. strain PCC 7942 (Golden and Stearns, 1988) and Synechococcus sp. strain PCC 7002 (Gingrich et al., 1992). Regions of high sequence identity occur in all membrane-spanning regions and in lumenal loops A and C. The major difference between the predicted IsiA products and CP43 consists of a large deletion in the fifth hydrophilic lumenal loop (the E loop, Bricker, 1990). Since the isiA gene is part of the operon encoding flavodoxin, transcription of this gene in response to iron stress was discussed in the section on flavodoxin expression. Under iron stress, isiA is transcribed into two messages, a monocistronic message containing only isiA and a dicistronic message that also contains the isiB gene for flavodoxin. In both Synechococcus sp. strain PCC 7942 and Synechococcus sp. strain PCC 7002, the monocistronic message is much more abundant than the dicistronic message. Superficially, this would appear to imply that there is a greater demand for IsiA than for flavodoxin. Because of the similarities between CP43 and the IsiA, it is tempting to surmise that under iron stress IsiA is capable of compensating for some as yet unknown iron-related function of CP43. Since a number of mutants have been created for psbC in Synechocystis sp. strain PCC 6803 (Vermaas et al., 1988; Yu and Vermaas, 1990; Rögner et al., 1991), one should be able to use these mutants to see if the isiA can functionally replace CP43 under iron stress. For example, a simpleminded experiment would test for the recovery of oxygen evolution in mutant cells grown under iron stress. However, preliminary experiments in this laboratory indicate that three psbC mutants (kindly provided by Wim Vermaas) did not recover oxygen evolution when grown heterotrophically in iron deficient media (Ferreira and Straus, 1994). Therefore, the function ofthe IsiA protein remains obscure, and the proposal, that the iron-stress-induced CPVI-4 complex serves as an auxiliary lightharvesting component that compensates for phycobilisome loss during iron stress, seems to be the best working hypothesis (Riethman and Sherman, 1988b).

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740 However, the observation that flavodoxin and IsiA are encoded in dicistronic operons in unicellular, non-nitrogen-fixing cyanobacteria but in separate operons in heterocystous, nitrogen fixing cyanobacteria is consistent with IsiA being involved in oxygen-evolving photosynthesis under iron stress (Leonhardt and Straus, 1994). Additionally, isolated PS II core complexes from CP43 minus mutants show blue shifts in their absorbance spectra similar to spectral shifts observed for iron-stressed wild type cells (Öquist, 1971; Guikema and Sherman, 1983; Rögner et al., 1991). This seemingly indicates that changes in the photosynthetic apparatus may involve a loss of CP43 and perhaps a compensatory addition of the IsiA protein. Clearly, more work is needed on this chlorophyll-binding, iron-stress-induced protein.

Catechol-type siderophores typically bind iron more tightly than hydroxamate types. For example, enterobactin, which forms the most stable of the known iron complexes, has a stability constant (Eq. 1) of while the hydroxamate siderophores, coprogen, ferrichrome and aerobactin have stability constants of and respectively. When expressed in the physiologically more meaningful value, pM (Eq. 2) catechol siderophores are still up to 10,000 times more efficient at binding iron than hydroxamate types; the pMs of enterobactin, coprogen, ferrichrome and aerobactin are 35.5, 27.5, 25.2 and 23.3 while the concentration of free in a solution that had a total iron concentration of at pH 7.4 would be about to (Raymond et al., 1984; Kerry et al., 1988).

C. Acquisition 1. Siderophores: Iron-Specific Chelators In response to conditions of iron deficiency most aerobic, and facultative anaerobic microbes produce iron-specific chelators called siderophores. Siderophores bind Fe3+ tightly, forming soluble complexes that are recognized by cellular transport systems. The detection, chemistry and function of microbial siderophores have been the subject of a substantial series ofreviews (Neilands, 1981, 1982, 1984; Hider 1984; Raymond et al. 1984; Braun and Winkelmann, 1987; Crosa, 1989; Matzanke, 1991; Winkelmann, 1991). For many years bacterial siderophores were categorized into two classes, hydroxamate-type and catechol-type siderophores, but the discovery of anguibactin from Vibrio anguillarum (Actis et al., 1986) introduced an intermediate-type of siderophore that contains both catecholate and hydroxamate structures. Since cyanobacterial research on siderophores has not yet approached the complexity ofthat being done in other microbes and since all the cyanobacterial literature just refers to hydroxymatetype or catechol-type siderophores, this review must necessarily focus on these two types. However, the number of iron-chelating biological molecules is growing at a fairly rapid rate and those interested in a recent assessment of this area are referred to the comprehensive treatise by Matzanke (1991). Figure 2 contains examples ofhydroxamate-type and catecholtype siderophores utilized by E. coli and some cyanobacteria.

2. Iron Transport in E. coli As one might expect, iron transport systems have been best studied in E. coli, in which the focus of research efforts and the ease ofgenetic manipulation not only led to the identification ofthe genes involved in iron transport but also led to the identification and isolation of fur, the gene responsible for regulation of iron-stress-induced genes. Although the E. coli system has been the subject of numerous reviews, it is worth describing briefly here because it probably illustrates many ofthe mechanisms one can expect to find in cyanobacteria and because the E. coli system may serve as a genetic tool to help unravel iron-stress regulation in cyanobacteria. Under conditions of iron stress, E. coli normally only produces enterobactin; however, some strains are also able to synthesize aerobactin, which is normally encoded on ColV plasmids. In spite of this relatively narrow spectrum of siderophore synthesis, E. coli actually possesses five iron-stress-induced, iron-transport systems: four siderophore systems and a citrate-mediated system. Receptor protein in the outer membrane for each system bind to the ferricsiderophore complexes which are too large to pass

Chapter 25 Iron Stress, fur Repressor, Gene Regulation

into the periplasmic space via the porin channels of the outer membrane. These receptors are: FepA which binds to ferric enterobactin; FhuA (TonA) which binds to ferrichrome; Iut which binds to ferric aerobactin; FhuE which can bind to either ferric coprogen or ferric rodotorulate; and FecA which binds to ferric dicitrate. However, transport into the cytoplasm is actually completed by only three different complexes composed of periplasmic and cytoplasmic membrane proteins, because ferrichrome, ferric aerobactin and ferric coprogen share the same complex. It should also be noted that, although many E. coli strains can only produce enterobactin, they are capable of transporting siderophores synthesized by other, very different organisms: e.g., the ferrichrome and coprogen produced by fungi. More details about the genes, proteins and mechanisms of iron transport may be found in a number of reviews (Braun and Winkel-

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mann, 1987; Bagg and Neilands, 1987; Crosa, 1989; Braun and Hantke, 1991).

3. Iron Transport in Cyanobacteria As indicated above, iron acquisition systems are understood in substantial detail in E. coli and other bacteria. In many ofthese systems siderophores have been structurally characterized, and various membrane-transport components have been identified. Unfortunately, comparatively little is known about cyanobacteria in this regard. Lange (1974) examined a variety of cyanobacteria for the ability to form natural chelators that promote growth at high pHs. Ofthe ten species examined, six were able to grow at high pHs in the absence of artificial chelators. In each case filtrates of these cyanobacteria could promote growth in the four species that could not produce their own chelators. Estep et al. (1975) were

742 the first to employ a standard biological test for siderophores in cyanobacteria; they showed that aqueous extracts of near-shore marine cyanobacteria were able to promote growth in the siderophore auxotroph Arthrobacter flavescens JG-9 (Burnham and Neilands, 1961). In the following year, Murphy et al. (1975) used the Csaky test to demonstrate chemically the production of hydroxyamate siderophores by cyanobacteria. Both Estep et al. (1975) and Murphy et al. (1976) suggested that it was the ability of cyanobacteria to sequester iron through the production and utilization of siderophores that allowed them dominate eucaryotic algae during cyanobacterial blooms. Although many cyanobacteria are capable of producing siderophores, many strains are not and depend on siderophores produced by bacteria or other cyanobacteria to continue growth in irondeficient environments (Lange, 1974). Other strains which produce their own siderophores may still utilize siderophores produced by other bacteria. By following the uptake of Goldman et al. (1983) showed that Anabaena sp. strain PCC 6411, that produces the siderophore schizokinen, was capable of utilizing aerobactin at a lower efficiency but was far less effective at using the trihydroxyamate siderophore ferrioxamine. Other strains of Anabaena sp. that did not produce hydroxyamate siderophores were much less efficient at using schizokinen to promote iron uptake. Those who so fastidiously maintain axenic cultures need to be reminded periodically that in nature cyanobacteria grow in the presence of other bacteria and may frequently use siderophores produced by other organisms, just as E. coli utilizes siderophores produced by other, very different, organisms. Siderophores are capable of binding to cupric ions as well as ferric ions; however, cyanobacteria do not transport the copper-siderophore complex. Therefore, siderophore secretion has the effect of detoxifying media containing copper (McKnight and Morel, 1980) and the ability of iron-stressed cyanobacteria to grow in the presence of toxic levels of copper has been used as an indication of siderophore production (Boyer et al., 1987). Clarke et al. (1987) showed that the growth of Anabaena sp. strain PCC 7120 was inhibited by the addition of 2.1 to copper and abolished by concentrations above However, the addition of schizokinen (see Fig. 2) to the media was able to alleviate the toxic effects ofcopper. In the absence of schizokinen copper was taken up by the

Neil A. Straus cells but when schizokinen was added to the media schizokinen bound copper remained in the media. Schizokinen (Fig. 2), which has been isolated from an Anabaena sp. and which is also produced by Bacillus megaterium, is the only cyanobacterial siderophore to be structurally characterized (Simpson and Neilands, 1976). However, Trick and Kerry (1992) recently demonstrated the production of two different and unique hydroxamate-type siderophores by Synechococcus sp. strain PCC 7942 and Anabaena variabilis. They showed that neither of these species produced catechol-type siderophores and that the molecular masses of the isolated siderophores were quite different from other hydroxamate siderophores, including schizokinen, aerobactin, ferrichrome and rhodotorulic acid. Using the commercial chelator EDDA (ethylene diamine di-o-hydroxyphenyl acetic acid) that has a high affinity for iron, Kerry et al. (1988) developed a bioassay to detect the presence of high-affinity, irontransport systems. By growing fourteen different strains of cyanobacteria in iron-replete media that contained different levels of EDDA, they were able to categorize the strains on the ability of their siderophores to compete with EDDA for available iron. They found three basic categories. Five of the strains could not survive low levels of EDDA and were assumed to lack a siderophoreproducing system. Five other strains were inhibited by intermediate levels of EDDA and were thought to produce siderophores of relatively moderate binding efficiency. The four remaining strains survived high levels of EDDA and were assumed to be capable of producing highefficiency siderophore systems. Recently, Brown and Trick (1992) reported that Oscillatoria tenius was able to sustain growth in iron replete medium containing of EDDA. They showed that this cyanobacterium was able to produce a catecholtype siderophore in addition to a hydroxamate- type siderophore. The production of these two siderophores is induced at different iron levels in the medium. They suggested that the production of the more efficient catechol-type siderophore accounted for the ability of this cyanobacterium to grow at high levels of EDDA and to thrive in low-iron ecosystems. They also suggested that the possession of the catechol-type siderophore system may be a common feature of other filamentous cyanobacteria that are often associated with highly alkaline, low-iron environments.

Chapter 25 Iron Stress, fur Repressor, Gene Regulation The EDDA test may be a rapid functional assay to differentiate cultures that produce only hydroxamatetype siderophores from those that are capable of producingcatechol-typesiderophores.Furthermore, after the addition of different siderophores to cyanobacteria that cannotproduce these siderophores, the same test could be used to assay for cyanobacteria that possessed transport systems for either or both siderophore types. Table 1 contains a list of cyanobacteria thathave been shown to produce siderophores by one or more tests. Biological tests include the ability to grow axenically under conditions of iron deprivation, the ability to survive copper toxicity, or the ability to promote growth ofbacterial siderophore auxotrophs. Chemical tests include the Csaky test for hydroxamates (Csaky, 1948; Gillam et al. 1981) and the catechol tests developed by Arnow (1937) and Rioux et al. (1983). The table includes species identified in the review by Boyer et al. (1987) and ones that have been added in the intervening time.

a. Siderophore Production and Nitrogen Fixation Hutchins et al. (1991) reported that siderophore production and nitrogen fixation appeared to be mutually exclusive processes in Anabaena sp. strain PCC 7120 cultivated in the absence ofiron. However, in a study involving controlled levels of iron in the medium, Kerry et al. (1988) showed that growth of Anabaena variabilis in the absence of a combined nitrogen source actually resulted in higher levels of siderophore production than growth in a medium containing nitrate. Cells grown without combined nitrogen appeared to have a much greater demand for iron, since siderophore production occurred at higher initial concentrations of iron in the medium. The relationship between iron stress and nitrogen fixation may be quite complex and will be an interesting one for further studies. In Klebsiella pneumoniae iron deficiency actually has a stimulatory effect on genes involved in nitrogen fixation in by inhibiting the repressive activity of NifL (Henderson et al., 1989). In any case, the apparently conflicting results for siderophore production and nitrogen fixation in different Anabaena sp. point out two difficulties in the literature on iron deprivation in cyanobacteria. Firstly, the limited number ofstudies in any one area are often done on different species; this makes comparisons difficult and generally slows the progress of research in this area. Secondly, many studies on iron stress, particularly earlier ones, used cells

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cultivated in the absence of iron; under these conditions cells may simply die. Some more standardized approach to controlling iron-stress conditions without approaching cell death must be employed. One approach to the study of iron-transport systems in cyanobacteria is to induce iron stress by including EDDA or other iron specific ligands into iron-replete culture media. Under these conditions, cyanobacteria experience iron stress but are able to utilize the iron in the medium by producing siderophores that can strip the iron complexed to EDDA in the medium. Scanlan et al. (1989) showed this by comparing the growth rates ofSynechococcus sp. strain PCC 7942 grown in iron-replete medium containing EDDA or in iron-deficient medium. The growth rate ofcells in iron-replete medium containing EDDA was similar to that of cells in iron-replete medium without EDDA. However, the cells in ironreplete medium containing EDDA were actually ironstressed and produced the outer membrane proteins that characterize the iron-stress condition. On the other hand, the cells in medium in which iron was limiting had a much slower growth rate and experienced severe symptoms of iron deprivation such as decreases in membranes, phycobilisomes and carboxysomes. IV. The Control of Gene Expression by Iron

A. The Fur Repressor of E. coli When faced with a reduced iron supply, E. coli and other bacteria activate a battery of genes whose products form an efficient iron-acquisition system. Iron-dependent regulation has also been demonstrated for many genes not involved in iron acquisition. Some of these genes, such as those for bacterial toxins, are indirectly related to iron acquisition because they cause the release of intracellular iron stores by killing host cells. Other iron-regulated genes, such as the genes for superoxide dismutases (Niederhoffer et al., 1989; Neiderhoffer et al., 1990; Hassan and Sun, 1992), form products that contain iron and therefore impinge on the iron inventory of the cell. Still others, such as the genes that control swarming in Vibrio parahemolyticus (McCarter and Silverman, 1989), have a less obvious connection to iron stress. Iron-regulated genes in E. coli and other bacteria are controlled by fur, a gene that codes for a DNA-

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binding, repressor protein (Crosa et al., 1989; Braun et al., 1990; Braun and Hantke, 1991). E. coli mutations that result in the simultaneous activation of all iron-transport membrane proteins map to the

Neil A. Straus

fur locus (Hantke, 1981). The DNA containing the fur locus has been cloned by its ability to rescue wild-type iron regulation in a fur mutant (Hantke, 1984). The gene has been sequenced and over-

Chapter 25 Iron Stress, fur Repressor, Gene Regulation produced from an expression vector (Schaffer et al., 1985; Wee et al., 1988). The fur gene codes for a protein of 148 amino acids containing twelve histidine residues. DNA binding experiments indicate that, in the presence of (or which is now used instead of because is less prone to oxidation), the Fur proteinbinds to two sequences in the promoter regions of two iron-regulated genes. These genes, iucA and cir, code for an aerobactin synthesis enzyme and the colicin I receptor proteins respectively (de Lorenzo et al., 1987, 1988; Griggs and Konisky, 1989). The sequences that were protected from DNAase I digestion were found to be similar to each other and to sequences in the promoter regions of other ironregulated genes. This led to the construction of a consensus sequence for Fur-binding (de Lorenzo et al., 1987). The function ofthis Fur-binding sequence was confirmed when it was shown that a synthetic oligonucleotide containing the consensus sequence could induce iron regulation in a ompF-lacZ fusion gene if it was inserted between the ompF promoter sequences and the lacZ gene (Calderwood and Mekalanos, 1988). Table 2 contains a list ofidentified and putative, Fur-binding sequences from ironregulated genes of different bacteria, including cyanobacteria. The table extends the list found in reviews by Braun et al. (1990), and Braun and Hantke (1991).

B. Iron-Regulated Genes in Cyanobacteria In a numberofcyanobacteria, the gene for flavodoxin expression is transcriptionally regulated under conditions ofiron stress. In Synechococcus sp. strain PCC 7942 the isiAB operon, that codes for a PS IIassociated, chlorophyll-binding protein and flavodoxin, was shown to be tightly regulated by iron concentration (Laudenbach et al., 1988, Laudenbach and Straus, 1988). An examination of sequences upstream from the transcriptional start site revealed a set of sequences that resembled Fur-binding sequences in E. coli (Table 2). The same iron-regulated operon has been isolated and sequenced from Synechococcus sp. strain PCC 7002. In both cases primer extension data identified the transcriptional start sites, and sequence data located potential Furbinding sequences in the operator region (Leonhardt and Straus, 1992, Table 2). The gene encoding flavodoxin from Anabaena sp. strain PCC 7119 has also been isolated and shown to be regulated by iron

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(Fillat et al., 1991). Unfortunately not enough upstream flanking sequence was reported to locate Fur-bindingsequences. Using antibodies to an iron-regulated membrane protein from Synechococcus sp. strain PCC 7942, Reddy et al. (1988) cloned a gene (irpA) that showed tight transcriptional regulation in response to iron deprivation. Although the transcriptional start site forthe gene was not determined, potential Fur-binding sequences were identified in sequences upstream from the coding region (Table 2). Insertional inactivation ofthis gene produced a mutant that was not able to survive in iron-stress medium and that showed increased sensitivity to the presence of a competing iron chelator (2,2'-dipyridyl) in ironreplete media. Since immunocytochemical studies placed IrpA in the cytoplasmic membrane, it appears IrpA is part ofan iron-transport system, although its localization indicates that it is probably not a receptor protein. Electrophoretic profiles of outer-membrane proteins from Synechococcus sp. strain PCC 7942 indicate that iron deprivation leads to the formation of proteins (92, 48–50, and 35 kDa) not found in cells grown in iron-replete conditions (Scanlan et al., 1989). If these proteins represent different siderophore receptors, this would be at least preliminary evidence for multiple iron-transport systems in Synechococcus sp. PCC strain 7942. Interestingly, cbpA, the gene for a carotenoidbinding, membrane protein in Synechococcus sp. PCC 7942, was reported to be transcriptionally repressed in conditions of iron deprivation (Reddy et al., 1989). As indicated in earlier sections, a number of symptoms of iron stress involve the loss of many different proteins (Sections III A, B); at least one of these proteins, ferredoxin, has been shown to be regulated at the post-transcriptional level (Van der Plas et al., 1988; Bovy et al., 1993). However, this is the first cyanobacterial report of regulation in which a gene is turned off at the transcriptional level in response to iron deficiency.

Acknowledgments I would like to thank Sam Beale and David Krogmann for sharing preprints of their respective chapters. Iron-stress research in this laboratory is supported by grants from the Natural Sciences and Engineering Research Council of Canada.

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748 Ferreira F and Straus NA (1994) Iron deprivation in cyanobacteria. J Appl Phycol 6: in press Fillat MF, Borrias WE and Weisbeck PJ (1991) Isolation and overexpression in Escherichia coli of the flavodoxin gene from Anabaena PCC 7119. Biochcm J 280: 187–191 Gasparich GE (1989) The Effects of Various Environmental Stress Conditions on Gene Expression in the Cyanobacterium Synechococcus sp. PCC 7002. Ph. D. thesis, The Pennsylvania State University Ghanotakis DF, de Paula JC, Demetriou DM, Bowlby NR, Petersen J, Babcock GT and Yocum CF (1989) Isolation and characterization of the 47 kDa protein and the D1-D2cytochrome b-559 complex. Biochim Biophys Acta 974: 44– 53 Gillam AH, Lewis AC and Andersen RJ (1981) Quantitative determination of hydroxamic acids. Anal Chem 53: 841–844 Gingrich JC, Gasparich GE, Saner K and Bryant DA (1990) Nucleotide sequence and expression of the two genes encoding D2 protein and the single gene encoding the CP43 protein of Photosystem II in the cyanobacterium Synechococcus sp. PCC 7002. Photosynth Res 24: 137–150 Golbeck JH and Bryant DA (1991) Photosystem I. In: Lee CP (ed) Current Topics in Bioenergetics, Vol 16, pp 83–177. Academic, New York Goldberg MB, Boyko SA and Calderwood SB (1990) Transcription regulation by iron of a Vibrio cholerae virulence gene and homology of the gene to the Escherichia coli Fur system. J Bacteriol 172: 6863–6870 Golden SS and Stearns GW (1988) Nucleotide sequence and transcript analysis of three Photosystem II genes from the cyanobacterium Synechococcus sp. PCC 7942. Gene 67: 85– 96 Goldman SJ, Lammers PJ, Berman MS and Sanders-Loehr J (1983) Siderophore-mediated iron uptake in different strains of Anabaena sp. J Bacteriol 156: 1144–1150 Gounaris K, Chapman DJ and Barber J (1989) Isolation and characterization of a D1/D2/cytochrome b-559 complex from Synechocystis 6803. Biochim Biophys Acta 973: 296–301 Griggs DW and Konisky J (1989) Mechanism for iron-regulated transcription of the Escherichia coli cir gene: metal-dependent binding of Fur protein to the promoters. J Bacteriol 171:1048– 1054 Guikema JA (1985) Fluorescence induction characteristics of Anacystis nidulans during recovery from iron deficiency. J Plant Nutr 8:891–908 Guikema JA and Sherman LA (1983) Organization and function of chlorophyll in membranes of cyanobacteria during iron starvation. Plant Physiol 73:250–256 Guikema JA and Sherman LA (1984) Influence of iron deprivation on the membrane composition of Anacystis nidulans. Plant Physiol 74: 90–95 Gounaris K, Chapman DJ and Barber J (1989) Isolation and characterization of a D1/D2/cytochrome b-559 complex from Synechocystis 6803. Biochim Biophys Acta 973: 296–301 Hantke K (1981) Regulation of ferric iron transport in Escherichia coli K12: Isolation of a constitutive mutant. Mol Gen Genet 182:288–292 Hantke K (1984) Cloning of the represser protein gene of ironregulated systems in Escherichia coli K12. Mol Gen Genet 197: 337–341 Hardie LP, Balkwill DL and Stevens SE Jr (1983a) Effects of

Neil A. Straus iron starvation on the physiology of the cyanobacterium Agmenellum quadruplicatum. Appl Environ Microbiol 45: 999–1006 Hardie LP, Balkwill DL and Stevens SE Jr (1983b) Effects of iron starvation on the ultrastructure of the cyanobacterium Agmenellum quadruplicatum. Appl Environ Microbiol 45: 1007–1017 Hassan HM and Sun HH (1992) Regulatory roles of Fnr, Fur, and Arc in expression of manganese-containing superoxide dismutase in Escherichia coli. Proc Natl Acad Sci USA 89: 3217–3221 Hauska G (1985) Organization and function of cytochrome complexes. In: Steinback KE, Bonitz S, Arntzen C and Bogorad L (ed) Molecular Biology of the Photosynthetic Apparatus, pp. 79–87. Cold Spring Harbor Laboratory Press, Cold Spring Harbor Hauska G, Hurt E, Gabellini N and Lockau W( 1983) Comparative aspects of quinol-cytochrome c/plastocyanin oxidoreductases. Biochim Biophys Acta 726: 97–133 Henderson N, Austin S and Dixon RA (1989) Role of metal ions in negative regulation of nitrogen fixation by the nifL gene product from Klebsiella pnewnoniae. Mol Gen Genet 216: 484–491 Hider RC (1984) Siderophore mediated absorption of iron. Struct Bonding 58: 25–88 Hurt EC, Hauska G and Shahak Y (1982) Electrogenic proton translocation by the chloroplast cytochrome complex reconstituted into phospholipid vesicles. FEES Lett 149:211– 216 Hutber GN, Hutson KG and Rogers LJ (1977) Effect of iron deficiency on levels of two ferredoxins and flavodoxins in a cyanobacterium. FEMS Microbiol Lett 1: 193–196 Hutchins DA, Rueter JG and Fish W (1991) Siderophore production and nitrogen fixation are mutually exclusive strategies in Anabaena 7120. Limnol Oceanogr 36: 1–12 Kerry A, Laudenbach DE and Trick CG (1988) Influence of iron limitation and nitrogen source on growth and siderophore production by cyanobacteria. J Phycol 24: 566–571 Lange W (1974) Chelating agents and blue-green algae. Can J Microbiol 20: 1311–1321 Laudenbach DE and Straus NA (1988) Characterization of a cyanobacterial iron stress-induced gene similar to psbC. J Bacteriol 170: 5018–5026 Laudenbach DE, Reith ME and Straus NA (1988) Isolation, sequence analysis, and transcriptional studies of the flavodoxin gene from Anacystis nidulans R2. J Bacteriol 170: 258–264 Leonhardt KG and Straus NA (1989) Sequence of flavodoxin from Anabaena variabilis 7120. Nucleic Acids Research 17: 4384 Leonhardt KG and Straus NA (1992) An iron stress operon involved in photosynthetic electron transport in the marine cyanobacterium Synechococcus sp. PCC 7002. J Gen Microbiol 138: 1613–1621 Leonhardt K and Straus NA (1994) Photosystem II genes isiA, psbDI and psbC in Anabaena sp. PCC 7120: Cloning, sequencing and transcriptional regulation in iron-stressed and iron-replete cells. Plant Mol Biol 24: 63–73 Matzanke BF (1991) Structures, coordination chemistry and functions of microbial iron chelates. In. Winkelmann G (ed) CRC Handbook of Microbial Iron Chelates, pp. 15–64. CRC Press, Boca Raton

Chapter 25 Iron Stress, fur Repressor, Gene Regulation Mahasneh IA (1991) Siderophore production in the Rivulariaceae, blue-green algae. Microbios 65: 97–104 Mayhem SG and Ludwig ML (1975) Flavodoxins and electron transferring flavoproteins. Enzymes (3rd Ed) 12: 57–118 McCarter L and Silverman M (1989) Iron regulation ofswarmer cell differentiation of Vibrio parahaemolyticus. J Bacteriol 171:731–736 McKnight DM and Morel FMM (1979) Release of weak and strong copper-complexing agents by algae. Limnol Oceanogr 24: 823–837 McKnight DM and Morel FMM (1980) Copper complexation by siderophores from filamentous blue-green algae. Limnol Oceanogr 25: 62–71 Miyazaki A, Shina T, Toyoshima Y, Gounaris K and Barber J (1989) Stoichiometry of cytochrome b-559 in Photosystem II. Biochim Biophys Acta 975: 142–147 Murphy TP, Lean DRS and Nalewajko C (1976) Blue-green algae: Their excretion of iron-selective chelators enables them to dominate other algae. Science 192: 900–902 Murphy TP, Hall KJ and Yesaki I (1983) Biogenic regulation of iron availability in an eutrophic hardwater lake. Sci Total Environ 28: 37–50 Nanba O and Satoh K (1987) Isolation of a Photosystem II reaction center consisting of D-1 and D-2 polypeptides and cytochrome b-559. Proc Natl Acad Sci USA 84: 109–112 Neilands JB (1981a) Microbial iron compounds. Ann Rev Biochem 50: 715–731 Neilands JB (1981b) Iron absorption and transport in microorganisms. Ann Rev Nutr 1:27–46 Neilands JB (1982) Microbial envelope proteins related to iron. Ann Rev Microbiol 36: 285–309 Neilands JB (1984) Methodology ofsiderophores. Struct Bonding 58: 1–24 Niederhoffer EC, Naranjo CM and Fee JA (1989) Relationship of superoxide dismutase genes, sodA and sodB, to the iron uptake (fur) regulon in Escherichia coli K-12. In: Hamer DN and Winge DR (ed) Metal Ion Homeostasis: Molecular Biology and Chemistry, pp 149–158. Liss, New York Niederhoffer EC, Naranjo CM, Bradley KL and Fee JA (1990) Control of Escherichia coli superoxide dismutase (sodA and sodB) genes by the ferric uptake regulation (fur) locus. J Bacteriol 172: 1930–1938 Öquist G (1971) Changes in pigment composition and photosynthesis induced by iron-deficiency in the blue-green alga Anacystis nidulans. Physiol Plant 25: 188–191 Öquist G (1974a) Iron deficiency in the blue-green alga Anacystis nidulans: Changes in pigmentation and photosynthesis. Physiol Plant 30:30–37 Öquist G (1974b) Iron deficiency in the blue-green alga Anacystis nidulans: Fluorescence and absorption spectra recorded at 77° K. Physiol Plant 31:55–58 Pakrasi HB, Riethman HC and Sherman LA (1985) Organization of pigment proteins in the Photosystem II complex of the cyanobacterium Anacystis nidulans R2. Proc Natl Acad Sci USA 82: 6903–6907 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 b-559 in Photosystem II. EMBO J. 7:325–332 Pakrasi HB, Diner BA, Williams JGK and Arntzen CJ (1989) Deletion mutagenesis of the cytochrome b-559 protein

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inactivates the reaction center of Photosystem II. Plant Cell 1:591–597 Pardo MB, Gomez-Moreno C and Peleato ML (1990) Effect of iron deficiency on ferredoxin levels in Anabaena variabilis PCC 6309. Arch Microbiol 153: 528–530 Peschek GA (1979) Nitrate and nitrite reductase and hydrogenase in Anacystis nidulans grown in Fe- and Mo-deficient media. FEMS Microbiol Lett 6: 371–374 Peschek GA and Schmetterer G (1982) Evidence for plastocytochrome f/b-563 reductase as a common electron donor to P700 and cytochrome oxidase in cyanobacteria. Biochem Biophys Res Comm 108: 1188–1195 Raymond KN, Muller G and Matzanke BF (1984) Complexation of iron by siderophores: A review of their solution and structural chemistry and biological function. Topics Curr Chem 123:49– 102 Reddy KJ, Bullerjahn GS, Sherman DM and Sherman LA(1988) Cloning, nucleotide sequence, and mutagenesis of a gene (irpA) involved in iron-deficient growth of the cyanobacterium Synechococcus sp. strain PCC7942. J Bacteriol 170: 4466– 4476 Reith ME, Laudenbach DE and Straus NA (1986) Isolation and nucleotide sequence analysis ofthe ferredoxin I gene from the cyanobacterium Anacystis nidulans R2. J Bacteriol 168:1319– 1324 Rhie G and Beale SI (1992) Biosynthesis of phycobilins: Ferredoxin-supported NADPH-independent heme oxygenase and phycobilin-forming activities from Cyanidium caldarium. J Biol Chem 267: 16088–16093 Riethman HC and Sherman LA (1988a) Immunological characterization of iron-regulated membrane proteins in the cyanobacterium Anacystis nidulans R2. Plant Physiol 88: 497–505 Riethman HC and Sherman LA (1988b) Purification and characterization of an iron stress-induced chlorophyll-protein from the cyanobacterium Anacystis nidulans R2. Biochim Biophys Acta 935: 141–151 Rioux C, Jordan DC and Rattray JBM (1983) Colorimetric determination of catechol siderophores in microbial cultures. Anal Biochem 133: 163–169 Rögner M, Chisholm DA and Diner BA (1991) Site-directed mutagenesis of the psbC gene of Photosystem II: Isolation and functional characterization of CP43-less Photosystem II core complexes. Biochemistry 30: 5387–5395 Sandmann G (1985) Consequences of iron deficiency on photosynthetic electron transport in blue green algae. Photosynth Res 6: 261–271 Sandmann G and Malkin R (1983) Iron-sulfur centers and activities of the photosynthetic electron transport chain in irondeficient cultures of the blue-green alga Aphanocapsa. Plant Physiol 73: 724–728 Sandmann G, Peleato ML, Fillat MF, Lazaro MC and GomezMoreno C (1990) Consequences of iron-dependent formation of ferredoxin and flavodoxin on photosynthesis and nitrogen fixation on Anabaena strains. Photosynth Res 119–125 Scanlan DJ, Mann NH and Carr NG (1989) Effect of iron and other nutrient limitations on the pattern of outer membrane proteins in the cyanobacterium Synechococcus PCC7942. Arch Microbiol. 152:224–228 Schaffer S, Hantke K and Braun V (1985) Nucleotide sequence of iron regulatory gene fur. Mol Gen Genet 200: 110–113

750 Sherman DM and Sherman LA (1983) Effect of iron deficiency and iron restoration on ultrastructure of Anacystis nidulans. J Bacteriol 156: 393–401 Simpson FB and Neilands JB (1976) Siderochromes in Cyanophyceae: Isolation and characterization ofschizokinen from Anabaena sp. J Phycol 12: 44–48 Smillie RM (1965a) Isolation of phytoflavin, a flavoprotein with chloroplast ferredoxin activity. Plant Physiol 40: 1124–1128 Smillie RM (1965b) Isolation of two proteins with chloroplast ferredoxin activity from a blue-green alga. Biochem Biophys Res Comm 20: 621–629 Takahashi Y, Hansson O, Mathis P and Satoh K (1987) Primary radical pair in the Photosystem II reaction center. Biochim Biophys Acta 893 49–59. Tollin G and Edmondson DE (1980) Purification and properties of flavodoxins. Meth Enzymol 69: 392–406 Trick CG and Kerry A (1992) Isolation and purification of siderophores produced by cyanobacteria, Synechococcus sp. PCC 7942 and Anabaena variabilis ATCC 29413. Curr Microbiol 24: 241–245 Troxler RF, Lin S and Offner GD (1989) Heme regulates expression of phycobiliprotein photogenes in the unicellular Rodophyte, Cyanidium caldarium. J Biol Chem 264: 20596– 20601 Uphoff TS and Welch RA (1990) Nucleotide sequencing of the Proteus mirabilis calcium-independent hemolysin genes (hpmA and hpmB) reveals sequence similarity with Serratia marcescens hemolysin genes (shlA and shlB). J Bacteriol 172: 1206–1216 Van den Berg CMG, Wong PTS and Chau YK (1979) Measurement of complexing materials excreted from algae and their ability to ameliorate copper toxicity. J Fish Res Bd Can 36: 901–905 Van der Plas J, De Groot RP, Woortman MR, Weisbeek PJ and

Neil A. Straus Van Arkel GA (1986) Coding sequence of a ferredoxin gene from Anacystis nidulans R2 (Synechococcus PCC 7942). Nucleic Acids Res 14: 7804 Van der Plas J, de Groot R, Woortman FC, Borrias M, Van Arkel G and Weisbeek P (1988) Genes encoding ferredoxins from Anabaena sp. PCC 7937 and Synechococcus sp. PCC 7942: Structure and regulation. Photosynth Res 18: 179–204 Vermaas WFJ and Ikeuchi M (1991) Photosystem II. In: Bogorad L and Vasil IK (ed) The Photosynthetic Apparatus: Molecular Biology and Operation, pp 25–111. Academic Press, San Diego Vermaas WFJ, Ikeuchi M and Inoue Y (1988) Protein composition of the Photosystem II core complex in genetically engineered mutants of the cyanobacterium Synechocystis sp. PCC 6803. Photosynth Res 17:97–113 Wee S, Neilands JB, Bittner ML, Hemming BC, Haymore BL and Seetharam R (1988) Expression, isolation and properties of Fur (ferric uptake regulation) protein of Escherichia coli) K12. Biol Metals 1:62–68 Winkelmann G (1991) Specificity of iron transport in bacteria andfungi. In: Winkelmann G(ed)CRC Handbook ofMicrobial Iron Chelates, pp 65–105. CRC Press, Bacon Raton 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 Yoch DC and Valentine RC (1972) Ferredoxins and flavodoxins of bacteria. Ann Rev Microbiol 26: 41–51 Yu J and Vermaas WFJ (1990) Transcript levels and synthesis of Photosystem II components in cyanobacterial mutants with inactivated Photosystem II genes. Plant Cell 2: 315–322 Zhou J, Gasparich GE, Stirewalt VL, de Lorimier R and Bryant DA (1992) The cpcE and cpcF genes of Synechococcus sp. PCC 7002. J Biol Chem 267: 16138–16145

Chapter 26 The Cyanobacterial Heat-Shock Response and the Molecular Chaperones Robert Webb The University of Texas at El Paso, Department of Biological Sciences, El Paso, Texas 79968, USA

Louis A. Sherman Purdue University, Department of Biological Sciences, Lilly Hall, West Lafayette, Indiana 47907, USA Summary I. Background A. Heat shock in E. coli B. Molecular Chaperone Concept 1. Definition 2. Bacterial Heat-Shock Proteins and Molecular Chaperones C. Thermal Tolerance and the Cellular Thermometer II. Functional Aspects, Protein Folding and Localization A. dnaK Mutants and Functional Domains of DnaK B. Molecular Chaperones and Protein Folding C. Successive Action of Molecular Chaperones D. Insertion and Translocation of Membrane Proteins E. Thylakoid Membrane Assembly III. Molecular Chaperones of the Cyanobacteria A. Heat-Shock Proteins of the Cyanobacteria B. Nucleotide-Binding Motif Sequence Comparisons C. Cyanobacterial Heat-Shock Genes D. Transcriptional Response of the Cyanobacteria to Heat Shock E. Mutational Studies of the Cyanobacterial Molecular Chaperones IV. Summary and Future Directions References

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Summary The studies of the heat-shock response, other stress responses and protein folding have become intimately intertwined during the past few years. The heat-shock response, which was first described in Drosophila melanogaster over 30 years ago (Ritossa, 1962), entails the rapid and selective increased expression of a large class of proteins. Surprisingly, this heat-shock response was described in Escherichia coli long after the phenomenon had been described in numerous eucaryotic organisms. Since that time, work in E. coli has been a very fertile field for an understanding of this response at the genetic and molecular biological levels. However, the key experiments which led to a synthesis of many different disciplines were the analysis of the assembly of ribulose 1,5-bisphosphate carboxylase/oxygenase (Rubisco) from chloroplasts. Research on the heat-shock proteins during the 1970s and 1980s demonstrated that such proteins were involved in protein-protein D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 751–767. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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interactions. Research on the assembly of Rubisco in E. coli demonstrated the absolute requirement for specific heat-shock proteins and helped lead to the concept of molecular chaperones. A voluminous literature has developed in each of these fields and a comprehensive review would be beyond the scope of this chapter. It is our intention to summarize the salient features of the heat-shock response and molecular chaperones and provide a listing of sufficient review articles for the interested reader. This chapter will concentrate on results and concepts as they relate to bacteria, chloroplasts and mitochondria and summarize the current thinking about the function of molecular chaperones. This will include models for the transport of proteins into and across membranes, since this is one important feature of life within a cyanobacterium. With this background in hand, a summary of what is known about the heat-shock response and its regulation in cyanobacteria will be presented. I. Background

A. Heat shock in E. coli All organisms are known to respond to heat by inducing the synthesis of the heat-shock proteins (hsp). This response appears to be one of the most highly conserved genetic systems known and has been shown to occur in members of the archaebacteria, eubacteria, plants and animals. The heat-shock regulon in E. coli consists of at least 18 proteins whose rates of synthesis are increased when cells are grown at higher than optimal temperatures (Van Bogelen et al., 1987). Five of the E. coli heat-shock proteins, including DnaK, DnaJ, GroEL, GroES, and GrpE, are now known to act as molecular chaperones and modulate many kinds of protein-protein interactions. The induction of these hsp is more a result of environmental perturbations than merely heat shock. Thus, in addition to temperature upshifts, carbon or phosphate starvation and exposure to ethanol or heavy metals can also induce these genes encoding these proteins (Van Bogelen et al., 1987). Antibiotics which interfere with ribosome function are also known to induce the expression of this regulon (Van Bogelen et al., 1990). It has now been shown that the heat-shock response can be modulated by one-carbon metabolism in E. coli (Gage and Neidhardt, 1993). A great deal of work has shown that the induction of the heat-shock proteins is under the control of a special positive activator, the product of the rpoH gene. This factor binds to the core RNA polymerase and permits the polymerase to recognize the heat-shock promoters (Grossman et al., 1984; Abbreviations: cpn – chaperonin; hsp – heat-shock protein; LHCP – light-harvesting chlorophyll a/b protein; Rubisco – ribulose 1,5-bisphosphate carboxylase/oxygenase; Ts – temperature-sensitive

Landick et al., 1984). Despite the extraordinary increase in transcriptional levels, it is thought that the increase in the amount of following exposure to high temperature is sufficiently rapid to fully account for the induction of the heat-shock proteins (Straus et al., 1987). The complex process of increasing the amount of is brought about by enhancing production of the rpoH message, by increasing the translation of existing message, and by stabilizing the existing which normally has a half-life of 1 min (Straus et al., 1987, 1990). Indeed, the relatively rapid turnover of is thought to be regulated by the heat-shock proteins DnaK, DnaJ and GrpE themselves (Tilly et al., 1983; Straus et al., 1987, 1990; Gamer et al., 1992). The precise mechanism by which a bacterial cell senses the stress conditions, and the signal which then leads to the increased transcription and translation of the rpoH gene is poorly understood. Since very little is known about putative heat-shock factors in cyanobacteria, this aspect of regulation will not be discussed in any detail. However, the overall involvement of denatured proteins, the functioning of molecular chaperones, and the possible involvement of molecular chaperones in gene regulation will be considered below.

B. Molecular Chaperone Concept 1. Definition An understanding of the function of hsp first requires an understanding of the somewhat complex terminology that has developed in the field. The term ‘molecular chaperone’ was first coined by Laskey et al. (1978) to describe the function of nucleoplasmin, which is involved in nucleosome assembly in Xenopus laevis eggs. The concept was extended by Ellis and colleagues (Ellis and Hemmingsen, 1989; Ellis, 1990a, b) who defined molecular chaperones as a

Chapter 26 Heat Shock and Molecular Chaperones group of unrelated cellular proteins that mediate the correct assembly of other polypeptides, but are not themselves components of the final structures (Ellis and Hemmingsen, 1989; Ellis, 1990a). There is a great deal of evidence which now suggests that the molecular chaperones actively and/or passively facilitate the folding of many monomeric polypeptides and the subunits of oligomeric proteins under physiological conditions. It should be noted that, although the molecular chaperones were initially implicated in the assembly of oligomeric structures, there is no evidence at present to indicate that they are directly involved in oligomer formation (Lorimer et al., 1993). Thus, we conclude that molecular chaperones are involved at the level of monomeric polypeptides. The chaperonins are a subset of the molecular chaperones. ‘Chaperonin’ is a term that was given to the groEL and groES gene products, which are also referred to as cpn60 and cpn 10, respectively. GroEL, hsp60 and cpn60 can be considered synonyms, as can DnaK and hsp70. The bacterial gene designations will be used for the remainder of this chapter.

2. Bacterial Heat-Shock Proteins and Molecular Chaperones The physical properties and putative functions of the bacterial heat-shock proteins are given in Table 1, which lists those genes known to be under the control of As indicated, a number of these genes had been identified with other functions prior to their designation as part of the heat-shock regulon. For example, the dnaK, dnaJ, grpE and groESL genes were originally discovered because mutations in these genes were shown to block bacteriophage assembly (Georgopoulos et al., 1990; Ellis and van der Vies, 1991; Zeilstra-Ryalls et al., 1991; Gatenby, 1992). Further genetic studies revealed that the groES and groEL genes were essential for bacterial growth (Fayet et al., 1989) and that these proteins were produced by cells at all times of their life cycle. However, following an appropriate stress, the cellular levels of GroEL could be increased from approximately 2% to virtually 10% of the cellular protein. There is now compelling biochemical data to indicate that GroEL and GroES interact functionally (Goloubinoff et al., 1989; Viitanen et al., 1990). The GroEL protein has a molecular mass of approximately 60 kDa and has been highly conserved throughout evolution. The E. coli GroEL protein is

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54% identical to yeast hsp60 and 48% identical to the Rubisco binding protein; this is substantial similarity, since both of those genes are encoded in the nucleus of their respective organisms. The native GroEL complex is composed of 14 identical subunits arranged in two layers with a seven-fold axis of symmetry. The GroEL oligomer readily dissociates into dimers which suggests that the dimeric form is involved in oligomerization (Gatenby, 1992). A great deal of research is currently underway to determine the three dimensional structure of the GroEL/GroES complex. Ishii et al. (1992) proposed two potential structures that could be differentiated by electron microscopy. Electron microscopic images of oligomeric GroEL viewed from above show a heptamer, with a rectangular profile with four stripes in side-view. It has been proposed that the functional complex consists of two GroEL heptamers stacked on top of each other (Ishii et al., 1992). GroES is thought to bind on top of one of the GroEL heptamers, thus resulting in a holo-chaperonin complex with one oligomeric GroEL and one oligomeric GroES. The GroEL complex has now been crystallized, but the crystals have so far demonstrated limited resolution (Spangfort et al., 1993). This appears to be due to the presence of two forms of GroEL monomers, as demonstrated by discrete bands on SDS polyacrylamide gels. While this work progresses, very interesting structural information continues to come from electron microscopy. Using the scanning transmission electron microscope and gold particles cross-linked to the target protein (dihydrofolate reductase), Braig et al. (1993) observed folding intermediates inside the central cavity of the GroEL 14-mer. If this conclusion is verified, the core provides a sequestered environment for folding and intermediates can be bound at multiple sites by the monomeric members of the ring. Although this finding is appealing, it is difficult to include schematically and, for purpose of illustration, we indicate that folding takes place on the surface of the GroEL oligomer (Fig. 1). A key function of GroEL is binding to other unfolded polypeptide chains. This is probably a critical aspect of chaperonin function. A critical experiment was performed by Bochkareva et al. (1988) who demonstrated through crosslinking experiments that GroEL associates with the unfolded forms of certain proteins. Importantly, this association was disrupted in the presence of ATP, but not in the presence of its nonhydrolyzable derivatives. They

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756 further demonstrated that GroEL does not associate with the folded forms of these polypeptides. The dnaK, dnaJ and grpE genes are known to work together and mutations in these genes have been shown to lead to similar phenotypes. These include: 1) overproduction of all hsp, even at permissive temperatures; 2) a block in host DNA and RNA syntheses at nonpermissive temperatures; and 3) a generalized defect in proteolysis (Georgopoulos et al., 1990; Gross et al., 1990). DnaK and the highly conserved hsp70s from virtually all other species appear to act in the monomeric form. These proteins have a very high level of identity, although sequence comparisons reveal that the amino-terminal two-thirds of the protein is more highly conserved than the carboxylterminal one-third (Neumann et al., 1989; Gross et al., 1990). DnaK and the hsp70s all have similar biochemical properties, including a high affinity ATP-binding site and a peptide binding site (Gething and Sambrook, 1992). Structural studies of mammalian cytosolic hsp70 have shown that an amino-terminal 44 kDa proteolytic fragment has ATPase activity and that this activity is not stimulated by protein binding (Chappell et al., 1987). The structure of the amino-terminal proteolytic fragment has been determined by X-ray crystallography (Flaherty et al., 1991) and has been shown to consist of a two-lobed domain with a deep cleft, which is the ATP-binding site. The structure of this site is very similar to the ATP-binding domain of G-actin (Flaherty et al., 1991) and many of the very highly conserved residues of the N-terminal domain are in this ATP-binding site. The carboxyl-terminal domain has not yet been structurally characterized, but two groups have used sequence comparisons to show that the carboxyl-terminus is similar to the wellcharacterized major histocompatibility complex class I antigeni-presenting molecule (Flajnik et al., 1991; Rippmann et al., 1991). Major histocompatibility complex class I molecules are known to bindpeptides in an extended conformation and it has been proposed that DnaK binds these extended coil formations; i.e., clathrin (Flaherty et al., 1990). Insight into DnaK function has come from both biochemical and genetic studies. The protein has a nonspecific 5'-exonuclease activity, which is inhibited by the alarmone ApppA that accumulates following a temperature-shift (Gatenby, 1992). A great deal of information has come from a mutant (dnaK756), in which the altered protein has been shown to be less

Robert Webb and Louis A. Sherman active against phosphates, more active against diand monophosphates and not inhibited by ApppA. These results may explain why dnaK756 bacteria overproduce heat-shock proteins at all temperatures (Georgopoulos et al., 1990). DnaK is also autophosphorylated, a property which can be inhibited in vitro by interaction with the GrpE protein. Interestingly, the autophosphorylation of DnaK756 protein is refractory to inhibition by GrpE. These are some of the miscellaneous results which lead to the conclusion that GrpE and DnaK interact in the folding pathway of nascent polypeptides. Another possible role for DnaK is protection or renaturation of other proteins during heat shock (Pelham, 1986). DnaK is also essential for the growth of E. coli at high temperatures and essential for cell viability during starvation for carbon. Finally, there is a great deal of evidence that DnaK is involved in the translocation of proteins across membranes, presumably by maintaining them in an unfolded state. This feature will be discussed in greater detail in the functional models to be described below.

C. Thermal Tolerance and the Cellular Thermometer An important question that relates to hsp function is how cells sense changes in temperature. Although the precise answer to this question is not fully understood, the involvement of DnaK as a cellular thermometer has been proposed and the arguments are summarized in Craig and Gross (1991) and in La Rossa and Van Dyk (1991). The model is related to the nature of protein-protein interactions of DnaK, DnaJ, and GrpE and the regulation of Thermal regulation by DnaK can be summarized as follows. Increasing the temperature would increase the substrates (i.e., unfolded segments of proteins) of DnaK and lead to more bound DnaK. Depletion of the pool of free DnaK would then be sufficient to induce the heat-shock response. The fundamental model has DnaK interacting directly with In this scenario, DnaK binds to and keeps it in a dormant state until heat shock signals the release of via a mechanism which may involve DnaK phosphorylation (Hartman et al., 1993). Alternatively, during steady-state growth, would be bound to DnaK, an interaction which could possibly facilitate the degradation of by exposing cleavage sites in the partially unfolded protein. Temperature upshift or a reduction in the pool of free DnaK would lead to the

Chapter 26 Heat Shock and Molecular Chaperones release of and thus prevent this type of degradation (Craig and Gross, 1991). Independent of the precise nature of the mechanism, it appears that the interaction of DnaK with is an important factor in preventing thermal denaturation and generating thermal tolerance. Recent evidence may also indicate the involvement of GroESL in thermal protection. Hartman et al. (1993) have demonstrated that GroEL can sequester proteins in an inactive, yet soluble and refoldable state (see Fig. 1). Since GroEL is also capable of phosphorylation, it is possible that a similar mechanism is at work with GroEL. At a minimum, this data indicates that GroESL will perform a protective role in vivo and likely increase the halflife of proteins, which otherwise might aggregate under physiological conditions. Although this function may protect proteins against inactivation, it would appear that DnaK is the most likely candidate for the actual cellular thermometer that regulates the expression of all heat-shock proteins. II. Functional Aspects, Protein Folding and Localization

A. dnaK Mutants and Functional Domains of DnaK Biochemical studies have demonstrated that the DnaK proteins possess ‘unfoldase’, ATPase and autophosphorylase activities. ATP hydrolysis by one portion of this protein appears to be translated into mechanical energy, changing the conformation of the protein and thus leading to the release of a polypeptide from the binding domain (Liberek, 1991). The first convincing evidence of a division between ATPase and autophosphorylase activity was provided by Cegielska and Georgopoulos (1989) who constructed genetically engineered, truncated forms of the DnaK protein. They showed that deletion of the carboxy end of the protein abolished autophosphorylation while leaving ATPase activity unaffected. As discussed (Flaherty et al., 1990), the solution of the crystal structure of the amino-terminal portion of a DnaK homolog has identified the catalytic pocket for ATP hydrolysis. Inspection of surrounding residues suggested that threonine residue 199 of the E. coli DnaK protein was the phosphate acceptor of the autophosphorylation reaction. This hypothesis was proven by McCarty and Walker (1991) using site-

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directed mutagenesis to change this threonine residue to either alanine, valine or aspartic acid. This work further suggested that the DnaK protein may serve as a cellular thermometer by directly sensing temperature changes, since both ATPase and autophosphorylase activities increase with increasing temperature. Analysis of mutant forms of DnaK has provided much information about specific domains and residues involved in the ATPase, autophosphorylase and protein-binding functions of these proteins as well as the relationship between these functions. A powerful approach to these questions is provided by the selection of bacterial strains with mutant DnaK phenotypes followed by thorough characterization of the mutant alleles. Miyazaki et al. (1992) sequenced the genes responsible for two widely studied phenotypes, both partially characterized for their inability to propagate phage; dnaK756(Ts), first described in 1971 (Georgopoulos and Hershkowitz), and dnaK7 (Ts), first described in 1979 (Itikawa and Ryu). The mutation responsible for dnaK7(Ts) was a one-base change resulting in a premature amber codon at amino acid number 150. The dnaK756(Ts) phenotype was shown to result from three A-for-G changes at nucleotides 95, 1364, and 1403, each resulting in aspartic acid residues rather than the glycine residues normally at these positions. The first of these changes is in the amino-terminal domain of DnaK (residue 32, still some distance from the ATPase catalytic site), whereas the other two changes affect amino acids numbers 455 and 468 of the carboxy-terminal third of this protein of 683 residues. It is not clear if a single change of this type, or some subset of these three changes, is responsible for the conformational change that generates the observed mutant phenotype. Selection for constitutive, high-level expression of DnaK by E. coli generated eleven mutants lacking various aspects of DnaK function (Wild et al., 1992). Dominant mutations which prevent propagation of phage always involved amino acids that had been implicated in ATP hydrolysis, either as part of the adenine binding pocket or as catalytic residues (e. g. Threonine 199). ATP hydrolysis was also shown to be essential for two functions of DnaK: the dissolution of denatured protein aggregates and the renaturation of heat-inactivated RNA polymerase (Skowyra et al., 1990). However, ATP hydrolysis may not be a part of all functions of DnaK . The constitutive expression of a human DnaK homolog, in which the ATP-

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binding domain had been deleted, was shown to protect rat cells from thermal stress (Li et al., 1992). This finding supports the idea that an additional function of DnaK is to act as a ‘macromolecular sponge’ (La Rossa and Van Dyk, 1991), removing misfolded proteins from folding kinetics considerations and preventing their aggregation. Another useful approach for identifying the functional domains of the molecular chaperones is the application of protein structure-prediction algorithms.The algorithm ‘Coils’(Lupas et al., 1991) was used to identify regions in the DnaK proteins from eucaryotic sources, E. coli and the cyanobacterium Synechococcus sp. strain PCC 7942 that were likely to form amphipathic helices (Webb and Sherman, 1992). These regions were identified in the carboxy-terminal third of the protein, a region of moderate to low amino acid sequence conservation. These structures are postulated to serve a role in either the binding of target polypeptides or the oligomerization of the molecular chaperones. Interestingly, the extension of this analysis to the GroEL proteins of E. coli and Synechococcus sp. strain PCC 7942 also revealed domains likely to form these structures. GroEL homologs are biochemically purified and appear to act as highly ordered oligomeric structures while the DnaK proteins are purified and appear to act as monomers. However, DnaK proteins may act as trimers during the disassembly of clathrin-coated vesicles. Electron micrographic studies (Heuser and Steer, 1989) identified DnaK trimers at the hub of clathrin triskelions. This idea is supported by the mutational studies of Wild et al. (1992) mentioned above. These workers reason that the dominant negative mutations that they have characterized in DnaK are typical of those seen for multimeric proteins. These types of mutations are thought to result from mixed oligomers in which mutant subunits affect the function of wild-type subunits in the complex. Mutational analysis of the amphipathic helices identified near the carboxy termini of the DnaK proteins may yield information about the potential for oligomerization of DnaK.

B. Molecular Chaperones and Protein Folding A great deal of effort has been expended to try to unravel the general processes by which chaperonins and DnaK mediate protein folding. The formalism summarized by Lorimer et al. (1992) and Gatenby

Robert Webb and Louis A. Sherman (1993) will be used below. The isomerization of proteins from the unfolded (U) state to the native (N) state involves the transient formation of folding intermediates (I). Thus, folding of proteins, as analyzed in vitro, can be represented by a two step process: Lorimer et al. (1992) represents the two-step process as the interaction of GroEL with the non-native states of proteins. This leads to the formation of a stable binary complex, which in turn suppresses aggregation but inhibits spontaneous folding to the final N-state. This is followed by release of the native protein from the binary GroEL complex. The initial fast step converts the unfolded polypeptide to the intermediate or molten-globule state (Kuwajima, 1989). The Istate is relatively stable and almost as compact as N, although it lacks the close packing of the secondary structural elements that are typical of the native state. The intermediate states are usually less soluble and have a propensity to aggregate. This is most likely due to exposed hydrophobic surfaces that are present in the I-state. In vitro this aggregation can be prevented or reduced by lowering the concentration of protein or lowering the temperature. In both cases this would suppress hydrophobic interactions and thus favor partitioning to the native state. However, such conditions are not possible within the cellular milieu and thus, molecular chaperones have evolved to help catalyze the formation of the native state. There is much recent evidence to indicate that GroEL both prevents aggregation and also inhibits the formation of the native state. GroEL has been shown to prevent aggregation of rhodanese (Martin et al., 1991; Mendoza et al., 1991), Rubisco (Goloubinoff et al., 1989; Viitanen et al., 1990), citrate synthase (Buckner et al., 1991), and αglucosidase. In addition, GroEL will also prevent the spontaneous folding of these; and other proteins into their native forms. It would appear that GroES is not absolutely required for the release process in vitro, but acts to increase the efficiency of the process. Nonetheless, the GroES-enhanced rates of release may be essential in vivo and account for the requirement of GroES for cell viability (Fayet et al., 1989). Interestingly, ATP hydrolysis may also not be essential for release but may be coupled to a requirement for rapid dissociation of the binary complex. The complexities and uncertainties involved in these various reactions are discussed in more detail in Hubbard and Sander (1991), Gatenby (1992), Georgopoulos (1992), and Lorimer et al. (1993).

Chapter 26 Heat Shock and Molecular Chaperones It is clear that DnaK and the hsp70 proteins function in a somewhat different manner than the chaperonins. In the first place, most evidence suggests that they function as 70 kDa monomers rather than as large oligomers. A series of possible mechanisms have been proposed for the functioning of this class of molecular chaperone: coating, threading and plucking. Coating occurs when DnaK molecules interact at many places along a protein to prevent misfolding or aggregation. The threading mechanism can explain the active unfolding of a protein chain. The chaperone complex bound to a section of protein backbone may slide along the chain ‘unraveling’ its folded conformation. A threading mechanism is consistent with the role of hsp70 in protein translocation. The protein to be translocated is converted into an unfolded form in order to be competent for translocation. An alternative to the threading process is called plucking, in which the ATP-bound form of the chaperone binds a polypeptide and releases it after ATP hydrolysis and likely undergoes a series of repeated binding, hydrolysis, release encounters. Thus, in many ways DnaK mediates an unfolded state which prevents aggregation and eventually permits correct folding to the native state.

C. Successive Action of Molecular Chaperones A model that demonstrates the involvement of molecular chaperones in protein folding is shown schematically in Fig. 1, as first described by Hartman et al. (1993). It would appear that there is a cascade of molecular chaperone activities that lead to the proper folding of nascent proteins: 1) DnaK interacts with extended polypeptide sequences emerging from the ribosome. After an appropriate length of the polypeptide is synthesized, it is then available for folding and can adopt the conformation of a collapsed folding intermediate; 2) DnaJ then binds to stabilize the complex; 3) folding is accompanied by the hydrolysis of ATP and the binding of GrpE; 4) the protein is transferred to GroEL; and 5) the native conformation is eventually acquired, mediated by GroEL in an ATP-dependent reaction. In this functional pathway, GroEL acts as the cellular catalyst of protein folding along with other molecular chaperones. The work of Langer et al. (1992) shows that the successive steps of these sequential chaperone reactions are highly coupled. DnaJ has a dual role in

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retarding the ATP-dependent release of DnaK and in directly contributing to the stabilization of the target protein. GrpE drives the reaction forward by coupling the release of the substrate protein from DnaK/DnaJ to the next step of the pathway, the association with GroEL. The DnaK ATPase is stimulated approximately 50-fold by DnaJ and GrpE acting synergistically (the latter facilitating ADP-ATP exchange), but productive release of the target protein from DnaK/DnaJ absolutely requires the presence of GroEL. For the folding pathway described here to be general, all the components should be considered essential genes. Interestingly, the dnaK gene is only quasi-essential, in that an E. coli deletion strain grows slowly at 30 °C and readily acquires suppressor mutations (Georgopoulos et al., 1990; Georgopoulos, 1992). GroEL can possibly function alone, although inefficiently. Alternatively, a second DnaK/DnaJ pair of proteins might exist. As we will describe below, two dnaK genes have been cloned and sequenced in Synechococcus sp. strain PCC 7942.

D. Insertion and Translocation of Membrane Proteins In bacteria, approximately 20% of the polypeptides synthesized are located partially or completely outside of the cytoplasm. The non-cytoplasmic proteins are even more significant in cyanobacteria when one includes all of the proteins located in the photosynthetic lamellae and within the lumen. Therefore, the cell must devote a significant effort to the insertion of membrane proteins and the translocation of proteins across membranes. Very little is known about these processes in cyanobacteria, but we now have two well-described systems that can act as guides for mechanisms that may exist in cyanobacteria. Not surprisingly, much has been learned about the general secretory pathway in E. coli and this work has recently been summarized in a comprehensive review (Pugsley, 1993). In addition, a great deal is now known about the translocation of proteins into the various compartments of mitochondria; this mechanism involves the molecular chaperones at each stage. This research in eucaryotic systems has been most valuable to a better understanding of the heat-shock proteins as molecular chaperones, and the mechanism of protein translocation into mitochondria will thus be presented in some detail.

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Most of our current understanding of bacterial secretion is based on the genetic approach pioneered by the laboratories of J. Beckwith and P. Bassford (Bieker et al., 1990). They isolated mutations that reduced protein export and which were identified by the accumulation of unprocessed presecretory proteins. This technique relies on the fact that certain presecretory hybrids are toxic to cells, when the export machinery is jammed by blocked translocation intermediates. Such mutants confer a Lac– phenotype due to the inability of the moiety to form a tetramer. Using this and related selection procedures, these individuals isolated numerous mutations in genes, which are referred to as sec genes, encoding components of the secretory pathway. These mutations are defined as conferring a conditional-lethal, pleiotropic export defects. More recent genetic and biochemical work has indicated that at least four of these sec genes (secD, secE, secF and secY) are intrinsic membrane proteins and comprise the translocation machinery in the membrane. SecA and SecB are cytoplasmic proteins, although SecA is known to interact with the membrane (Kumamoto, 1990; Oliver et al., 1990). In fact, SecA has been identified as a docking protein, which helps bring the ribosomally-bound protein to the membrane for eventual insertion or translocation. SecB is now recognized as a pilot protein, which helps escort the secretory protein to the membrane. It has been shown that SecB binds to multiple sites on polypeptides and appears to recognize proteins that fold rather slowly. In all regards, SecB is basically a general molecular chaperone and is similar to DnaK in many regards. However, SecB is not induced by heat-shock conditions or by other stresses. Thus, in E. coli, SecB binds to nascent proteins and acts as a molecular chaperone or pilot and escorts the target protein to the membrane. It then docks with and transfers the protein to the SecA protein. In turn, SecA transfers the protein to the SecE/Sec Y complex, resulting in either translocation across or insertion into the membrane (Pugsley, 1993). Research to this date has not implicated the heat-shock proteins in this process, except under conditions where proteins such as SecB are mutated. However, the eucaryotic relatives of DnaK and GroEL have most certainly been implicated in translocation into the mitochondrion (Ostermann et al., 1989; Geli and Glick, 1990; Kang et al., 1990; Glick and Schatz, 1991; Gething and Sambrook, 1992; Glick et al., 1992). Most proteins destined for

Robert Webb and Louis A. Sherman the mitochondria are made on free cytoplasmic polysomes and are imported posttranslationally. Since proteins in their native conformation would not cross the mitochondrial membrane, it is long been recognized that it was necessary to prevent premature folding of precursor proteins by cytosolic ‘antifolding’ proteins. Indeed, these are now known to be the molecular chaperones and they bind to newly synthesized precursors and prevent mature portions from assuming the native conformation (Glick and Schatz, 1991). Recent work has indicated that hsp70 molecules assume this function and operate both in the cytoplasm as well as in the mitochondrial matrix. A model which summarizes current ideas concerning the mechanism of protein translocation into mitochondria is described in Fig. 2. DnaK-like molecules bind to the polypeptide chain to maintain a relatively unfolded conformation. Such an unfolded conformation seems to be necessary to provide competence for translocation. The protein to be translocated then interacts with a receptor which is part of the mitochondrial translocation complex; details of this translocation complex can be found in the above-mentioned reviews and that of Sollner et al. (1992). It is generally thought that the aminoterminal presequence is inserted into the outer membrane and translocated across both the inner and outer membranes, exploiting the energy of the membrane potential across the inner membrane. The DnaK-like molecules in the mitochondrial matrix then bind tightly to the precursor protein. This binding appears to take place at a number of sites as translocation progresses, possibly preventing movement back toward the cytoplasm. The unfolded precursor protein is then transferred to GroEL structures during translocation. GroEL finally mediates protein folding, insertion into the inner mitochondrial membrane, or translocation into the intermembrane space. The diagram in Fig. 2 indicates that there are likely multiple steps which require hydrolysis of ATP. It is thought that GroES is required during these processes to modulate or coordinate the ATPase activity of each GroEL protomer, perhaps to prevent all sites discharging simultaneously which would lead to premature release of only partially folded molecules. With GroEL and GroES interacting, this would prolong the contact between the chaperonin complex and the substrate until the nascent protein no longer was recognized by GroEL and was released. The details of the action of DnaK and GroEL in bacterial

Chapter 26 Heat Shock and Molecular Chaperones

insertion and translocation have not yet been demonstrated and no such work has yet been attempted in cyanobacteria. Nonetheless, the finding of multiple dnaK and groEL genes in Synechococcus sp. strain PCC 7942 and Synechocystis sp. strain PCC 6803, respectively, may point to their involvement in multiple functions, including membrane protein insertion and protein translocation.

E. Thylakoid Membrane Assembly The precise geometry of bound cofactors and protein complexes is essential for the efficient function of photosynthetic membranes, since distance shifts of Ångstroms can seriously affect the coupling of energy or electron carriers. The insertion of individual proteins into thylakoid membranes, the binding of

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cofactors, and the assembly of functional complexes each occur in a highly ordered fashion. In Chlamydomonas sp. the PS II core subunits CP47, D1 and D2 have been shown to be synthesized concertedly (de Vitry et al., 1989). The addition of cofactors such as pheophytin, chlorophyll, quinone and iron to nascent D1 protein emerging from the ribosome occurs concomitantly with the insertion of this protein into the membrane (Kim et al., 1991). A number of soluble protein factors have been identified which are required for the import of proteins into chloroplasts and the insertion of nascent polypeptides into thylakoid membranes. A GroEL homolog was shown to form stable complexes with both subunits of Rubisco, the of ATP synthase and the light harvesting chlorophyll a/b protein (LHCP) of PS II upon their import into

762 chloroplasts (Lubben et al., 1989). The hydrophobia LHCP protein is synthesized on cytosolic ribosomes and must remain soluble in the cytoplasm and chloroplast stroma prior to its final localization. This protein was shown to require ATP and a soluble extract of chloroplasts for efficient insertion into isolated thylakoid membranes (Fulson and Cline, 1988). An unidentified, soluble protein of 65 kDa from chloroplast extracts was shown to participate in this process (Fulson and Cline, 1988). The 25 kDa LHCP protein was isolated from chloroplast stroma as part of a soluble 120 kDa complex (Payan and Cline, 1991). The non-LHCP components of this complex co-migrated in denaturing polyacrylamide gels with the 65 kDa protein previously shown to be necessary for its insertion into isolated thylakoids (Payan and Cline, 1991). However, immunoprecipitation experiments demonstrated that this 65 kDa protein was not a homolog of GroEL (Payan and Cline, 1991). More recently, a chloroplast homolog of DnaK was shown to be associated with the LHCP protein (Yalovsky et al., 1992). The presence of this DnaK-like protein facilitated the integration of LHCP into isolated thylakoid membranes, and depletion of this protein from chloroplast extracts correlated with decreased LHCP insertion activity (Yalovsky et al., 1992). It appears very likely that both the DnaK and GroEL molecular chaperones are necessary for the insertion of proteins into thylakoid membranes. The results discussed above are reminiscent of the more clearly established situation found in the assembly of mitochondrial protein complexes, namely that successive actions of GroEL and DnaK homologs are necessary for these processes (Osterman et al., 1989). GroEL acts in stabilizing newly synthesized polypeptides and preventing their aggregation, while DnaK may act directly at the site of the insertion of these proteins into thylakoid membranes. The conference of membrane-insertion competence upon proteins is most easily visualized as the maintenance of proteins in unfolded or partially folded states by the molecular chaperones, although the precise nature of the molecular interactions for these processes have not been determined. Immunocytochemical studies have localized a large fraction of cellular GroEL protein to the thylakoid membranes of vegetative cells of Anabaena sp. strain PCC 7120 (Jager and Bergmann, 1990). This phenomenon has also been observed for the thylakoid membranes of Synechococcus sp. strain PCC 7942

Robert Webb and Louis A. Sherman (R. Webb and L. Sherman, unpublished results). This is a necessary condition for the involvement of GroEL in photosynthetic membrane assembly. The majority of the immunogold label’ appears to be associated with the outermost thylakoid membrane in cells grown under both iron-sufficient and iron-limited conditions, as though this leaflet was the site of thylakoid membrane assembly. When the cells were subjected to heat stress, many more immunogold particles could be found in the cytosol.

III. Molecular Chaperones of the Cyanobacteria

A. Heat-Shock Proteins of the Cyanobacteria The cyanobacteria respond to heat shock in a manner very similar to that seen in more thoroughly studied bacterial systems such as E. coli. Within minutes of a shift in temperature, general protein synthesis is repressed while a subset of proteins is overexpressed. Many members of this subset are common to all of the cyanobacteria studied to date. These isotopic protein labeling investigations have been performed by a number of groups (Borbely et al., 1985; Bhagwat and Apte 1989; Webb et al., 1990; Lehel et al., 1992; Blondin et al., 1993), and their results are summarized in Table 2. An examination of this compiled data reveals that the set of proteins expressed in response to this stimulus can be grouped into four size classes: small (11–4 kDa), intermediate (40–50 kDa), large (55–80 kDa) and those greater than 90 kDa. Only a few of these proteins have been unambiguously identified. The DnaK protein of Synechocystis sp. strain PCC 6803 (Lehel et al., 1992) and the GroEL proteins of both Synechococcus sp. strain PCC 7942 (Webb et al., 1990) and Synechocystis sp. strain PCC 6803 (Lehel et al., 1992) were identified using antibodies. The cyanobacterial species examined in detail each expressed proteins of 65 kDa which did not seem to have homologs in the set of proteins expressed by E. coli. One possibility is that the 65 kDa protein seen in Synechococcus sp. strain PCC 7942 represents the second copy of DnaK that has been cloned from this cyanobacterium (R. Webb and L.A. Sherman, unpublished results). Moreover, there is not a direct, one-to-one correspondence in the set of heat-shock proteins observed in the closely related Synechococcus sp. strains PCC 6301 and PCC 7942. This is likely due to slight differences in the protocols

Chapter 26 Heat Shock and Molecular Chaperones

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was shown to interact with the and phosphates of ATP. Aspartic acid residue 8 chelates the metal ion of the nucleotide-metal complex while glycine residue 10 abuts the in a way that would be difficult for an amino acid with a larger side chain. The hydroxyl groups from threonine numbers 11 and 12 form hydrogen bonds to the phosphate oxygens. The ATPase catalytic site is equally highly conserved. Neuman et al. (1989) first proposed the region of the hsp70 consensus between amino acids numbers 219 and 223 as the catalytic site based on sequence similarity to the protein kinases. This suggestion was supported by the crystallographic work of Flaherty et al. (1991) who extended this comparison to include actin. Aspartic acid residue 206 (a histidine in actin) serves as the proton acceptor in the ATPase reaction and, as mentioned, threonine 204 is the site of DnaK autophosphorylation. This region is identical in the bacterial and cyanobacterial species discussed above. These species also show greater than 90% sequence identity in comparison to the crystallized Hsc70 sequence in the area of the adenine binding pocket. This pocket is formed by one section of a long helix (glutamic acid residue 268 to arginine 272 of the hsc70 sequence), a helix (glycine 339isoleucine 343) and aspartic acid residue 366 (Flaherty et al., 1991).

C. Cyanobacterial Heat-Shock Genes used to induce this response but may further suggest differences in the regulation of the expression of the different proteins.

B. Nucleotide-Binding Motif Sequence Comparisons As discussed, there is a fairly high degree of amino acid sequence identity in pairwise comparisons of members of either the GroEL or DnaK families. Comparisons of the ATPase catalytic sites of DnaK proteins from eucaryotic, bacterial and cyanobacterial sources reveal extraordinary sequence conservation related to function. Amino acid residues 6 to 16 from E. coli, Bacillus subtilis, Synechocystis sp. strain PCC 6803 (Chitnis and Nelson, 1991) and both DnaK proteins of Synechococcus sp. strain PCC 7942 (R. Webb and L.A. Sherman, unpublished results) are identical to the Hsc70 sequence crystallized by Flaherty et al. (1991). This region

Thus far, only the genes for cyanobacterial GroEL, GroES and DnaK homologs from Synechococcus sp strain PCC 7942 and Synechocystis sp. strain PCC 6803 have been cloned and sequenced. Interestingly, multiple copies of the genes for these molecular chaperones are being found. groES and groEL genes constitute an operon in Synechococcus sp. strain PCC 7942 (Webb et al., 1990) and low-stringency Southern hybridizations failed to detect other copies of these genes. However, Synechocystis sp. strain PCC 6803 possesses two groEL genes. One stands alone (Chitnis and Nelson, 1991) while the other is again bicistronic with groES (Lehel et al., 1993). Conversely, one dnaK gene has been identified in Synechocystis sp. strain PCC 6803 (Chitnis and Nelson, 1991) while two copies of dnaK have been found in Synechococcus sp. strain PCC 7942 (R. Webb and L.A. Sherman, unpublished results). The significance of this situation is unclear but presents interesting possibilities. The cyanobacteria must direct proteins to more compartments than most

764 other eubacteria due to the presence of thylakoid membranes. It is tempting to speculate that different versions of the molecular chaperones may be associated with and function within these specialized compartments. This is analogous to the situation found in eucaryotic cells where slightly different GroEL homologs function in the mitochondrial lumen and endoplasmic reticulum.

D. Transcriptional Response of the Cyanobacteria to Heat Shock Examination of the transcriptional response of the cyanobacteria to environmental stresses has begun using the available homologous gene probes. The groESL operon of Synechococcus sp. strain PCC 7942 encodes a bicistronic 2.4-kb transcript which accumulated to 120-fold higher levels within 20 min after cells were shifted from 30 to 45 °C (Webb et al., 1990). The abundance of this transcript decreases to baseline levels after 60–120 min. Very similar kinetics are observed for the expression groESL operon of Synechocystis sp. strain PCC 6803 (Lehel et al, 1993), the second groEL gene and dnaK gene of this cyanobacterium (Chitnis and Nelson, 1991) and the two dnaK genes of Synechococcus sp. strain PCC 7942 (R. Webb and L.A. Sherman, unpublished results). The promoter region of the Synechococcus sp. strain PCC 7942 groESL operon shares some similarity with the E. coli consensus heat-shock promoter at both the –10 and –35 positions and differs significantly from other promoters identified in this cyanobacterium (Webb et al., 1990). This suggests that an alternative RNA polymerase subunit is involved in the induction of this operon. It should be noted that the recA gene of Synechococcus sp. strain PCC 7002 also possesses a promoter with strong sequence similarity to the promoters of E. coli (Murphy et al., 1990). Subsequent investigations have demonstrated that RecA protein levels increased after heat shock (V. L. Stirewalt and D. A. Bryant, unpublished results).

E. Mutational Studies of the Cyanobacterial Molecular Chaperones The use of mutant bacterial strains continues to be an enormously powerful tool for the elucidation of protein function in vivo. This approach is only now becoming possible for the study of molecular chaperone function in the cyanobacteria as their

Robert Webb and Louis A. Sherman genes have only recently been identified and sequenced. A mutant strain of Synechococcus sp. strain PCC 7942, in which groEL expression does not respond to heat shock, has been constructed (R. Webb, unpublished results). Constitutive groEL expression, sufficient to allow cellular growth, results from the read-through transcription of a kanamycin resistance cassstte-groEL fusion in the Synechococcus sp. strain PCC 7942 chromosome. When these mutant cells are incubated for 12 h at 45 °C, they appear longer and wider than cells grown at 30 °C or wildtype cells at either temperature. This is similar to the phenotype observed for groEL(Ts) mutants of E. coli which form filaments at the non-permissive temperature. In nature, light intensities incident on cyanobacteria at mid-day can approach 2000 This intensity is known to adversely affect the function of PS II centers by a phenomenon known as photoinhibition. Recovery from photoinhibition has been shown to require the synthesis and assembly of new PS II reaction centers. The mutant Synechococcus sp. strain exhibits a growth rate comparable to that of the wild-type at but grows increasingly poorly as the light intensity is increased to potentially photoinhibitory intensities. The mutant strain grows at 33 to 56% of the wild-type rate under illumination of and 16 to 40% of the wild-type rate when subj ected to It is hypothesized that under these conditions there is increased cellular demand for GroEL protein to aid the reassembly of damaged PS II centers in the thylakoid membranes. When considered together with the finding of GroEL protein in association with cyanobacterial thylakoid membranes, the failure to increase GroEL synthesis could decrease the efficiency of this repair process and result in poor growth at these light intensities. Support for this idea awaits further investigation.

IV. Summary and Future Directions There is a wealth of literature detailing the action of the molecular chaperones in mitochondrial systems and E. coli. By comparison, the information available from cyanobacterial systems is rather limited. This knowledge base does not need to increase by being derivative but rather should exploit the unique opportunities presented by the use of a bacterial system with more compartments and protein destinations than the bacteria studied to date. Information gained by studying photosynthetic

Chapter 26 Heat Shock and Molecular Chaperones membrane assembly should be directly applicable to studies of all membrane systems. Two lines of research seem to hold the most promise at this time. It should be possible to develop an in vitro membrane assembly system from cyanobacterial material. This system would allow direct approaches to uncovering the set of cofactors and proteins necessary for the addition of prosthetic groups to membrane proteins during the assembly of functional membrane-bound complexes. Cyanobacterial systems are also valuable for investigations into the functions performed by products of the different copies of the molecularchaperone genes. Cyanobacterial cells and their membranes are more easily manipulated than isolated organelles for the study of secretion, membrane assembly and protein localization and importantly, cyanobacteria can be genetically engineered allowing the use of site-directed or regulatory mutants in physiological studies. References Bhagwat AA and Apte SK (1989) Comparative analysis of proteins induced by heat shock, salinity, and osmotic stress in the nitrogen-fixing cyanobacteriuin Anabaena sp. strain L-31. J Bacteriol 171: 5178–5189 Bieker KL, Phillips GJ and Silhavy TJ (1990) The sec and prl genes of Escherichia coli. J Bioenerg Biomembr 22:291 –310 Blondin PA, Kirby PJ and Barnum SR (1993) The heat-shock response and acquired thermotolerance in three strains of cyanobacteria. Curr Microbiol 26: 79–84 Bochkareva ES, Lissin NM and Girshovich AS (1988) Transient association of newly synthesized unfolded proteins with the heat-shock GroEL protein. Nature 336: 254–257 Borbely GG, Suranyi G, Korcz A and Palfi Z (1985) Effect of heat shock on protein synthesis in the cyanobacterium Synechococcus sp. strain PCC 6301. J Bacteriol 161: 1125– 1130 Braig K, Simon M, Furuya F, Hainfeld JF and Horwich AL (1993) A polypeptide bound by the chaperonin GroEL is localized within a central cavity. Proc Natl Acad Sci USA 90: 3978–3982 Buckner J, Schmidt M, Fuchs M, Jaenicke R, Rudolph R, Schmid FX and Kiefhaber T (1991) GroE facilitates refolding of citrate synthase by suppressing aggregation. Biochemistry 30:1586– 1591 Cegielska A and Georgopoulos C (1989) Functional domains of the Escherichia coli DnaK heat-shock protein as revealed by mutational analysis. J Biol Chem 264: 21112–21130 Chappell TG, Konforti BB, Schmid SL and Rothman JE (1987) The ATPase core of a clathrin uncoating protein. J Biol Chem 262: 746–751 Chitnis PR and Nelson N (1991) Molecular cloning of the genes encoding two chaperone proteins of the cyanobacterium Synechocystis sp. PCC 6803. J Biol Chem 266: 58–65

765 Craig EA and Gross CA (1991) Is hsp70 the cellular thermometer? Trends Biochem Sci 16: 135–140 Craig EA, Gambill BD and Nelson RJ (1993) Heat-shock proteins: Molecular chaperones of protein biogenesis. Microbiol Rev 57:402–414 de Vitry C, Olive J, Drapier D, Recouvrer M and Wollman F-A (1989) Posttranslational events leading to the assembly of Photosystem II protein complex: A study using photosynthesis mutants from Chlamydomonas reinhardtii. J Cell Biol 109: 991–1006 Ellis RJ (1990a) Molecular chaperones: The plant connection. Science 250: 954–959 Ellis RJ (1990b) The molecular chaperone concept. Seminars Cell Biol 1: 1–9 Ellis RJ and Hemmingsen SM (1989) Molecular chaperones: Proteins essential for the biogenesis of some macromolecular structures. Trends Biochem Sci 14: 339–342 Ellis RJ and van der Vies SM (1991) Molecular chaperones. Annu Rev Biochem 60: 321–347 Fayet O, Ziegelhoffer T and Georgopoulos C (1989) The groES and groEL heat-shock gene products of Escherichia coli are essential for bacterial growth at all temperatures. J Bacteriol 171: 1379–1385 Flaherty KM, Deluca-Flaherty C and McKay DB (1990) Three dimensional structure of the ATPase fragment of a 70K heatshock cognate protein. Nature 346: 623–628 Flaherty KM, McKay DB, Kabash W and Holmes K (1991) Similarity of the three-dimensional structure of actin and the ATPase fragment of a 70 kDa heat-shock cognate protein. Proc Natl Acad Sci USA 88: 5041–5045 Flajnik M, Canel C, Kramer J and Kasahara (1991) Hypothesis: Which came first, MHC class I or class II? Immunogenetics 33: 295–300 Fulson DR and Cline K (1988) A soluble protein factor is required in vitro for membrane insertion of the thylakoid precursor protein, pLHCP. Plant Physiol 88: 1146–1153 Gage DJ and Neidhardt FC (1993) Modulation of the heat-shock response by one-carbon metabolism in Escherichia coli. J Bacteriol 175: 1961–1970 Gamer J, Bujard H and Bukau B (1992) Physical interaction between heat-shock proteins DnaK, DnaJ and GrpE and the bacterial heat-shock transcription factor Cell 69: 833–842 Gatenby AA (1992) Protein folding and chaperonins. Plant Mol Biol 19:677–687 Geli V and Glick B (1990) Mitochondrial protein import. J Bioenerg Biomembr 22: 725–751 Georgopoulos C (1992) The emergence of the chaperone machines. Trends Biochem Sci 17: 295–299 Georgopoulos CP and Hershkowitz I (1971) Escherichia coli mutants blocked in lambda DNA synthesis. In: A.D. Hershey (ed) The Bacteriophage Lambda, pp 553–564. Cold Spring Harbor Laboratory Press, Cold Spring Harbor Georgopoulos C, Ang D, Liberek K and Zylicz M (1990) Properties of the Escherichia coli heat-shock proteins and their role in bacteriophage growth. In: Morimoto R, Tissieres A, and Georgopoulos C (eds) Stress Proteins in Biology and Medicine, pp 191–221. Cold Spring Harbor Laboratory Press, Cold Spring Harbor Gething M-J and Sambrook J (1992) Protein folding in the cell. Nature 355: 33–45 Glick B and Schatz G (1991) Import of proteins into mitochondria.

766 Annu Rev Genet 25: 21–44 Click B, Beasley EM and Schatz G (1992) Protein sorting in mitochondria. Trends Biochem Sci 17: 453–459 Goloubinoff P, Christeller JT, Gatenby AA and Lorimer GH (1989) Reconstitution of active dimeric ribulose bisphosphate carboxylase from an unfolded state depends on two chaperonin proteins and Mg-ATP. Nature 342: 884–889 Gross CA, Straus D, Erickson JW and Yura T (1990) The function and regulation of heat-shock proteins in Escherichia coli. In: Morimoto R, Tissieres A, and Georgopoulos C (eds) Stress Proteins in Biology and Medicine, pp 167–189. Cold Spring Harbor Laboratory Press, Cold Spring Harbor. Grossman AD, Erickson JW and Gross CA (1984) The htpR gene product of E. coli is a sigma factor for heat-shock promotors. Cell 38: 383–390 Hartman DJ, Surin BP, Dixon NE, Hoogenraad NJ and Høj PV (1993) Substoichiometric amounts of the molecular chaperones GroEL and GroES prevent thermal denaturation and aggregation of mammalian mitochondrial malate dehydrogenase in vitro. Proc Natl Acad Sci USA 90: 2276–2280 Heuser J and Steer CJ (1989) Trimeric binding of the 70 kDa uncoating ATPase to the vertices of clathrin triskelia: A candidate intermediate in the vesicle uncoating reaction. J Cell Biol 109: 1457–1466 Hubbard TJP and Sander C (1991) The role of heat shock and chaperone proteins in protein folding: Possible molecular mechanisms. Protein Eng 4: 711–717 Ishii N, Taguchi MS and Yoshida M (1992) Structure of holochaperonin studied with electron microscopy. Oligomeric cpn 10 on top of two layers of cpn60 rings with two stripes each. FEBS Lett 299: 169–174 Itikawa H and Ryu RY (1979) Isolation and characterization of a temperature sensitive dnaK mutant of Escherichia coli B. J Bacteriol 138:339–344 Kang P-J, Ostermann J, Shilling J, Neupert W, Craig EA and Pfanner N (1990) Requirement for hsp70 in the mitochondrial matrix for translocation and folding of precursor proteins. Nature 348: 137–143 Kim J, Klein PG and Mullet JE (1991) Ribosomes pause at specific sites during synthesis of membrane bound chloroplast reaction center protein D1. J Biol Chem 266:14931–14938 Kumamoto CA (1990) SecB protein: A cytosolic export factor that associates with nascent exported proteins. J Bioenerg Biomembr 22: 337–352 Kuwajima K (1989) The molten globule state as a clue for understanding the folding and cooperativity of globular-protein structure. Proteins 6: 87–103 Landick R, Vaughn V, Lau ET, VanBogelen RA, Erickson JW and Neidhardt FC (1984) Nucleotide sequence of the heatshock regulatory gene of E. coli suggests its protein product may be a transcriptional factor. Cell 38: 175–182 Langer T, Lu C, Echols H, Flanagan J, Hayer MK and Hartl F-U (1992) Successive action of DnaK, DnaJ and GroEL along the pathway of chaperone-mediated protein folding. Nature 356: 683–689 La Rossa RA and Van Dyk TK (1991) Physiological roles of the DnaK and GroE stress proteins: Catalysts of protein folding or macromolecular sponges? Mol Microbiol 5: 529–534 Laskey RA, Honda BM, Mills AD and Finch JT (1978) Nucleosomes are assembled by an acidic protein which binds histones and transfers them to DNA. Nature 275: 416–420

Robert Webb and Louis A. Sherman Lehel C, Wada H, Kovacs E, Torok Z, Gombos Z, Horvath I, Murata N and Vigh L (1992) Heat-shock protein synthesis of the cyanobacterium Synechocystis sp. PCC 6803: Purification of the GroEL-related chaperonin. Plant Mol Biol 18: 327–336 Lehel C, Loss D, Wada H, Gyorgyei J, Horvath I, Kovacs E, Murata N and Vigh L (1993) A second groEL-like gene, organized in a groESL operon is present in the genome of Synechocystis sp. PCC 6803. J Biol Chem 268: 1799–1804 Li GC, Ligeng LL, Liu RY, Rehman M and Lee WMF (1992) Heat-shock protein hsp70 protects cells from thermal stress even after deletion of its ATP-binding domain. Proc Natl Acad Sci USA 89: 2036–2040 Liberek K, Skowyra D, Zylicz M, Johnson C and Georgopoulos C (1991) The Escherichia coli DnaK chaperone, the 70 kDa heat-shock protein eukaryotic equivalent, changes conformation upon ATP hydrolysis, thus triggering its dissociation from a bound target protein. J Biol Chem 266: 14491–14496 Lorimer GH, Todd MJ and Viitanen PV (1993) Chaperonins and protein folding: Unity and disunity of mechanisms. Phil Trans R Soc Lond B 339: 297–304 Lubben TH, Donaldson GK, Viitanen PV and Gatenby AA (1989) Several proteins imported into chloroplasts form stable complexes with the GroEL-related chloroplast molecular chaperone. Plant Cell 1:1223–1230 Lupas A, Van Dyk M and Stock J (1991) Predicting coiled coils from protein sequences. Science 252: 1162–1164 Martin J, Langer T, Boteva R, Schramel A, Horwich AL and Hartl F-U (1991) Chaperonin-mediated protein folding at the surface of groEL through a ‘molten globule’-like intermediate. Nature 352: 36–42 McCarty JS and Walker GC (1991) DnaK as a thermometer: Threonine-199 is site of autophosphorylation and is critical for ATPase activity. Proc Natl Acad Sci USA 88: 9513–9517 Mendoza JA, Rogers E, Lorimer GH and Horowitz, PM (1991) Chaperonins facilitate the in vivo folding of monomeric mitochondrial rhodanese. J Biol Chem 266: 13044–13049 Miyazaki T, Tanaka S, Fujita H and Itikawa H (1992) DNA sequence analysis of the dnaK gene of Escherichia coli B and of two dnaK genes carrying the temperature-sensitive mutations dnaK7(Ts) and dnaK756(Ts). J Bacteriol 174: 3715–3722 Murphy RC, Gasparich GE, Bryant DA and Porter RD (1990) Nucleotide sequence and further characterization of the Synechococcus sp. strain PCC 7002 recA gene: Complementation of a cyanobacterial recA mutation by the Escherichia coli recA gene. J Bacteriol 172: 967–976 Neumann D, Nover L, Parthier B, Rieger R, Scharf K-D, Wollgiehn R and Nieden UZ (1989) Heat shock and other stress response systems of plants. Biolog Zentral 108: 1–156 Neidhart FC and Van Bogelen RA (1987) Heat-shock responses. In: Neidhart FC, Ingraham JL, Low KB, Magasanik B, Schaechter M, and Umbarger HE (eds) Escherichia coil and Salmonella typhimurium : Cellular and Molecular Biology, pp 1334–1345. American Society for Microbiology, Washington, DC. Oliver DB, Cabelli RJ and Jarosik GP (1990) SecA protein: Autoregulated initiator of secretory precursor protein translocation across the E. coli plasma membrane. J Bioenerg Biomembr 22: 311–336 Ostermann J, Horwich AL, Neupert W and Hartl F-U (1989) Protein folding in mitochondria requires complex formation with hsp60 and ATP hydrolysis. Nature 341: 125–130

Chapter 26 Heat Shock and Molecular Chaperones Payan L and Cline K (1991) A stromal protein factor maintains the solubility and insertion competence of an imported thylakoid membrane protein. J Cell Biol 112: 603–613 Pelham HRB (1986) Speculations on the functions of the major heat shock and glucose-regulated proteins. Cell 46: 959–961 Pugsley AP (1993) The complete general secretory pathway in gram-negative bacteria. Microbiol Rev 57: 50–108 Rippmann F, Taylor W, Rothbard J and Green NM (1991) A hypothetical model for the peptide binding domain of hsp70 based on the peptide binding domain of HLA. EMBO J 10: 1053–1059 Ritossa F (1962) A new puffing pattern induced by temperature shock and DNP in Drosophila. Experientia 18: 571–73 Skowyra D, Georgopoulos C and Zylicz M (1990) The E. coli dnaK gene product, the hsp70 homolog, can reactivate heatinactivated RNA polymerase in an ATP hydrolysis-dependent manner. Cell 62: 939–944 Söllner T, Rassow J, Wiedmann M, Schlossmann J, Keil P, Nuepert W and Pfanner N (1992) Mapping of the protein import machinery in the mitochondrial outer membrane by crosslinking of translocation intermediates. Nature 355:84–87 Spangfort MD, Surin BP, Oppentocht JE, Weibull C, Carlemalm E, Dixon NE and Svensson LA (1993) Crystallization and preliminary X-ray investigation of the Escherichia coli molecular chaperone cpn60 (GroEL). FEBS Lett 320: 160– 164 Straus D, Walter WA and Gross CA (1987) The heat-shock response of E. coli is regulated by changes in the concentration of Nature 329: 348–351 Straus D, Walter WA and Gross CA (1990) DnaK, DnaJ and GrpE heat-shock proteins negatively regulate heat-shock gene expression by controlling the synthesis and stability of Genes Dev 4: 2202–2209 Tilly K, McKittrick N, Zylicz M and Georgopoulos C (1983) The

767 dnaK protein modulates the heat-shock response of Escherichia coli. Cell 34: 641–646 Van Bogelen RA, Kelley PM and Neidhardt FC (1987) Differential induction of heat shock, SOS, and oxidation stress regulons and accumulation of nucleotides in Escherichia coli. J Bacteriol 169:26–32 Van Bogelen RA, Hutton ME and Neidhardt FC (1990) Geneprotein database of Escherichia coli K-12: Edition 3. Electrophoresis 11: 1131–1166 Viitanen PV, Lubben TH, Reed J, Goloubinoff P, O’Keefe DP and Lorimer GH (1990) Chaperonin-facilitated refolding of ribulosebisphosphate carboxylase and ATP hydrolysis by chaperonin 60 (groEL) are dependent. Biochemistry 29: 5665–5671 Webb R and Sherman LA (1992) Chaperones classified. Nature 359: 485–486. Webb R, Reddy KJ and Sherman LA (1990) Regulation and sequence of the Synechococcus sp. strain PCC 7942 groESL operon, encoding a cyanobacterial chaperonin. J Bacteriol 172: 5079–5088 Wild J, Kamath-Loeb A, Ziegelhoffer E, Lonetto M, Kawasaki Y and Gross CA (1992) Partial loss of function mutations in DnaK, the Escherichia coli homologue of the 70 kDa heatshock proteins, affect highly conserved amino acids implicated in ATP binding and hydrolysis. Proc Natl Acad Sci USA 89: 7139–7143 Yalovsky S, Paulsen H, Michaeli D, Chitnis P and Nechustai R (1992) Involvement of a chloroplast HSP70 heat-shock protein in the integration of a protein (light-harvesting complex protein precursor) into the thylakoid membrane. Proc Natl Acad Sci USA 89: 5616–5619 Zeilstra-Ryalls J, Fayet O and Georgopoulos C (1991) The universally conserved GroE (Hsp60) chaperonins. Annu Rev Microbiol 45: 301–325

Chapter 27 Heterocyst Metabolism and Development C. Peter Wolk MSU-DOE Plant Research Laboratory, Michigan State University, East Lansing, Ml 48824, USA

Anneliese Ernst Lehrstuhl für Physiologie und Biochemie der Pflanzen, Universität Konstanz, D7750 Konstanz, Germany

Jeff Elhai Dept. of Biological Sciences, Florida International University, Miami, FL 33199, USA Summary I. What is a Heterocyst? II. Genetic Tools III. Metabolism of Mature Heterocysts A. Requirements for Anaerobic Fixation of 1. Nitrogenase Genes and Neighboring DNA 2. Electron Donor to Nitrogenase 3. Organic Electron Donors 4. Intercellular Movement of Carbon Compounds 5. Conduits for Intercellular Movement 6. Photosystem I: Source of ATP and Reductant 7. Do Heterocysts Have a Photosystem II? B. Additional Requirements for Aerobic Fixation of 1. Introduction 2. Barrier to Permeation of Oxygen 3. The Respiratory Apparatus of Heterocysts 4. Abundant Synthesis of Nitrogenase 5. Antioxidative Agents C. Control of Nitrogenase Activity 1. Introduction: Control of Synthesis of Nitrogenase 2. Metabolic Control 3. Modification of Dinitrogenase Reductase D. Other Metabolism 1. Metabolism of Fixed Nitrogen a. Assimilation and Export b. Synthesis of Nitrogenous Metabolites and Macromolecules 2. Metabolism of 3. Metabolism of IV. The Differentiation Process A. Introduction B. Candidate Principles 1. During the Course of Differentiation, Genes are Expressed in an Ordered Sequence 2. The Expression of Genes at One Stage Depends Upon Gene Products that Appear at a Previous Stage D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 769–823. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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3. Regulation of Expression Occurs Primarily, But Not Exclusively, at the Level of Transcription C. Very Early Responses to Nitrogen Deprivation D. HetR Plays a Critical Role Early in Differentiation E. Genes Affecting the Maturation of Proheterocysts F. Deposition of the Heterocyst Envelope 1. Mutant Phenotypes 2. Heterocyst Envelope Polysaccharide 3. Envelope Glycolipids G. Developmental Control of Nitrogenase Activity V. Pattern Formation and Perpetuation A. The Normal Pattern B. Natural and Induced Variations on the Normal Pattern 1. Failure to Differentiate 2. Differentiation Only at the Termini of Filaments 3. Formation of Strings of Heterocysts 4. Phenotypes Not Reported C. Models of Pattern Formation D. HetR, Part 2: The Role in Pattern Formation E. Possible Signals of Positional Information VI. Relationship of Diverse Differentiation Processes in Cyanobacteria Acknowledgments References

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Summary Heterocysts are differentiated cells that are specialized for fixation of in an aerobic environment. In heterocysts in the light, Photosystem I generates ATP, but no photosynthetic production of takes place. Instead, reductant moves into heterocysts from vegetative cells. In return, fixed nitrogen moves from heterocysts to vegetative cells. In neither case is there certainty about the identity of the traffic molecules. Pathways of electron-donation to have been extensively investigated, but their in-vivo importance remains to be critically tested. Nitrogenase in heterocysts is protected from inactivation by by a variety of means, principally by enhanced respiration and by a barrier, the heterocyst envelope, to entry of However, the respiratory apparatus and the biosynthetic processes that result in synthesis of the barrier have been little studied. The detailed mechanisms underlying metabolic, environmental, and developmental control of nitrogenase are under investigation. Studies of heterocyst development are being greatly facilitated by recent advances in the genetics of Anabaena sp. An autoregulated gene, hetR, that is activated shortly after nitrogenstepdown is critical for the differentiation of heterocysts. Two enigmas remain to be answered: how is it determined which cells will differentiate; and, after differentiation is initiated, what intercellular interactions and intracellular mechanisms regulate the progression of the differentiation process? An evolutionary and biochemical relationship between the processes leading to the formation of heterocysts and akinetes is suggested.

I. What is a Heterocyst? Perhaps 2.2 billion years before the writing of this chapter (Cloud, 1972; Nagy, 1974), our world was anaerobic, and autotrophic microorganisms that could Abbreviations: Anabaena cylindrica (F) – the Fogg strain; Anabaena cylindrica (W) – Anabaena cylindrica strain ATCC 29414; DAB – diaminobenzidine; DBMIB – dibromomethylisopropylbenzoquinone; HQNO – 2-(n-heptyl)-4-hydroxyquinoline N-oxide; PEP – phosphoenolpyruvate

gain their energy from sunlight, their carbon and (often) their nitrogen from the atmosphere, and their reductant from water were in the ascendancy. As they grew and flourished, they polluted the world with a noxious byproduct, oxygen, derived from photooxidation of water. Although inhibitory to many biochemical reactions, oxygen was seldom as detrimental as it was to the process of nitrogen fixation, because of the exceptionally negative reducing potentials required for that process. Some of these autotrophic microorganisms adapted to the

Chapter 27 Heterocyst Metabolism and Development newly changed environment by making their nitrogen-fixing enzymes (nitrogenases) only when oxygen happened to be absent. Other such microorganisms found that although it was necessary to stop nitrogen fixation during the day, when their intracellular oxygen concentration was highest, they could fix nitrogen during the night, when their intracellular was lowest. Yet others met the challenge of increased atmospheric oxygen by modifying a small percentage of their cells for the task of fixing nitrogen (Stewart and Lex, 1970; Mullineaux et al., 1981; Gallon, 1981, 1992; Fay, 1992). In the last of these groups, physiological differentiation was accompanied by morphological differentiation (Fig. 1). The nitrogen-fixing cells are known as heterocysts (‘hetero’ = other), from the time when their function was unknown (Fritsch, 1991). Heterocysts were presumably not the first differentiated microbial cells; one may speculate that Clostridium sp. had already learned to sporulate,

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and perhaps even cyanobacteria had developed the ability to make akinetes, in response to a limitation of energy or nutrients. However, this was differentiation with a difference: Whereas the role of spores, akinetes and other dormant types of cells is to promote the survival of the individual cell until certain nutrients become more abundant, the role of heterocysts is to provide a needed nutrient immediately to all cells in the filament. The relationship between heterocysts and vegetative cells is thus mutualistic: The heterocysts provide the nitrogen that is needed by the organism, leaving to the other cells the task of acquiring carbon. Heterocysts, whose essential nature involves metabolic exchange, evolved in filamentous cyanobacteria. They are unknown in unicellular cyanobacteria, and have not been reported to differentiate in unicellular mutants of filamentous species. Optimal proliferation when fixed nitrogen is limiting requires coordination of the differentiation of some vegetative cells into N2-fixing heterocysts,

772 while others continue to fix and replicate. It remains a challenge to elucidate how it is determined which vegetative cells are to differentiate into heterocysts, and how they do so. That nitrogen fixation is restricted to heterocysts in aerobically grown cultures of Anabaena sp. (Fay et al., 1968) was shown by a variety of indirect means, culminating in the direct demonstration of recovery of a large fraction of nitrogenase activity and of a very large fraction of nitrogenase in isolated heterocysts of A. variabilis (Peterson and Wolk, 1978b; reviewed by Wolk, 1982). More recently, localization of nitrogenase in heterocysts was shown by immunoelectronmicroscopy (Murry et al., 1984a; Bergman et al., 1986; Braun-Howland et al., 1988; Rai et al., 1989; Bergman and Rai, 1989) and, for the product of a particular nif operon, by in vivo transcriptionalreporting(Fig. 2; Elhai andWolk, 1990). It appears likely, but has not been established, that in some strains of heterocyst-forming cyanobacteria, but not others, nitrogenase is active in all vegetative cells under anaerobic conditions (see Section III A, 1). In order to avoid inactivation of nitrogenase by oxygen, heterocysts stop synthesizing oxygen, and limit the rate of entry of that gas. Presumably because the van der Waal’s radii of nitrogen and oxygen are similar, 1.5 Å and 1.4 Å, respectively, evolution apparently found no way to permit to enter while excluding However, a barrier that consists of laminae of long-chain lipid molecules, and that is surrounded in turn by a protective layer of polysaccharide, reduces the permeability of both gases to the extent that the that enters still suffices for the needs of the organism for nitrogen, while the that enters can be reduced to water by respiration (Walsby, 1985). At the current atmospheric ratio of approximately this strategy requires that a substantial fraction of the electron flux generated by photosynthesis throughout the cyanobacterial filament be devoted to the process of reducing to and to (Turpin et al., 1985; Murry and Wolk, 1989). Because photosynthetic water-splitting, the ultimate source of the electrons used for fixation, cannot continue in heterocysts, it must be replaced by movement of reductant from (photosynthetic) vegetative cells to heterocysts. In addition, there must be movement of fixed nitrogen from heterocysts to vegetative cells. Therefore, some channel(s) must remain open between the two types of cells. Although

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entry of around the periphery of the heterocyst can be largely eliminated by the envelope layers, no such barrier may be present in the polar region in order not to block the channel for movement of nutrients. Such a channel should therefore have a cross-sectional area no larger than required to support the needed flux of nutrients; and respiration, especially next to those channels, is required to maintain microaerobiosis within the heterocysts. In fact, a ‘honeycomb’ of membranes rich in oxidative enzymes is located in the region of the heterocysts near the channels (see Section III B, 2). Nature having invented a microaerobic cell for an aerobic world, one wonders to what uses it may be put, in addition to nitrogen fixation. It is our guess that the process of heterocyst differentiation evolved after, and was in part based on, the process of akinete differentiation, but that is an open question. The polysaccharide of heterocyst envelopes has been found in the envelopes of akinetes (Cardemil and Wolk, 1979), and the glycolipids of heterocyst envelopes have been found in a preparation of akinetes (Soriente et al., 1993; see also Sutherland et al., 1985). If major envelope

Chapter 27 Heterocyst Metabolism and Development constituents were borrowed from akinete differentiation, then a key new element in the evolution of the heterocyst envelope may have been the method of maintaining metabolic junctions open to vegetative cells while largely blocking ingress of gases. Even that element may represent an adaptation of the differentiation process of akinetes, because in those strains in which only a small proportion of vegetative cells differentiates into akinetes, enlarging akinetes may derive nutrients from other cells (Wolk, 1964). This review will begin with a discussion of the functioning of mature, nitrogen-fixing heterocysts, will then describe the process of differentiation that leads to a nitrogen-fixing cell, and finally will discuss what is known of the processes that lead to the spacing of heterocysts. Because the work of the last decade will be emphasized, the interested reader is encouraged also to return to earlier reviews (e.g., Wolk, 1973, 1982; Haselkorn, 1978; Adams and Carr, 1981a) as well as to contemporary reviews that cover some of the same material but from different perspectives (Adams, 1992a; Fay, 1992; Gallon, 1992; Haselkorn and Buikema, 1992; Buikema and Haselkorn, 1993; Tandeau de Marsac and Houmard, 1993).

II. Genetic Tools Many of the results discussed in this review derived from recent technical advances in the area of the genetics of heterocyst-forming cyanobacteria. Most of the relevant techniques were developed using Anabaena sp. strain PCC 7120 (ATCC 27893), but have in some cases been extended to other strains. These techniques are outlined briefly below and are discussed more thoroughly in Chapter 19. DNA is most often introduced into Anabaena sp. from Escherichia coli, by conjugation mediated by plasmid RP-4 and its close relatives (Wolk et al., 1984), although electroporation is a usable alternative (Thiel and Poo, 1989). It is ordinarily critical to circumvent the effect of the restriction endonucleases of the recipient strain (Wolk et al., 1984; Elhai and Wolk, 1988; Thiel and Poo, 1989). The DNA introduced may replicate independently, for example by making use of the cyanobacterial replicon pDU1 (Wolk et al., 1984), or may integrate into the genome by homologous recombination (Golden and Wiest, 1988). Double recombination can be used to replace or intercept targeted genes (Golden and Wiest, 1988);

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the conditionally lethal gene, sacB, has been shown to be helpful for selection of double recombinants (Cai and Wolk, 1990). Certain derivatives (Wolk et al., 1991) of transposon Tn5 provide a powerful means to identify, in Anabaena sp., genes of interest in such a way that their subsequent cloning is facilitated. Mutants have also been obtained upon treatment with chemical mutagens (Currier et al., 1977; Buikema and Haselkorn, 1991a) and upon irradiation with ultraviolet light (Wolk et al., 1988); the genes affected can be cloned by complementation with cosmid libraries (Wolk et al., 1988; Buikema and Haselkorn, 1991a). The fusion of promoters to reporter genes enables the experimenter to monitor promoter activity easily under different conditions. Both luxAB (encoding luciferase) and lacZ (encoding have been used to report on the activity of promoters regulated during nitrogen deprivation in Anabaena sp. (Elhai and Wolk, 1990). Detection of luciferase activity can be sufficiently sensitive to allow measurement of the relative strength of transcription from a promoter of choice in individual cells along a filament, in vivo (Elhai and Wolk, 1990; Wolk et al., 1993; see Fig. 2). Reporter genes have also been placed in transposons to facilitate the creation of a wide variety of promoter-reporter fusions (Wolk et al., 1991, 1993; C. P. Wolk, unpublished results). A physical map of the 7.13-Mbp genome of Anabaena sp. strain PCC 7120 has been constructed on the basis of pulsed-field gel electrophoresis of fragments generated by rarely cutting restriction endonucleases (Bancroft et al., 1989; see Chapter 19); recent improvements permit mapping to within about 20 kbp (Kuritz et al., 1993). Close to 2% of the genome of this strain has been sequenced. Isolated heterocysts that are metabolically very active, and in particular have high nitrogenase activity, can be isolated from Anabaena variabilis strain ATCC 29413 (Peterson and Wolk, 1978b) and Anabaena sp. strain CA (Smith et al., 1988), but have not yet been isolated from Anabaena sp. strain PCC 7120. Anabaena variabilis strain ATCC 29413 is also known as Anabaena sp. strain PCC 7937 and was derived from UTEX 1444, a strain originally designated as Anabaena flos-aquae. Hereafter in this chapter, Anabaena variabilis will refer to strain ATCC 29413 unless explicitly stated otherwise. Gene replacement is possible with this strain (Maldener et al., 1991; Toelge et al., 1991; Mannan et al., 1991),

774 but the strain has not served as a source of DNA suitable for pulsed-field gel electrophoresis, nor has it proven amenable to transposon mutagenesis. Since potential viral vectors (Khudyakov and Gromov, 1973; Hu et al., 1981) have not been proven capable of transduction, transfer of genes from one strain of Anabaena sp. to another now proceeds via E. coli. Anabaena sp. strain PCC 7120 can transfer part or all of its a megaplasmid to a different strain (Muro-Pastor et al., 1994), but whether it can also transfer chromosomal DNA is unknown.

III. Metabolism of Mature Heterocysts

A. Requirements for Anaerobic Fixation of 1. Nitrogenase Genes and Neighboring DNA To fix and to maintain a capacity to fix under anaerobic conditions, heterocysts require the products of nif genes (this section), reductant (sections III A, 2–5) and ATP (Section III A, 6). Nitrogen fixation is catalyzed by dinitrogenase, a dimer of the polypeptides encoded by nifD (Lammers and Haselkorn, 1983; Golden et al., 1985) and nifK (Mazur and Chui, 1982), which is supplied with electrons by dinitrogenase reductase, which is a dimer of the polypeptide encoded by nifH (Mevarech et al., 1980). These genes (nifH, nifD and nifK) are contiguous and form an operon in the chromosomes of heterocysts. Dinitrogenase reductase contains a single 4Fe-4S cluster that can be irreversibly oxidized by Dinitrogenase contains four 4Fe-4S centers, that are arranged in two clusters, and two copies of FeMo-co, which is an iron- and molybdenumcontaining cofactor, all of which add to the lability of the enzyme. Dinitrogenase and dinitrogenase reductase together comprise nitrogenase. The nitrogenases of various diazotrophic organisms exhibit a high degree of sequence similarity at the amino acid level, and often on the DNA level (Ruvkun and Ausubel, 1980). The nitrogenase of heterocysts may, therefore, resemble in its three-dimensional structure the models constructed from X-ray crystallographic data of Azotobacter vinelandii nitrogenase (Kim and Rees, 1992; Georgiaidis et al., 1992; see Chapter 16), and it may exhibit detailed enzymatic properties similar to those observed with the enzymes of Klebsiella aerogenes or Azotobacter vinelandii (Thorneley and Lowe, 1985).

C. Peter Wolk, Anneliese Ernst & Jeff Elhai The known genes in the nif region (Fig. 3) of heterocyst DNA in Anabaena sp. strain PCC 7120 include (in order, and starting upstream from nifH) rbcS-rbcL (encoding the small and large subunits of ribulose bisphosphate carboxylase/oxygenase; Curtis and Haselkorn, 1983, and Nierzwicki-Bauer et al., 1984b; the rbcLS genes were localized near the nif genes by Herrero and Wolk, 1986; Golden et al., 1988; and Brusca et al., 1989); nifB, fdxN, nifS, and nifU (Mulligan and Haselkorn, 1989); nifH, nifD, and nifK; and (3' from nifK), nifE, nifN, nifX, an unidentified open reading frame, and nifW (Haselkorn, 1992; Haselkorn and Buikema, 1992); two open reading frames of unknown function whose products enhance, but are not required for, the nitrogen-fixation process (Borthakur et al., 1990); and, about 7 kbp from nifK, fdxH (Böhme and Haselkorn, 1988), which encodes a special ferredoxin (see Section III A, 2, below and Chapter 12) that is the donor of electrons to dinitrogenase reductase in heterocysts. The named nif genes are homologues of corresponding Klebsiella aerogenes nif genes; nifB, nifN and nifE are thought to be involved in the synthesis of FeMo-co, whereas the roles of nifS, nifU, nifX and nifW have not been defined. In Nostoc

Chapter 27 Heterocyst Metabolism and Development commune, a gene denoted glbN, found sandwiched between nifU and nifH, encodes a hemoprotein that has been detected only after protracted (24–56 h) incubation under conditions of microaerobiosis and nitrogen deprivation (Potts et al., 1992). A multifunctional chaperonin (GroEL protein) has been localized by immunoelectron microscopy in heterocysts as well as in vegetative cells of Anabaena sp. strain PCC 7120 (Jäger and Bergman, 1990; the groEL gene encoding a related protein was cloned from Synechococcus sp. strain PCC 7942 by Webb et al., 1990; see Chapter 26) and may well function in proper folding of nitrogenase (Govezensky et al., 1991). Whereas the genes mentioned in the previous paragraph are clustered in the chromosome of Anabaena sp. strain PCC 7120 (Haselkorn, 1992; Haselkorn and Buikema, 1992), a nifH homolog, (Robinson and Haselkorn, 1985), of unknown function is present ca. 0.6 Mbp distant from the cluster (Kuritz et al., 1993). Other strains of Anabaena sp. contain additional copies of nif genes (Hirschberg et al., 1985; Herrero and Wolk, 1986; Golden et al., 1988; Thiel, 1993). Expression of at least one of these is first observed, under aerobic conditions, after about 8.5 h of nitrogen deprivation (Wealand et al., 1989), presumably in maturing heterocysts (see Peterson and Wolk, 1978b). Under anaerobic conditions, nitrogenase mRNA is detectable within 1.5–2 h of nitrogen deprivation, in filaments that lack any morphological signs of differentiation (Helber et al., 1988a). It remains to be determined whether the two activities, distinguished by their kinetics of synthesis and their relate to different locations, heterocysts and vegetative cells, respectively, and to different copies of nif genes. Azotobacter vinelandii is known to encode, in addition to a Mo-containing nitrogenase, so-called alternative nitrogenases that contain either vanadium (V) or neither Mo nor V (Bishop and Joerger, 1990). Both are more than the Mo-containing nitrogenase, and unlike that nitrogenase, both reduce partially to rather than exclusively to Anabaena variabilis (Kentemich et al., 1988, 1991; Thiel, 1993), and evidently certain other cyanobacteria (Bortels, 1940; Chan et al., 1991), are able to grow on in the absence of Mo and presence of V, or in the absence of both, and reduce in part to and so appear to produce at least a third nitrogenase. Genes of the V-dependent nitrogenase, vnfD and vnfK, of Anabaena variabilis have been

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identified by sequence homology with the respective Azotobacter vinelandii genes (Thiel, 1993). Whether these genes are active in vegetative cells, in heterocysts, or in both has yet to be determined. For reasons that are unclear, the nif operons in the vegetative cells of Anabaena sp. strain PCC 7120 are interrupted by two excision elements (also known as excisons) that are removed from the chromosome of Anabaena sp. strain PCC 7120 late during heterocyst differentiation (Golden et al., 1985, 1988). Excision is a necessary prelude to transcription of nifHDK as a complete operon from a promoter 5' from nifH (Haselkorn et al., 1983), and to transcription ofnifBfdxN-nifS-nifU as a complete operon from a promoter 5' from nifB (Mulligan and Haselkorn, 1989). The 11-kbp excison that is present within, and close to the 3' end of, the coding sequence of nifD within vegetative cells of Anabaena sp. strain PCC 7120 has been extensively sequenced (Lammers et al., 1986, 1990); it contains at least five open reading frames. One (xisA) is located close to nifK and is transcribed on the opposite DNA strand; xisA encodes the site-specific recombinase, or excisase, that catalyzes excision of the element (Lammers et al., 1986). Excision takes place by a site-specific recombination between 11-bp directly repeated sequences that flank the insert (Lammers et al., 1986). The insert is released as a stable, circular molecule of DNA (Golden et al., 1985). Site-specific inactivation of xisA did not prevent transcription of nifHD, but eliminated rearrangement of the 11-kbp element, so that transcription failed to reach nifK, thereby leaving the strain unable to fix (Golden and Wiest, 1988). In addition, deletion of the nifH promoter did not prevent normal excision of the 11kbp element (Golden et al., 1991). Another of the open reading frames in the excison shows homology to cytochrome P450 (Lammers et al., 1990). The gene, fdxN, that is interrupted by the second, 55-kbp excison encodes a bacterial type 4Fe-4S ferredoxin of unknown function that is also found adjacent to nifB in certain other diazotrophs (Mulligan et al., 1988). Excision of this element occurs by recombination within 5-bp, directly repeated sequences (Golden et al., 1987). The corresponding site-specific excisase is encoded by a gene,; xisF, that is located proximal to nifS (Carrasco et al., 1994; Golden et al., 1992); interruption ofxisF results in a Fix¯ phenotype (Kuritz et al., 1993). The occurrence of the 55-kbp excison within fdxN

776 and of the 11-kbp excison within nifD is by no means ubiquitous within the heterocyst-forming cyanobacteria: the 55-kbp element is absent from Anabaena variabilis (Herrero and Wolk, 1986; Golden et al., 1988); and the 11-kbp element, although present in some other strains of the genera Anabaena, Nostoc and Calothrix (Meeks et al., 1988; Brusca et al., 1989; Franche and Cohen-Bazire, 1987; Kallas et al., 1983), is absent from the major symbiont of Azolla sp. (Nierzwicki-Bauer and Haselkorn, 1986; Meeks et al., 1988; see also Franche and CohenBazire, 1987), and Fischerella sp. strain ATCC 27929 (Saville et al., 1987), as well as from diverse fixing cyanobacteria that lack heterocysts (Barnum and Gendel, 1985; Kallas et al., 1985; Apte and Thomas, 1987). In fact, after experimental elimination of the 11-kbp element from its genome, Anabaena sp. strain PCC 7120 still showed normal growth and heterocyst differentiation in a medium free of fixed nitrogen (Brusca et al., 1990). Despite the non-essentiality of the 11-kbp excision element, and the probable non-essentiality of the 55-kbp element, in Anabaena sp. strain PCC 7120 under conditions that have been tested, they remain regulatory elements in the sense that transcription of the genes encoding dinitrogenase depends upon their excision. Recently, a third rearrangement has been identified that takes place upon nitrogen stepdown, within the SalD fragment of the chromosome of Anabaena sp. strain PCC 7120 (Matveyev et al., 1994). A 10.5kbp element is excised from within the hupL gene, which encodes the large subunit of a membranebound NiFe uptake hydrogenase. The element contains a presumptive site-specific recombinase gene denoted xisC that shows sequence similarity to xisA. The hupL element is flanked by 16-bp direct repeats that are not similar to the recombination sites for the nifD or fdxN elements (J. Golden, personal communication).

2. Electron Donor to Nitrogenase Two plant-type [2Fe-2S] ferredoxins were found in heterocysts of Anabaena variabilis (Böhme and Schrautemeier, 1987a). One, found only in heterocysts, is encoded by the gene fdxH (Böhme and Haselkorn, 1988; Schrautemeier and Böhme, 1992), while the other is similar, and perhaps identical, to the ferredoxin observed in vegetative cells and

C. Peter Wolk, Anneliese Ernst & Jeff Elhai encoded by petF (Alam et al., 1986; see Chapter 12). The products oftranslation ofthe two genes differ in 47 out of 98 amino acids. The occurrence of an additional, bacterial-type ferredoxin in Anabaena variabilis was recently reported (Yakunin and Gogotov, 1993); it may possibly be encoded by fdxN (Section III A, 1). The heterocyst-specific [2Fe-2S] ferredoxin encoded by fdxH mediates electron transport to nitrogenase in the light much more efficiently than does the ferredoxin from vegetative cells, and is able to mediate electron transfer from glucose 6-phosphate or isocitrate to nitrogenase in the dark, whereas the vegetative cell ferredoxin cannot (Schrautemeier and Böhme, 1985). The electron donor to nitrogenase under Fe-replete conditions is for that reason considered (Schrautemeier and Böhme, 1985; Böhme and Schrautemeier, 1987a) to be the product of the structural gene fdxH (Böhme and Haselkorn, 1988). The difference in ability to donate electrons to nitrogenase is attributable to a group of positively charged residues characteristic of FdxH that may facilitate interaction with dinitrogenase reductase (Schmitz et al., 1993b). Transcripts of fdxH are first observed at 18 h after nitrogen-stepdown (Böhme and Haselkorn, 1988), about the same time that nitrogenase activity is first observed. The nitrogenase isolated from Azotobacter vinelandii has an much lower than the corresponding value of–290 mV for pyridine nucleotides in that organism. For that reason, a mechanism dependent on membrane energization was proposed to stabilize the potential of flavodoxin, the electron donor to nitrogenase, at a value more negative than –290 mV (Haaker and Klugkist, 1987). No such mechanism is required for nitrogenase in heterocysts as shown by the facts that (i) the activity of that nitrogenase can be supported, in cell-free reactions, by carbohydrates and related metabolites (Section III A, 3), and (ii) so long as an ATPgenerating system is present, electron transport to nitrogenase is not influenced by uncouplers (Schrautemeier et al., 1984). The vegetative-cell-type ferredoxin in heterocysts is thought to function in cyclic electron transport through Photosystem I (Böhme and Haselkorn, 1988; see Chapter 10) and to connect the pool of NADP(H) with the reducing side of PS I, thereby helping to balance the demands of pyridine nucleotideconsuming and -producing processes (see Section III A, 6). Reduction of the heterocyst-specific

Chapter 27

Heterocyst Metabolism and Development

ferredoxin by Photosystem I or by the action of ferredoxin: oxidoreductase may be the normal process in the light and in the dark, respectively. The non-photochemical reductive pathway is discussed in Section III A, 3, and photochemical reduction is discussed in Sections III A, 6 and III D, 1. The physiological consequences of inactivating fdxH have not been reported, but there is indirect evidence that under conditions of iron deficiency, this ferredoxin may be replaceable in Anabaena sp. strain PCC 7120 by a flavodoxin (Leonhardt and Straus, 1989). An mRNA for pyruvate-flavodoxin oxidoreductase (identified by sequence comparison), the product of nifJ, is produced specifically under

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those conditions (Bauer and Haselkorn, 1992; Schmitz et al., 1993), and Synechococcus sp. flavodoxin can reduce the nitrogenase from Anabaena cylindrica (F) (Bothe, 1970). Such a replacement may be possible also in Anabaena variabilis; however, Anabaena sp. strain ATCC 29211 does not appear to make flavodoxin (Sandmann et al., 1990; see also Schrautemeier and Böhme, 1984).

3. Organic Electron Donors (see Fig. 4) Because photosynthetic water-splitting in heterocysts would be incompatible with nitrogenase activity, it appears highly likely that the reduction of and

778 in heterocysts is supported by organic metabolites from vegetative cells (see Section III A, 4). Alternatives include reduced inorganic solutes such as reduced proteins, or solid-state transfer via membranes, but there is no substantive support for any of these possibilities. A variety of metabolites canbe oxidized by dehydrogenases, with concomitant reduction of or The NADPH that results can reduce ferredoxin, or transfer electrons to PS I or to the respiratory electron transport chain, in each case via ferredoxin: oxidoreductase, whereas the NADH that results can transfer electrons to PS I or to the respiratory electron transport chain (see Chapters 10 and 13). As described below, the three metabolites that are primary candidates for electron donor to in heterocysts are glucose6-phosphate and 6-phosphogluconate (both of which are oxidized by the oxidative pentose phosphate cycle, the first two steps of which are catalyzed by glucose-6-phosphate dehydrogenase and 6-phosphogluconate dehydrogenase), and isocitrate (oxidized by isocitrate dehydrogenase). Reduction of is enzymatically coupled to the oxidation of glyceraldehyde 3-phosphate and malate by the corresponding dehydrogenases. In addition, pyruvate has the capacity toreduce ferredoxin directly in a reaction catalyzed by pyruvate:ferredoxin oxidoreductase. Enzymes of the oxidative pentose phosphate cycle are highly active in heterocysts (see Wolk, 1982; Udvardy et al., 1984). These enzymes were first found to be active in heterocysts of Anabaena cylindrica (W) at approximately. 60- to 70-fold higher specific activity than in vegetative cells (Winkenbach and Wolk, 1973), and in the presence of their substrates and of heterocyst ferredoxin were shown to support high rates of acetylene reduction by heterocyst homogenates (Schrautemeier et al., 1984; Böhme and Schrautemeier, 1987b). Glycolysis probably does not contribute greatly to catabolism of hexose phosphates in heterocysts, because the activity of phosphofructokinase, a key enzyme of that pathway, is low in heterocysts (Böhme, 1987). In homogenates of vegetative cells and of heterocysts, glucose 6-phosphate dehydrogenase is inhibited by 5 mM NADPH and by reduced thioredoxin (Papen et al., 1983; Cossar et al., 1984; Ip et al., 1984; Udvardy et al., 1984) and reactivated by oxidized glutathione (Udvardy et al., 1984; Cossar et al., 1984). These effects are thought to prevent a futile cycle of photosynthesis and dissimilation of carbohydrate in the vegetative cells, but whether

C. Peter Wolk, Anneliese Ernst & Jeff Elhai they operate in heterocysts is unclear. The thioredoxin used was shown by immunoelectronmicroscopy to be present to some extent in heterocysts, although its relative concentrations in vegetative cells and heterocysts appear to vary from strain to strain (Cossar et al., 1985a; Dai et al., 1992). In contrast, reduced thioredoxin activates NADP-dependent isocitrate dehydrogenase (Papen et al., 1983). Anaerobic homogenates of heterocysts, upon supplementation with various substrates and with and as cofactors, support nitrogenase activity in the dark with greater activity than in the light (Schrautemeier et al., 1984; Böhme and Schrautemeier, 1987b). It was concluded that the oxidative pentose phosphate pathway is functional in intact heterocysts in the light and in the dark. Whether the triose phospihates produced by the oxidative pentose phosphate cycle can be further metabolized by glycolysis within heterocysts depends upon the availability of glyceraldehyde-3-P dehydrogenase. That enzyme, whose availability was previously controversial (see Wolk, 1982), appears now to be abundantly present in heterocysts of Anabaena cylindrica (F), Anabaena variabilis, and Anabaena sp. strain PCC 7119 (Papen et al., 1986b), where it can also serve as a source of NADH and of phosphoenolpyruvate (PEP). It is stimulated by thiols but not by thioredoxin-m of the same strain (Papen et al., 1986b; see Chapter 24). Cyanobacteria in general (Stanier and Cohen-Bazire, 1977), and heterocysts in particular (Newer et al., 1983), are unable to oxidize sugars completely by the combined action of glycolysis and the tricarboxylic acid cycle, because they lack two enzymes of that cycle: succinate dehydrogenase and dehydrogenase. Nonetheless, the PEP formed can be carboxylated to form oxaloacetate or metabolized by pyruvate kinase to form pyruvate (Neuer and Bothe, 1983). The pyruvate, in turn, can be carboxylated to form malate or oxidized by ferredoxin in the presence of coenzyme A to form acetylcoenzyme A, and reduced ferredoxin (Neuer and Bothe, 1982, 1983, 1985). Activities of NADHlinked malate dehydrogenase (see also Kovacheva, 1980), yielding oxaloacetate, and citrate synthase and aconitase, yielding isocitiate, were also observed. However, because the measured activities were ‘low’ (for example, isolated heterocysts reduced acetylene, in the presence of pyruvate, with an activity less than 2% of the activity of intact filaments: Neuer and Bothe, 1985; see, however, Böhme, 1987),

Chapter 27 Heterocyst Metabolism and Development the physiological significance of the pathway studied as a source of either electrons or ammonium acceptor for nitrogen fixation was quite unclear (Neuer and Bothe, 1983). Measured activities of isocitrate dehydrogenase, yielding and NADPH, were higher than the activities of the enzymes just discussed, and activities ofisocitrate dehydrogenase were much higher in heterocysts than in vegetative cells in Anabaena cylindrica (F) and Anabaena sp. strain PCC 7119 (Kami et al., 1982; Kami and TelOr, 1983; Papen et al., 1983). Isocitrate, or a combination of pyruvate, CoA, and oxaloacetate, supported relatively high nitrogenase activity by extracts of heterocysts (Böhme, 1987), although as discussed in the last paragraph, production of those substrates within heterocysts may be limiting. Like the oxidation of glucose 6-phosphate by glucose 6phosphate dehydrogenase, the oxidation ofisocitrate by isocitrate dehydrogenase is stimulated by rapid removal of the products and (or) oxidation of the reduced cofactor, and is inhibited by a high ratio of NADPH to NADP (Karni and Tel-Or, 1983). It is, therefore, problematic to conclude that isocitrate dehydrogenase in heterocysts serves as electron donor, via NADPH, to ferredoxin and thence to nitrogenase, or as precursor of and thereby of glutamate. Heretofore, our knowledge of heterocyst catabolism has been based on the measurement of the responses of the nitrogenase activity of homogenates to the addition ofsubstrates and on the determination of enzymatic activities. Both approaches involve substantial uncertainty: in homogenates, effectors may be unnaturally diluted, and enzymatic activities in vitro can underestimate or overestimate metabolic throughputs in vivo. The ppc gene encoding PEP carboxylase has been cloned and sequenced from Anabaena sp. strain PCC 7120 (Luinenburg and Coleman, 1992). Moreover, now that a heterologous probe is available from a unicellular cyanobacterium for the zwf gene that encodes glucose 6-phosphate dehydrogenase (Scanlan et al., 1992), it may soon be possible to clone the corresponding gene from Anabaena sp. With such clones in hand, it should be possible to test (by inactivation or by use of celltype-specific antisense constructs) the roles of the corresponding enzymes in electron donation in heterocysts of Anabaena sp. The hope is that such procedures, when applied to a wide variety of enzymes, will help to clarify which enzymes and

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pathways are of major importance in heterocysts.

4. Intercellular Movement of Carbon Compounds Pulse-chase experiments demonstrated unequivocally that from fixed by vegetative cells moves into heterocysts and is metabolized there (Wolk, 1968). Whatever the carbon-containing substance(s) is (or are) that move into heterocysts, theypresumably are a source of electrons for nitrogen fixation, of carbon skeletons for assimilation of fixed nitrogen, and of building blocks for envelope materials in the heterocyst. To date, four kinds ofexperiments appear to contribute to the identification of the substances that move from vegetative cells. Firstly, and already alluded to, there are demonstrations of very high specific activity ofcertain enzymes, dehydrogenases of the oxidative pentose phosphate cycle and apparently also isocitrate dehydrogenase, in heterocysts. Whereas one can argue that there might be reason for excess capacity of an enzyme whose job is to catalyze the reduction of ferredoxin by NADPH, it still seems likely that the substrates for those enzymes should be present in abundance. Secondly, a study was made of radioactive metabolites present in whole filaments of Anabaena cylindrica (F), and in heterocysts isolated rapidly from those filaments in the cold, after decreasing durations (300, 120, and 20 seconds) of assimilation of by intact filaments. After 20 sec of fixation, aspartate, alanine, and glutamate accounted for a much higher percentage of the total radioactivity in heterocysts than in filaments. Glucose 6-phosphate held a much smaller, but significant fraction of the total (4.1%; 33% of the total in filaments). During 30-min incubations, heterocysts themselves metabolized principally to aspartic acid (23%) and glutamine (35%; Jüttner, 1983). Thirdly, certain anabolic or catabolic enzymes may be restricted to one cell type or the other. For example, Schilling and Ehrnsperger (1985) found that in Anabaena variabilis, sucrose synthase is associated almost exclusively with vegetative cells, whereas alkaline invertase, which cleaves sucrose, appeared to be present almost exclusively in heterocysts. Fourthly, the metabolism of certain exogenous substances by isolated heterocysts has been determined. For example, when isolated heterocysts of Anabaena cylindrica (F) were incubated in the light for 30 min under an atmosphere of and alanine, glutamate, and pyruvate were each

780 metabolized extensively to glutamine (85%, 92%, and 50%, respectively), and glucose to glucose6-phosphate (59%), fructose-6-phosphate (21%) and glutamate (21%) (Jüttner, 1983). Because it is clear from the results of Jüttner (1983) and Neuer and Bothe (1983) that heterocysts are capable of anaplerotic, reactions, there seems no unequivocal way to distinguish what fraction of the radioactivity found in heterocysts after short periods of fixation of by filaments was due tofixationdirectly by heterocysts. Moreover, Jüttner’sexperimentshave an intrinsicdifficultythat metabolites can be detected as having moved into heterocysts only to the extent that their metabolic pools in vegetative cells have become extensively labeled by the end of the period of pulse-labeling. The authors of this chapter conclude that the above results support as a working hypothesis, but by no means prove, the interpretation that (i) sucrose moves from vegetative cells to heterocysts; (ii) sucrose is transformed to hexose phosphates in the heterocysts; and (iii) hexose phosphates are metabolized by the oxidative pentose phosphate cycle, and in part converted, by glycolysis and by reactions of the partial citric acid cycle, to and thence to glutamate and glutamine. The movement of nitrogen-containing metabolites between vegetative cells and heterocysts will be discussed in Section III D, 1.

5. Conduits for Intercellular Movement Via what conduits do nutrients move between heterocysts and vegetative cells? In different cyanobacteria, heterocysts exhibit one, two or (sometimes) three pores which provide a route, unblocked by the glycolipid layer of the envelope, for movement of hydrophilic substances into and out of heterocysts. The pore contains a ‘neck’ region of reduced diameter at the end of which the heterocyst plasmalemma is narrowly separated from the plasmalemma of the adjacent cell. The region of proximity is not unlike that seen in a late stage in the separation ofsister vegetative cells in Mastigocladus sp. (Nierzwicki et al., 1982). Within that region are structures called microplasmodesmata that appear to join adjacent cells (Lang and Fay, 1971; Giddings and Staehelin, 1978; Roussard-Jacquemin, 1983). However, one does not know that intercellular movement of nutrients takes place within the microplasmodesmata, rather than separately through

C. Peter Wolk, Anneliese Ernst & Jeff Elhai the end membranes of the two adjacent cells with diffusion between those two membranes. It seems feasible that intercellular transport is impaired in some mutants that are incapable of aerobic fixation of and whose heterocyst protoplasts are shrunken (i.e., non-turgid) or vacuolate (Ernst et al., 1992), and in trichlorfon-treated cultures whose heterocysts have a similar appearance (Orús and Marco, 1991). Investigation of these mutants could provide crucially needed information about the mechanism of intercellular interactions. It will also be of great interest to learn whether permeases present at the radial periphery of vegetative cells are present at their intercellular junctions and at the junctional area of heterocysts. It seems clear that heterocysts isolated with the techniques of the 1960’s and early 1970’s were either (to borrow a term from Stanier et al., 1971) ‘microbial corpses’, or were only sufficiently intact to retain proteins but were extremely porous to metabolites of low molecular weight (see Wolk, 1979). Combinations of relatively gentle measures for disrupting vegetative cells (Peterson and Wolk, 1978b; Smith et al., 1988), together with a choice of what may be hardier strains, led to the isolation of heterocysts that were far more intact and metabolically proficient. Heretofore, all isolated heterocysts have had a limited duration of reducing activity (e.g., 25-fold decrease in 11 h; Jensen et al., 1986). That they slowly hemorrhage their metabolites seems unlikely because they can be stored at 0 °C for 12 hours without loss of such activity (Jensen et al., 1986). Perhaps they require nutrients with which they have not been provided. It is unclear whether isolated heterocysts would be able to take up all nutrients that they might need. That is, it is possible that their intercellular conduits become sealed upon disjunction from vegetative cells. Heretofore, the investigation of metabolic needs of isolated heterocysts has focused on the effect of metabolites on immediate acceleration of nitrogenase activity rather than on whether metabolites might increase the duration of that activity. Privalle and Burris (1984) found that sucrose, D-glucose, D-glucose 6-phosphate, and D-fructose 1,6-bis-phosphate each increased the rate of reduction of by isolated heterocysts of Anabaena sp. strain PCC 7120 about 4-fold in the presence of argon. D-erythrose increased that rate 10-fold but unlike the sugars just mentioned, was inhibitory above about 5 mM and also inhibited the otherwise

Chapter 27 Heterocyst Metabolism and Development strongly stimulatory effect of added The significance of these effects of D-erythrose, striking though they are, is regrettably unclear (see also Privalle, 1984; Jensen et al., 1986). Inasmuch as added nutrients are having some effect, some uptake is clearly possible; perhaps multiple kinds of nutrients are needed. This point will also be addressed in Section III D, 1 below.

6. Photosystem I: Source of ATP and Reductant Heterocysts contain PS I. In several strains of Anabaena and Nostoc sp., the ratio of total chlorophyll per PS I reaction center in heterocysts is 1/3 to 2/3 as great as in vegetative cells (Alberte et al., 1980; Almon and Böhme, 1980; Peterson et al., 1981b; Houchins and Hind, 1984). Under anaerobic conditions in the dark, heterocysts of Anabaena variabilis do not support significant nitrogenase activity. The activity of heterocysts isolated from that strain that had been grown with fructose in the light was increased by low or (much more) by incubation in the light. Unlike the situation with heterocysts isolated from photoautotrophically grown cultures (Peterson and Wolk, 1978b), no stimulation of that activity was observed with indicating that the heterocysts were replete with stored carbohydrate (Jensen et al., 1986). The effect of light was evidently to satisfy the ATP requirement of nitrogenase by photophosphorylation (see also Janaki and Wolk, 1982). In fact, it has been directly demonstrated that both light and low but increasing increase the concentration of ATP and decrease the concentration of AMP in isolated heterocysts (Ernst et al., 1983). PS I-driven photophosphorylation could involve cyclic (Almon and Böhme, 1982; see also Wolk, 1982) or linear electron flow (see Chapter 10). In cyclic electron flow electrons would pass from the reducing side of PS I to the oxidizing side of that photosystem. That linear flow is possible is shown by the observations that the use of or NADH as electron donor for reduction of by heterocyst homogenates (Schrautemeier et al., 1984) and frozenand-thawed heterocysts provided with ATP (Houchins and Hind, 1982; Houchins, 1985) is dependent upon light to photoreduce ferredoxin. In contrast, NADPH can reduce ferredoxin in the dark as well as in the light. is generated by nitrogenase (Simpson and Burris, 1984); NADH, by the oxidation of glyceraldehyde phosphate and malate; and NADPH, by the

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oxidation of glucose 6-phosphate, 6-phosphogluconate, and isocitrate (see above). Electrons from and NADH are transferred to PS I via, respectively, a membrane-bound hydrogenase (Peterson and Wolk, 1978a; Houchins and Burris, 1981c; see Section III C, 4) and a membranebound pyridine nucleotide dehydrogenase. Such a dehydrogenase, possibly a peripheral subcomplex of an NAD(P)H-plastoquinone oxidoreductase (Berger et al., 1991, 1993; Mi et al, 1992), has been purified from Anabaena variabilis (Alpes et al., 1989; Berger et al., 1991, 1993; see Chapter 13). Transfer of electrons from NADPH appears to be dependent upon soluble ferredoxin: oxidoreductase and ferredoxin (Houchins and Hind, 1982; Hawkesford et al., 1983; Schrautemeier et al., 1984; Houchins, 1985; Fillat et al., 1990). Dibromomethylisopropylbenzoquinone (DBMIB, an inhibitor of plastoquinone-mediated reactions at the cytochrome complex) inhibits the donation of electrons from NADPH to in the light but not in the dark. These observations indicate that the pathway of electron transfer from NADPH to nitrogenase is changed by light, presumably by a change in the properties of ferredoxin: oxidoreductase (Schrautemeier et al., 1984; Fillat et al., 1991). This enzyme may exhibit post-translational modification (Schluchter and Bryant, 1992) and different principal molecular forms in heterocysts and vegetative cells (Rowell et al., 1981). Electrons donated to PS I by NADH, and NADPH in the light pass through (i) a (membranebound) pool of plastoquinone molecules; (ii) a cytochrome complex; and (iii) cytochrome (or plastocyanin), that can transfer electrons either to PS I or to a respiratory cytochrome (see Section III B, 3 and Chapter 13). The principal evidence for the involvement of plastoquinone molecules is that the reactions are inhibited by DBMIB (see above; Schrautemeier et al., 1984, 1985; Houchins and Hind, 1983b). The stoichiometries of components of the cytochrome complex (cytochrome and cytochrome and P700 (at the reaction center of PS I) are similar in vegetative cells and heterocysts (Almon and Böhme, 1980; Houchins and Hind, 1984). The oxidation of quinones by the cytochrome complex converts electron transport from twoelectron carriers (quinones) to one-electron, metalloprotein carriers. In this complex in heterocysts, two cytochromes operate that differ in kinetic

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behavior, midpoint potential and susceptibility to the inhibitor 2-(n-heptyl)-4-hydroxyquinoline Noxide (HQNO). The kinetics of oxidation and reduction of the cytochromes established that plastoquinone is oxidized in a two-stepprocess known as a Q-loop (Houchins and Hind, 1983a,b, 1984). Transfer of electrons through the cytochrome complex leads to the pumping of protons (Houchins, 1985), and thence to the generation of ATP (Almon and Böhme, 1982). Vegetative cell-type ferredoxin reduced by PS I can mediate two processes in heterocysts: (i) cyclic electron flow around Photosystem I (see above, but see Chapter 10), with attendant phosphorylation; and (ii) a transhydrogenation reaction between and NADH-producing and NADPH-consuming processes (Schrautemeier et al., 1985; Böhme and Schrautemeier, 1987a). Cyclic electron flow mediated by the vegetative cell-type ferredoxin and linear electron flow mediated by the heterocystspecific ferredoxin would compete for electrons from PS I. How that competition is regulated is unknown, but a remarkable homeostasis in cellular concentration of ATP was observed when nitrogenase activity was varied by changing the light intensity or concentration of or by adding DBMIB (Ernst and Böhme, 1984).

Thiel et al. (1990) probed Western blots of proteins from heterocysts of Nostoc sp. strain ATCC 29150 with antibodies to five major reaction-center proteins of Photosystem II, and found evidence of the presence of all five of the proteins, and also observed the occurrence of a peak of fluorescence characteristic of chlorophyll bound to one of those proteins. The content of phycobiliproteins in mature heterocysts varies from essentially zero (Thomas, 1970) to up to at least 40% of that of vegetative cells (Peterson et al., 198la). Moreover, convincing evidence was obtained for the presence of not just phycobiliproteins, butphycobilisomes, in heterocysts ofAnabaena variabilis (Ke et al., 1983). However, the available evidence indicates that to a large extent, light energy absorbed by these phycobilisomes is transferred to PS I (Peterson et al., 1981a) rather than, as is normally the case in vegetative cells, principally to PS II. [It has recently become clear that under appropriate light conditions, phycobilisomes in vegetative cells also transfer energy to PS I (see Chapter 6).] In sum, parts of PS II appear to be present, but whether they function without production of remains unknown.

7. Do Heterocysts Have a Photosystem II?

1. Introduction

During the differentiation of heterocysts, evolution is lost and pigments associated with PS II may be lost (see Wolk, 1982, for review). Although the sensitivity of nitrogenase leads to the expectation that heterocysts would not produce heterocysts might nonetheless retain a modified PS II that is able to receive electrons from an organic (or inorganic) donor (see Garlick et al., 1977, but also Giddings et al., 1981; Fry et al., 1984; Pistorius and Gau, 1986; Bockholt et al., 1991). Whether a high-potential form of cytochrome a component of the PS II reaction center, is present in heterocysts was controversial a decade ago (Wolk, 1982) and remains controversial (Papageorgiou and Isaakidou, 1981; Houchins and Hind, 1984). Braun-Howland and Nierzwicki-Bauer (1990), working with Anabaena azollae endosymbiotic in Azolla caroliniana, found that each of three antisera specific for different epitopes of the 32-kDa protein of the reaction center of PS II showed an abundance of that protein in heterocysts of all stages of differentiation. In addition,

Additional biochemical requirements are imposed by the need to fix in an aerobic environment. It is important to realize that the interior of heterocysts that are functioning to fix nitrogen in an aerobic environment is microaerobic, and cannot be anaerobic. Walsby (1985) pointed out that because ofthe physical similarities ofmolecules of and (see Section I), there is no obvious way that the former could be enabled to enter a cell while the latter is wholly excluded. By observing the collapse of gas vacuoles within heterocysts in response to increases in pressures of gases in the overlying gas phase, he estimated experimentally that the molar flux of into a heterocyst that is in a medium in equilibrium with air is between 0.5 and 1.0 of the corresponding molar flux of Any biochemical mechanism for removing (reacting with) the that enters has a non-zero and so in the presence of a constant influx of camnot render the interior of the heterocyst anaerobic. Elhai and Wolk (1991) made an initial, very crude estimate of the intracellular

B. Additional Requirements for Aerobic Fixation of

Chapter 27 Heterocyst Metabolism and Development concentration of in heterocysts as on the order of based on the response of luciferase, an requiring enzyme, in heterocysts to procedures that were thought to increase intracellular greatly. For comparison, is present at about in water that is in equilibrium with air at 1 atm and 30 °C. Gallon et al. (1992) used the of fluorescence of 4-[1-pyrenyl] butyric acid to measure the in cyanobacterial cells, but did not apply their technique to heterocysts. Therefore, to maintain nitrogenase activity in an aerobic milieu, the heterocyst requires (1) a barrier to diffusion of gases (discussed in Section III B, 2); (2) a respiratory apparatus to reduce that enters (see Section III B, 3); (3) a means to replace nitrogenase that is inactivated (see Section III B, 4); and (4) mechanisms of the same sort that vegetative cells have to protect against the deleterious effects of oxygen on photosynthetically active cells (see Section III B, 5). In addition, they have need ofa response to non-steady-state increases in originating from the fact (pointed out by Minchin, 1986) that the intracellular can change if, upon transition from light to dark, the availability of reductant within the heterocysts diminishes (see Section III C, 3). The

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general subject of the oxygen relations of nitrogen fixation in cyanobacteria has been the subject of recent reviews (Fay, 1992; Gallon, 1992), to which the reader is referred for more extensive discussion.

2. Barrier to Permeation of Oxygen Walsby (1985) deduced that the major pathway of gaseous diffusion into the heterocyst is its envelope (Fig. 5), rather than its terminal pore(s). He concluded on theoretical grounds, and Haury and Wolk (1978) and Murry and Wolk (1989) added experimental support, that the principal barrier to entry of is the glycolipid layer in the envelope of the heterocyst, and that the role of the polysaccharide layer is principally to provide a protective casing around the layer of glycolipids. The initial structural studies ofheterocyst envelope glycolipids (Bryce et al., 1972; Lambein and Wolk, 1973) led to the determination that they consisted of combinations of (i) a sugar moiety that is predominantly glucose (~90%), and to a lesser extent galactose (~10%), (ii) a hydrophobic portion that consists of C-26 (Anabaena cylindrica (W) glycolipids I and III) and C-28 (Anabaena cylindrica

784 glycolipids II and IV) polyhydroxy hydrocarbons with an sometimes a c-3, and sometimes an 3 hydroxyl, and (iii) a sugar linked to c-1 of the lipid moiety by either an ester linkage (Anabaena cylindrica glycolipids I and II) or an ether linkage (Anabaena cylindrica glycolipids III and IV). Subsequent studies showed the presence of two additional lipids in Anabaena sphaerica, one of which appeared (on the basis ofretention time in the gas chromatograph) also to be present in Nostoc muscorum and the other, in Nostoc linckia (Lorch and Wolk, 1974). Recent studies (Davey and Lambein, 1992a,b) extend the list of heterocyst glycolipids. To date, only those from Anabaena cylindrica have been directly localized to the envelope of the heterocyst (Winkenbach et al., 1972), where they serve to limit the rate of entry of (see Section III B, 1). A recent report (Soriente et al., 1992) demonstrates clearly that a heterocyst-type glycolipid from Nodularia harveyana has a lipid moiety consisting of the 3-keto derivative of 1,25hexacosanediol (Fig. 6a), and yet, because of the reactivity resulting from the 3-keto group, gives results that are consistent with all ofthe data obtained for Anabaena cylindrica glycolipid I. Both epimers that would have resulted from hydroxylation at the C-3 position, hexacosanediol and 3S,25R-hexacosanediol, were observed (Fig. 6b,c). In view of the rarity of glycolipid esters, it seems likely that the aglycones of Anabaena cylindrica glycolipids I and II are 3-keto alcohols rather than acids; the authors concur with Soriente et al. (1992) that their structures should be reinvestigated. Cyanospira rippkae contains the longer heterocyst glycolipids octacosanediol and keto-3R-octacosanol (Soriente et al., 1993). The heterocyst envelope polysaccharide in Anabaena cylindrica (W) has been very extensively defined (Cardemil and Wolk, 1976, 1979; Fig. 7A) and the corresponding (but somewhat different: see Fig. 7B,C) polysaccharides from Anabaena variabilis strain ATCC 29413 and Cylindrospermum licheniforme strain ATCC 29412 defined by composition and linkage analysis (Cardemil and Wolk, 198la). The results are consistent with the following interpretation: the polysaccharides consist of concatemers of an oligosaccharide that has a tetrasaccharide backbone, some of the glucosyl residues at the

C. Peter Wolk, Anneliese Ernst & Jeff Elhai

reducing end of that tetrasa.ccharide are substituted on the 4-position with apentasaccharide, either xylose or arabinose; some of the central glucosyl residues are substituted on the 6-position with galactose; the third glucose residue is substituted with glucose on the 2- and the 4-position, or neither; and the mannose is substituted (in Anabaena cylindrica, only partially) on the 2-position. It is unknown, but seems likely, that the ‘fibrous layer’ observed during early stages of envelope deposition in heterocysts of Anabaena cylindrica (F; Lang and Fay, 1971) consists of an uncompacted, perhaps minimally substituted, form of this polysaccharide. Excess permeability of the heterocyst envelope would increase the expenditure of photosynthetically generated reducing power that would have to be expended to reduce the that enters with the if the of the interior of the heterocyst is to be maintained at a very low value. Rippka and Stanier (1978) observed that the glycolipid layer of the heterocyst envelope fails to form under anaerobic conditions. The mechanism underlying this effect is uncertain; perhaps there is a requirement of for the hydroxylation of the envelope glycolipids; alternatively or in addition, there may be an intracellular oxygen-sensor. Murry et al. (1984b) determined that the of nitrogenase activity decreased in vivo, and the apparent for acetylene increased, with increasing values (from 15 to ofthe concentration of in equilibrium with the medium in which the heterocysts had been induced. It therefore appeared that the barrier to penetration of gases is adjusted in response to the external and perhaps so as to establish some particular intracellular Such a variation in thickness of the heterocyst envelope, and especially

Chapter 27 Heterocyst Metabolism and Development

of the glycolipid layer of the envelope, increasing with ambient has been directly demonstrated (Kangatharalingam et al., 1992). In Klebsiella aerogenes the products of the nifAL operon regulate the transcription of the rest of the nif genes in response to and to the concentration of In normal heterocysts, a combination of the envelope and active respiration largely exclude from the vicinity of nitrogenase, so that there may be no need to have nif transcription sensitively regulated by In fact, nitrogenase synthesis continues in heterocysts ofmutant strains at values ofintracellular sufficient to inactivate the enzyme essentially totally (Ernst et al., 1992; see also Peat et al., 1988). To date, no analog of the nifAL regulatory system has been found in Anabaena sp. strain PCC 7120, in which nitrogenase appears to be restricted to heterocysts. Such a system might be more readily expected in a strain such as Anabaena variabilis, in which nitrogenase may be present in vegetative cells

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under anaerobic conditions (Helber et al., 1988a; see Section III A, 1).

3. The Respiratory Apparatus of Heterocysts Anabaena variabilis can grow in the dark, fixing nitrogen, with a growth rate approximately onefourth that in the light (Wolk and Shaffer, 1976; Jensen, 1983; see also Anderson and McIntosh, 1991). It follows that in heterocysts under these conditions, respiration provides ATP, and reduced pyridine nucleotides (or pyruvate) reduce ferredoxin, at rates that are probably about 4-fold lower than the corresponding rates in the light. Walsby (1985) has pointed out that because enters heterocysts more slowly than does (see Section III B, 1) fixation can be rate-limited in the dark by the resulting limitation on the rate of respiratory generation of ATP. Respiration in heterocysts has, therefore, a dual function: removal of deleterious to the

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activity of nitrogenase and, at least in the dark, generation of ATP. The costs of fixing nitrogen include the reductant (and energy) for fixation and assimilation, the reductant to reduce that entersheterocysts together with and the ‘capital expense’ of making a heterocyst, costs that can approximate half of the photosynthetically generated electron flux within the filament (Turpin et al., 1985; Murry and Wolk, 1989). Uptake of by diazotrophic cultures is biphasic, showing major components of respiration with low and high affinity for (Fig. 8); nitrogenase activity in the dark correlates with the activity of the low-affinity oxidase, whereas cultures grown in the presence of fixed nitrogen show only the activity of the high-affinity system (Jensen and Cox, 1983). The activity with low affinity for oxygen was attributed to an oxidase in heterocysts, acting behind a diffusion barrier established by the heterocyst envelope (Jensen and Cox, 1983). Mutants with defects in synthesis ofthe heterocyst envelope do not show the low-affinity oxidase activity (Murry and Wolk, 1989). The heterocysts have so great a respiratory capacity that, although they represent a small minority of the cells of a filament, their contribution to respiration by the filament approximately equals (Jensen and Cox, 1983), or substantially exceeds (Murry and Wolk, 1989), the respiratory rate of a nitrate-grown filament. (A lightdependent consumption of oxygen by heterocysts [Smith et al., 1986] is, however, ofdubious biological significance, because it is stimulated by treatment with temperatures of 85-100 °C.) The liberation of by filaments, pre-labeled with and incubated in the presence of saturating amounts of is less inhibited by light than is the case for non-fixing filaments, presumably because heterocysts maintain a high rate of catabolism of sugar in the light, as a source of reducing equivalents for respiration and nitrogen fixation (Scherer and Böger, 1982). The membranes of vegetative cells of cyanobacteria show a differentiation between an (intracytoplasmic) thylakoidal membrane system that contains both photosynthetic and respiratory complexes, and a (peripheral) cytoplasmic membrane with respiratory capacity (see Chapter 13). Heterocysts appear to have, in addition, a third kind of membrane–one that is largely or exclusively devoted to respiration (Fig. 5). Photomicrographs show a

C. Peter Wolk, Anneliese Ernst & Jeff Elhai

paucity of absorption of chlorophyll toward the poles of heterocysts, in the region of so-called ‘honeycomb’ membranes (Braun-Howland et al., 1988), and this is precisely the region where, in the dark, the product of oxidation of diaminobenzidine, which reacts with hemoproteins, is localized (Murry et al., 1981). However, no spectroscopic evidence has been found that there are two pools ofcytochromes: one oxidized by oxidase and the other by PS I (Houchins and Hind, 1983a; however, see Chapter 10). Incorporation of into the membrane sulfolipids of mature heterocysts in filaments incubated with suggests that membrane synthesis or, more likely, turnover occurs in those cells (Giddings et al., 1981). Cytochrome reduced, by the cytochrome complex, is thought to donate its electrons either to PS I or to respiratory cytochrome oxidase, in cyanobacterial vegetative cells (Peschek, 1987; also

Chapter 27 Heterocyst Metabolism and Development see Chapters 10, 12, and 13) and in heterocysts (Böhme and Almon, 1983; Böhme and Ernst, 1984; Houchins and Hind, 1983a; Houchins, 1985). Rates of oxidation of cytochrome c by cytoplasmic and thylakoid membranes isolated from vegetative cells of Anabaena variabilis were 35 and 46 nmol (min mg whereas the corresponding rates for membranes isolated from heterocysts were 350 and 2400 nmol (Wastyn et al., 1988). These rates were further stimulated approximately 6-fold by the non-ionic detergent, n-octyl glucoside. Both immunoblotting experiments (Wastyn et al., 1988), absorption properties, and other characteristics of the cytochrome c oxidase highly purified from heterocysts of that same strain (Häfele et al., 1988) indicated that the oxidase is of an type, the only type of cytochrome oxidase thus far detected in cyanobacteria (Peschek et al, 1982; Peschek, 1987). Remarkably, despite the differences in activity, probing of immunoblots of thylakoid membranes of vegetative cells and heterocysts by antisera against cytochrome of Paracoccus denitrificans showed similar amounts of antigenic material (per protein) in the two types of cells (Wastyn et al., 1988). It appears that factors other than the concentration of the terminal oxidase limit the oxidase activity ofthe different types of membrane. Although immunocytochemical localization studies with antibodies against the oxidase apparently have not been reported, such studies with antibodies against cytochrome showed that it was concentrated specifically near the (peripheral) cytoplasmic membrane of heterocysts (Serrano et al., 1990). If there is indeed only one type of cytochrome oxidase, how it is partitioned between different membrane systems remains unresolved. Mutants whose heterocysts are defective in respiration, like those in which the physical barrier to the entry of oxygen into heterocysts is defective, would be expected to exhibit oxygen-sensitive nitrogen fixation. Anabaena sp. strain PCC 7120 mutant (Ernst et al., 1992), whose rate ofuptake of is less than 20% of that of other mutants studied, appears to be a mutant of the former type. Remarkably, mutations affecting nifE (mutant M61) and nifN (mutant LD2; Ernst et al., 1992), genes that in Klebsiella aerogenes are involved in the synthesis of the FeMo-cofactor of nitrogenase, also led to the formation of oxygen-sensitive nitrogenase activity. However, because the nitrogenase of those two

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mutants catalyzed reduction of beyond in part to their increased sensitivity to oxygen presumably resulted from a modification of nitrogenase rather than from a defect in respiration.

4. Abundant Synthesis of Nitrogenase Bands of iron-containing protein that migrate with dinitrogenase and dinitrogenase reductase are highly conspicuous in nondenaturing electrophoretograms of proteins extracted from heterocysts ofAnabaena variabilis and are not observed in corresponding electrophoretograms of proteins from vegetative cells of the same filaments or of filaments (Peterson and Wolk, 1978b; Fig. 9). That is, nitrogenase proteins are major proteins in heterocysts (see also Fleming and Haselkorn, 1974; Paerl, 1982; Smith et al., 1987). In the presence of chloramphenicol (considered sufficient to stop protein synthesis), the half-life of acetylene-reducing activity decreased with increasing (11.7 h at 0% 5.8

788 h at 20% and 3.8 h at 50% all at 0.03% as did material cross-reacting with antibodies against dinitrogenase (Murry et al., 1983a, and M. Murry, personal communication). These results indicate that turnover of nitrogenase is stimulated by It has also been directly demonstrated, by use of luciferase as a transcriptional reporter, that nitrogenase structural genes are specifically transcribed in heterocysts (Elhai and Wolk, 1990), and it has been shown that isolated heterocysts synthesize nitrogenase (Janaki and Wolk, 1982). It can be concluded, therefore, that ambient levels of do inactivate nitrogenase within heterocysts, and that in order to maintain nitrogen-fixing capacity, compensatory de novo transcription of nif genes and synthesis of the enzyme take place in heterocysts.

5. Antioxidative Agents Because oxygen that enters heterocysts can be reduced by PS I in those cells, giving rise to superoxide radicals that can be highly toxic, it is no surprise that heterocysts contain superoxide dismutase. The relative amounts of superoxide dismutase in heterocysts and vegetative cells appear to differ from organism to organism: less in heterocysts of Anabaena azollae freshly isolated from Azolla sp.(Canini et al., 1991), approximately equal (and distributed throughout the heterocyst) in Anabaena cylindrica as determined by immunogold labeling (Canini et al., 1992; see also Daday et al., 1977, and Henry et al., 1978), and much more in heterocysts of Anabaena variabilis (Bagchi et al., 1991). Because the products of dismutation of superoxide radicals are hydrogen peroxide (which is also toxic) and oxygen, one might expect that where there is superoxide dismutase there will be catalase; and although not detected in either cell type in Anabaena variabilis (Bagchi et al., 1991), it was found in roughly equal amounts in heterocysts and vegetative cells in Anabaena cylindrica (Henry et al., 1978) and in Anabaena sp. strain PCC 7119 (TelOr et al., 1986). Ascorbate peroxidase, an alternative means of removing was present at approximately equal specific activities in heterocysts and vegetative cells of Anabaena sp. strain PCC 7119 (Tel-Or et al., 1986) and in vegetative cells of Anabaena variabilis (although only when cultures had been bubbled with air), but was not detected in heterocysts of the latter strain, although they did contain glutathione and glutathione

C. Peter Wolk, Anneliese Ernst & Jeff Elhai peroxidase (as did its vegetative cells, at comparable concentration and specific activity; Bagchi et al., 1991). Comparable activities of glutathione reductase were found in the two types of cells of Anabaena sp. strain PCC 7119 (Kami et al., 1984; Serrano et al., 1984). No antioxidative agent specific to heterocysts has been identified.

C. Control of Nitrogenase Activity

1. Introduction: Control of Synthesis of Nitrogenase If nitrogenase is present and active, the extent of its activity is regulated by the availability ofsubstrates, i.e., ATP, reductant and (see Section III C, 2). can reduce the amount of active enzyme by irreversible oxidation or by eliciting a reversible inactivation (see Section III C, 3). Some nitrogenase of Anabaena variabilis appears to be under environmental control: under strictly anaerobic conditions, but not in the presence of nif transcripts and nitrogenase activity were found after only 1.5-2 and 3 h of nitrogen deprivation, respectively. Even after 15 h of nitrogen-deprivation, few proheterocysts and essentially no mature heterocysts were seen. Transcription of nitrogenase in ammonium-grown filaments was prevented by ammonium, but not by nitrate, glutamine or glutamate (Helber et al., 1988a; for the regulation of uptake of nitrogenous substances, see Chapter 16). In contrast, the nifHDK operon of Anabaena sp. strain PCC 7120 was expressed neither in the presence nor absence of unless heterocysts were formed (Elhai and Wolk, 1990; Ernst et al., 1992; see, however, Spence and Stewart, 1987). Moreover, promoter fusions of with the structural genes of luciferase, luxAB, showed that in both Anabaena sp. strain PCC 7120 and Anabaena sp. strain PCC 7118, a strain that forms no heterocysts and expresses nitrogenase only under strictly anaerobic conditions (Rippka and Stanier, 1978), light is emitted only from non-contiguous cells. In the case ofAnabaena sp. strain PCC 7120, these cells can be identified as heterocysts. It has been proposed that transcription of nifHDK in the latter two strains is under developmental control (Elhai and Wolk, 1990; see Section IV G). After addition of carbamyl-phosphate or of to anaerobically induced Anabaena variabilis (Helber et al., 1988b) or Anabaena sp. strain PCC 7120 (Haselkorn et al., 1983), the amount of nifHD mRNA

Chapter 27 Heterocyst Metabolism and Development decreases, but whether in response to diminished transcription, enhanced turnover, or both, is unknown. Evidence has been presented that ammonium inhibits nitrogen fixation by heterocyst-bearingfilamentsby inhibiting heterocyst formation and, under certain physiological conditions, by an effect on enzymatic activity, rather than by regulating nitrogenase biosynthesis in heterocysts (Murry et al., 1983b; Ernst et al., 1990a; see also Section III C, 3 and Chapter 16).

2. Metabolic Control In heterocysts, ATP is formed by PS I and by respiration; in consequence, the nitrogenase activity of heterocysts is ATP-limited under anaerobic conditions in the dark (Ernst et al., 1983). It is thought that PS I-stimulated electron transport to nitrogenase is coupled to phosphorylation, and that as a result, ATP does not limit nitrogenase activity in the light (Ernst and Böhme, 1984). Whether ATP limits nitrogenase activity in dark, aerobic conditions has not been determined, but this could depend on the availability of reductant. The nitrogenase activity of cultures under aerobic conditions in the dark correlates strongly with the previous history of illumination of those cultures (Fay, 1976; Ohmori, 1984). Cultures with large amounts of carbohydrate reserves, and heterocysts isolated from them, have high endogenous nitrogenase activity that cannot be stimulated by hydrogen (Kumar et al., 1983; Smith et al., 1985; Jensen et al., 1986). The time-course of accumulation of glycogen has been described by Ernst et al. (1984; see also Ernst and Böger, 1985). In contrast, stimulates the nitrogenase activity of cultures that lack significant stores of glycogen, and heterocysts isolated from such cultures have a low endogenous nitrogenase activity that can be stimulated by The photosynthetic activity of vegetative cells supports the nitrogenase activity of such cultures in the light essentially directly, as shown by the observation that DCMU, an inhibitor of photosynthetic fixation of is strongly inhibitory (e.g., Ernst et al., 1990b).

3. Modification of Dinitrogenase Reductase A reversible posttranslational modification of dinitrogenase reductase (NifH) that is accompanied by cessation of enzymatic activity is observed (i) upon exposure of cultures of two strains of Anabaena

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sp. to a very high concentration of or to ammonia at high pH (Smith et al., 1987; Reich and Böger, 1989; see also Pienkos et al., 1983, and Reich et al., 1986, 1987); (ii) when synthesis of carbohydrate is blocked in cultures that lack reserves of glycogen and are in equilibrium with air; and (iii) for cultures that are grown with dark-light cycles, during the periods of darkness (Ernst et al., 1990a). The supply of carbohydrates to heterocysts is presumed to affect the ability of those cells to reduce that enters them, and the treatments that lead to modification of NifH at ambient may affect that supply. It was therefore proposed that the signal for modification is correlated with an increase in the internal of heterocysts. For example, activated species of oxygen that are formed upon transfer of electrons to the that accumulates inside heterocysts might be involved in the regulation (Bagchi et al., 1991). The modification coordinates the assimilation of nitrogen with the assimilation of carbon (Ernst et al., 1990a) and may (the idea has not been directly tested) help to reduce the loss of nitrogenase during periods when the electron flux from vegetative cells to heterocysts is inadequate to prevent a buildup of in the lattercells. A similarly responsive modification of NifH is observed in diazotrophic cyanobacteria that do not form heterocysts (Stal and Bergman, 1990; Ohki et al., 1991, 1992; Brass et al., 1992). The mechanism and the metabolic activities that control the modification of dinitrogenase reductase in cyanobacteria are not known. A posttranslational regulation of nitrogenase activity was first observed in Rhodospirillum rubrum, and was shown to result fromADP-ribosylationofarginine-101ofonesubunit of the dimeric protein in response to such environmental signals as the presence of fixed nitrogen and the availability of light (reviewed by Ludden and Roberts, 1989). This arginine residue is located in a highly conserved region of the enzyme at which dinitrogenase reductase and dinitrogenase may make contact during electron transfer (Kim and Rees, 1992). In Rhodospirillum rubrum, the ADP-ribosylation that inactivates the enzyme and the hydrolytic cleavage of ADP-ribose that activates the enzyme are catalyzed by two highly specific enzymes (Ludden and Roberts, 1989). In contrast to what has been observed with Rhodospirillum rubrum, modification of dinitrogenase reductase in cyanobacteria is observed only in the presence of (Ernst et al., 1990a) and affects both subunits of the enzyme. The modified dinitrogenase reductase of Rhodospirillum

790 rubrum cross-reacts with an antibody that is specific for arginine-bound ADP-ribose, but the enzyme from Anabaena variabilis does not, and the modifying enzymes of Rhodospirillum rubrum neither activate nor inactivate isolated nitrogenase from Anabaena variabilis (Durner et al., 1994). The mechanism of modification of cyanobacterial dinitrogenase reductase remains to be elucidated.

D. Other Metabolism 1. Metabolism of Fixed Nitrogen a. Assimilation and Export Enzymological data and labeling studies with (reviewed by Wolk, 1982; also see Chapter 16) provide compelling evidence that is assimilated by the glutamine synthetase-glutamate synthase pathway in Anabaena. This is a two-step, cyclic pathway. Glutamine synthetase catalyzes the formation of glutamine from ammonium and glutamate, with the expenditure of ATP. Glutamate synthase, also called glutamine oxoglutarate aminotransferase (GOGAT), catalyzes the formation of two molecules of glutamate from one molecule, each, of glutamine and with ferredoxin as reductant. Nitrogen assimilated by heterocysts can be transferred to vegetative cells, stored in the heterocysts, or even lost from the filaments (Wolk et al., 1974; Paerl, 1984; Newton and Cavins, 1985; Subramanian and Shanmugasundaram, 1986). The evidence is substantial (albeit not as compelling) that the glutamine synthetase that assimilates the produced from is resident in the heterocyst; the heterocyst then releases glutamine, or a derivative of it, to adjacent vegetative cells (reviewed by Wolk, 1982). Glutamine synthetase is present at fairly similar levels in heterocysts and vegetative cells, as shown both by enzymatic assay (Wolk, 1982) and by immunoelectron microscopic localization (Bergman and Rai, 1989; Rai et al., 1989; Renström-Kellner et al., 1990). The location of glutamate synthase is controversial. Thomas et al. (1977) failed to detect glutamate synthase in heterocysts of Anabaena cylindrica (W). Gupta and Carr (198la) observed the formation of from by extracts of heterocysts of Anabaena cylindrica (F), and ascribed the reaction to glutamate synthase. Had they observed

C. Peter Wolk, Anneliese Ernst & Jeff Elhai the formation of from ketoglutarate, the conclusion would have been secure; but the reaction that they observed could have been due to glutaminase. Rai et al. (1982), working with A. variabilis, obtained the same results as had Gupta and Carr (1981 a), but attributed them to glutaminase; the small amount of glutamate produced from labeled was attributable to other cells contaminating the purified heterocysts. Häger et al. (1983), assaying extracts of heterocysts isolated from Anabaena sp. strain PCC 7119, observed glutamate synthase activities of 2–5 nmols ketoglutarate converted (min mg values which must be compared with glutamine synthetase activities in the same heterocyst preparations of about 50 nmol (min mg and which are in any case much lower than typical nitrogenase activities for isolated heterocysts. In general, heterocysts appear to conserve their reductant for nitrogenase: they lack the reductive pentose phosphate (Calvin) cycle (see Wolk, 1982; Cossar et al., 1985b), and do not reduce either (Giddings et al., 1981) or (Kumar et al., 1985; Rai and Bergman, 1986); and cyanobacterial glutamate synthase is dependent on reduced ferredoxin (Lea and Miflin, 1975; Häger et al., 1983; Marques et al., 1992). It would, therefore, be premature to conclude from a weak activity in one strain that the glutamate synthase-catalyzed step in assimilation of is normally localized in heterocysts. If glutamate synthase is present only in vegetative cells, then prior to being further metabolized, glutamine must move from heterocysts to vegetative cells (Thomas et al., 1977) If, on the other hand, glutamate synthase is involved in the assimilation of within heterocysts (as; depicted provisionally in Fig. 4), one might anticipate finding that its other substrate, is also synthesized in heterocysts. is derived in metabolism from the action of isocitrate dehydrogenase. Bothe and coworkers (Neuer and Bothe, 1982; Papen et al., 1983) have reported much higher activities of isocitrate dehydrogenase in heterocysts than in vegetative cells in Anabaena sp. strain PCC 7119 (the properties of the enzyme were the same from the two types of cells) and in Anabaena cylindrica (F), with exceedingly low values in the vegetative cells of the latter organism. Further analysis showed that vegetative cells of A. cylindrica contain a heatlabile, non-dialyzable, factor that can completely inhibit the isocitrate dehydro-

Chapter 27 Heterocyst Metabolism and Development genase activity of extracts of heterocysts (Neuer et al., 1983). Perhaps isocitrate dehydrogenase activity was not absent from, but merely inhibited in extracts of, vegetative cells. It is therefore unclear whether, in vivo, isocitrate dehydrogenase is active or inactive in vegetative cells, and, in turn, whether synthesis of glutamate by glutamate synthase is dependent upon provision of from heterocysts. In Jüttner’s (1983) experiments with Anabaena cylindrica (F) described in Section III A, 4, the majority ofradioactivity recovered from heterocysts after the shortest period (20 s) of pulse-labeling of filaments with was in the form of aspartate, while less than 6% was present as sugars and sugar phosphate. A series of different experiments by Lawrie et al. (1976) led to a related result. Cultures, also of Anabaena cylindrica (F), were nitrogendeprived by 18 h of incubation in the light under (80%:20%:0.04%, v/v), exposed to for 30 min, supplemented with or and the radioactivity of their soluble metabolites then measured as a function of time. Sugars and sugar phosphates were the most extensively labeled; and in the presence of glutamine was extensively labeled, with aspartate close behind; but upon addition of aspartate was by far the most extensively labeled nitrogenous metabolite. Aspartate, which receives its nitrogen in a transamination reaction from glutamate (Meeks et al., 1977; see also Wolk et al., 1976) is therefore an additional candidate for a carrier of fixed nitrogen from heterocysts to vegetative cells. Whether the dicarboxylic acid precursor of aspartate would come from the vegetative cells or from the glycolytic intermediates of heterocysts remains to be determined.

b. Synthesis of Nitrogenous Metabolites and Macromolecules Are the additional amino acids needed for synthesis of proteins made within the heterocysts or imported from vegetative cells? With one possible exception, six enzymes of arginine biosynthesis were approximately as active in heterocysts as in vegetative cells in Anabaena sp. strain PCC 7120 and in Anabaena cylindrica (F), whereas two enzymes of arginine catabolism were less active in the heterocysts (Gupta and Carr, 1981b). Approximately equal concentrations of ornithine transcarbamylase, one of the enzymes involved in arginine biosynthesis, in the two types ofcells was confirmed by immunoelectron

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microscopy (Lindblad, 1992). Thus, it seems highly probable that heterocysts can make their own arginine from basic building blocks. Isolated, metabolically active heterocysts were able to incorporate into cysteine and glutathione, but not into methionine (Giddings et al., 1981); however, incorporation into cysteine may have been due to an exchange reaction rather than to de novo synthesis of the amino acid. The finding of comparable activities of alanine dehydrogenase, glutamate-pyruvate transaminase, and glutamate-oxaloacetate transaminase in vegetative cells and heterocysts (Stewart et al., 1975) suggests that at least alanine and aspartic acid can be made in heterocysts. Jüttner (1983) found, in heterocysts isolated after 20 sec of assimilation of by filaments, that 16% of the label was in alanine, and that isolated heterocysts metabolized very extensively (fully halfof the added label) to (85%) and glutamate (13%), presumably via intermediates of the interrupted tricarboxylic acid cycle; he therefore proposed that alanine was being imported into the heterocysts. However, because very little radioactivity was found in alanine in whole filaments, it seems just as likely that alanine was generated in the heterocysts. In sum, there is only very fragmentary knowledge of the ability of heterocysts to make amino acids. Material accumulated in large amounts in the neck region at the poles of heterocysts was shown (Lang et al., 1972) to be cyanophycin, a high-molecularweight copolymer of arginine and aspartic acid that serves as a nitrogenous reserve in cyanobacteria (Simon, 1971). Gupta and Carr (1981 c) found that both the synthetase that forms the polymer and the enzyme that degrades it to arginyl-aspartate dipeptides are present at much higher specific activity in heterocysts than in vegetative cells of Anabaena sp. strain PCC 7120 and in Anabaena cylindrica (F). Autoradiography of intact filaments of Cylindrospermum licheniforme that had been prelabeled for 3 h with showed that synthesis of RNA proceeded at similar rates in vegetative cells, and in immature and mature heterocysts (Van de Water and Simon, 1984). Heterocysts in intact Anabaena variabilis filaments, but not in isolated form, accumulated label from into tRNA and rRNA (Lynn and Ownby, 1987), but there is no evidence that isolated heterocysts transcribe mRNA. It is clear from use of luciferase as a reporter (Elhai and Wolk, 1990; see also Northern analyses of RNA encoding heterocyst-specific proteins [Haselkorn et

792 al., 1983; Mulligan and Haselkorn, 1989]) that mRNA is made within mature heterocysts in situ in filaments, and that the resulting mRNA is translated in isolated heterocysts (see Section III B, 4). It may be that the principal reason that isolated heterocysts eventually lose nitrogenase activity is that they have stopped synthesizing RNA. Heterocysts can phosphorylate ADP to generate ATP (see Section III A, 6), but whether they can make purines and pyrimidines is unknown. An attempt should be made to determine whether adding all four nucleotide triphosphates, or precursors of them, stimulates RNA synthesis by isolated heterocysts. Therearebothsimilarities anddifferencesbetween the sets of polypeptides synthesized by heterocysts in situ in filaments and by isolated heterocysts (Janaki and Wolk, 1982); but clearly, both synthesize a wide range of polypeptides. According to DNA-excess RNA hybridization techniques, about 45% of the genome is expressed in heterocysts (cf. 65% in vegetative cells), and 15–25% of the genome encodes heterocyst-specific transcripts. Moreover, the latter appeared to be present at higher abundance than transcripts that were common to the two types of cells (Lynn et al., 1986). Because even 15% (~1 Mb of DNA) exceeds by a factor of about ten the coding capacity that would appear to be needed specifically for differentiation and nitrogen fixation (see sections III, A 1 and IV), one wonders whether the large number ofheterocyst-specific transcripts may relate not to those processes, but rather to metabolic responses to microaerobiosis within the heterocysts.

2. Metabolism of Mature heterocysts lack ribulose bisphosphate carboxylase/oxygenase (see review by Wolk, 1982; Cossar et al., 1985b; Rai et al., 1989; see, however, Bergman and Rai, 1989). If, as seems likely from the high specific activities of the enzymes in question, the oxidative pentose phosphate cycle and (or) isocitrate dehydrogenase are (is) used to generate reducing equivalents for nitrogen fixation, one result is to generate a large amount of and being more polar than and would presumably have greater difficulty moving out of heterocysts (unless there are carriers for it, or truly open conduits; see Fig. 4). generated by decarboxylation of isocitrate by isocitrate dehydrogenase could be stoichiometrically recycled for the synthesis of

C. Peter Wolk, Anneliese Ernst & Jeff Elhai additional oxaloacetate (and thence isocitrate) from PEP or pyruvate if the requisite enzymes are sufficiently active (see Section III A, 3), but generated by decarboxylation ofglucose 6-phosphate and 6-phosphogluconate must leave the heterocyst lest the acidify the cell, inactivating cellular metabolism. In fact, a large release of attributable to the nitrogen-fixation process has been demonstrated (Scherer et al., 1982). A gene putatively encoding carboxysomal carbonic anhydrase (Fukuzawa et al., 1992; Yu et al., 1992) and a gene whose product is involved in transport of inorganic carbon have been cloned from unicellular cyanobacteria (Ogawa, 1992). It would be of great interest to determine whether homologous genes are active in heterocysts, and whether vectorial transport of inorganic carbon occurs and is oriented outward rather than inward at their junction to vegetative cells. It has long been known (Allen and Arnon, 1955; Apte and Thomas, 1980, 1983, 1984) that there is a greater requirement for under fixing conditions than during growth with fixed nitrogen, and it is known that in Anabaena variabilis strain M-3 (Kaplan et al., 1984) and other cyanobacteria (Miller et al., 1990), transport of requires millimolar concentrations of whereas transport of requires only micromolar concentrations of However, sodium deficiency curtails photosynthesis of Anabaena sp. strain PCC 7119 within 2 h but nitrogenase activity (reduction of only after 8 h (Maeso et al., 1987). Nonetheless, it should be determined whether protracted nitrogenase activity ofheterocysts is dependent upon transport of

3. Metabolism of is the most effective known exogenous electron donor to in illuminated, isolated heterocysts (Peterson and Wolk, 1978b). In addition, uptake of hydrogen probably constitutes part of the protection of nitrogenase within heterocysts against inactivation by There are two bases for this statement. Firstly, reduction of by (the oxyhydrogen, or Knallgas, reaction: provides extensive protection against of nitrogenase in isolated heterocysts (Peterson and Wolk, 1978b). Secondly, nitrogenase generates at least one mol of per mol of fixed (Simpson and Burris, 1984), and the molar flux of into heterocysts

Chapter 27 Heterocyst Metabolism and Development is roughly half of that of (see Section III B, 1). The oxyhydrogen reaction, using internally generated hydrogen, therefore has the potential to reduce much of the that enters the heterocyst, provided that the nitrogenase is adequately active (Walsby, 1985). Reductant, and even part ofthe energy expended by the ATPase activity of dinitrogenase reductase, are thereby conserved if the is not lost by diffusion from a suspension of cells. Cyanobacterial hydrogenases (see below) appear to be nickeldependent enzymes (Zhang et al., 1984; Pederson et al., 1986; Almon and Böger, 1984; see also Hausinger, 1987). Under conditions ofhigh-temperature stress, permitted to provide protective reductant to two heterocyst-forming cyanobacteria (Pederson et al., 1986). Addition of to a suspension of Anabaena sp. strain CA abolished net hydrogen production (Zhang et al., 1984) indicating that, at least in the presence of (Spiller et al., 1978), loss of by diffusion can be prevented by the activity of uptake hydrogenase. Like nitrogenase, hydrogenase is an enzyme native to anaerobic (or microaerobic) conditions, and is in a sense a portal to the study of microaerobiosis. There is an extensive literature on hydrogen metabolism in cyanobacteria (see reviews by Lambert and Smith, 1981, and Houchins, 1984; see Chapter 12), but that literature also illustrates our general lack of information about cyanobacterial life in the absence (or near-absence) of oxygen. Strains of Anabaena sp. have both an ‘uptake’ hydrogenase which is irreversibly inactivated by oxygen and is, at least under aerobic conditions of culture, restricted to heterocysts (Peterson and Wolk, 1978a; Houchins and Burris, 1981a,b; see also Eisbrenner et al., 1981), and a ‘reversible’ hydrogenase which is present in vegetative cells as well as in heterocysts (and is termed reversible because it can catalyze the evolution as well as the uptake of hydrogen; but it is also reversible in the sense that it is reversibly inactivated by oxygen; Houchins and Burris, 1981a,b). The immediate physiological acceptor is known for neither of these types of hydrogenase, nor is the physiological donor known for the reversible hydrogenase. Microaerobic conditions of growth, and even anaerobic preincubation of extracts, greatly increase the activity (presumably in vegetative cells) of the reversible, but not ofthe uptake, hydrogenase ofeither Anabaena sp. strains PCC 7119 or PCC 7120 (Houchins and

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Burris, 1981a; Papen et al., 1986a). An antibody that was raised against a soluble, reversible hydrogenase of Synechococcus sp. strain PCC 6301 and another that was raised against a membrane-bound uptake hydrogenase of Alcaligenes latus both recognize proteins in vegetative cells and in heterocysts (Kentemich et al., 1989; Lindblad and Selstedt, 1990), but may possiblybe reacting with the same protein(s). A gene encoding a reversible cyanobacterial hydrogenase has not been cloned with certainty (see, however, Ewart et al., 1990), whereas a gene encoding an uptakehydrogenase was identified serendipitously because of its presence at the recombination site of an excision element (Section III A, 1). The numbers of hydrogenase genes and hydrogenases present, and their regulation, remain to be determined. IV. The Differentiation Process

A. Introduction After outlining a possible framework for understanding the changes that underlie differentiation in Anabaena sp., this chapter will present a discussion ofthe progression ofdifferentiation from a vegetative cell to a mature heterocyst, here defined as a cell capable ofaerobic fixation of A discussion of the process of pattern formation, i.e., the development ofheterocyst spacing, will be left to the end. Despite the increasing availability of appropriate probes in the form ofcloned and sequenced genes, and defined antibodies, no systematic study of the progression of differentiation at any but an ultrastructural level has been performed (Wilcox et al., 1973b). In order to create a time-line of the process of differentiation (Fig. 10), it is therefore necessary to piece together data from different strains. Fortunately, extensive genetic study of one strain (Anabaena sp. strain PCC 7120) has provided sufficient information to organize the combined data. In that strain, certain unidentified genes respond to nitrogen-stepdown within 20 min (Section IV C); a developmentally critical gene, hetR, starts to be activated within 2 h, and is active in spaced cells within 3.5 h (Section IV D); between about 4 and 7 h, a gene (hetP) whose insertional inactivation blocks the formation of proheterocysts (Section IV D), another (devA) that is needed for their maturation (Section IV E) and a gene (hepA) that is involved in deposition of the envelope of the

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heterocyst (Section IV F) are first activated; at about 18 h, one or two DNA rearrangements take place, and activity of genes that are involved in electron transfer to is first observed (Section IV G); and aerobic nitrogen fixation is under way within 24 h.

B. Candidate Principles One of the two major enigmas facing analysis of heterocyst development is: what regulates the transcription of development-related genes after heterocyst differentiation has been initiated? It has been estimated that about one hundred genetic loci

C. Peter Wolk, Anneliese Ernst & Jeff Elhai

are required for sporulation in Bacillus sp. (Losick et al., 1989) and in Myxococcus sp. (Kroos et al., 1986). It would be surprising if heterocyst differentiation required a greatly different number. What little is known about the genetic events underlying the differentiation of heterocysts may be organized by referring to three principles ([l]–[3], below) that have emerged from the study of differentiation in other eubacteria [Myxococcus sp., Kroos and Kaiser (1987); Streptomyces sp., Chater (1989);Caulobacter sp., Bryan et al. (1990), and especially Bacillus subtilis, Errington (1993)].

Chapter 27 Heterocyst Metabolism and Development

1. During the Course of Differentiation, Genes are Expressed in an Ordered Sequence Figure 10 illustrates this principle with reference to several genes that are induced during the differentiation of heterocysts. Mutants of Anabaena sp. strain PCC 7120 can, apparently, be similarly arrayed: see Table 1. Almost two dozen additional mutations with transposons at different loci have been identified that are incapable of aerobic fixation of (Ernst et al., 1992), but the expression of only a few of the genes directly affected has been studied. It has long been appreciated that proteins appear and disappear at characteristic times during heterocyst differentiation (Fleming and Haselkorn, 1974). However, few have been characterized.

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2. The Expression of Genes at One Stage Depends Upon Gene Products that Appear at a Previous Stage Studies in other bacteria of epistatic relationships amongst developmentally controlled genes have produced trees of genetic dependencies that give a broad picture of the regulatory strategies used to control differentiation in those organisms. The premise that heterocyst differentiation is governed by an interlocking series of regulatory steps rests on several observations: results (see Section IV G) concerning the expression of the nitrogenase structural genes, nifHDK; the observation that expression of hepA (Holland and Wolk, 1990) and of the genes mutated in M7 and P2 (Ernst et al., 1992),

796 depends upon the integrity of hetR (Black et al., 1993; Y. Cai, I. Maldener, and C. P. Wolk, unpublished results); and the findings of Bradley and Carr (1977) that maturation of prohetcrocysts of Anabaena cylindrica (F) is aborted by treatment with ammonium, rifampicin, proflavin or fluorouracil only up to 8 to 10 h after nitrogen-stepdown (see also Van de Water and Simon, 1984), and that development to proheterocysts is aborted by several of those same substances only up to 1 to 2 h after nitrogen stepdown. One may conjecture that these two periods correspond to the times of activation of envelopebiosynthetic genes and of hetR, respectively. Even if heterocyst differentiation does depend upon a series of regulatory steps, there are several ways in which genetic dependencies might be organized (Fig. 11). Nitrogen-deprivation might evoke a developmental program of gene expression

C. Peter Wolk, Anneliese Ernst & Jeff Elhai only in cells that are destined to become heterocysts (Fig. 11 A, B). Several heretofore characterized genes that are induced in response to nitrogendeprivation (hepA, hetR, and nifH) are expressed in well-spaced cells that are probably presumptive heterocysts. No currently available information distinguishes whether heterocyst differentiation involves a linear developmental pathway (Fig. 11 A) or a branched program (Fig. 11 B). For example, a pathway ofgene expression that leads to the synthesis and functioning of nitrogenase may branch off from the pathway of morphological differentiation. This possibility is consistent with observations that Anabaena sp. strain PCC 7118 can express nitrogenase activity in spaced cells despite there being little or no visible differentiation of heterocysts (Elhai and Wolk, 1990; see also Spence and Stewart, 1987). It appears likely that nitrogen deprivation evokes a program of gene expression in non-differentiating cells as well as in those that are differentiating (Fig. 11C). Two proteins are thought to be active predominantly or exclusively in vegetative cells of filaments: sucrose synthase (Schilling and Ehrnsperger, 1985) and an inhibitor of isocitrate dehydrogenase (Papen et al., 1983). A putative modifier of the activity ofglutamine synthetase may be a third such example, because glutamine synthetase is regulated differently by ammonium depending upon whether or not the filaments have been deprived of fixed nitrogen (Orr and Haselkorn, 1982). In addition, gene tln6 (Wolk et al., 1991; Y. Cai and C. P. Wolk, unpublished results), although not required for differentiation, has been found to be expressed specifically in all cells except those that are differentiating. If gene expression in vegetative cells does differ from gene expression in vegetative cells grown on fixed nitrogen, then the regulatory basis forthe difference mustbe elucidated.

3. Regulation of Expression Occurs Primarily, But Not Exclusively, at the Level of Transcription There is growing evidence that transcriptional control is an important mode of regulation during heterocyst differentiation. Fifteen to 25% ofthe coding capacity of the Anabaena variabilis genome is expressed in heterocysts, but not in filaments grown aerobically with combined nitrogen (Lynn et al., 1986). During the course ofdevelopment, several genes are regulated at a transcriptional level (see Fig. 10). These include

Chapter 27 Heterocyst Metabolism and Development tln2 and tln6 (no known function; Wolk et al., 1991); cpcBA and apcAB operons (encoding the major phycobiliproteins phycocyanin and allophycocyanin; Wealand et al., 1989); hetR (Black et al., 1993; required early in heterocyst differentiation: Buikema and Haselkorn, 1991b); hepA (Wolk et al., 1993; required for polysaccharide biosynthesis or stability; Wolk et al., 1988); and nifHDK (Elhai and Wolk, 1990). Several other characterized genes are also probably controlled at the level of transcription, as judged by changes in the abundance of mRNA. These genes include glnA (encoding glutamine synthetase; active, but transcribed from different promoters, in heterocysts and vegetative cells; Tumer et al., 1983); sigB and sigC (Brahamsha and Haselkorn, 1992; these genes encode sigma factors, modular components of RNA polymerase that are responsible for the recognition ofpromoter sequences (see Helmann and Chamberlain, 1988, and Chapter 20); and several genes expressed late in differentiation that are related to the synthesis or function of nitrogenase (Böhme and Haselkorn, 1988; Mulligan and Haselkorn, 1989; Borthakur et al., 1990; Golden et al., 1991). If transcriptional control is responsible for differences between vegetative cells and heterocysts, one would expect to find that the activity of certain promoters is confined to one cell type or the other. Indeed, this has been shown to be true for (expressed only in heterocysts) and (expressed only in vegetative cells) (Elhai and Wolk, 1990). Two other promoters, (Black et al., 1993) and (Wolk et al., 1993) show localized expression during the course of heterocyst differentiation. It is very likely, but at present not known with certainty, that the cells expressing these two promoters are en route to becoming heterocysts. All examined promoters of Anabaena sp. strain PCC 7120 known to be expressed in vegetative cells contain typical E. coli-like –10 sequences (see Chapter 20) and are recognized in vitro by RNA polymerase isolated from vegetative cells (Schneider et al., 1991). However, this polymerase is not able to recognize a promoter, that is active in filaments only under conditions that elicit the formation of heterocysts (Schneider et al., 1987). glnA and all other genes that are known to be expressed in heterocysts and whose transcriptional start sites have been identified, have at least one transcriptional start site preceded by a sequence that has little similarity to E. coli-like promoters. Several are known to have

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multiple transcriptional start sites, and thus multiple promoters: glnA (Tumer et al., 1983); atpBE (encoding subunits of the proton-translocating ATP synthase; Curtis, 1987); psbB (encoding a PS II protein; Lang and Haselkorn, 1989); sigA (encoding the major sigma factor of Anabaena sp. strain PCC 7120; Brahamsha and Haselkorn, 1991), and hetR (Buikema and Haselkorn, 1992). These alternative promoters are differentially active, depending upon the nitrogen status ofthe filament. Promoter switching has been observed also in Bacillus sp., as a strategy to maintain expression of genes that are required under different conditions (Wang and Doi, 1987; Chibazakura et al., 1991). A striking feature of regulation in other bacteria that differentiate is that the expression of sets of genes is directed by an ordered appearance of sigma factors (Losick and Stragier, 1992; Apelian and Inouye, 1990; Brun and Shapiro, 1992). It is possible that switching of sigma factors is an integral part of transcriptional control over heterocyst differentiation as well. Cyanobacteria possess an RNA polymerase that is similar in most respects to that of other bacteria, except that whereas the subunit substructure ofthe RNA polymerase ofother eubacteria is the homolog of the subunit from diverse cyanobacteria is comprised of two polypeptides from separately transcribed genes (Schneider et al., 1987; Schneider and Haselkorn, 1988; Xie et al., 1989; Xie and Potts, 1991; Bergsland and Haselkorn, 1991). The RNA polymerase is associated with several sigma factors (Brahamsha and Haselkorn, 1991, 1992; Tanaka et al., 1992; L. Caslake and D. A. Bryant, personal communication). At least two novel sigma factors, encoded by the sigB and sigC genes, appear during heterocyst differentiation of Anabaena sp. strain PCC 7120 (Brahamsha and Haselkorn, 1992). These sigma factors are not required for heterocyst differentiation, since mutants in which sigB, sigC, or both are disrupted grow normally on Both mutants exhibit modified colony morphology, however, so that the expression of those two sigma factors is not without effect. The non-differentiating cyanobacterium Synechococcus sp. strain PCC 7002 has at least five sigma factors that bear strong sequence similarity to the major sigma factors of other eubacteria (e.g., Escherichia coli or Bacillus subtilis L. Caslake, T. Gruber, and D.A. Bryant, personal communication). These observations further illustrate that the occurrence of multiple sigma factors

798 need not be related to morphological differentiation. Bacteria frequently sense and react to environmental changes by means of conserved twocomponent systems (Stock et al., 1990). In such systems, one component senses some physiologically relevant signal, such as nitrogen status, and is stimulated to phosphorylate a second component. The second component, depending upon its phosphorylation state, differentially affects some physiological response, often one that is transcriptionally regulated. Protein phosphorylation and dephosphorylation in response to environmental stimuli have been observed in cyanobacteria (Allen and Holmes, 1986; Mann et al., 1991), but it is not known whether any of the target proteins directly affects transcription. Proteins PatA and PatB are involved in the patternization of heterocyst formation (Liang et al., 1992, 1993). PatA strongly resembles the regulatory component of such two-component systems, particularly in the positioning ofan aspartate residue at which such proteins are phosphorylated (Liang et al., 1992). However, PatA does not contain a known DNA-binding motif; moreover, neither a corresponding sensory component nor any promoters that are regulated by that protein have yet been identified. PatB has such a DNA-binding motif, but is not known to be phosphorylated (Liang et al., 1993). Post-transcriptional regulation of RNA and posttranslational modification ofproteins play an integral role in controlling differentiation in other bacteria (Burbulys et al., 1991; Strauch and Hoch, 1992; Losick and Stragier, 1992). Cyanobacteria are certainly capable ofsuch regulatory processes (Allen and Holmes, 1986; Mann et al., 1991;Kulkarni et al., 1992; Bovy, 1993a,b; Sobczyk et al., 1993). The disruption ofone gene (pknA) that evidently encodes a protein kinase decreases the frequency of heterocysts (Zhang, 1993), but it is unclear whether theactionofthe proteindirectlyaffectsdifferentiation or whether the observed effect is an indirect response to a change in rate of growth.

C. Very Early Responses to Nitrogen Deprivation Certain genes that lack a discernible developmental phenotype respond with increased transcription within 20 min of nitrogen-stepdown (Y. Cai and C. P. Wolk, unpublished). Any of these genes that respond to the intracellular concentrations of nitrogenous metabolites might provide a means of

C. Peter Wolk, Anneliese Ernst & Jeff Elhai monitoring those concentrations after stepdown or of visualizing gradients of concentration of those metabolites within filaments. For a fuller account of the mechanisms by which nitrogen-fixing cyanobacteria respond initially to nitrogen deprivation, the reader is referred to Chapter 16. The production of phosphorylated derivatives of guanosine is a common response of bacteria to nutrient stress, producing the stringent response, a set of economies that includes the reduction of biosynthesis ofstable RNA (Cashel and Rudd, 1987). Mutants of Bacillus sp. (Ochi et al., 1981) and Streptomyces sp. (Ochi, 1986) defective in this response are unable to differentiate normally under physiological conditions. Anabaena sp. has been reported to produce ppGpp within 30 minutes of nitrogen deprivation (Akinyanju and Smith, 1979, 1982, 1987; however, the phenomenon was not observed by Adams et al., 1977). There has been no report of a classical stringent response in cyanobacteria; and unlike non-diazotrophic bacteria, nitrogen-fixing cyanobacteria might react to nitrogen deprivation as a temporary condition rather than one that warrants long term economies. Within two hours of nitrogen deprivation, activation of hetR (Buikema and Haselkorn, 1991b) has begun (Black et al., 1993; see sections IV, D and V, D), proteolysis has measurably increased (Wood and Haselkorn, 1980;Thiel, 1990), and proteolytically derived amino acids are excreted (Fay, 1969; Ownby et al., 1979; Fleming and Haselkorn, 1974; Thiel, 1990). At least two proteases take part (Wood and Haselkorn, 1979) in the major reorganization of proteins that has been shown (Fleming and Haselkorn, 1974) to occur during heterocyst differentiation. One of these, a serine protease, has been isolated and characterized (Lockau et al., 1988), and by means of a null mutant has been shown not to be essential for heterocyst formation under laboratory conditions (Maldener et al., 1991). Major targets for a second protease, but not when slowly growing cultures are subjected to nitrogenstepdown (Thiel, 1990), are the phycobiliproteins (Wood and Haselkorn, 1980), that may initially comprise over 60% of the soluble protein within the cell (Bogorad, 1975). It is probable that transcription ofthe apcAB (encoding allophycocyanin) and cpcBA (encoding phycocyanin) genes ceases; cpcBA mRNA reappears once nitrogen-deprivation is alleviated, initially by protein breakdown and later by fixation of (i.e., after mature heterocysts appear; Wealand

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et al., 1989). Isolated, mature heterocysts of Anabaena variabilis may have less then 5% as much cpc mRNA as do vegetative cells under such conditions (Johnson et al., 1988) or considerably more than that (Section III A, 7). Cyanophycin, an arginine-aspartate copolymer (Simon, 1971), constitutes a second principal nitrogen reserve that is mobilized rapidly when nitrogen-depletion becomes severe (Mackerras et al., 1990).

D. HetR Plays a Critical Role Early in Differentiation Non-fragmenting mutants that show no obvious signs of differentiation (the phenotype) have been isolated on several occasions (Wilcox et al., 1975; Rippka and Stanier, 1978; Mishra and Tiwari, 1986; Spence and Stewart, 1987; Buikema and Haselkorn, 1991a; Ernst et al., 1992). Anabaena sp. strain PCC 7118, although can differentiate to some extent, as judged by patterned loss of phycobiliprotein fluorescence (Elhai and Wolk, 1990). Mutants with lesions in hetR are the best characterized of the nonfragmenting mutants. The gene was cloned by complementation, and sequenced. The protein that it encodes lacks any known DNA-binding motifs and was not found to show similarity to any previously sequenced protein (Buikema and Haselkorn, 1991b). The gene is significantly induced within two hours after nitrogen deprivation, and at least 20-fold within several hours in well separated cells (probably those that are becoming heterocysts; Black et al., 1993; Section IV B, 3). It remains active in mature heterocysts (Fernández Piñas and Wolk, 1994). In Nostoc ellipsosporum, a hetR mutation also blocks the differentiation of akinetes, even under conditions where akinetes normally form but heterocysts do not, and hetR remains active in mature akinetes (Leganés et al., 1994). The role of hetR is not, therefore, specific to heterocyst formation. Induction of hetR in Anabaena sp. strain PCC 7120 requires that the gene be intact (Fig. 12), suggesting that the encoded protein participates in a positive regulatory circuit that controls the expression of hetR (Black et al., 1993). As might be expected by the requirement of an intact hetR gene for heterocyst maturation, hepA (Black et al., 1993) and the gene mutated in M7 (see Section IV E and Table 1; Y. Cai and C. P. Wolk, unpublished results) are induced only if hetR is functional. Because the effect of hetR on the activation of these two genes is long delayed,

it is apt to be very indirect. The properties of hetR are consistent with the idea that it acts as a master switch of heterocyst differentiation. Positive autoregulation is unusual, perhaps because it magnifies small differences in expression and effectively restricts the promoter to only two levels of activity. This characteristic is, however, precisely what is desired in a control mechanism that serves to stabilize developmental decisions. Positive autoregulation is seen in the control of lysogeny [coliphage lambda gene cI(Ptashne, 1992)], ultimate steps in Bacillus sp. differentiation [activation of sigG (Sun et al., 1992) and of sigK (Kroos et al., 1989)], commitment to conidiophore development in Aspergillus niger [brlA (Adams et al., 1988; Timberlake, 1991)], and segmental pattern formation in Drosophila melanogaster[eve(Jiang et al., 1991)]. In each of these cases, autoregulation functions to ensure continued expression of the gene when the initial, inducing conditions no longer remain. It is not essential for HetR, the protein encoded by hetR, to interact directly with DNA to be part of an autoregulatory circuit. It might, for example, interact

800 with a different DNA-binding entity, presumably protein but conceivably RNA, although a legion of less direct mechanisms is possible. As noted above, physiological experiments of Bradley and Carr (1977) indicated that a developmental landmark occurs after approximately 1–2 h of nitrogen deprivation. The activation of the hetR gene may be that landmark. Three additional, non-fragmenting Het¯ mutants (Ernst et al., 1992), denoted N10 (mutated in hetN; Black and Wolk, 1994), P6 (mutated in hetP; Fernández-Piñas et al., 1994), and C3 (mutated in hetC; Khudyakov and Wolk, 1994) have been identified, and will be discussed further in Section V B, 3. The hetN and hetP genes have been mapped distant from hetR (Kuritz et al., 1993; see Chapter 19), whereas hetC maps ca. 1100 bp 5' from hetP, and is in the same orientation. There is no reason to think that all genes have been identified that, if mutated, would result in a non-fragmenting Her phenotype.

E. Genes Affecting the Maturation of Proheterocysts Proheterocysts, cells that are en route to completion of differentiation as mature heterocysts, pass through six morphologically distinguishable stages prior to maturation (Wilcox et al., 1973b; reviewed by Wolk, 1982). Although one does not know what regulates developmentally active genes once cells have differentiated to the stage of being proheterocysts, a process is available to screen for relevant regulatory genes, and a possible example of such a regulatory gene has been identified. Some mutants that are unable to fix nitrogen under aerobic conditions mutants; see Table 1 A, B) fail to complete morphological differentiation. Such developmental arrest may correspond specifically to an effect on envelope formation phenotype; see Table 1 A, B), or may more generally involve aborted maturation of the heterocyst protoplast. In the latter case, developmental arrest may occur due either to a lesion in the developmental regulatory program, or to a lesion in the nutritional interactions that are required to support that program. Diaminobenzidine (DAB) is normally oxidized by, and thereupon forms a dark brown precipitate in, the honeycomb membranous region of the heterocyst protoplast before its oxidation is observed in vegetative cells. Mutants (denoted see Table 1 A, B) that fail to oxidize

C. Peter Wolk, Anneliese Ernst & Jeff Elhai diaminobenzidine may be defective in a specific hemoprotein required for oxidation of DAB. Alternatively and more generally, those mutants may be defective in formation of the honeycomb membranes in which oxidation of DAB takes place. There is no reason to think that a mutant need be or that a mutant need be so that a phenotype that results from a single mutation appears to correspond to a mutation that pleiotropically, but specifically, affects heterocyst maturation.Three mutants have been identified and M7; Ernst et al., 1992), and the last of them has been extensively studied (Maldener et al., 1994). Reconstruction of the mutation gives rise to the same phenotype, showing that the phenotype is due to the presence of the transposon at a single site in the chromosome. Because the heterocysts of this mutant (unlike those of some other mutants) have neither vacuoles nor a shrunken protoplast, the morphology of the mutant gives no reason to suggest that the heterocysts of M7 have experienced an interruption of substrate supply from vegetative cells. The sequence of the gene that is interrupted in that mutant, however, shows homology to various permeases, suggesting that such an interruption, rather than an alteration in regulation, may be the basis of the phenotype.

F. Deposition of the Heterocyst Envelope 1. Mutant Phenotypes The heterocyst envelope consists of a layer of polysaccharide surrounding a layer of glycolipid laminae, which in turn surrounds a wall layer presumably corresponding to that of normal vegetative cells. That the underlying wall is ‘normal’ is supported by a few, and crude, chemical determinations (Dunn et al., 1971), and is consistent with low-resolution electron micrographs (Lang, 1965; Lang and Fay, 1971; Wilcox et al., 1973b). What few results there are suggest that amino acids are incorporated into the wall early during the differentiation process, and that slow synthesis or turnover of muramic acid (N-acetylglucosamine, a constituent of peptidoglycan) occurs throughout the process. How the external envelope layers can be deposited through the preexisting wall remains unclear, but there is abundant precedent for deposition of extracellular polysaccharides by bacteria (Leigh and

Chapter 27 Heterocyst Metabolism and Development Coplin, 1992). A large number of pertinent genes has been transposon-tagged (Ernst et al., 1992), and two from Anabaena sp. strain PCC 7120 have been cloned by complementation (Wolk et al., 1988). Only one of these has been studied to any substantial extent. This gene, originally called hetA (Holland and Wolk, 1990) but later renamed hepA (Ernst et al., 1992), is required for synthesis or stabilization of the polysaccharide layer ofthe heterocyst envelope. It is first activated at 5 to 7 h after nitrogen-stepdown, apparently specifically in cells that are differentiating. By 10 h after nitrogen-stepdown, about when morphological differentiation first becomes clear by transmission light microscopy, it is transcribed perhaps two orders ofmagnitude more strongly than before induction (Wolk et al., 1993). The proximal cause of this burst of transcription is not known, although it is known that it is dependent on activation of hetR several hours earlier (Black et al., 1993). Other mutants (Currier et al., 1977; Wolk et al., 1988; Ernst et al., 1992; mutants NF30; EF113; and N9, and M8, respectively) show, as abnormalities of normal envelope polysaccharide deposition in heterocysts, (1) thick and thin areas of deposition; (2) slight, pole-localized synthesis; (3) barrel-stave deposition, in which a cylindrical envelope is not closed at the ends; (4) envelopes formed but detached; and (5) aberrant pore structure. Each of these phenotypes implies the existence of a mutable morphogenetic determinant. The identification, regulation, and coordination of the corresponding genes all await elucidation, and those that have been transposon-tagged (Ernst et al., 1992) or that have already been complemented (Wolk et al., 1988) are immediately accessible to study. Although it is known that certain of these mutations are not clustered (hepB, mutated in EF113, and hepA are separated by > 0.9 Mb; Kuritz et al., 1993), it seems likely that others will be, but no systematic attempt has yet been made to look for contiguous, related functions in cloned DNA that complements mutations. There is extensive recent interest, and advances, in the genetic engineering of higher plants for production ofnovel lipids (Ohlrogge et al., 1991), so that identification ofthe genes involved in heterocyst envelope glycolipid synthesis would be of much interest. Two UV-induced mutants (EF114 and EF122) of Anabaena sp. strain PCC 7120 that were unable to fix nitrogen under aerobic conditions showed no synthesis of heterocyst envelope

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glycolipid and had a corresponding circumferential gap in their envelopes just inside of the envelope polysaccharide (Wolk et al., 1988). These mutants have not yet been complemented. Nine genetically distinguishable, transposon-generated mutants (Ernst et al., 1992) proved deficient in envelope glycolipids. Among the nine, most showed pleiotropic effects in the polysaccharide structure, while mutants P2 and E96 seem the best candidates for effects specific to production of envelope glycolipid. These mutants also await analysis. It must be borne in mind that the apparent absence (or presence ofonly trace amounts) of envelope glycolipids in some such mutants may be attributable not to non-formation ofthe glycolipids, but to their loss from the filaments in the absence of a solid barrier of polysaccharide (Murry and Wolk, 1989; see also Garcia-Gonzalez et al., 1991).

2. Heterocyst Envelope Polysaccharide Many cyanobacteria, including strains of Anabaena sp., produce copious amounts of extracellular polysaccharide under certain conditions of growth, and in Anabaena cylindrica (W), the monosaccharide composition of the extracellular polysaccharide (glucose, mannose, galactose, xylose) in the heterocyst envelope (73 ± 5 : 21 ± 4 : 3 ± 1 : 4±2) shows a familial resemblance to the monosaccharide composition of the extracellular polysaccharide of vegetative cells (47 ± 3 : 25 ± 3 : 6 ± 1 : 21 ± 3) (Dunn and Wolk, 1970), although the former polysaccharide is deposited as a dense layer and the latter polysaccharide is diffluent. However, whereas the details of the heterocyst envelope polysaccharide have been extensively studied (Section III B, 2), no analysis of the structure of the vegetative cell extracellular polysaccharide has been (to our knowledge) presented for any cyanobacterium, despite an abundance of compositional analyses (Wolk, 1973)! There is, therefore, no way of even guessing whether, beyond synthesis ofthe constituent sugars, synthesis of heterocyst envelope polysaccharide represents a modification of a process that can take place in vegetative cells or represents a wholly different process. Isolated heterocysts of Anabaena variabilis, incubated with metabolize it to and arabinose, and incorporate the sugars into their envelopes with glycosidic linkages characteristic of their envelope polysaccharide (Cardemil and Wolk,

802 1981b). Evidence obtained suggested that phosphoglycolipids, presumptively polyisoprenol monophosphate glycolipids, are involved as carriers of glycosyl residues between sugar nucleotides and a lipid-diphosphate-linked, forming oligosaccharide. Completed oligosaccharide units would then be donated to an elongating polysaccharide chain. If the polysaccharides from Anabaena cylindrica (W) and Anabaena variabilis are taken as models, and if one assumes that systems for generating activated and lipid-linked sugars are shared with vegetative cells, then four heterocyst-specific enzymes would be required to generate the backbone; six enzymes would be required to attach side chains to the backbone; one to concatemerize the fully formed oligosaccharide repeating subunits; and one (see Cardemil and Wolk, 1981b), a phosphatase, for resynthesis of lipid-linked oligosaccharide. Loss ofan enzyme required for synthesis or concatemerization of the backbone would be expected to give rise to a phenotype such as that of mutant M22 (Ernst et al., 1992), in which the heterocyst envelope is missing. On the other hand, the loss of an enzyme required to add a side-branch to that backbone, as part of the synthesis of the oligosaccharide repeating subunit, might reduce the non-covalent interactions of polymerized polysaccharide, and so lead to a diffluent envelope, as in mutants M64 (Ernst et al., 1992) and EF116 (Wolk et al., 1988). The product of the hepA gene (Holland and Wolk, 1990) that is mutated in EF116 is, in fact, most closely related to ATPdependent inner membrane transport proteins (R. Jones, personal communication). The polysaccharide layer of the envelope of preexisting heterocysts also decomposes, and the envelope glycolipids are lost, when cultures of Anabaena sp. strain PCC 7119 are incubated in the absence of boron (Garcia-Gonzalez et al., 1991), a condition which impairs growth of that organism only when it is fixing (GarciaGonzález et al., 1988). These results suggest that the heterocyst envelope is maintained intact by complexing with boron, although it is certainly possible that, in addition, boron is involved in biosynthesis of the envelope.

3. Envelope Glycolipids Despite the importance of the glycolipid layer as a barrier to entry of oxygen and the availability of pertinent mutants, the biochemistry of biosynthesis of envelope glycolipids remains virtually unin-

C. Peter Wolk, Anneliese Ernst & Jeff Elhai vestigated. A gene, hglK, that is required for this biosynthesis has been cloned and sequenced (Buikema and Haselkorn, 1993). Extensive incorporation of from into the heterocyst envelope glycolipid fraction of Anabaena cylindrica (F) occurs after imposition of conditions of nitrogen deprivation (Krepski and Walton, 1983). Transcripts that encode biotin carboxyl carrier protein, a subunit of acetyl coenzyme A carboxylase, increase in abundance upon nitrogen stepdown (Gornicki et al., 1993). If heterocyst-specific, this subunit may be active in the pathway of biosynthesis of envelope glycolipid. If, as is possible, heterocyst glycolipids, once deposited, do not turn over, a heterocyst that forms with a dense layer of glycolipid will not subsequently be able to increase the permeability of its envelope.

G. Developmental Control of Nitrogenase Activity The Anabaena sp. strain PCC 7120 structural genes for nitrogenase (nifH, nifD, and nifK) and for the heterocyst-specific ferredoxin (fdxH) that conveys electrons to nitrogenase (see Sections III A,1 and 2) are first transcribed between about 18 and 24 h after nitrogen-stepdown (Böhme and Haselkorn, 1988; Golden et al., 1991). This is very close to the time that nitrogenase activity is first measured under aerobic conditions. Fusions to luxAB, encoding bacterial luciferase, have shown that transcription from the nifH promoter is specific to heterocysts. It is not known why the promoter of the nifHDK operon is first utilized at about 18 h, nor why fdxH is first transcribed at that time; but it is known that transcription from the nifHDK promoter extends to nifK only after the 11-kb excision element located within nifD (Section III A, 1) is excised (Golden et al., 1991)–again at about 18 h (Golden et al., 1985). The excision of that element depends upon activation of the xisA gene, that encodes the excisase of that element. The xisA gene is itselftranscribed so weakly and (or) ephemerally that its transcript has not yet been detected. Upstream from xisA is a region that regulates its excision (Brusca et al., 1990). A proteinaceous DNA binding factor, denoted VF1 and later renamed BifA, has been identified that binds upstream from xisA and is a candidate for a regulator oftranscription of that gene (Ramasubramanian et al., 1991). BifA also binds strongly to sequences upstream from glnA,

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near the nif-like promoter (Tumer et al., 1983), and weakly to the promoter regions of nifH and rbcLS (Chastain et al., 1990). The gene encoding BifA, also referred to as NtcA, has been cloned and sequenced (Wei et al., 1993; Frías et al, 1993); the protein was found to exhibit sequence similarity to NtcA, a nitrogen-responsive regulatory protein from Synechococcus sp., as well as to a family of bacterial regulators typified by the cAMP-binding protein CRP (VegaPalas et al., 1992). A role for BifA in genetic regulation remains to be demonstrated, but preliminary results indicate that mutants lacking BifA require ammonia for growth (like mutants) and fail to progress beyond the proheterocyst stage after nitrogen deprivation (J. Golden, personal communication; E. Flores, personal communication).

V. Pattern Formation and Perpetuation

A. The Normal Pattern When long, nitrogen-replete filaments of Anabaena sp. are deprived of nitrogen, heterocyst formation is initiated simultaneously at many positions along the filaments. The frequency distribution of the number of vegetative cells between developing heterocysts rises to a maximum as spacing goes from 1 to about 9 cells, and then falls (Wolk and Quine, 1975; Fig. 13). In contrast, the frequency distribution of the number (n) of cells between randomly selected, differentiating cells in a linear array is a monotonically decreasing exponential function ofthe form where f is the fraction of cells that are differentiating (see Wolk, 1967). The fundamental fact of the pattern of distribution of heterocysts is, therefore, that those cells are not randomly spaced. During subsequent growth on new heterocysts appear, more or less equidistant from preexisting heterocysts, and so the pattern is perpetuated. The simplest model that has been proposed (Fay et al., 1968) to account for the perpetuation of the pattern is that a product of fixation acts, as does exogenously supplied or a derivative of it, as an inhibitor of heterocyst formation. A possible alternative model is that vegetative cells produce a readily diffusible stimulator of heterocyst formation, with heterocysts acting as localized sinks for that stimulator, and reaction with acting as a metabolic sink for the same stimulator. However, the results of surgical experiments with Cylindrospermum licheniforme

were inconsistent with this model (Wolk and Quine, 1975). Occasionally, two new heterocysts form in the interval between preexisting, distant heterocysts, giving rise to three inter-heterocyst intervals of approximately equal length. It appears that the simplest models are inadequate, alone, to account for this observation, which is suggestive of a process that may occur on a larger scale during de novo pattern formation (see Section V C). Minimum doubling times vary from about 4 to 24 h in different strains; and as a correlation during growth on between strains as well as for a particular strain, the average number of vegetative cells per heterocyst tends to increase as the doubling time increases (Fogg, 1949; Antarikanonda and Lorenz, 1982). The lesser fraction of heterocysts presumably reflects a lesser need for fixation of dinitrogen (it is not just that the nitrogenase activity per heterocyst decreases) as well as a decreased expenditure of photosynthate on the production of

804 heterocysts. Because the pattern of spaced heterocysts is evident at the very earliest stages of morphological differentiation, long before heterocysts are mature, pattern formation upon nitrogen stepdown does not, at least initially, depend on nitrogen fixation (Neilson et al., 1971). In fact, a pattern is present much earlier: fusions of luxAB, encoding luciferase, to (Wolk et al., 1993) and to (Black et al., 1993) establish that transcription from these promoters takes place in well-spaced cells after only 7.5 h, and even 3.5 h, of nitrogen-deprivation, respectively, long before any morphological change is observed. The second major enigma facing analysis of heterocyst development is: how is it determined which cells will differentiate?

B. Natural and Induced Variations on the Normal Pattern Unusual patterns of positions of heterocysts have been observed in mutants, as well as in wild-type strains that have been subjected to certain laboratory manipulations. Analysis of these variations may shed light on the mechanisms underlying normal pattern formation.

C. Peter Wolk, Anneliese Ernst & Jeff Elhai have been isolated (Buikema and Haselkorn, 199la; Ernst et al., 1992), but in some of these mutants, fragmentation to filament lengths below which differentiation fails to take place in the wild-type strain occurs only long after 24 h, an interval of time that should suffice for differentiation to be evident (J. Elhai, T. Black, A. Ernst and C. P. Wolk, unpublished observations). The two phenotypes, fragmentation and failure to differentiate, may both be due to the presence of aberrant intercellular junctions that preclude normal intercellular communication. Fragmenting mutant 522 of Anabaena sp. strain PCC 7120 is incapable of aerobic fixation of dinitrogen; the gene that complements the mutation has been identified and sequenced (Buikema and Haselkorn, 1993); the product of its translation resembles no known protein. The isolation ofmutants that remain intact, but show little or no sign of differentiation, even after protracted deprivation for fixed nitrogen, has often been reported (Singh et al., 1972; Wilcox et al., 1975; Currier et al., 1977; Spence and Stewart, 1987; Buikema and Haselkorn, 1991a; Ernst et al., 1992). In such cases, it is unclear whether a spacing pattern is absent, or has only not been made manifest.

1. Failure to Differentiate

2. Differentiation Only at the Termini of Filaments

Studies involving fragmentation of filaments (Wolk, 1967; Wolk and Quine, 1975) or surgical ablation of specific cells (Wilcox et al., 1973b) suggest that a minimum number of adjacent vegetative cells may be required for heterocyst differentiation to proceed. In fact, regression of proheterocysts was observed in small fragments originally containing a proheterocyst (Wilcox et al., 1973b). The molecular basis of a minimal filament length to stabilize heterocyst formation is not known, but the phenomenon suggests that there could be a nutritional requirement for differentiation, that cells that might differentiate are able to measure the availability ofa required nutrient, and that provision of such a nutrient (or nutrients) might enable differentiation when it would not otherwise occur. Alternatively, perhaps a minimum number of vegetative cells is required to inactivate a morphogenetically active substance that inhibits development, in order that such a substance not accumulate within a presumptive heterocyst (see sections V, C–E). Mutants that fragment and fail to differentiate

Strains of Anabaena sp. and Nostoc sp. form intercalary heterocysts that are connected to the vegetative cells to either side by means of pore structures (Section III A, 5). Some strains develop terminal heterocysts, that possess only one terminal pore, either occasionally as in Anabaena sp. strain PCC 7120 and Anabaena variabilis, or regularly, as in Anabaena siamensis, a strain that grows in short filaments terminated by two heterocysts (Antarikanonda and Lorenz, 1982). Anabaena siamensis appears to fulfill the classical description of an Anabaenopsis sp.(Geitler, 1932). Species of Cylindrospermum sp. naturally form long filaments that have terminal heterocysts and lack intercalary heterocysts (Geitler, 1932). In Anabaenopsis sp. and certain strains of Cylindrospermum sp., instead of single cells differentiating as the norm, pairs of contiguous cells differentiate simultaneously, head to head. Each cell of the pair differentiates as a onepored (terminal) heterocyst; the two then separate and interrupt the filament. In Cylindrospermum licheniforme strain ATCC 29412, on the other hand,

Chapter 27 Heterocyst Metabolism and Development a single cell distant from, and approximately bisecting the interval between, two heterocysts dies, whereupon the adjacent cells differentiate into terminal heterocysts. In both of these situations, it is therefore normal for filaments to have only one-pored heterocysts, one at each end. The tapering filaments of members ofthe genera Gloeotrichia and Rivularia bear only a single, one-pored heterocyst at the broader end. Germlings of akinetes of Anabaena sp. normally form one terminal heterocyst and then, after further growth, form a second terminal heterocyst at the other end, before progressing to the normal pattern of differentiation of intercalary heterocysts (Fay et al., 1968; Sutherland et al., 1979; Skill and Smith, 1987). Mutants of Anabaena cylindrica (F) and Anabaena sp. strain PCC 7120 have been reported that no longer form intercalary heterocysts but do form terminal heterocysts on long filaments (Wilcox et al., 1975; Liang et al., 1992). Because of their low ratio of heterocysts to vegetative cells, mutant strains of this type grow slowly on dinitrogen. Revertants and pseudorevertants of the Anabaena cylindrica mutant were also reported (Wilcox et al., 1975); in addition to regaining the ability to form intercalary heterocysts, the pseudorevertants frequently produced pairs of adjacent heterocysts. The Anabaena sp. strain PCC 7120 gene, denoted patA, whose mutant form was responsible for the phenotype, was cloned by complementation of its point mutation, and sequenced (Liang et al., 1992). The interchangeability between the wild-type form and the phenotype of a patA mutant suggests that the mechanisms underlying the two morphologies do not differ radically. It is possible that the biochemical difference between the normal and patA phenotypes relates to a difference in the mobility of (i.e., the permeability to) a morphogenetically active substance along the filament, or to the stability (or turnover) of such a substance (see sections V, C and V, E). Thus, analysis of the patA mutant and of other possible mutants with similar phenotypes may help in the identification of a putative morphogenetically active substance. Such interpretations would appear more likely if the gene analyzed proves to be active throughout the filament.

3. Formation of Strings of Heterocysts Strings of heterocysts have been observed in some strains. Contiguous heterocysts occur naturally in

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Mastigocladus laminosus (Nierzwicki-Bauer et al., 1984a; Stevens et al., 1985). Strings of proheterocysts are also observed in filaments of Anabaena cylindrica (F) when nitrogen deprivation is imposed on an old culture, although only one member of the string normally proceeds to maturity (Wilcox et al., 1973a). Brief exposure to light of high intensity just before or after nitrogen-stepdown also increases the number of paired heterocysts that form (Adams and Carr, 1981b). Intercalary heterocysts, induced by nitrogen stepdown, often occur as multiples in Cylindrospermum licheniforme, which otherwise forms only terminal heterocysts (Wolk and Quine, 1975). Wild-type strains that form no strings of heterocysts can give rise to mutants that do form such strings (Wolk, 1982), and a similar phenotype is seen by secondary mutation ofmutant strains that make only terminal heterocysts (Wilcox et al., 1975) or no heterocysts at all (Ernst et al., 1992). Entire filaments of certain mutants can transform into heterocysts by sequential differentiation and detaching of cells seriatim along the filament (e.g., mutant NF12 of Anabaena variabilis: Currier et al., 1977; these heterocysts are nearly all one-pored). Perhaps the most informative case in which multiple heterocysts occur under conditions is that of Anabaena sp. strain PCC 7120 carrying extra copies ofhetR on a derivative of plasmid pDU1 (Buikema and Haselkorn, 1991b). Heterocysts are produced by such a strain even when it is grown on nitrate. An intact copy of hetR is required to see the effect, thus excluding the possibility that a regulatory region of the gene is titrating an inhibitor of heterocyst differentiation. Like the wild-type form of hetR, the wild-type form of hetP (Section IV D) gives rise to strings (clusters) of heterocysts when it, together with contiguous DNA, is present in supernumerary copies in wild type Anabaena sp. (Fernández-Piñas et al., 1994). However, hetP is activated hours later than hetR and is not autoregulatory; and unlike the case with hetR, an insertional mutation in hetP has no effect on akinete formation (see section VI). The close proximity of hetC and hetP, and the fact that transposon mutants of both are suggests that the two genes may be functionally related. Sequencing of hetC (Khudyakov and Wolk, 1994) has shown that the product ofits translation is similar throughout a 2-kbp region of the gene to bacterial ATP-binding cassette transporters that are involved in the export of proteins (Fath and Kolter, 1993).

806 The identity and role of the protein presumptively transported by the hetC product are clearly of great importance for pattern formation in Anabaena sp. One speculative possibility is that the substrate protein is, or regulates the intercellular movement of, an inhibitor of differentiation (see Section V C). A mutation that suppresses the phenotype of mutant N10 (hetN) leads to the formation of grouped (clustered) heterocysts. Whereas attempts to reconstruct mutants (hetR), P6 and C3 consistentlyconferred the original phenotype(Black and Wolk, unpublished results; Fernández-Piñas et al., 1994; Khudyakov and Wolk, 1994), attempts to reconstruct theN10 mutation with various insertions have yielded, in addition to strains that are strains thatproduce multiple contiguous heterocysts! This variability may be due to transcriptional interactions between the hetN gene and an open reading frame that is transcribed on the opposite strand from the hetN gene and that ends just 42 bp away from the 3' end of the latter. The gene hetN and contiguous genes show strong homology to NAD(P)H-dependent oxidoreductases that are involved in the biosyntheses of fatty acids, Nod factor, and polyketides, leading to the suggestion (Black and Wolk, 1994) that a secondary metabolite may be involved in the regulation of the pattern of heterocyst spacing. Treatments with certain compounds have been described that produce strings ofadjacent heterocysts in filaments growing on dinitrogen: rifampicin (an inhibitor of RNA polymerase; Wolk and Quine, 1975), at a low concentration; mitomycin (an inhibitor of DNA replication; Tyagi, 1974); 7-azatryptophan (Mitchison and Wilcox, 1973; Wolk, 1982; Chen et al., 1987a; Adams, 1992b); and a few other amino acid analogs (Wolk, 1982). Rifampicin and 7-azatryptophan may also produce heterocysts in the presence of normally inhibitory concentrations of fixed nitrogen (Dikshit et al., 1979; Chen et al., 1987a). The action of 7-azatryptophan may be due to a general starvation for amino acids owing to inhibition of glutamate synthase (Chen et al., 1987a,b) or to synthesis of defective protein (Bottomley et al., 1980), among other possibilities. The fact that Anabaena sp. strain 1F showed no such response to 7-azatryptophan (Chen et al., 1987b), illustrates the potential for considerable variation between strains with regard to their responses to these effectors. Formation of strings of heterocysts also follows

C. Peter Wolk, Anneliese Ernst & Jeff Elhai from (i) exposure of Anabaena variabilis to 1 mM cyclic AMP (cAMP; Smith and Ownby, 1981), that being a concentration attained by the cells of a different strain of Anabaena sp. within 1 h of transfer to nitrogen-deficient medium (Francko and Wetzel, 1981); (ii) growth ofcertain strains that are resistant to amino acid analogs and release amino acids into the extracellular milieu (Kerby et al., 1987); (iii) incubation with a high-molecular-weight fraction of Difco neo-peptone B118 (Sharma, 1984); and (iv) certain mutations, for example in mutant M7 of Wilcox et al. (1975). In short, a wide diversity of effectors induces string formation. It appears that het genes and genes whose mutated forms give rise to an M7-like phenotype both regulate the formation of strings of heterocysts, but with opposing tendencies. That is, the former stimulate, and the latter suppress, that phenotype. There seems to be a subtle balance between the differentiation of isolated and clustered heterocysts, fairly easily tipped toward the formation of strings of heterocysts.

4. Phenotypes Not Reported No strain of Anabaena sp. or Nostoc sp. has, to our knowledge, been reported to exhibit under any condition a random distribution of heterocysts. Therefore, the non-randomness of heterocyst spacing may not be readily perturbed. Canavanine, an analog of arginine, was reported (i) to block heterocyst differentiation totally, perhaps owing to its toxic effect on growth (Kumar and Kumar, 1981); and (ii) to induce random formation of akinetes (Nichols et al., 1980). Since patterns of differentiation of akinetes and heterocysts are closely linked (Geitler, 1925, p. 199), this highly unusual absence of pattern of akinetes commends a study of the effect of lower concentrations of canavanine on heterocyst pattern formation. Although short strings of heterocysts have been observed to form simultaneously, and all cells of a filament to differentiate sequentially (Section V B, 3), mutants have not been found in which whole filaments differentiate simultaneously. However, envelope deposition involves nearly a doubling of cell mass (Dunn and Wolk, 1970) despite a cessation of photosynthesis, so that a search for such mutants, which could be highly useful experimentally, should involve efforts to predict and provide requisite nutrition for the differentiation process.

Chapter 27 Heterocyst Metabolism and Development

C. Models of Pattern Formation A priori, two extreme types of mechanism might explain the development and maintenance of pattern in filaments of heterocyst-forming cyanobacteria. On the one hand, the positions of heterocysts might be determined solely by cell lineage. An influence of lineage on differentiation in Anabaena sp. is illustrated by the observation that when cells of Anabaena catenula are deprived of nitrogen, cell division becomes asymmetric, and in this case, heterocysts develop only from the smaller ofthe two daughter cells (Mitchison and Wilcox, 1972). Cell lineage cannot be the sole determining factor, because the positions at which heterocysts appear can be manipulated experimentally, by fragmentation of the filaments (Wolk, 1967; Wolk and Quine, 1975), or by selectively breaking filaments at sites at which differentiation is being initiated (Wilcox et al., 1973a). Upon fragmentation of filaments, most fragments will initiate differentiation of a heterocyst by the time that they are four cells in length, and virtually all by the time that they are five cells in length (Wolk, 1967; van de Water and Simon, 1982; Jüttner, 1983), unless a heterocyst is already present or developing (Wolk, 1967; Wolk and Quine, 1975). An alternative extreme is that the spacing between heterocysts might be governed completely dynamically, for example by interactions of diffusible substances that inhibit or promote differentiation in a concentration-dependent manner (Haselkorn, 1978; Wolk, 1989). In this context, the possibility that intercellular communication takes the form of electrical potentials that are transmitted along the filaments (Chailakhyan et al., 1982; Häder, 1987) has not been evaluated. Electrical signals could provide a means for one cell to determine a kind of average metabolic vigor of the other cells in its filament. It is possible that the true mechanism of pattern formation lies somewhere between the two extremes noted. That is, a provisional pattern may be laid down as a consequence of cell-to-cell differences in nutrition, position in the cell cycle, size, or some other parameter, and then subsequently modified by interactions between cells. The leech, Theromyzon rude, for instance, is able to modify a provisional pattern determined by lineage in response to positional cues (Keleher and Stent, 1990; see also Horvitz and Sternberg, 1991). Two models to explain spacing patterns have been

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devised that rely on cell-cell interactions by diffusible substances (Wolk, 1989, 1991; Fig. 14). One, the ‘altruistic’ model (Fig. 14A), envisions that cells that reach a threshold level of starvation initiate the process of differentiation, and simultaneously release a diffusible, differentiation-inhibiting substance. This substance could be a nitrogenous substance from a store unavailable to uncommitted cells, or a purely informational molecule. The second, ‘selfish’ model (Fig. 14B; see Wolk, 1989) calls on cells in the filament to respond to nutrientdeprivation in a manner typical of bacteria and, in some cases, cyanobacteria (Thiel, 1988 [deprivation for phosphate]; Jeanjean and Broda, 1977 [deprivation for sulfate]), i.e., by inducing uptake systems of high affinity for the scarce nutrient. This effect might also propagate along a filament, as cells successively become depleted of nitrogenous metabolites by a nitrogen-

808 deprived neighbor. The model posits that cells from which nutrients are being pumped to vegetative cells on either side become especially nitrogen-poor, and are thereupon activated to become heterocysts. Highaffinity amino acid uptake systems in cyanobacteria have been reported (Flores and Muro-Pastor, 1988; Herrero and Flores, 1990; Xu and McAuley, 1990). The idea, inherent in the first model, that inhibitory fields emanate from differentiating cells, has greatly influenced the interpretation of experimental results (Wolk, 1967; Wilcox et al., 1973a,b; Wolk and Quine, 1975; Mitchison et al., 1976; Buikema and Haselkorn, 1991b). The second model illustrates that the available experimental results are equally consistent with the view that the cells that differentiate are not the first that sense nitrogen-deprivation. Much progress has been made in the development of mathematical models of de novo formation of biological patterns, such as segmentation in early embryogenesis of Drosophila melanogaster (Meinhardt, 1986), that are applicable to analyses of lateral inhibition in a filamentous organism (Turing, 1952; Gierer and Meinhardt, 1972; Meinhardt, 1982, 1986; Oster and Murray, 1989; see also Wilcox et al., 1973a). The models require two critical components: (1) an inhibitor and (2) an activator. The inhibitor must diffuse rapidly and inhibit the function of the activator. The activator must diffuse much more slowly if at all, and must increase both its own production (i.e., it is autoregulated) and the production of the inhibitory substance. These simple assumptions can lead to the prediction ofa transition from random fluctuations in an otherwise homogeneous initial state to a combination of localized activation of the activator and stable gradients of the diffusible inhibitor. The general idea is thatpositive fluctuations in production of the activator lead both to further production of the activator, which remains localized, and to production of the inhibitor, which diffuses rapidly to adjacent cells and inhibits them from inducing the activator. As a consequence, secondary induction can occur only at points distant from the site of initial induction. The diffusible inhibitor might be identified as a nitrogenous substance released by committed cells, as suggested for the ‘altruistic’ model in the preceding paragraph and in Fig. 14A, or purely symbolic molecule (see Section V E). The idea that HetR may serve as the requisite autoregulated activator is discussed below. Alternatively, the process described in this paragraph

C. Peter Wolk, Anneliese Ernst & Jeff Elhai could take place after the ‘selfish’ process of pattern formation, if both occur.

D. HetR, Part 2: The Role in Pattern Formation Wilcox et al. (1973a,b) showed convincingly (i) that at least in certain instances, it is not an isolated cell, but instead a string (or group or cluster) of cells that initiates differentiation; and (ii) that one cell of the string completes differentiation while the others revert to a vegetative state, presumably as a result of some kind of competition. However, if a differentiating cell produces an inhibitor that deters the differentiation of adjacent cells, why does the cell not inhibit its own differentiation? This question has been of central importance to understanding of cyanobacterial pattern formation from the outset. Recent results, particularly regarding hetR, justify a speculative but concrete answer to this question— one that fits well with general models of lateral inhibition (discussed above) and with a model put forth by Wilcox et al. (1973a). From the ability of HetR to act as an autocatalytic activator (Black et al., 1993; Section IV D), one would predict that a plasmid -borne hetR gene acts by increasing the level of HetR sufficiently in some cells to ‘flip a developmental switch.’ If HetR is equated with the autoregulated activator integral to the activator/inhibitor model described in the previous section, then pattern formation as well as differentiation would depend on its function. On the other hand, if HetR is a switch that responds passively to a preexistent pattern, then it should be possible to discern either that pattern, or a patterned response to it, in hetR mutants. Either way, the autoregulatory properties of the hetR gene, and the model, provide a framework for understanding the absence of self-inhibition by differentiating cells. How the high activity of hetR in differentiating cells would regulate competition within strings is unclear, although one possibility is that abundantly produced HetR might in some way act counter to an inhibitor (e.g., compete with the inhibitor, or compete with a protein that senses the level of inhibitor). Not only developing heterocysts within strings, but also solitary developing heterocysts in filaments that are exposed to combined nitrogen, can regress (Bradley and Carr, 1977; Adams and Carr, 1981a, 1989; and see also Wilcox et al., 1973b); it will be of great interest to learn whether experimental manipulation of the rate of transcription

Chapter 27 Heterocyst Metabolism and Development of hetR can control regression. It should be kept in mind that hetR might be centrally (and negatively) involved in the resolution of strings to single differentiating cells, and yet uninvolved, or involved only peripherally, in generation of the primary pattern.

E. Possible Signals of Positional Information The models described above predict the existence of an inhibitory signal that passes from cell to cell. The critical signal may be an intrinsic characteristic of starvation (e.g., scarcity of an amino acid), or a purely symbolic molecule (e.g., ppGpp). Many responses are elicited by nitrogen deprivation; the task is to determine which are, and which are not, intimately related to the process of pattern formation. If the signal is an amino acid or derivative, it is likely to originate from one of the two macromolecular reserves that provide nitrogen to filaments during deprivation: phycobiliproteins and cyanophycin. As fixed nitrogen in the medium is gradually used, the level of cyanophycin in a filament increases dramatically (Mackerras et al., 1990). When the external nitrogen source has been completely exhausted, the store of cyanophycin is consumed. The enzymes, cyanophycin synthetase and cyanophycinase, that make and degrade the polymer, respectively, have a much higher level of activity in heterocysts than in vegetative cells (Gupta and Carr, 1981 c), and measurements from filaments that have proheterocysts but lack heterocysts are consistent with the view that these enzymatic activities are also high in differentiating cells. These observations fit in well with the diffusibleinhibitor model presented above (Fig. 14A) and with the conjecture (Section III D, 1) that aspartate, one of the two amino acid constituents of cyanophycin, may be of importance in intercellular transfer of derived fixed nitrogen. The first cell that reaches a critical stage of nitrogen starvation would be expected to switch over from cyanophycin accumulation to cyanophycin degradation. The diffusion of the resulting arginine and aspartate (or of other derivative compounds) may prevent adjacent cells from activating their own catabolism of cyanophycin. The sites of secondary heterocyst formation would therefore be distant from the first cells to have begun breaking down cyanophycin. Whether or not cyanophycin plays a central role in nitrogen flow through differentiating filaments, arginine is not likely to be the diffusible inhibitor

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called for by some of the models discussed above, because heterocyst formation is not inhibited by increases in the intracellular concentration of arginine to a level higher than that measured in ammoniumgrown filaments (Thiel and Leone, 1986). Glutamine was once proposed as a possible diffusible inhibitor (Thomas et al., 1977), but this possibility is suspect for similar reasons (Thiel and Leone, 1986; Chen et al., 1987a). It must be borne in mind, however, that the amino acid pool measurements that argue against specific nitrogenous substances as diffusible inhibitors are average values. It is conceivable that the concentrations of these substances in particular cells in the filament may differ greatly from the average. It also remains possible that it is not the concentration of any individual amino acid (or of amino acids in general) that inhibits heterocyst differentiation but rather a ratio of an amino acid, representing the N-status of the cell, and a carboxylic acid, representing the C-status of the cell, that is decisive, as it is in the ntr system of enteric bacteria (Magasanik and Neidhardt, 1987). Signal transduction commonly proceeds through small, purely symbolic molecules, such as cAMP and ppGpp, even in procaryotes (Botsford and Harman, 1992; Cashel and Rudd, 1987). In that light, it is interesting that the level of cAMP rises markedly, albeit slowly, in Anabaena sp. in response to nitrogen starvation (Hood et al., 1979; Francko and Wetzel, 1981), caused perhaps by a decrease in the activity of cAMP phosphodiesterase (Ownby and Kuenzi, 1982), and that exogenous cAMP induces aberrant spacing of heterocysts (Smith and Ownby, 1981). As in eucaryotes, the production of cAMP by adenylate cyclase is controlled by very low levels of calcium and is greatly stimulated by a calmodulin from Anabaena sp. (Bianchini et al., 1990). Calmodulin was localized by immunoelectron microscopy in both heterocysts and vegetative cells (Pettersson and Bergman, 1989), but with no role ascribed to it in either type of cell. While the well known role of cAMP in patterned development of Dictyostelium discoideum (Kimmel and Firtel, 1991) may invite speculation on a morphogenetic role in Anabaena sp., other hypotheses about the role of cAMP, such as that it monitors the energy-state of the cell, are no less tenable (Ohmori et al., 1988; Ohmori, 1989; Onek and Smith, 1992). The great progress made in the genetic analysis of Anabaena sp. should not prevent one from recognizing that genetic activity is an intracellular

810 phenomenon, whereas spacing, like nutritional cooperation, depends on intercellular interactions. Neither a genetic nor a biochemical approach alone is likely to solve the problem ofhow pattern emerges. The challenge is how to exploit the powerful genetic tools now available to aid in the biochemical analysis of a signal transduction system that is spatially inhomogeneous.

VI. Relationship of Diverse Differentiation Processes in Cyanobacteria Several observations support the hypothesis (Geitler, 1925; Wolk, 1964) that the differentiation of heterocysts and the differentiation of akinetes are related processes: (1) the envelopes of akinetes (spores) of Anabaena cylindrica (W) and Anabaena variabilis have the same polysaccharide that is characteristic of the envelopes of the heterocysts of the same species; (2) akinetes and heterocysts are both non-dividing cells; and (3) in certain taxa, akinetes often or occasionally form where a heterocyst would be expected to form (Fig. 15). Because, in Nostoc ellipsosporum, a hetR mutation blocks the differentiation ofboth heterocysts and akinetes at an early stage (Section IV D), the two types of differentiation appear to share an early step or regulatory process. Concordantly, introduction of a

C. Peter Wolk, Anneliese Ernst & Jeff Elhai hepA mutation (which affects heterocyst envelope formation in Anabaena sp. strain PCC 7120) into Anabaena variabilis, disrupted the formation of the envelope of both heterocysts and akinetes in the latter strain (F. Leganés and C. P. Wolk, unpublished results). In contrast, a hetP insertional mutation in Nostoc ellipsosporum blocks heterocyst formation but not the formation of akinetes (Leganés et al., 1994). Therefore, an insertional mutation in hetP appears to affect heterocyst formation after the two pathways of differentiation diverge. Because it seems that the need to sporulate would have preceded, in evolutionary time, the need to fix dinitrogenaerobically, heterocystdifferentiation may have built upon the regulation and/or metabolism underlying akinete formation, rather than vice versa. Sequences of cells can differentiate into akinetes, which are (by definition) non-motile, and also into hormogonia, a motile cell stage that excludes heterocyst formation (Geitler, 1932; Rippka et al., 1979, defined hormogonia differently). These alternative life styles of cyanobacteria remain very little explored (see Nichols and Adams, 1982; Damerval et al., 1991; also see Chapter 28). It is anticipated that their study, facilitated by the recent advances in cyanobacterial genetics, will also help to illuminate the processes underlying heterocyst differentiation.

Chapter 27 Heterocyst Metabolism and Development Acknowledgments A. E. is a member of the Sonderforschungsbereich 248 of the University of Konstanz and supported by DFG grant Er158/1-1. Work from the Wolk laboratory is supported by the U.S. Department of Energy under grant DE-FG02-90ER20021 and by NSF grant IBN 9118152.

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820 Acta 634: 237–248 Peterson RB, Shaw ER, Dolan E and Ke B (1981b) A photochemically active heterocyst preparation from Anabaena variabilis. Photobiochem Photobiophys 2: 79–84 Pettersson A and Bergman B (1989) Calmodulin in heterocystous cyanobacteria: biochemical and immunological evidence. FEMS Microbiol Lett 60: 95–100 Pienkos PT, Bodmer S andTabita FR (1983) Oxygen inactivation and recovery of nitrogenase activity in cyanobacteria. J Bacteriol 153: 182–190 Pistorius EK. and Gau AE (1986) A possible model for the organic prosthetic group of the water-splitting complex in the cyanobacterium Anacystis nidulans. FEBS Lett 206: 243–248 Potts M, Angeloni SV, Ebel RE and Bassam D (1992) Myoglobin in a cyanobacterium. Science 256: 1690–1692 Privalle LS (1984) Effects of D-erythrose on nitrogenase activity in whole filaments of Anabaena sp. strain 7120. J Bacteriol 160: 794–796 Privalle LS and Burris RH (1984) D-Erythrose supports nitrogenase activity in isolated Anabaena sp. strain 7120 heterocysts. J Bacteriol 157: 350–356 Ptashne M (1992) A genetic switch (2nd edition). Blackwell Scientific Publishers, Oxford Rai AN and Bergman B (1986) Modification of metabolism in heterocysts of the cyanobacterium Anabaena 7120 (ATCC27893). FEMS Microbiol Lett 36:133–137 Rai AN, Rowell P and Stewart WDP (1982) Glutamate synthase activity of heterocysts and vegetative cells of the cyanobacterium Anabaena variabilis Kütz. J Gen Microbiol 128: 2203–2205 Rai AN, Borthakur M, Singh S and Bergman B (1989) AnthocerosNostoc symbiosis: immunoelectronmicroscopic localization of nitrogenase, glutamine synthetase, phycoerythrin and ribulose-l,5-bisphosphate carboxylase/oxygenase in the cyanobiont and the cultured (free-living) isolate Nostoc 7801. J Gen Microbiol 135: 385–395 Ramasubramanian TS, Wei T-F and Golden JW (1991) Sequence specific DNA-bindingfactorsfrom vegetativecells ofAnabaena PCC 7120. Abstr 195A, Abstr 7th Intl Symp Photosynthetic Prokaryotes, Amherst, MA Reich S and Böger P (1989) Regulation ofnitrogenase activity in Anabaena variabilis by modification of the Fe protein. FEMS Microbiol Lett 58: 81–86 Reich S, Almon H and Böger P (1986) Short-term effect of ammonia on nitrogenase activity of Anabaena variabilis (ATCC29413). FEMS Microbiol Lett 34: 53–56 Reich S, Almon H and Böger P (1987) Comparing short-term effects of ammonia and methylamine on nitrogenase activity in Anabaena variabilis (ATCC 29413). Z Naturforsch 42c: 902–906 Renström-Kellner E, Rai AN and Bergman B (1990) Correlation between nitrogenase and glutamine synthetase expression in the cyanobacterium Anabaena cylindrica. Physiol Plant 80: 12–19 Rippka R and Stanier RY (1978) The effects of anaerobiosis on nitrogenase synthesis and heterocyst development by Nostocacean cyanobacteria. J Gen Microbiol 105: 83–94 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 Robinson SJ and Haselkorn R (1985) Abstr 5th Intl Symp

C. Peter Wolk, Anneliese Ernst & Jeff Elhai Photosynthetic Prokaryotes, p 246, Grindelwald, Switzerland Roussard-Jacquemin M (1983) Étude ultrastructurale de la différentiation des hétérocysts chez la cyanobactérie, Anabaena cylindrica Lemm. Can J Microbiol 29: 1564–1575 Rowell P, Diez J, Apte SK and Stewart WDP (1981) Molecular heterogeneity offerredoxin: oxidoreductase from the cyanobacterium Anabaena cylindrica. Biochim Biophys Acta 657: 507–516 Ruvkun GB and Ausubel FM (1980) Interspecies homology of nitrogenase genes. Proc Natl Acad Sci USA 77: 191–195 Sandmann G, Peleato ML, Fillat MF, Lázaro MC and GómezMoreno C (1990) Consequences of the iron-dependent formation offerredoxin and flavodoxin on photosynthesis and nitrogen fixation on Anabaena strains. Photosynth Res 26: 119–125 Saville B, Straus N and Coleman JR (1987) Contiguous organization of nitrogenase genes in a heterocystous cyanobacterium. Plant Physiol 85: 26–29 Scanlan DJ, Newman J, Sebaihia M, Mann NH and Cart NG (1992) Cloning and sequence analysis of the glucose-6phosphate dehydrogenase gene from the cyanobacterium Synechococcus PCC 7942. Plant Mol Biol 19: 877–880 Scherer S and Böger P (1982) Respiration of blue-green algae in the light. Arch Microbiol 132: 329–332 Scherer S, Stürzl E and Böger P (1982) Interaction of respiratory and photosynthetic electron transport in Anabaena variabilis Kütz. Arch Microbiol 132: 333–337 Schilling N and Ehrnsperger K (1985) Cellular differentiation of sucrose metabolism in Anabaena variabilis. Z Naturforsch 40c: 776–779 Schluchter WM and Bryant DA (1992) Molecularcharacterization 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 Schmitz O, Kentemich T, Zimmer W, Hundeshagen B and Bothe H (1993a) Identification of the nifJ gene coding for pyruvate:ferredoxin oxidoreductase in dinitrogen-fixing cyanobacteria. Arch Microbiol 160: 62–67 Schmitz S, Schrautemeier B and Boehme H (1993b) Evidence from directed mutagenesis that positively charged amino acids are necessary for interaction of nitrogenase with the [2Fe-2S] heterocyst ferredoxin (FdxH) from the cyanobacterium Anabaena sp. PCC 7120. Mol Gen Genet 240: 455–460 Schneider GJ and Haselkorn R (1988) RNA polymerase subunit homology among cyanobacteria, other eubacteria, and archaebacteria. J Bacteriol 170: 4136–4140 Schneider GJ, Tumer NE, Richaud C, Borbely G and Haselkorn R (1987) Purification and characterization ofRNA polymerase from the cyanobacterium Anabaena 7120. J Biol Chem 262: 14633–14639 Schneider GJ, Lang JD and Haselkorn R (1991) Promoter recognition by the RNA polymurase from vegetative cells of the cyanobacterium Anabaena 7120. Gene 105: 51–60 Schrautemeier B and Böhme H (1984) Different functions assigned to NAD(H) and NADP(H) in light-dependent nitrogen fixation by heterocysts of Anabaena variabilis. FEMS Microbiol Lett 25: 215–218 Schrautemeier B and Böhme H (1985) A distinct ferredoxin for nitrogen fixation isolated from heterocysts of the cyanobacterium Anabaena variabilis. FEBS Lett 184: 304–308 Schrautemeier B and Böhme H (1992) Coding sequence of a

Chapter 27 Heterocyst Metabolism and Development heterocyst ferredoxin gene (fdxH) isolated from the nitrogenfixing cyanobacterium Calothrix sp. PCC 7601. Plant Mol Biol 18: 1005–1006 Schrautemeier B, Böhme H and Böger P (1984) In vitro studies on pathways and regulation of electron transport to nitrogenase with a cell-free extract from heterocysts of Anabaena variabilis. Arch Microbiol 137: 14–20 Schrautemeier B, Böhme H and Böger P (1985) Reconstitution of a light-dependent nitrogen-fixing and transhydrogenase system with heterocyst thylakoids. Biochim Biophys Acta 807: 147–154 Serrano A, Rivas J and Losada M (1984) Studies on glutathione reductase activity in the cyanobacterium Anabaena sp. strain 7119. Adv Photosynth Res 2: 711–714 Serrano A, Giménez P, Scherer S and Böger P (1990) Cellular localization of cytochrome in the cyanobacterium Anabaena variabilis. Arch Microbiol 154: 614–618 Sharma P (1984) Heterocyst and akinete induction with altered pattern in Anabaena cylindrica, caused by neo-peptone. Arch Microbiol 139: 196–201 Simon RD (1971) Cyanophycin granules from the blue-green alga Anabaena cylindrica: a reserve material consisting of copolymers of aspartic acid and arginine. Proc Natl Acad Sci USA 68: 265–267 Simpson FB and Burris RH (1984) A nitrogen pressure of 50 atmospheres does not prevent evolution of hydrogen by nitrogenase. Science 224: 1095–1097 Singh RN, Singh SP and Singh PK (1972) Genetic regulation of nitrogen fixation in blue-green algae. In: Desikachary TV (ed) Taxonomy and Biology of Blue-Green Algae, pp 264–268. University of Madras, India Skill SC and Smith RJ (1987) Synchronous akinete germination and heterocyst differentiation in Anabaena PCC 7937 and Nostoc PCC 6720. J Gen Microbiol 133: 299–303 Smith DL, Patriquin DG, Dijak M and Curry GM (1986) The effect of light-dependent oxygen consumption on nitrogenase activity in Anabaena cylindrica. Can J Bot 64: 1843–1848 Smith G and Ownby JD (1981) Cyclic AMP interferes with pattern formation in the cyanobacterium Anabaena variabilis. FEMS Microbiol Lett 11: 175–180 Smith RL, Kumar D, Zhang X, Tabita FR and Van Baalen C (1985) and metabolism by isolated heterocysts from Anabaena sp. strain CA. J Bacteriol 162: 565–570 Smith RL, Van Baalen C and Tabita FR (1987) Alteration ofthe Fe protein of nitrogenase by oxygen in the cyanobacterium Anabaena sp. strain CA. J Bacteriol 169: 2537–2542 Smith RL, Van Baalen C and Tabita FR (1988) Isolation of metabolically active heterocysts from cyanobacteria. Meth Enzymol 167: 490–495 Sobczyk A, Schyns G, Tandeau de Marsac N and Houmard J (1993) Transduction of the light signal during complementary chromatic adaptation in the cyanobacterium Calothrix sp. strain PCC 7601: DNA binding proteins and modulation by phosphorylation. EMBO J 12: 997–1004 Soriente A, Sodano G, Gambacorta A and Trincone A (1992) Structure of the ‘heterocyst glycolipids’ of the marine cyanobacterium Nodularia harveyana. Tetrahedron 48: 5375– 5384 Soriente A, Gambacorta A, Trincone A, Sili C, Vincenzini M and Sodano G (1993) Heterocyst glycolipids of the cyanobacterium Cyanospira rippkae. Phytochem 33: 393–396

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Spence DW and Stewart WDP (1987) Heterocystless mutants of Anabaena PCC7120withnitrogenaseactivity.FEMS Microbiol Lett 40: 119–122 Spiller H, Ernst A, Kerfin W and Böger P (1978) Increase and stabilization of photoproduction of hydrogen in Nostoc muscorum. Z Naturforsch 33c: 541–547 Stal LJ and Bergman B (1990) Immunological characterization of nitrogenase in the filamentous, non-heterocystous cyanobacterium Oscillatoria limosa. Planta. 182: 2887–2891 Stanier RY and Cohen-Bazire G (1977) Phototrophic prokaryotes: the cyanobacteria. Ann Rev Microbiol 31: 225–274 Stanier RY, Kunisawa R, Mandel M and Cohen-Bazire G (1971) Purification and properties of unicellular blue-green algae (order Chroococcales). Bacteriol Rev 35: 171–205 Stevens SE, Nierzwicki-Bauer SA and Balkwill DL (1985) Effect of nitrogen starvation on the morphology and ultrastructure of the cyanobacterium Mastigocladus laminosus. J Bacteriol 161: 1215–1218 Stewart WDP and Lex M (1970) Nitrogenase activity in the bluegreen alga Plectonema boryanum strain 594. Arch Mikrobiol 73:250–260 Stewart WDP, Haystead A and Dharmawardene MWN (1975) Nitrogen assimilation and metabolism in blue-green algae. In: Stewart WDP (ed) Nitrogen Fixation by Free-Living Microorganisms, pp 129–158. Cambridge Univ Press, Cambridge Stock JB, Stock AM and Mottonen JM (1990) Signal transduction in bacteria. Nature 344: 395–400 Strauch MA and Hoch JA (1992) Sporulation in prokaryotes and lower eukaryotes. Curr Opin Genet Devel 2: 799–804 Subramanian G and Shanmugasundaram S (1986) Uninduced ammonia release by the nitrogen-fixing cyanobacterium Anabaena. FEMS Microbiol Lett 37: 151–154 Sun D, Cabrera-Martinez RM and Setlow P (1991) Control of transcription of the Bacillus subtilis spoIIIG gene, which codes for the forespore-specific transcription factor J Bacteriol 173: 2977–2984 Sutherland JM, Herdman M and Stewart WDP (1979) Akinetes of the cyanobacterium Nostoc PCC 7524: macromolecular composition, structure and control of differentiation. J Gen Microbiol 115: 273–287 Sutherland JM, Stewart WDP and Herdman M (1985) Akinetes of the cyanobacterium Nostoc PCC 7524: morphological changes during synchronous germination. Arch Microbiol 142: 269–274 Tanaka K, Masuda S and Takahashi H (1992) Multiple rpoD-related genes of cyanobacteria. Biosci Biotech Biochem 56: 1113-1117 Tandeau de Marsac N and Houmard J (1993) Adaptation of cyanobacteria to environmental stimuli: new steps towards molecular mechanisms. FEMS Microbiol Rev 104: 119–190 Tel-Or E, Huflejt ME and Packer L (1986) Hydroperoxide metabolism in cyanobacteria. Arch Biochem Biophys 246: 396–402 Thiel T (1988) Phosphate transport and arsenate resistance in the cyanobacterium Anabaena variabilis. J Bacteriol 170: 1143– 1147 Thiel T (1990) Protein turnover and heterocyst differentiation in the cyanobacterium Anabaena variabilis. J Phycol 26: 50–54 Thiel T (1993) Characterization of genes for an alternative nitrogenase in the cyanobacterium Anabaena variabilis. J

822 Bacteriol 175: 6276–6286 Thiel T and Leone M (1986) Effect of glutamine on growth and heterocyst differentiation in the cyanobacterium Anabaena variabilis. J Bacteriol 168: 769–774 Thiel T and Poo H (1989) Transformation of a filamentous cyanobacterium by electroporation. J Bacteriol 171: 5743– 5746 Thiel T, Hartnett T and Pakrasi HB (1990) Examination of Photosystem II in heterocysts of the cyanobacterium Nostoc sp. ATCC 29150. In: Murata N (ed) Current Research in Photosynthesis, Vol 1, pp 291–294. Kluwer, Dordrecht Thomas J (1970) Absence of the pigments of Photosystem II of photosynthesis in heterocysts ofa blue-green alga. Nature 228: 181–183 Thomas J, Meeks JC, Wolk CP, Shaffer PW, Austin SM and Chien W-S (1977) Formation of glutamine from and by heterocysts isolated from Anabaena cylindrica. J Bacteriol 129: 1545–1555 Thorneley RNF and Lowe DJ (1985) Kinetics and mechanisms of the nitrogenase enzyme system. In: Spiro TG (ed) Molybdenum Enzymes, pp 221–284. Wiley & Sons, New York Timberlake WE (1991) Temporal and spatial controls of Aspergillus development. Curr Opin Genet Devel 1: 351–357 Toelge M, Ziegler K, Maldener I and Lockau W (1991) Directed mutagenesis of the gene psaB of Photosystem I of the cyanobacterium Anabaena variabilis ATCC 29413. Biochim Biophys Acta 1060: 233–236 Turner NE, Robinson SJ and Haselkorn R (1983) Different promoters for the Anabaena glutamine synthetase gene during growth using molecular or fixed nitrogen. Nature 306: 337– 342 Turing AM (1952) The chemical basis of morphogenesis. Phil Trans Roy Soc Lond B 237: 37–72 Turpin DH, Layzell DB and Elrifi IR (1985) Modeling the C economy of Anabaena flos-aquae. Estimates ofestablishment, maintenance, and active costs associated with growth on and Plant Physiol 78: 746–752 Tyagi VVS (1974) Effects of some inhibitors of DNA- and protein-synthesis on heterocyst formation in the blue-green alga Anabaena doliolum. Beitr Biol Pflanzen 50: 177–183 Udvardy J, Borbely G, Juhasz A and Farkas GL (1984) Thioredoxins and the redox modulation of glucose-6-phosphate dehydrogenase in Anabaena sp. strain PCC 7120 vegetative cells and heterocysts. J Bacteriol 157: 681–683 Van de Water SD and Simon RD (1982) Induction and differentiation of heterocysts in the filamentous cyanobacterium Cylindrospermum licheniforme. J Gen Microbiol 128: 917– 925 Van de Water S and Simon RD (1984) Heterocyst differentiation in Cylindrospermum licheniforme: studies on the role of transcription. J Gen Microbiol 130: 789–796 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. Molec Microbiol 6: 1853–1859 Walsby AE (1985) The permeability of heterocysts to the gases nitrogen and oxygen. Proc R Soc Lond B 226: 345–366 Wang L-F and Doi RH (1987) Promoter switching during development and the termination site of the operon of Bacillus subtitis. Molec Gen Genet 207: 114–119

C. Peter Wolk, Anneliese Ernst & Jeff Elhai Wastyn M, Achatz A, Molitor V and Peschek GA (1988) Respiratory activities and cytochrome oxidase in plasma and thylakoid membranes from vegetative cells and heterocysts of the cyanobacterium Anabaena ATCC 29413. Biochim Biophys Acta 935: 217–224 Wealand JL, Myers JA and Hirschberg R (1989) Changes in gene expression during nitrogen starvation in Anabaena variabilis ATCC 29413. J Bacteriol 171: 1309–1313 Webb R, Reddy KJ and Sherman LA (1990) Regulation and sequence of the Synechococcus sp. strain PCC7942 groESL operon, encoding a cyanobacterial chaperonin. J Bacteriol 172: 5079–5088 Wei T-F, Ramasubramanian TS, Pu F and Golden JW (1993) Anabaena sp. strain PCC 7120 bifA gene encoding a sequencespecific DNA-binding protein cloned by in vivo transcriptional interference selection. J Bacteriol 175: 4025–4035 Wilcox M, Mitchison GJ and Smith RJ (1973a) Pattern formation in the blue-green alga, Anabaena. I. Basic mechanisms. J Cell Sci 12: 707–725 Wilcox M, Mitchison GJ and Smith RJ (1973b) Pattern formation in the blue-green alga, Anabaena. II. Controlled proheterocyst regression. J Cell Sci 13: 637–649 Wilcox M, Mitchison GJ and Smith RJ (1975) Mutants of Anabaena cylindrica altered in heterocyst spacing. Arch Microbiol 103: 219-223 Winkenbach F and Wolk CP (1973) Activities of enzymes of the oxidative and the reductive pentose phosphate pathways in heterocysts of a blue-green alga. Plant Physiol 52: 480–482 Winkenbach F, Wolk CP and Jost M (1972) Lipids of membranes and of the cell envelope in heterocysts of a blue-green alga. Planta 107: 69–80 Wolk CP (1964) Experimental studies on the development of a blue-green alga. Ph. D. Dissertation, Rockefeller Institute, New York Wolk CP (1967) Physiological basis of the pattern of vegetative growth of a blue-green alga. Proc Natl Acad Sci USA 57:1246– 1251 Wolk CP (1968) Movement of carbon from vegetative cells to heterocysts in Anabaena cylindrica. J Bacteriol 96: 2138– 2143 Wolk CP (1973) Physiology and cytological chemistry of bluegreen algae. Bacteriol Rev 37: 32–101 Wolk CP (1979) Intercellular interactions and pattern formation in filamentous cyanobacteria. In: Subtelny S and Konigsberg IR (eds) Determinants of Spatial Organization, pp 247–266. Academic Press, NY Wolk CP (1982) Heterocysts. In: Carr NG and Whitton BA (eds) The Biology of Cyanobacteria, pp 359-386. Blackwell Scientific Publishers, Oxford Wolk CP (1989) Alternative models for the development of the pattern of spaced heterocysts in Anabaena (Cyanophyta). Pl Syst Evol 164: 27–31 Wolk CP (1991) Genetic analysis of cyanobacterial development. Curr Opin Genet Devel 1: 336–341 Wolk CP and Quine MP (1975) Formation of one-dimensional patterns by stochastic processes and by filamentous blue-green algae. Develop Biol 46: 370–382 Wolk CP and Shaffer PW (1976) Heterotrophic micro- and macrocultures of a nitrogen-fixing cyanobacterium. Arch Microbiol 110: 145–147 Wolk CP, Austin SM, Bortins J and Galonsky A (1974)

Chapter 27 Heterocyst Metabolism and Development Autoradiographic localization of after fixation of labeled nitrogen gas by a heterocyst-forming blue-green alga. J Cell Biol 61: 440–453 Wolk CP, Thomas J, Shaffer PW, Austin SM and Galonsky A (1976) Pathway of nitrogen metabolism after fixation of labeled nitrogen gas by the cyanobactcrium, Anabaena cylindrica. J Biol Chem 251: 5027–5034 Wolk CP, Vonshak A, Kehoe P and Elhai J (1984) Construction of shuttle vectors capable of conjugative transfer from Escherichia coli to nitrogen-fixing filamentous cyanobacteria. Proc Natl Acad Sci USA 81: 1561–1565 Wolk CP, Cai Y, Cardemil L, Flores E, Hohn B, Murry M, Schmetterer G, Schrautemeier B and Wilson R (1988) Isolation and complementation of mutants of Anabaena sp. strain PCC 7120 unable to grow aerobically on dinitrogen. J Bacteriol 170: 1239–1244 Wolk CP, Cai Y, and Panoff J-M (1991) Use of a transposon with luciferase as a reporter to identify environmentally responsive genes in a cyanobacterium. Proc Natl Acad Sci USA 88: 5355– 5359 Wolk CP, Elhai J, Kuritz T and Holland D (1993) Amplified expression of a transcriptional pattern formed during development of Anabaena. Molec Microbiol 7: 441–445 Wood NB and Haselkorn R (1979) Proteinase activity during heterocyst differentiation in nitrogen-fixing cyanobacteria. In: Cohen GH and Holzer H (eds) Limited Proteolysis in Microorganisms, pp 159–166. US DHEW Publication No.

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(NIH) 79–1591, Bethesda, MD Wood NB and Haselkorn R (1980) Control of phycobiliprotein proteolysis and heterocyst differentiation in Anabaena. J Bacteriol 141: 1375–1385 Xie W-Q and Potts M (1991) Gene cluster rpoBC1C2 in cyanobacteria does not constitute an operon. Arch Biochem Biophys 284: 22–25 Xie W-Q, Jäger K and Potts M (1989) Cyanobacterial RNA polymerase genes rpoC1 and rpoC2 correspond to rpoC of Escherichia coli. J Bacteriol 171: 1967–1973 Xu P and McAuley PJ(1990) Uptake of amino acids by the cyanobacterium Anabaena ATCC 27893. New Phytol 115: 581–585 Yakunin AF and Gogotov IN (1993) Natural electron donors for the nitrogenase reaction in the cyanobacterium Anabaena variabilis. Microbiology (Engl transl) 62: 58–62 Yu J-W, Price GD, Song L and Badger MR (1992) Isolation of a putative carboxysomal carbonic anhydrase gene from the cyanobacterium Synechococcus PCC7942. Plant Physiol 100: 794–800 Zhang C-C (1993) A gene encoding a protein related to eukaryotic protein kinases from the filamentous heterocystous cyanobacterium Anabaena PCC 7120. Proc Natl Acad Sci USA 90: 11840–11844 Zhang X, Tabita FR and Van Baalen C (1984) Nickel control of hydrogen production and uptake in Anabaena spp. strains CA and 1F. J Gen Microbiol 130: 1815–1818

Chapter 28 Differentiation of Hormogonia and Relationships with Other Biological Processes Nicole Tandeau de Marsac Unité de Physiologie Microbienne, Département de Biochimie et Génétique Moléculaire, Institut Pasteur, 28 rue du Docteur Roux, 75724 Paris Cedex 15, France

Summary I. Introduction II. Occurrence of Hormogonia Among Cyanobacteria III. Factors Modulating the Production of Hormogonia A. Nitrogen and Osmotic Effect B. Phosphorus, Iron and Other Nutrients C. The Effect of Light IV. Morphological, Ultrastructural, Biochemical and Genetic Changes During the Differentiation of Hormogonia A. Adhesion Properties B. Morphology and Ultrastructural Properties C. Genome Copy Number D. Gas Vesicle Genes E. Other Genes V. Relationships of Hormogonium Differentiation with Other Biological Processes A. Hormogonia and Phycobiliprotein Synthesis B. Electron Transport Chain and Cell Differentiation C. Nitrogen Assimilation and Cell Differentiation D. Sigma Factors VI. Hormogonia and Symbiosis VII. Further Prospects Acknowledgments References

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Summary Several filamentous heterocystous and non-heterocystous cyanobacteria can differentiate hormogonia that are distinguishable from vegetative trichomes by cell shape and, in some species, by cell motility and the presence of gas vesicles. Numerous environmental factors, including light and nutrients, can stimulate or inhibit hormogonium differentiation. In contrast to heterocysts, hormogonia are a transient cell form; they regenerate vegetative trichomes after a few rounds ofcell division. In symbiotic associations, hormogonia are the infective units. A factor synthesized by the plant partner could be responsible for the stimulation of the hormogonium differentiation and could thus enhance the establishment of the symbiosis. Some strains belonging to the genera Nostoc, Tolypothrix and Calothrix display a complex developmental cell cycle that includes both the differentiation of gas-vacuolated hormogonia, heterocyst differentiation and

D. A. Bryant (ed): The Molecular Biology of Cyanobacteria, pp. 825–842. © 1994 Kluwer Academic Publishers. Printed in The Netherlands.

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complementary chromatic adaptation. In Calothrix sp. strains PCC 7601 and PCC 7504, these three types of adaptation depend on two different photoreceptor systems. The control of both hormogonium and heterocyst differentiation depends on the Photosynthetic electron transport chain. Hormogonium differentiation is likely to be induced by a net oxidation ofthe plastoquinone pool, while heterocyst differentiation is stimulated by its net reduction. In contrast, complementary chromatic adaptation appears to be controlled by a photoreversible pigment, but this photoreceptor has not yet been characterized. Upon transfer from a nitrate- to an ammoniumcontaining medium, glutamine synthetase activity decreases by approximately 50%; synthesis of phycoerythrin decreases, while that of phycocyanin increases; methionine sulfoximine, an inhibitor of glutamine synthetase, prevents hormogonium differentiation and has an opposite effect on heterocyst differentiation. Based on these results, the cyanobacterial the counterpart of the regulatory protein from enteric bacteria that is involved in the control of both glutamine synthetase synthesis and activity, is proposed to play a role in the coordination of hormogonium and heterocyst differentiation, and complementary chromatic adaptation. These adaptation mechanisms most probably occur through cascades of phosphorylation/dephosphorylation of effectors and the interplay of specific sigma factors. I. Introduction Cyanobacteria display a wide variety of morphogenetic and developmental processes that can lead to the formation of baeocytes, which are minute reproductive cells; akinetes, resting cells often referred to as spore-like structures; hormogonia that are involved in the dispersal of the species in their natural habitats; and heterocysts, cells that are specialized for nitrogen fixation (Wolk, 1982, 1991; Rippka and Herdman, 1985; Adams, 1992; Tandeau de Marsac and Houmard, 1993). All four different cell types imply complex, and very fascinating, differentiation processes worth study in these Photosynthetic procaryotes. However, during the last decade, most studies have been conducted on the differentiation of heterocysts (see Chapter 27, this book) and to a far lesser extent on that of hormogonia. This review will focus on the molecular advances towards an understanding of the differentiation of hormogonia, but will also summarize some basic information on the occurrence of hormogonia and on the various factors presently known to influence their production. Finally, this differentiation process will be considered in a more general context, in relationships with heterocyst formation and with diverse adaptations to environmental stimuli, such as light intensity/spectral quality or nutrient availability, as well as the role of hormogonia in symbiosis.

Abbreviations: A3C – azetidine-3-carboxylic acid; DCMU – 3-(3,4-dichlorophenyl)-1,1dimethylurea; DBMIB – 2,5dibromo-3-methyl-p-benzoquinone

II. Occurrence of Hormogonia Among Cyanobacteria The term ‘hormogonium’ was initially used by phycologists to designate short motile trichomes liberated from immotile ensheathed parental trichomes, but has been extended by Rippka et al. (1979) to encompass motile or immotile trichomes of cells, that are often gas-vacuolated and devoid of heterocysts, with a cell shape and size distinguishable from the vegetative cells. Some cyanobacteria belonging to the genera Lyngbya or Leptolyngbya (section III described by Rippka et al., 1979) or Scytonema (section IV) produce hormogonia which are not gas-vacuolated and are morphologically very similar to vegetative trichomes but shorter in length. The motility of hormogonia from strains belonging to section III is not very conspicuous. In contrast, hormogonia from Chlorogloeopsis sp. (Fig. 1A–C) and Fischerella sp. (also called Mastigocladus sp.; section V) are generally highly motile and not gasvacuolated, but are composed of small cylindrical cells, easily distinguished from vegetative trichomes. The velocity of hormogonia from Mastigocladus laminosus on a surface has been calculated at between 1.7 and (Hernandez-Muniz and Stevens, 1987). Under nitrogen-deficient conditions, both terminal and intercalary heterocysts usually differentiate in late hormogonia from Chlorogloeopsis sp. (Fig. 1B), while they are almost exclusively intercalary in those from Fischerella sp. (Rippka et al., 1979). In mature vegetative trichomes, heterocysts are terminal, lateral or intercalary in these two cyanobacterial strains. The most typical gas-vacuolated hormogonia, with cells smaller in

Chapter 28 Hormogonium Differentiation

size than those of vegetative trichomes, are found amongst heterocystous cyanobacteria belonging to the genera Nostoc, Calothrix and Tolypothrix (section IV; see Figs. 2A–C, 3A–C, 4A–D and 5A–B). In these strains, hormogonia represent a transient morphological stage in the developmental cell cycle. Differentiation commences with several rounds of rapid cell divisions and may be in some species accompanied by expression of gas vesicles and motility. Subsequently hormogonia generally lose motility and gas vacuolation as they start regenerating vegetative trichomes. Late hormogonia differentiate two terminal heterocysts (Nostoc sp.) or only one

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terminal heterocyst (Scytonema, Calothrix and Tolypothrix sp.) undernitrogen deficient conditions. In the mature vegetative state, intercalary heterocysts are produced in both Nostoc and Scytonema sp. (Rippka et al., 1979; Rippka, 1988; Castenholz and Waterbury, 1989; see Figs. 3B, 4A and 9B). Cyanobacterial taxonomy has been to date primarily based upon the determination of morphological characters, among which hormogonium production (Nostoc sp ) or its absence (Anabaena sp.) has been used as a discriminatory property. Unfortunately, this criterion has proved unsatisfactory since strains can loose their capacity to differentiate hormogonia or be recalcitrant to hormogonium production under laboratory conditions. As a consequence of DNA/DNA reassociation experiments performed by Lachance (1981), and DNA/ DNA hybridization experiments and restriction pattern analyses using the gvpA gene encoding the structural gas vesicle protein from Calothrix sp. strain PCC 7601 as a probe (Damerval et al., 1989), one can now distinguish between species belonging to the genera Anabaena or Nostoc. For example, strains PCC 6411, PCC 7119 and the widely studied strain PCC 7120, originally described as Nostoc sp. and reassigned to the Anabaena genus on the basis of their inability to differentiate hormogonia (Rippka et al., 1979), as well as Anabaena sp. strain PCC 7118, belong to the same gvpA hybridization group as Nostoc sp. strains PCC 6705 and PCC 6719. As these results confirmed the high degree of interrelatedness previously established on the basis of DNA/DNA reassociation experiments (Lachance, 1981), these four Anabaena strains are now reassigned

Nicole Tandeau de Marsac

to the genus Nostoc (Rippka and Herdman, 1992). A second gene, gvpC, involved in gas vesicle formation in Calothrix sp. strain PCC 7601 (see paragraph 4 D; Damerval et al., 1987) is present in the six Nostoc (Anabaena) strains mentioned above, but absent in the Nostoc strains PCC 6720, PCC 73102 and PCC 8009 (Damerval et al., 1989). The presence or absence of the gvpC gene is thus not of taxonomic significance. Interestingly, the presence of the gvpC gene correlates with a rapid and abundant

Chapter 28 Hormogonium Differentiation production of gas vesicles as it occurs during the production of hormogonia in a number of Nostoc and Calothrix species (Damerval et al., 1989).

III. Factors Modulating the Production of Hormogonia Several environmental factors have been described that induce or inhibit the differentiation of hormogonia. In 1844, Thuret first described that a transfer of freshly harvested colonies of Nostoc verrucosum into a bowl of water induces the production of swarming, motile short trichomes within 2 – 3 days (Thuret, 1844). It is indeed established that the resuspension in distilled water of trichomes of several strains, or their transfer to fresh medium,causeshormogoniumproduction; however, if the inoculum utilized for subculturing is greater than 5–10% of the culture volume, no differentiation occurs (for reviews, see Lazaroff, 1973; Rippka and Herdman, 1985; Herdman and Rippka, 1988). Based on these observations, it has been proposed that cells in the stationary phase of growth synthesize and release an inhibitor of hormogonium differentiation. Although the chemical nature of the inhibitor is unknown, it has been shown to be dialyzable and inactivated by autoclaving (Herdman and Rippka, 1988).

A. Nitrogen and Osmotic Effect Removal of from the culture medium triggers the formation of gas-vacuolated hormogonia in Nostoc muscorum (Nostoc sp. strain PCC 6719) (Armstrong et al., 1983), and in Calothrix PCC 7601 and PCC 7504 (Herdman and Rippka, 1988). In Nostoc muscorum, a replacement of by equiosmolar concentrations of NaCl, KCl or prevents the induction while glucose or sucrose do not. These results indicate that the induction of gasvaculolated hormogonia in Nostoc muscorum does not result from nitrogen starvation and that ionic solutes, in contrast to non-ionic ones, prevent this induction. Moreover, if the removal of is combined with an increase of light intensity, the induction is larger than the sum ofeither one ofthese factors taken separately (Armstrong et al., 1983). In Calothrix sp. strain PCC 7601, replacement of by an equiosmolar concentration of NaCl does not prevent hormogonium formation and a short period

829 (2 h) of nitrogen starvation is sufficient to trigger the differentiation process (Herdman and Rippka, 1988). Consequently, hormogonium production in Calothrix sp. strain PCC 7601 can apparently result from rapid changes in nitrogen metabolism, rather than from an osmotic effect. In support of this hypothesis, methionine sulfoximine, a glutamate analogue that inhibits glutamine synthetase, prevents the differentiation of hormogonia. Furthermore, the level of the glutamine synthetase transcripts is transiently lowered during hormogonium differentiation in both Calothrix sp. strains PCC 7601 and PCC 7504 (G. Guglielmi and T. Damerval, unpublished; see further discussion in Section V). A cold shock (transfer ofthe trichomes to 4–10 °C) can also lead to a rapid induction of hormogonium production in these strains (Herdman and Rippka, 1988).

B. Phosphorus, Iron and Other Nutrients In some Calothrix sp. strains, iron-deficient cultures produce hormogonia upon readdition ofiron (Douglas et al., 1986; B. A. Whitton,personal communication). Although gas vesicles form in some of these strains, these hormogonia are often atypical with the colorless hair persisting in hair-forming strains. Similarly, in various heterocystous strains cultivated under phosphorus limitation, hormogonium induction occurs upon phosphorus repletion (Wood et al., 1986; Castenholz and Waterbury, 1989; Mahasneh et al. 1990; Whitton, 1992; B. A. Whitton, personal communication). In several Calothrix sp. strains, the cessation of hormogonium formation appears to be correlated with the induction of phosphatase (phosphomonoesterase and phosphodiesterase) activities, implying that fluctuations in ambient phosphate concentrations in natural environments could alternately lead to the formation of hormogonia or to the development of phosphatase activities (Islam and Whitton, 1992). If this hypothesis is correct, Calothrix sp. strain PCC 7601 is not representative of natural populations since, although this strain is transiently able to differentiate hormogonia, it does not express extracellular phosphatase activity under conditions of phosphorus limitation (B. A. Whitton, personal communication). Calothrix sp. strain PCC 7601 may indeed have lost its capacity to produce extracellular phosphatase activity during subculturing in phosphorus-rich culture media under laboratory conditions. In several Calothrix sp. strains it has also been shown that azetidine-3-carboxylic acid (A3C),

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a gametocide which reduces the rate of cell division in Chlorella sp. but does not affect cell size increase (Kelly, 1991), inhibits hormogonium differentiation at a concentration which still allows growth (B. A. Whitton, personal communication). By using different concentrations of A3C, it is possible to dissociate an effect on cell division from one on gas vesicle formation, since gas vesicles are synthesized when the concentration of A3C is reduced to while hormogonia devoid of gas vesicles can be produced at concentration of A3C up to (B. A. Whitton, personal communication). In the thermotolerant cyanobacterium Mastigocladus laminosus UTEX 1931, massive hormogonium production has been observed to occur at 45 °C, 6 to 10 h after transfer from a liquid culture to the surface of a medium solidified with 1.5% agar (Hernandez-Muniz and Stevens, 1987). Water solidified with agarose could not sustain the production of motile hormogonia and the presence ofglucose was not essential. appeared to be the only external factor required if the inoculum was taken from an actively growing culture, but light was also necessary if initial cultures were in stationary phase. Changes in environmental factors, such as deprivation of sucrose, elevated temperature and transition to darkness trigger hormogonium differentiation in Chlorogloea fritschii (Evans et al., 1976). However, massive production of hormogonia similar to that observed with Nostoc muscorum is never obtained in this strain, as it always simultaneously generates single cells.

C. The Effect of Light A substance from white-light- or red-light-grown cultures of Nostoc commune 584 and Nostoc muscorum A, whose activity was lost after autoclaving, has been reported to promote differentiation of motile trichomes in non-motile cultures incubated in darkness in the presence of glucose (for a review, see Lazaroff and Vishniac, 1961; Robinson and Miller, 1970; Lazaroff, 1973). This stimulating substance has never been characterized, but the determination of the effects of different light wavelengths on hormogonium production has been further investigated. The transfer of colonies of apparent aseriate cells of Nostoc muscorum A (Nostoc sp. strain PCC 7906) from darkness to red light causes the induction of motile hormogonia inde-

Nicole Tandeau de Marsac pendently of the presence or absence of sugar in the culture medium, but green light reverses the red light effect (Lazaroff, 1966,1972; for reviews, see Lazaroff and Schiff, 1962; Lazaroff, 1973). The action spectrum for the photoinduction of hormogonium differentiation shows a sharp peak in the red region with a maximum at 650 nm, while it displays a much broader curve in the green region for photoreversal of the red light effect. Differentiation occurs in a rather synchronous fashion but synchrony is progressively lost after one turn of the developmental cell cycle. A similar response to the spectral quality of the light is observed with Calothrix sp. strains PCC 7601 and PCC 7504 and has been the subject of recent detailed studies (Tandeau de Marsac et al., 1988; Herdman and Rippka, 1988; Damerval et al., 1991; Campbell et al., 1993). Transfer to a fresh culture medium initiates the differentiation of hormogonia in Calothrix sp. strain PCC 7601, but during the first 12 h after the induction the succession of events that leads to mature hormogonia is strongly dependent upon light conditions (Damerval et al., 1991). No differentiation occurs if exponentially growing cells are transferred into a fresh medium and incubated at 25 °C in the dark in the presence or in the absence of glucose, whatever the heterotrophic or photoheterotrophic conditions used for the precultures. In contrast, if cells are transferred to dim or bright (1 to red illumination for 24 h, 100% differentiation is obtained. Using white, blue, purple, yellow, green and red lights, at equivalent Photosynthetic photon flux density, it has been shown that green light alone or in combination with other wavelengths inhibits hormogonium differentiation, while red light activates differentiation. When both green and red lights are provided, the rate ofdifferentiation is greatly reduced but increases with an increasing ratio of red to green light. IV. Morphological, Ultrastructural, Biochemical and Genetic Changes During the Differentiation of Hormogonia

A. Adhesion Properties A number of benthic filamentous cyanobacteria producing hormogonia, Anabaenopsis circularis (PCC 6720), Calothrix desertica (PCC 7102), Fremyella diplosiphon and Plectonema boryanum

Chapter 28 Hormogonium Differentiation (PCC 6306) have been tested for their capacity to adhere to solid surfaces (Fattom and Shilo, 1984). Determination ofthe hydrophobic characteristics of the cell envelopes of both mature vegetative trichomes and hormogonia showed that vegetative trichomes are hydrophobic and thus display adhesion properties, while hormogonia are initially hydrophilic. However, hormogonia become hydrophobic within 48 h after formation when incubated in the light, but remain hydrophilic ifthey are incubated in the dark or in the light in the presence of chloramphenicol or 3-(3,4-dichlorophenyl)-l,l dimethylurea (DCMU; an inhibitor of Photosystem II). This indicates that the properties of the cell envelopes depend on the cell type and their physiological state, and changes require both protein synthesis and light energy. Such a transformation process would certainly play an important role in the detachment of the progeny from the benthic interface and dispersal in natural habitats.

B. Morphology and Ultrastructural Properties In Mastigocladus laminosus, motile hormogonia are composed of an average of 13.6 narrow cells that possess ultrastructural characteristics resembling those of cells located at the tips of the lateral branches in the trichomes (Nierzwicki et al., 1982; Balkwill et al., 1984; Nierzwicki et al., 1984a,b; HernandezMuniz and Stevens, 1987). Narrow cells (nearly uniform, long and cylindrical) display typical features of cells rapidly growing and dividing, and differ from wide cells (large, pleiomorphic and rounded up) in containing more ribosomes, more nuclear material, numerous cyanophycin and polysaccharide granules, but fewer centrally located carboxysomes and peripherally located lipid bodies. In narrow cells, the thylakoids are usually peripheral and less dispersed within the cytoplasm than in wide cells. In Nostoc muscorum A, cells from light-grown hormogonia are longer and thinner than those of non-motile vegetative trichomes grown in the light or those of dark-grown aseriate colonies (Lazaroff 1972; Ginsburg and Lazaroff, 1973; for reviews, see Lazaroff and Vishniac, 1962,1964; Lazaroff, 1973). Thylakoids in the cytoplasm of Nostoc sp. hormogonial cells form whorls rather than parietal stacks. In some cyanobacteria, hormogonia can be distinguished from vegetative trichomes by their motility and gas vacuolation, two properties that also contribute to the dispersal of the species. Motility is generally correlated with the presence of fimbriae

831 (or pili-like structures) whose number and spatial distribution on the outer surface of the cell wall appear to correspond to that of pores in the peptidoglycan layer (Guglielmi and Cohen-Bazire, 1982a,b). Pores could represent structures allowing fimbriae to cross the peptidoglycan layer or correspond to the anchoring sites of fimbriae on the murein sacculus. Fimbriae are abundantly present and arranged in a peritrichous manner on the outer cell surface of hormogonial trichomes from both the Nostoc sp. symbiont of the lichen Peltigera canina (Dick and Stewart, 1980) and from Calothrix sp. strain PCC 7601 (Damerval et al., 1991). These structures, approximately 7 nm in diameter and up to 3 or long in Nostoc sp. or Calothrix sp., respectively, differ from all of the major fimbrial types from heterotrophic bacteria (Dick and Stewart, 1980; Damerval et al., 1991). Figure 6 shows purified pores (Fig. 6B) and fimbriae (Fig. 6C) in the peptidoglycan layer of hormogonial cells from Calothrix sp. strain PCC 7601 (Damerval et al., 1991). Fimbriae have been biochemically charac-

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terized only from the unicellular Synechocystis sp. strains PCC 6701 (Guglielmi and Cohen-Bazire, 1982b)andCB3 (Vaara et al., 1984). However, it has been shown that the synthesis of fimbriae and the differentiation of hormogonia are prevented by chloramphenicol in Calothrix sp. strain PCC 7601 (G. Guglielmi and R. Rippka, unpublished results).

C. Genome Copy Number Hormogonium formation in Calothrix sp. strain PCC 7601 results from rapid cell divisions, approximately three cycles within 22 h, without cell elongation or DNA synthesis (Rippka and Herdman, 1985; Herdman and Rippka, 1988). This process implies that cell size diminishes (about 7-fold as calculated on a protein content basis) and that vegetative cells initially contain multiple copies of the genome. The genome size of Calothrix sp. strain PCC 7601 is unknown, but based on that of the closely related strain Calothrix sp. strain PCC 7101 Lachance, 1981), the mean DNA content corresponds to 24 genome equivalents for a vegetative cell and 3.4 for an hormogonial cell. This lower number of genome equivalents could be advantageous for genetic manipulations as selection for a specific mutational change would require less replication cycles for complete segregation of the mutated gene copy.

D. Gas Vesicle Genes In procaryotic organisms, gas vacuoles consist of aggregates of gas vesicles that are hollow, rigid, cylindrical structures closed at each extremity by a conical cap (for reviews, see Walsby, 1972,1981a,b, 1987; Walsby and Hayes, 1989). Made up of a monolayer of hydrophobic protein molecules forming ribs 4.5 nm in width, these structures are totally impermeable to water but permeable to gases that rapidly diffuse in and out, and provide cells with buoyancy. During the past 20 years, gas vesicles have been the subject of a number of biochemical, physiological and ecological studies in various cyanobacteria, while the molecular aspects of their synthesis during hormogonium differentiation has been studied in Calothrix sp. strain PCC 7601 and to a lesser extent in Calothrix sp. strain PCC 7504 (Tandeau de Marsac et al., 1985,1988; Csiszàr et al., 1987; Damerval et al., 1987, 1991). In Nostoc muscorum (Nostoc sp. strain PCC 6719), it has been

Nicole Tandeau de Marsac shown that the addition of chloramphenicol, which arrests the synthesis of the gas vesicle protein, accelerates the disappearance of gas vesicles due to the collapse of the preformed structures by rising turgor cell pressure, but no further studies of gas vesicle synthesis have been performed (Armstrong et al., 1983). Four genes involved in the formation of gas vesicles have been characterized in Calothrix sp. strain PCC 7601: gvpA1 and gvpA2 that correspond to a duplication of the structural gene encoding an identical GvpA protein; gvpC, a gene that forms an operon With gvpA1 and gvpA2 and encodes a protein of 162 amino acid residues whose presence has been correlated with strains producing gas vesicles rapidly and abundantly, and which, according to Walsby and Hayes (1989), may strengthen the gas vesicle structure against collapse pressure by attaching to the outside of the gas vesicle structure; and gvpD, a gene encoding a 7.5 kDa protein of unknown function, but whose sequence is 82% identical to GvpA (Tandeau de Marsac et al., 1988; Damerval et al., 1989; Walsby and Hayes 1989). The gvpA1A2C operon is transcribed as three mRNA species of 0.3 kb (gvpA1), 0.8 kb (gvpA1A2) and 1.4kb (gvpA1A2C). A fourth mRNA species of 0.6 kb has been ascribed to gvpD which is transcribed as a monocistronic unit. In addition, a 0.4 kb-long RNA species corresponds to an antisense RNA starting from the 3' end of the gvpA2 gene and ending in the gvpA1 gene (Csiszàr et al., 1987). The role of this antisense RNA is still unknown, but it has been proposed that it could prevent translation and/or modify mRNA stability by forming an homologous duplex with the three transcripts from the gvpA1A2C operon. In the course of the differentiation of hormogonia in Calothrix sp. strain PCC 7601, the transcription of the gvp genes starts 1.5 h after the induction by transfer of the cells to fresh culture medium and incubation under red light, and it becomes maximal after 6 h (Damerval et al., 1991). From 9 to 12 h, degradation of the gvp mRNAs commences and after 12 h gvp transcripts are no longer detected. If after 3 h of incubation under red light, differentiating hormogonia are exposed to green light, the differentiation process immediately stops and remains at the stage of cell division, no gas vesicles are synthesized, the transcription of the corresponding gvp genes is arrested, but the preexisting transcripts persist for at least 10 h without showing as much

Chapter 28 Hormogonium Differentiation degradation as those from cells continuously incubated underred light. Although it is tempting to hypothesize that the antisense RNA plays a role in the control of the expression of the gvp genes under green light, there is currently no experimental evidence to support this assumption. If cells are transferred to green light between 6 and 12 h after the induction of hormogonium differentiation under red light, the differentiation process stops at a stage that depends on the time when green light is applied but, after 12 h under red light, the differentiation process can proceed and fully differentiated hormogonia are produced even if green light is applied (Damerval et al., 1991). When the gvp genes are highly expressed, i. e. between 3 and 6 h, the transcription of the genes encoding phycobiliproteins, namely the and subunits of allophycocyanin, phycocyanin-1, phycocyanin-2 (specifically transcribed in cells under red light) is specifically arrested (Damerval et al., 1991). However, transcripts corresponding to the green light specific phycoerythrin genes remain detectable at low levels until 9 h after hormogonium induction. In contrast to the genes encoding other phycobiliproteins, the transcription of the phycoerythrin genes does not restart after the gas vesicle genes are switched off in cells incubated under continuous red light.

E. Other Genes The transcription of the genes encoding the two subunits of the ribulose bisphosphate carboxylase, first enzyme of the Calvin cycle, is not greatly affected during hormogonium differentiation (Damerval et al., 1991). In Calothrix sp. strain PCC 7601, the glnA gene encoding glutamine synthetase is transcribed as two mRNA species 1.8 and 1.6 kb long. The 1.8kb transcript is the most abundant mRNA species for at least 24 h after the induction of hormogonium differentiation under red light, while the 1.6-kb mRNA species predominates in cells incubated under green light during the same period of time (D. Campbell, unpublished). Such a change inthe relative abundance of the glnA transcripts is also clearly visible depending on the nitrogen source: The 1.8-kb mRNA species predominates in ammonium-grown cells and the 1.6-kb species in nitrate-grown cells (Elmorjani et al., 1992). Thus, similar effects are observed in cells grown in the presence of ammonium under white light or in cells differentiating hormogonia under red light. The Calothrix sp. strain

833 PCC 7601 glnA transcripts have notyet been mapped, but different start sites for transcription, as well as different regulatory sequences and effectors, including sigma factors, are possibly involved in the control of the expression of the glnA gene, depending on changes in environmental parameters, such as light and nitrogen sources. Preliminary results indicate that indeed changes occur in expression of a sigma factor during the first hours of the differ entiation of hormogonia (D. Campbell and A.-M. Castets, unpublished results). Figure 5 shows the morphological and ultrastructural changes that occur in Calothrix sp. strain PCC 7601 from the initiation of hormogonium differentiation to the regeneration of mature vegetative trichomes. Figure 7 summarizes the different stages of development and the time course of the expression of some genes during the differentiation process (Damerval et al., 1991).

V. Relationships of Hormogonium Differentiation with Other Biological Processes Numerous studies have been performed on complementary chromatic adaptation in Calothrix sp. strain PCC 7601 and it is now well known that green light specifically promotes the transcription of the cpeBA operon encoding the and subunits of phycoerythrin, while red light specifically promotes that of the cpcB2A2H2I2D2 operon encoding the and subunits of the phycocyanin-2 and the associated linker polypeptides (for reviews, see Grossman et al., 1988, 1993; Grossman, 1990; Tandeau de Marsac et al., 1988; Tandeau de Marsac, 1991; Tandeau de Marsac and Houmard, 1993; see also chapter 21 in this book). At least three proteins bind to specific DNA sequences upstream from either one of these two operons: RcaA and RcaB for the cpeBA operon in green light-grown cells; RcaD for the cpcB2A2H2I2D2 operon in red light-grown cells (Sobczyk et al., 1993; A. Sobczyk and J. Houmard, unpublished results). The affinity of RcaA and RcaD for their target DNA depends on their phosphorylation state indicating that both transcriptional and posttranslational controls come into play during complementary chromatic adaptation and that phosphorylation/dephosphorylation of protein effectors, as found in numerous bacterial regulatory systems (Stock et al., 1989, 1990), may represent a means to transduce the light signal in the cascade of

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events leading from signal perception to the physiological response in this light adaptation process. Neither the photoreceptor involved in complementary chromatic adaptation nor the intermediate mediator molecules in the signal transduction pathway have been identified. Physiological and biochemical studies indicate, however, that the photoreceptor, like the phytochrome of Photosynthetic eucaryotes, can exist in two interconvertible forms, P and P', whose action maxima are situated at 540 nm and 640 nm, respectively, i. e. at shorter wavelengths than the (660 nm) and (730 nm) forms of the phytochrome (for reviews, see Grossman et al., 1988 and 1993; Tandeau de Marsac et al., 1988; Grossman, 1990; Tandeau de Marsac, 1991; Tandeau de Marsac and Houmard, 1993; see also chapter 21 in this book).

A. Hormogonia and Phycobiliprotein Synthesis The fact that both the differentiation of hormogonia and the modulation of phycobiliprotein synthesis during complementary chromatic adaptation respond to similar wavelengths in the visible light spectrum leads to the question whether the two phenomena depend on the same or on two different photoperception systems in Calothrix sp. strain PCC 7601. Although hormogonium production is activated by red illumination around 640 nm and inhibited by

Nicole Tandeau de Marsac

green illumination around 540 nm, major differences exist in the effects of the spectral light quality on phycobiliprotein synthesis and hormogonium formation (Damerval etal., 1991). Firstly, continuous red illumination, even as low as of is required during the first 12 h of the differentiation process, while short flashes of red light can trigger the synthesis of phycocyanin-2. Secondly, no hormogonia are produced in the dark even in the presence of an exogenous carbon source, while dark-adapted cells possess a phycobiliprotein composition identical to that of cells grown under red light with an abundant synthesis of phycocyanin2, but nophycoerythrin. Thirdly, as mentioned earlier, the transcription of all the genes encoding phycobiliproteins, including the red light-specific phycocyanin-2 and the green light-specific phycoerythrin genes, is arrested during the differentiation process.

B. Electron Transport Chain and Cell Differentiation By using inhibitors of the thylakoid electron transport chain, DCMU and DBMIB (2,5-dibromo-3-methylp-benzoquinone), it has been recently shown that the opposing effects of red and green light on hormogonium differentiation arise through differential excitations of Photosystems I and II (Campbell

Chapter 28 Hormogonium Differentiation et al., 1993). The presence of DCMU, which blocks reduction of plastoquinone, and of light of any spectral quality promotes the differentiation of hormogonia, irrespective of the light regime used for the preculture. In contrast, DBMIB, which inhibits oxidation of plastoquinone, blocks this differentiation process, including transcription of genes encoding gas vesicles. The presence of DBMIB increases the frequency of heterocyst differentiation (Campbell et al., 1993). This is, in fact, consistent with earlier observations that both heterocyst differentiation and their associated nitrogen fixation capacity are promoted by green light in Anabaena azollae (Wu et al., 1982), Nostoc sp. (Wyman and Fay, 1987) and Calothrix sp. strain PCC 7601 (T. Damerval and G. Guglielmi, unpublished results). Thus, the two inhibitors have opposing effects on cell differentiation, as do red and green illumination. However, DCMU or DBMIB do not qualitatively alter the pattern of expression of the genes encoding phycobiliproteins. This indicates that the control of cell differentiation does not depend on the same photoperception system as complementary chromatic adaptation, but on a system involving changes in the redox state of a component of the electron transport chain, most probably the plastoquinone pool whose net oxidation under red light (or DCMU) or net reductionundergreen light (orDBMIB) would induce hormogonium or heterocyst differentiation, respectively (Campbell et al., 1993). Although this photoperception system differs from that involved in complementary chromatic adaptation, the regulation of phycobiliprotein synthesis and cell differentiation cannot be totally independent and must share at least some step(s) in their respective signal transduction pathways, since the transcription of the genes encoding phycobiliproteins stops, at least transiently, during hormogonium differentiation (Damerval et al., 1991). Such interconnections could be achieved through the synthesis of sigma factors and/or that of auxiliary transcriptional effectors whose affinity for DNA can be modulated by phosphorylation, as it is thought to occur for the transcription of the phycocyanin-2 and phycoerythrin genes in response to red and green illumination, respectively (Sobczyk et al., 1993; A. Sobczyk and J. Houmard, unpublished results), and/or through the modulation of enzyme activities by post-translational modifications. This also holds for the interactions expected to exist between the nitrogen regulatory system and the control of cell differentiation on the one hand and the

835 regulation ofphycobiliprotein synthesis on the other hand.

C. Nitrogen Assimilation and Cell Differentiation The recent discovery of a cyanobacterial (GlnB) highly similar to its bacterial counterpart (Tsinoremas et al., 1991), has opened a new way for investigating the regulatory control of nitrogen assimilation and more generally the relationships between nitrogen and carbon metabolism. In enteric bacteria the is uridylylated or deuridylylated when the intracellular ratio of glutamine: ketoglutarate is low or high, respectively, and the differential modification state ofthe protein controls both the activity and the synthesis of glutamine synthetase, a key enzyme in nitrogen assimilation (Reitzer and Magasanik, 1987; Magasanik, 1993). In the cyanobacteria Calothrix sp. strains PCC 7601 and PCC 7504, glutamine synthetase activity is low in cells grown in the presence of ammonium while it is high in the presence ofnitrate. Moreover, the total amount of glnA transcripts is generally lower in ammonium- than in nitrate-grown cells (Elmorjani et al., 1992; S. Liotenberg and D. Campbell, unpublished results). These results are in good agreement with those of Mérida et al. (1990) who showed that both the activity and the amount of glutamine synthetase in ammonium-grown cells of Calothrix sp. strain PCC 7601 decrease to approximately 40% of the levels in nitrate-grown cells. The question then arises whether the synthesis and activity of glutamine synthetase is controlled via differential modification of in cyanobacteria, as occurs in enteric bacteria. In the unicellular cyanobacterium Synechococcus sp. strain PCC 7942, the modification state of has been shown to depend on the nitrogen status of the cells as in enteric bacteria, although the protein is modified by a seryl-phosphorylation and not by uridylylation (Forchhammer and Tandeau de Marsac, 1994). In addition, in Synechococcus sp. strain PCC 6301, the protein is more abundantly modified in nitrate-grown cells incubated under light which excites Photosystem II than under light which excites Photosystem I (Tsinoremas et al., 1991). Thus, the modification state of in cyanobacteria may depend on the nitrogen status of the cells and also on their Photosynthetic activity which, together with the respiratory metabolism, governs the intracellular content of In Calothrix sp. strains PCC 7601 and PCC 7504,

Nicole Tandeau de Marsac

836

hormogonium differentiation can proceed in the presence offixed nitrogen in the culture medium and is a transient phenomenon that affects all the cells within a trichome. Conversely, heterocyst differentiation is repressed by ammonium and to a lesser extent by nitrate, and is an irreversible process that affects only a few cells. Methionine sulfoximine blocks the differentiation of hormogonia, while addition of glutamine to a methionine sulfoximinetreated culture restores the differentiation process (G. Guglielmi, unpublished results). Methionine sulfoximine has an opposite effect on heterocyst differentiation as it induces the formation of groups of contiguous heterocysts in various cyanobacteria, even in the presence of ammonium, but this effect is reversed by glutamine (For reviews, see Wolk, 1982; Adams, 1992). Finally, in Calothrix sp. strain PCC 7504, hormogonium differentiation is proportional to the intracellularglutamine concentration (possibly the glutamine: ratio) and correlates with the presence of in its unmodified form, while heterocyst differentiation is inversely proportional to glutamine concentration and correlates with modified (D. Campbell, unpublished results). Based on these results it is hypothesized that the protein connects changes in the intracellular nitrogen: carbon ratio to the induction of cell differentiation processes in these cyanobacteria. However, the effects of the spectral light quality on cell differentiation (this chapter, Section V B) probably act via a parallel pathway, since red or green light do not detectably alter glutamine pools or modification (D. Campbell, unpublished results). In summary, nitrogen metabolism plays a key role in both differentiation processes as the primary regulator of heterocyst differentiation and has an important influence on hormogonium differentiation. The protein is likely to be involved in these regulatory processes, possibly as an intermediary component in the transduction pathways or more likely as part of a parallel pathway. There certainly exist interactions and cross-talk at some steps that allow cells to coordinate these regulatory events. Another level of regulation that must be integrated in the complex network ofinteractions in Calothrix sp. strains PCC 7601 and PCC 7504 is the transient change in phycobiliprotein content that follows a transfer ofthe cells from a nitrate- to an ammoniumcontaining culture medium. Under such conditions, the synthesis of phycoerythrin decreases, while the

synthesis of phycocyanin tends to increase, giving the ammonium-grown culture a pigmentation reminiscent of a red light-grown one. This regulation operates mainly at the transcriptional level (S. Liotenberg, unpublished results).

D. Sigma Factors A search for specific sigma factors of RNA polymerase and determination of their role in the different regulatory mechanisms is an area of research worthy of pursuit as it will provide important clues for understanding the interconnections between these different adaptation processes in cyanobacteria (for additional discussion of RNA polymerase and transcriptional regulation, see Chapter 20). Three genes encoding sigma factors, sigA, sigB and sigC have been characterized from Nostoc (Anabaena) sp. strain PCC 7120 (Brahamsha and Haselkorn, 1991,1992) and from Synechococcus sp. strain PCC 7002 (L. Caslake and D. A. Bryant, personal communication), and one, sigA, has been characterized from Calothrix sp. strain PCC 7601 (A.-M. Castets, unpublished results). The predicted amino acid sequences of sigA from the Nostoc and Calothrix sp. are 97% identical in sequence. The Nostoc sp. sigA gene that encodes the 52 kDa principal sigma factor of vegetative-cell RNA polymerase is transcribed as two main mRNA species 2.2 and 1.7 kb long. The abundance of the 1.7-kb transcript remains fairly constant under both nitrogen-replete and nitrogendeplete conditions, whereas that of the 2.2-kb transcript increases following removal of fixed nitrogen (Brahamsha and Haselkorn, 1991). The Nostoc (Anabaena) PCC 7120 sigB and sigC genes encode putative sigma factors of 38,431 and 47,459 daltons, respectively, that share a high degree of similarity with SigA (Brahamsha and Haselkorn, 1992). Transcripts from both sigB and sigC are expressed only under nitrogen starvation, but in addition the expression of sigC, in contrast to sig B, is also specifically induced under sulfur starvation. The sigA gene from Calothrix sp. strain PCC 7601 hybridizes to at least five mRNA species among which one species appears to be more highly expressed during the first 8 h following hormogonium induction (D. Campbell and A.-M. Castets, unpublished results).

Chapter 28 Hormogonium Differentiation

837

VI. Hormogonia and Symbiosis

Some filamentous nitrogen-fixing cyanobacteria can establish symbiotic associations with various plants, including cycads (gymnosperms), Gunnera sp. (angiosperms), Anthoceros sp., Blasia sp. and Phaeoceros sp. (bryophytes), and Azolla sp. (pteridophytes), the marine diatomRhizosolenia sp., or with fungi to form lichens (Stewart et al., 1980; Meeks, 1988; Lindblad and Bergman, 1990). Among the cyanobacterial genera most commonly encountered in symbiosis are the hormogonium forming strains of the genera Nostoc (Figs. 8, 9), Calothrix, Scytonema and Fischerella. The presence of hormogonia has been noted on apical meristems of Azolla sp. stems, on the stem glands of Gunnera sp. (Fig. 10), in the Nostoc sp. colonies of Blasia and Anthoceros species. Reconstitution of associations in the laboratory has been successful only withNostoc spp. and Gunnera, Blasia, Phaeoceros and Anthoceros species. Based on reconstirution experiments, it is clear now that motile hormogonia represent a developmental stage of the symbiotic cyanobacteria that is important for the establishment of symbiosis (Campbell and Meeks, 1989; for reviews, see Stewart et al., 1980; Peters and Meeks, 1989; Meeks, 1990; Bergman et al., 1992). However, even if the gliding motility displayed by hormogonia is a prerequisite

for infection, some other properties of the Nostoc strains may be required for the infection process as some nitrogen-fixing Nostoc strains, capable of forming motile hormogonia, were unable to reconstitute an association with Anthoceros sp. (Enderlin and Meeks, 1983) and with Gunnera sp. (Bonnett and Silvester, 1981). The eukaryotic partners also participate in the stimulation of hormogonium formation under environmental conditions of nitrogen limitation. In Gunnera chilensis and G. manicata,

838

Nostoc sp. hormogonia, actively gliding through the channels towards the gland interior, might be attracted by phenolic compounds known to be present in the glands (Johansson and Bergman, 1992). Such compounds can indeed induce a chemotactic response in symbiotic bacteria (Munoz Aguilar et al., 1988). However, there exists no current evidence ofa specific chemotactic response in either Gunnera or Anthoceros sp. infected by Nostoc strains (Bergman et al., 1992; Meeks, 1990). It has been suggested that the production of hormogonia in the Anthoceros/Nostoc sp. associations is likely to be mediated by extracellular products synthesized by the bryophyte (Campbell and Meeks, 1989; for a review, see Meeks, 1990). Such a conclusion was based on the following observations: induction by a culture medium conditioned by the plant; suppression ofthe effect by autoclaving the conditioned medium; induction in the absence of combined nitrogen, but no inhibition in its presence; induction under environmental conditions – darkness or excess nutrients – which are usuallynoteffectiveinfree–livingstrains. Aputative inducing factor of low molecular mass, rather unstable upon storage and heat labile, has been found to act on Nostoc sp. strain UCD 7801 (Campbell and Meeks, 1989). Its chemical nature has not been determined, but this compound, whose production is inhibited in the presence of ammonium ions, is not identical to luteolin or acetosyringone, since these higher plant metabolites do not cause induction of hormogonium formation in this Nostoc strain. The hormogonia of symbiotic and non-symbiotic Nostoc strains are short, straight trichomes with small cells, tapered at each end of the trichome, lack heterocysts and do not increase in biomass. They posses carboxysomes, cyanophycin granules, lipid bodies and a large amount of glycogen reserves. While these hormogonia do not fix nitrogen, they assimilate and at a rate approximately 65% of that of the vegetative trichomes, but the specific activities of glutamine synthetase and ribulose bisphosphate carboxylase remain unchanged. Intracellular gas vesicles have not been seen by light microscopy or in thin sections of hormogonial cells (for reviews, see Meeks, 1990; Bergman et al., 1992). However, as shown by Damerval et al. (1989) in a survey of thirty cyanobacterial strains, this does not necessarily means that these strains lack the genetic information for the structural gas vesicle protein. Three days after the inoculation of Gunnera sp. with Nostoc sp., abundant

Nicole Tandeau de Marsac hormogonia accumulate on the glands before they move into the gland channels (Johansson and Bergman, 1992). In the Anthoceros sp. association, the timing of Nostoc sp. hormogonium differentiation is similar to that observed for Calothrix sp. strain PCC 7601. Approximately 100% hormogonium differentiation is obtained within 12 h and the hormogonial stage lasts for less than 48 h, followed by the differentiation of heterocysts and reinitiation of cell growth (Campbell and Meeks, 1989). Symbiotic Nostoc sp. trichomes usually contain a much higher percentage of heterocysts (up to 30 – 80% of cells) than the free-living cultures of the same strain, although some heterocysts may not be functional (for reviews, see Meeks, 1990; Bergman et al., 1992). While symbiotic vegetative cells contain cyanophycin granules, phycobilisomes, carboxysomes and glycogen granules, such inclusions appear to be absent from heterocysts. However, the presence or absence of phycoerythrin in heterocysts of the symbiont in Anthoceros sp. remains a matter of debate since, essentially all cyanobacterial cells in laboratory reconstitution emit a bright red color clearly visible by epifluorescence microscopy upon excitation at 560 nm (Meeks, 1990), but phycoerythrin has not been detected by immunoelectronmicroscopy of field samples (Rai et al., 1989). The regulation of assimilation in cyanobacterial symbionts differs depending on the type of association: regulation has been shown to be exerted at a posttranslational level in the Anthoceros sp. association (Joseph and Meeks, 1987); no decrease in either the specific activity or the relative level of glutamine synthetase protein has been observed in the cycads (Lindblad and Biergman, 1986) and the Gunnera sp. association (Bergman et al., 1992); enzyme synthesis has been reported to be lower in some lichens (Rai, 1990) and in Azolla association (Orr and Haselkorn, 1982; Lee et al., 1988; BraunHowland and Nierzwicki-Bauer, 1990) relative to free-living cyanobacteria. VII. Further Prospects Although recent studies on hormogonium differentiation have provided deeper insights into the molecular events involved, many aspects of this process remain unclear. Honnogonia from both freeliving and symbiotic cyanobacteria share common structural and physiological features, as well as

Chapter 28 Hormogonium Differentiation genetic properties that require more thorough investigation. In symbiotic associations, hormogonia primarily represent a means for infecting the plant partner. Therefore, symbiotic cyanobacteria may have evolved additional specific regulatory systems to respond to the plant stimuli that are worth elucidation together with the relationships between the different partners in the association. Controversial results on the occurrence of inhibitor(s) or of molecule(s) stimulating hormogonium differentiation have been obtained that require clarification. Such compounds might be involved in several key steps in the differentiation of hormogonia: i) the initial triggering or modulation of the response of the cells to environmental factors; ii) the propagation of the differentiation process along the cells in the trichomes; iii) and/or the control of the timing of the regeneration of vegetative trichomes. A search for mutants affected in the formation of hormogonia would also be useful, as their biochemical and genetic analysis would help identify the genes and proteins required for both the structural changes and the regulatory circuitry involved in hormogonium differentiation. At present, the only mutant available is SF33, a derivative of Calothrix sp. strain PCC 7601 (also called Fremyella diplosiphon strain UTEX 481) which grows as shorter trichomes than the parental strain (J. Cobley, personal communication). This mutant has lost the capacity to differentiate both hormogonia and heterocysts, but has retained the property of undergoing complementary chromatic adaptation in response to green and red illumination. The molecular events that occur during the differentiation of hormogonia can be studied as parts of a process that lead to morphological, physiological and.genetic changes within each cell along a trichome. However, since this transient differentiation process always precedes further adaptations to variations in environmental parameters, it is of even more interest to understand the network of interactions between hormogonium differentiation and other adaptation processes, such as heterocyst differentiation and chromatic adaptation. In this respect, the different aspects of the differentiation of hormogonia worthy of further examination include: i) the mechanism(s) of perception of the various environmental signals, in particular in response to the spectral light quality and to the availability of the nitrogen sources; ii) the different steps in the cascade of events that, in the absence of combined nitrogen, leads either to heterocyst differentiation via an hormogonial stage

839 (under red light) or directly to heterocyst differentiation (under green light); iii) the interactions between the regulatory mechanisms controlling cell differentiation, nitrogen assimilation and chromatic adaptation. We now have good evidence that the differentiation of both hormogonia and heterocysts in Calothrix sp. strains PCC 7601 and PCC 7504 depends on the redox state of a component of the Photosynthetic electron transport chain (most probably plastoquinone) rather than on a phytochrome-like photoreversible pigment which, although not yet characterized, is believed to control complementary chromatic adaptation in cyanobacteria. These Calothrix strains thus possess at least two photoperception systems. Both photoresponsive signal transduction pathways are likely to be connected to the nitrogen regulatory system involving the protein. These interactions could occur through specific effectors (sigma factors, cis/trans effectors, etc.) and through protein kinases/phosphatases, as for example those required to phosphorylate/ dephosphorylate the RcaA and RcaD proteins, two putative transcriptional activators for the genes encoding phycoerythrin under green light and phycocyanin-2 under red light, respectively (Sobczyk et al., 1993; A. Sobczyk and J. Houmard, unpublished results). Other transcriptional regulators have been recently discovered: RcaC in the Calothrix mutant SF33 (Chiang et al., 1992) and PatA in Nostoc (Anabaena) sp. strain PCC 7120 (Liang et al., 1992), which are involved in the control of the complementary chromatic adaptation and in the regulation of the pattern formation of heterocysts, respectively. All of these response regulators share functional or structural similarities with those from two-component regulatory systems in eubacteria. One can thus anticipate that phosphorylation cascades are indeed parts of the signal transduction pathways leading to both cell differentiation and chromatic adaptation in cyanobacteria.

Acknowledgments I gratefully acknowledge Pr. J. Meeks and Drs. D. Campbell and J. Houmard for careful reading of the manuscript and very helpful discussions. I also thank Prs. B. Bergman and J. Meeks, Drs. C. Johansson, G. Guglielmi and R. Rippka for providing photomicrographs, Pr. B. A. Whitton for communicating

840 data prior to publication and J. Lefèbvre for her help in the preparation of the manuscript.

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842 Stock JB, Stock AM and Mottonen JM (1990) Signal transduction in bacteria. Nature 344: 395–400 Tandeau de Marsac N (1991) Chromatic adaptation by cyanobacteria. In: Bogorad L and Vasil IK (eds) Cell Culture and Somatic Cell Genetics of Plants, Vol 7B, pp 417–446. Academic Press Inc., New York Tandeau de Marsac N and Houmard J (1993) Adaptation of cyanobacteria to environmental stimuli: New steps towards molecular mechanisms. FEMS Microbiol Rev 104: 119–190 Tandeau de Marsac N, Mazel D, Bryant DA and Houmard J (1985) Molecular cloning and nucleotide sequence of a developmentally regulated gene from the cyanobacterium Calothrix PCC 7601: A gas vesicle protein gene. Nucl Acids Res 13: 7223–7236 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 Tsinoremas NF, Castets A-M, Harrison MA, Allen JF and Tandeau de Marsac N (1991) Photosynthetic electron transport controls nitrogen assimilation in cyanobacteria by means of posttranslational modification of the glnB gene product. Proc Natl Acad Sci USA 88: 4565–4569 Thuret G (1844) Note sur le mode de reproduction du Nostoc verrucosum. Ann Sci Nat Bot Biol Vég 2: 319–323 Vaara T, Ranta H, Lounatmaa K and Korhonen TK (1984) Isolation and characterization of pili (fimbriae) from Synechocystis CB3. FEMS Microbiol Lett 21: 329–334 Walsby AE (1972) Structure and function of gas vacuoles. Bacteriol Rev 36: 1–32

Nicole Tandeau de Marsac Walsby AE (198la) Cyanobacteria: Planktonik gas-vacuolate forms. In: Starr MP, Stolp H, Trüper HG, Balows A and Schlegel HG(eds) The Prokaryotes, pp 224–235. SpringerVerlag, Berlin Walsby AE (1981b) Gas-vacuolate bacteria (apart from cyanobacteria). In: Starr M.P., Stolp H, Trüper HG, Balows A and Schlegel HG (eds) The Prokaryotes, pp 441–447. SpringerVerlag, Berlin Walsby AE (1987) Mechanisms of buoyancy regulation by planktonic cyanobacteria with gas vesicles. In: Fay P and Van Baalen C (eds) The Cyanobacteria, pp 377–392. Elsevier, Amsterdam Walsby AE and Hayes PK (1989) Gas vesicle proteins. Biochem J 264: 313–322 Whitton BA (1992) Diversity, Ecology, and Taxonomy of the Cyanobacteria. In: Mann NH and Carr NG (eds) Photosynthetic Prokaryotes, pp 1–51. Plenum Press, New York Wolk CP (1982) Heterocysts. In: Carr NG and Whitton BA (eds) The Biology of Cyanobacteria, pp 359–386. Blackwell Scientific Publications, Oxford Wolk CP (1991) Genetic analysis of cyanobacterial development. Curr Opin Genet Develop 1: 336–341 Wood P, Peat A and Whitton BA (1986) Influence ofphosphorus status on fine structure of the cyanobacterium (blue-green alga) Calothrix parietina, Cytobios 47: 89–99 Wu GL, Zhong ZP, Bai KZ, Wang FZ and Cui C (1982) The effects of light quality on the growth and development of Anabaena azollae. Acta Bot. Sin. 24: 46–53 Wyman M and Fay P (1987) Acclimation to the natural light climate. In: Fay P. and Van Baalen C. (eds) The Cyanobacteria, pp 347–376. Elsevier, Amsterdam

Organism Index A Aglaothamnion neglectum 76, 78, 156, 184, 186, 203, 204 Agmenellum quadruplicatum strain PR-6 (also see Synechococcus sp. strain PCC 7002) 33, 167, 411, 422 Alcaligenes eutrophus 107, 446, 448 Alcaligenes latus 399, 793 Amaranthus hybridus 585, 594, 603, 605 Anabaena 28, 31, 490 Anabaena azollae 9, 460, 696, 782, 783, 788, 835 Anabaena azollae-Azolla caroliniana 460 Anabaena catenula 807 Anabaena cylindrica 398, 399, 411, 420, 453, 494, 503, 505, 508, 509, 510, 655, 723–725, 777–779, 783–785, 788 Anabaena cylindrica (Fogg strain) 771, 791, 794, 796, 802, 805 Anabaena cylindrica (Wolk strain) 790, 810 Anabaena cylindrica PCC 7122 15, 16 Anabaena cylindrica strain ATCC 29414 771 Anabaena cylindrica strain CCAP 1403/2a 718 Anabaena flos-aquae 278, 392, 544, 655, 773 Anabaena siamensis 804 Anabaena sphaerica 386, 784 Anabaena sp. 5, 7–10, 132, 157, 161, 382, 388, 393, 497, 654, 660, 663, 666, 770, 772, 798, 807, 809, 828 Anabaena sp. strain 1F 505, 806 Anabaena sp. strain ATCC 29151 388, 736 Anabaena sp. strain ATCC 29211 388, 736, 777 Anabaena sp. strain CA 456 Anabaena sp. strain M-131 589, 591, 598, 599 Anabaena sp. strain PCC 6309 735,736 Anabaena sp. strain PCC 6411 742 Anabaena sp. strain PCC 7118 796, 799, 828 Anabaena sp. strain PCC 7119 150, 271, 283 284, 387–391, 423, 495, 718–722, 724, 726, 736, 737, 745, 778, 779, 788, 790, 792, 802 Anabaena sp. strain PCC 7120 (ATCC 27893) 4, 8, 9, 34, 40, 100, 124, 145, 146, 157–159, 161, 164, 165, 192, 194, 196, 197, 199, 200, 202, 204, 220, 290, 364, 366, 383–388, 402, 414, 415, 417–420, 422, 426, 441, 447, 455, 457, 460, 461, 478, 490, 492, 494–497, 500, 502, 503, 506, 508–510, 537, 570, 572, 584, 585, 588, 589, 592–605, 620-623, 626–629, 631, 633, 634, 635, 694–697, 718–726, 737, 742, 743, 762, 772–780, 785–789, 792, 794–806, 811–819, 823. See also Nostoc sp. strain PCC 7120 Anabaena sp. strain PCC 7122 20 Anabaena sp. strain UTEX 1444 773 Anabaena variabilis 158, 159, 173, 198, 200, 267, 278, 341, 385, 386, 392–396, 414–428, 461, 475, 476, 483–485, 494, 496, 497, 499, 504, 506, 510, 532, 561, 567, 570, 666, 736, 743, 771, 773, 775–791, 796, 799–806, 810 Anabaena variabilis strain ATCC 29413 (PCC 7937) 9, 74, 264, 271, 288, 323, 337, 339, 340, 342, 344, 348, 350, 388, 392, 393, 395, 421, 424–426, 495–499, 504, 588, 592, 593, 598, 602, 605, 695, 724, 773, 784, 794 Anabaena variabilis strain M-3 792 Anabaena variabilis strain PCC 6309 388 Anabaenopsis circularis 830 Anabaenopsis sp. 461, 804

Anacystis nidulans See also Synechococcus sp. strain PCC 6301 and 7942 10, 688, 696, 697 Anotrichium tenue 166 Antithamnion sp. 78, 100 Anthoceros punctatus 457, 460 Anthoceros sp. 837, 838 Aphanizomenon 28 Aphanizomenon flos-aquae 225, 275, 385 Aphanocapsa sp. strain PCC 6714 655 See also Synechocystis sp. strain PCC 6714 Aphanothece sacrum 383, 385 Arabidopsis thaliana 70, 220, 285, 459, 538 Arthrospira 19 Arthrospira jenneri 125, 411 Arthrospira sp. strain PCC 7345 19 Arthrospira sp. strain PCC 8005 14–16, 18, 19 Aspergillus niger 799 Astasia longa 95, 107 Azolla caroliniana 782 Azolla nilotica 9 Azolla sp. 5, 8, 9, 10, 776, 788, 837, 838 Azotobacter vinelandii 345, 492, 774–776

B Bacillus sp. 794–799 Bacillus sp. strain PS3 261, 264, 266, 271, 272 Bacillus licheniformis 82 Bacillus stearothermophilus 74 Bacillus subtilis 73, 74, 82, 525, 615, 528–532, 615, 616, 621, 622, 627,631,634,763, 794 Bacillus thermoproteolyticus 345 Bacillus thuringiensis 601 Blasia sp. 837 Bradyrhizobium japonicum 272, 290, 529, 531 Brassica napus 443 Bumilleriopsis filiformis 81, 695

C Calothrix 5, 7, 10, 825, 827–829, 837 Calothrix desertica 830 Calothrix membranacea 394 Calothrix mutant SF33 839 Calothrix parietina 663, 734 Calothrix sp. 44, 144, 776 Calothrix sp. strain PCC 7101 832 See also Tolypothrix sp. strain PCC 7101 632 Calothrix sp. strain PCC 7102 15, 16 Calothrix sp. strain PCC 7504 829, 830, 832, 835, 839 Calothrix sp. strain PCC 7601 10, 75, 144–146, 151, 152, 155, 158–160, 164, 165, 167, 173, 178, 179, 181, 183, 184, 188, 192, 196, 198, 200–202, 210, 344, 348, 350, 385, 506, 589, 592, 598, 599, 602, 603, 630, 634, 635, 642–653,666, 694, 828–839

844

Organism Index

Capsicum annuum 78 Caulobacter sp. 794 Chamaesiphon sp. strain PCC 7430 15, 16, 20 Chlamydia trachomatis 82 Chlamydomonas eugametos 105 Chlamydomonas moewusii 100, 105, 106 Chlamydomonas reinhardtii 70, 95, 99, 100, 102, 106, 225, 266, 270, 277, 279, 285, 286, 289, 291, 292, 302, 337, 393, 394, 450, 471, 484, 485, 524, 525, 528, 544, 545, 547, 695, 721, 724, 761

Chlorarachnion sp. 92 Chlorella pyrenoidosa 680 Chlorella sp. 830 Chlorella vulgaris 524 Chlorobium limicola 333 Chlorobium limicola f. thiosulfatophilum 321, 330 Chlorobium phaeobacteroides 321 Chlorobium sp. 271, 364 Chlorobium vibriforme 321 Chlorobium vibrioforme f. thiosulfatophilum NCIB 8 525 Chloroflexus aurantiacus 142, 215 Chlorogloea fritschii 830 Chlorogloeopsis 10, 826 Chlorogloeopsis fritschii 453 Chlorogloeopsis fritschii strain CCAP 1411/16 723 Chlorogloeopsis fritschii strain PCC 6718 14–16, 20 Chlorogloeopsis sp. HTF (strain PCC 7518) 12–16, 20 Chondrus crispus 108, 388 Chromatium sp. 272 Chroococcidiopsis 8 Chroococcus 7 Chroomonas sp. 76, 163, 183, 184, 188 Clostridium butylicum 386 Clostridium pasteurianum 337 Clostridium sp. 771 Coccochloris peniocystis 453 Corynebacterium nephridii 717 Cryptomonas maculata 188, 189 Cryptomonas sp. 98, 100, 107 Cryptomonas sp. strain 71, 73, 78, 80–82, 188, 190, 203, 539 Cryptomonas sp. strain CS-24 204 Cyanidium caldarium 78, 81, 107, 166, 167, 387, 532, 534, 535, 537, 695, 735 Cyanophora paradoxa 15, 16, 19, 66–84, 95, 103, 111, 140, 142, 144, 155, 159, 169, 170, 200, 203, 206, 208–211, 279, 340, 397, 453, 460, 539, 545, 601, 621, 695 Cyanospira rippkae 784 Cyanothece 39, 490 Cylindrospermum 10 Cylindrospermum licheniforme 791, 803 Cylindrospermum licheniforme strain ATCC 29412 784, 785, 804 Cylindrospermum sp. 804 Cylindrospermum sp. strain PCC 7417 15, 16 Cylindrotheca sp. 78 Cylindrotheca sp. strain N1 446, 447

Desulfovibrio gigas 345, 398, 531 Desulfovibrio sp. 401 Dictyostelium discoideum 809 Dictyota dichotoma 78 Drosophila melanogaster 799, 808 Dunalliela bardawil 571

E Enteromorpha prolifera 389, 393 Epifagus virginiana 94, 95 Erwinia herbicola 78, 571–573 Erwinia uredovora 77, 571–573 Escherichia coli 12, 15–17, 37, 73, 74, 77, 78, 80, 82, 96, 261, 275, 276, 280, 284, 285, 287, 290, 345, 364, 368, 371, 373, 393, 419, 427, 443, 482, 506, 524, 525, 528, 529, 531, 532, 535–538, 544, 547, 548, 582, 584, 585, 588–591, 594, 602– 605, 614, 616, 617, 620–632, 634–637, 702, 705, 716, 740, 754, 763 Euglena gracilis 70, 73, 75, 79, 81, 96,105, 528–530, 539, 540 Euglena gracilis strain UTEX 753 720

F Fischerella 10, 490, 826, 837 See also Mastigocladus Fischerella muscicola 394 Fischerella sp. (Mastigocladus laminosum) 146, 598 Fischerella sp. strain ATCC 27929 776 Fischerella sp. strain PCC 7414 14, 15, 16, 20, 123 Fremyella diplosiphon See Calothrix sp. strain PCC 7601

G Gastroclonium coulteri 143, 186, 191, 203 Geosiphon pyriforme 67 Glaucocystis nostochinearum 67, 68, 82, 453 Glaucosphaera vacuolata 67 Gloeobacter sp. strain PCC 8105 18 Gloeobacter violaceus 143, 177, 206, 209, 288, 411, 426 Gloeobacter violaceus strain PCC 7421 15–17, 125 Gloeocapsa alpicola 411, 561 Gloeocapsa sp. strain PCC 73106 15, 16 Gloeochaete wittrockiana 67, 68 Gloeothece 490, 491 Gloeothece sp. strain PCC 6501 15, 16 Gloeothece sp. strain PCC 6909 428, 497 Gloeotrichia 805 Gracilaria tenuistipata 107 Gracilaria verrucosa 108 Griffithsia pacifica 143, 208 Gunnera chilensis 837 Gunnera manicata 837 Gunnera sp. 9, 837, 838

D Denticula sp. 67 Dermocarpa sp. strain PCC 7437 15, 16, 20 Dermocarpa violaceae 411

H Heliobacillus mobilis 321, 330, 333

Organism Index Heliobacterium chlorum 93, 103, 271, 272, 321 Heliobacterium gestii 321 Hyphomorpha perrieri 771

845

Lactobacillus leichmannii 726 Lactococcus lactis 82 Lemna gibba 286, 297 Lemna purpusilla 289 Lepidium sativum L. 546 Leptolyngbya 826 Limnothrix redekei strain Mef 6705 15–18, 21 Lyngbya 826 Lyngbya sp. 400 Lyngbya sp. strain PCC 7419 14, 15, 16, 19 Lyngbya–Plectonema–Phormidium (LPP) 10, 490

Nostoc muscorum 272, 384, 385, 423, 829–832 Nostoc muscorum strain ATCC 29151 424 Nostoc sp. 265, 383, 771, 776, 838 Nostoc sp. strain ATCC 27896 589 Nostoc sp. strain ATCC 29133 589 Nostoc sp. strain ATCC 29150 782 Nostoc sp. strain MAC 155, 200, 201, 374, 383–385, 416, 420, 422, 438, 457, 589, 736 See also Nostoc sp. strain PCC 8009 Nostoc sp. strain PCC 6411 828 Nostoc sp. strain PCC 6705 828 Nostoc sp. strain PCC 6719 828, 829, 832 Nostoc sp. strain PCC 6720 828 Nostoc sp. strain PCC 7119 828 Nostoc sp. strain PCC 7120 4, 15, 16, 836, 839 See also Anabaena sp. strain PCC 7120 Nostoc sp. strain PCC 7121 285 Nostoc sp. strain PCC 73102 15, 16, 399, 455, 828 Nostoc sp. strain PCC 7524 584 Nostoc sp. strain PCC 7906 263, 268, 270, 272–276, 278, 280, 287–300, 302, 422 Nostoc sp. strain PCC 8009 288, 323, 343, 347, 349, 350, 352, 374, 418, 421, 424, 425, 589, 598, 736, 828 Nostoc sp. strain UCD 7801 456, 460, 838 Nostoc verrucosum 387, 829

M

O

Macrozamia sp. 455 Mantoniella squamata 104 Marchantia polymorpha 15, 16, 19, 75, 93, 95, 103, 324, 343, 482, 545 Mastigocladus sp. See also Fischerella sp. Mastigocladus laminosus 129, 140, 145–161, 164–176, 181, 183, 187, 188, 191–215, 326, 374, 415, 417, 663, 805, 831 See also Fischerella sp. Mastigocladus laminosus strain UTEX 1931 830 Mastigocladus sp. 780, 826 ‘Mastigocladus sp. HTF’ strain PCC 7518 20 Mastigocoleus testarum 771 Methanococcus vannielii 82 Microcoleus sp. strain l0mfx 15, 16, 19 Microcoleus sp. strain PCC 7420 15, 16, 19 Microcystis aeruginosa 225, 278, 397, 399, 415, 421, 574 Microcystis sp. strain PCC 7005 10 Mycoplasma capricolum 82 Myxococcus sp. 794 Myxosarcina sp. strain PCC 7312 16, 20 Myxosarcina sp. strain PCC 7313 15

Ochromonas danica 103, 105, 106 Odontella sinensis 76, 100 Olisthodiscus luteus 450 OS (Ocotopus Springs isolate)–V–L–13 15–17 OS(Ocotopus Springs isolate)–V–L–16 15–17 OS(Ocotopus Springs isotate)–VI–L–4 15–17 OS(Ocotopus Springs isolate)–VI–L–8 15–17 Oscillatoria 7, 490, 491 Oscillatoria agardhii 561, 566, 574 Oscillatoria amphigranulata CCCNZ–concert–Oa 15, 16, 18, 19 Oscillatoria limnetica 15, 16, 19, 285, 398, 400 Oscillatoria redekei 17 Oscillatoria sp. 19, 30 Oscillatoria sp. strain PCC 6304 15, 16, 18, 20 Oscillatoria sp. strain PCC 7105 15, 16, 19 Oscillatoria sp. strain PCC 7515 15, 16, 19 Oscillatoria splendida 574 Oscillatoria tenius 742 Oscillatoria terebriformis 285

K Klebsiella aerogenes 774, 785, 787 Klebsiella pneumoniae 495

L

P N Neurospora crassa 78, 272, 274, 532, 570–572 Nodularia 10, 28 Nodularia harveyana 784 Nodularia sp. strain PCC 73104 15, 16, 20 Nostoc 5, 7–10, 490, 825, 827– 829, 831, 835, 837, 838 Nostoc commune 564, 567, 661, 774 Nostoc commune strain UTEX 584 496, 620, 830 Nostoc cycas strain PCC 7422 9 Nostoc ellipsosporum 799, 810

Paracoccus denitrificans 264, 266, 272, 279, 284, 290, 419, 427, 787 Paulinella chromatophora 66, 67 Pavlova lutherii 78, 80, 82 Peliaina cyanea 67 Peltigera canina (lichen) 831 Peptococcus aerogenes 345 Phaeoceros sp. 837 Phormidium 7, 10, 18 Phormidium ectocarpi 19 Phormidium ectocarpi strain CCAP 1462/5 15, 16

Organism Index

846 Phormidium ectocarpi strain N182 15, 16, 19 Phormidium ectocarpi strain PCC 7375 7, 15, 16, 20 Phormidium foveolarum 394, 491 Phormidium fragile strain PCC 7376 15, 16, 20 Phormidium laminosum 129, 220, 225, 324, 326, 396, 509, 564, 566, 567 Phormidium luridum 414, 415, 423, 426, 428, 560 Phormidium minutum strain D5 15, 16, 19 Phormidium persicinum 126, 127, 144, 151, 215 ‘Phormidium persicinum’ strain CCAP 1462/5 7 Phormidium sp. strain C86 151 Phormidium sp. strain PCC 7375 See Phormidium ectocarpi strain PCC 7375 Porphyra purpurea 539 Phorphyridium cruentum 144, 179, 208 Plasmodium sp. 110 Plectonema 10, 31, 490 Plectonema boryanum 95, 278, 394, 415, 417, 418, 422, 426, 491, 494, 496, 497, 499, 544, 545, 584, 589–593, 663, 830 Plectonema boryanum strain PCC 73110 10, 15, 16, 19 Plectonema boryanum strain UTEX 594 584 Plectonema sp. strain PCC 73110 495 Pleurocapsa minor 411 Pleurocapsa sp. strain PCC 7321 15, 16, 20 Polysiphonla boldii 203 Porphyra nereocystis 683 Porphyra purpurea 70, 73, 76–79, 83, 100, 101, 107, 203, 397, 535 Porphyra sp. 81 Porphyra umbilicalis 350, 534 Porphyra yezoensis 680, 686, 688 Porphyridium aerugineum 78, 144, 203, 208, 209 Porphyridium cruentum 127, 134, 143, 150, 155, 165, 169, 176, 178, 183, 184, 188–191, 197, 198, 206, 208, 210–216, 397, 679, 682, 683, 686, 688 Porphyridium sordidum 173, 185–187 Prochlorococcus marinus 14–17, 42, 49, 51, 53, 54, 103, 547 Prochloron didemni 49–51, 54, 92, 103 Prochloron sp. 7, 8, 10, 14–16, 50, 51–57, 59, 60, 62 Prochlorothrix hollandica 8, 14–16, 19, 49–62, 100, 103, 105, 220, 290, 325, 522, 566, 695, 696, 698, 712–714 Propionibacterium freudenreichii 526 Propionigenium modestum 368 Proteus vulgaris 12 Pseudanabaena 10, 21, 490 Pseudanabaena foveolarum 394 Pseudanabaena galeata 17 Pseudanabaena galeata strain CCCOL–75–PS 15, 16, 18 Pseudanabaena sp. strain M2 574 Pseudanabaena sp. strain PCC 6903 8, 15–17 Pseudanabaena sp. strain PCC 7403 8, 10, 15, 16, 18, 21 Pseudanabaena sp. strain PCC 7409 8, 21, 164, 181, 200, 202, 584, 631, 645, 647 Pseudomonas denitrificans 529, 548 Pseudomonas sp. 272 Pylaiella littoralis 70, 107 Pyrenomonas salina 82

R Rhizobium meliloti 525 Rhizobium sp. 261

Rhizolenia 30 Rhizosolenia sp. 837 Rhodella violacea 102, 144, 152, 169, 185, 203, 205, 210–212 Rhodobacter capsulatus 77, 78, 133, 263, 264, 267, 270, 272– 275, 277, 279, 289–294, 296–299, 301, 302, 539–542, 545, 546, 548, 571–573, 659 Rhodobacter sphaeroides 264, 266, 267, 269, 271, 272, 275–277, 279, 282, 289, 292, 294, 298, 299, 440, 446, 448, 451, 452, 521, 527–530, 540, 546, 548, 722 Rhodopseudomonas capsulatus 419 See Rhodobacter capsulatus Rhodopseudomonas viridis 272, 276, 329 Rhodospirillum rubrum 264, 269, 272, 277, 279, 280, 297, 375, 398, 440, 441, 446, 452, 479, 546, 789 Richelia intracellularis 28, 30 Rivularia 805

S Saccharomyces cerevisiae 37, 78 Salmonella typhimurium 77, 507, 529, 547, 661 SAR100 (Sargasso Sea isolate) 15–17 SAR139 (Sargasso Sea isolate) 15–17 SAR6 (Sargasso Sea isolate) 15–17 SAR7 (Sargasso Sea isolate) 15–17 Sauromatum guttatum 427 Scenedesmus obliquus 233, 393, 524, 546, 547 Scytonema 826, 828, 837 Scytonema sp. strain PCC 7110 15, 16 Spirulina 19 Spirulina maxima 275, 277, 278, 383, 385, 394, 398, 503 Spirulina platensis 277, 383, 394, 505, 529, 564, 566, 574 Spirulina sp. 390, 391, 422, 562, 660, 666 Spirulina sp. strain PCC 6313 15, 16, 19 Staphylococcus carnosus 82 Streptomyces coelicolor 616, 622 Streptomyces sp. 794, 798 Sulfolobus acidocaldarius 271, 272 Synechococcus 5, 7, 27–30, 32, 34, 35, 490, 491, 655 Synechococcus cedrorum 655 Synechococcus elongatus 241, 244, 245, 323, 339, 344, 348, 561, 567 Synechococcus elongatus strain CCAP 1497/1 7 Synechococcus leopoliensis 654 Synechococcus lividans II 103 Synechococcus lividans III 103 Synechococcus lividus 278, 335 Synechococcus lividus strain CCCY7C-S 15, 16, 19 Synechococcus sp. 8, 69, 129, 131, 177, 178, 202, 283, 286, 323, 324, 326, 327, 330, 335, 336, 337, 339, 340, 342, 343, 346, 347, 350, 352, 388, 560, 565, 654, 660, 661, 663, 666, 777, 803 Synechococcus sp. (marine) 655 Synechococcus (Marine Cluster-A (MC-A)) 28 Synechococcus sp. (PIM9–S1E) 160 Synechococcus sp. strain DC2 631 Synechococcus sp. strain L-1402-1 505 Synechococcus sp. strain NIBB 1059 122 Synechococcus sp. strain PCC 6301 See also Anacystis nidulans 7, 10, 15, 16, 19–21,75, 96, 104, 123, 127, 134, 143, 144, 146, 150, 154, 155, 159, 160, 194, 200, 201, 204, 241, 288, 323, 335–337, 344, 345, 347, 350, 364, 385, 389, 390, 392, 396, 397, 399, 411, 412, 414, 415, 417–429, 440–442, 444,

Organism Index 446–456, 458, 461–463, 475, 476, 498, 501, 508, 509, 522, 526, 527, 529, 532, 542, 548, 549, 590, 654, 666, 682, 707, 725, 734–736, 762, 793, 835 Synechococcus sp. strain PCC 6301 mutant AN112 160 Synechococcus sp. strain PCC 6307 566 Synechococcus sp. strain PCC 6311 574 Synechococcus sp. strain PCC 6716 265, 280, 374, 375, 398 Synechococcus sp. strain PCC 6908 735 Synechococcus sp. strain PCC 7002 20, 33, 34, 42, 43, 76, 79, 107, 129, 145, 146, 150, 155, 156, 159, 160, 162, 164, 167, 171, 172, 173, 174, 176, 192, 193, 194, 196, 197, 198, 200, 202, 204, 270, 272, 274, 275, 276, 278, 279, 283, 285, 286, 288, 289, 290, 323, 324, 327, 328, 330, 333, 337, 339, 340, 341, 342, 343, 345, 346, 347, 348, 350, 352, 353, 382, 387, 390, 411, 414, 416, 417, 418, 419, 422, 451, 455, 502, 506, 509, 526, 536, 537, 565, 584, 590, 592, 596, 599, 600, 601, 602, 604, 621, 622, 633, 663, 655, 665, 682, 683, 694, 696, 734, 735, 737, 738, 739, 764, 797, 836 Synechococcus sp. strain PCC 7335 20 Synechococcus sp. strain PCC 7942 12, 33, 34, 37, 39, 40, 41, 82, 95,96, 122–124, 146, 150, 160, 191, 194, 278, 284, 382, 383, 387, 390, 392, 394–396, 414, 424, 425, 429, 457, 458, 461, 469, 472, 475, 476–481, 492, 498–506, 510, 529, 530, 564–574, 590–593, 596, 597, 601–603, 621, 622, 628–631, 634, 654, 659–668, 694–710, 715, 718–721, 724–726, 734, 735, 737–739, 742, 743, 745, 758–764, 775, 835 Synechococcus sp. strain PCC 7943 590 Synechococcus sp. strain RF–1 461, 491, 497 Synechococcus sp. strain UTEX 625 418, 505, 509 Synechococcus sp. strain WH 7803 8, 30, 31, 33–44, 166, 574 Synechococcus sp. strain WH 7805 17, 166 Synechococcus sp. strain WH 8008 42 Synechococcus sp. strain WH 8012 34 Synechococcus sp. strain WH 8018 33, 34, 42 Synechococcus sp. strain WH 8020 36, 37, 44, 166, 177, 178, 181, 183, 184, 186, 189, 202, 537 Synechococcus sp. strain WH 8101 39 Synechococcus sp. strain WH 8103 17, 34, 36, 37, 42, 166, 177, 178, 180, 202 Synechococcus vulcanus 107, 225–227, 229, 323, 337, 340–344, 348, 350, 384, 397, 699 Synechococcus vulcanus Copeland 220, 221 Synechocystis (Aphanocapsa) 7, 10, 29

847 Synechocystis sp. 14, 132, 336, 725 Synechocystis sp. strain CB 3 832 Synechocystis sp. strain PCC 6308 7, 15, 16, 453, 456, 590, 666, 668 Synechocystis sp. strain PCC 6701 181, 644, 647, 832 Synechocystis sp. strain PCC 6714 123, 125, 126, 132, 134, 265, 271, 283, 288, 396, 415, 417, 418, 419, 420, 421, 422, 423, 426, 456, 566, 567, 570, 571, 573, 590, 687, 695, 696, 697, 699, 707, 709, 710, 735, 736. See also Aphanocapsa sp. strain PCC 6714. Synechocystis sp. strain PCC 6803 42, 74, 95, 100, 101, 107, 128– 131, 220, 223, 225, 228, 230, 232, 237, 239, 265, 270, 272, 274–279, 284–286, 289, 323, 326, 327, 330, 335–350, 352, 364, 386, 388, 392, 393, 395, 397, 398, 412, 418, 419, 422, 424, 425, 427, 451, 453, 461, 471, 475–477, 479, 481, 482, 503–510, 523–527, 545, 564, 571, 573, 574, 584, 590–592, 595, 599–604, 621, 694–699, 702, 706, 708–710, 724, 737, 738, 761–764 Synechocystis sp. strain PCC 6906 15, 16 Synechocystis sp. strain WH 8501 39, 166, 178

T Thermus sp. 44, 261 Thermus thermophilus 271, 272 Theromyzon rude (leech) 807 Thiobacillus neopolitanus 478 Tolypothrix 825, 827, 828 Tolypothrix sp. 394 Tolypothrix sp. strain PCC 7101 644, 648 See also Calothrix sp. strain PCC 7101 Trichodesmium 28, 30, 40, 490 Trichodesmium sp. 11, 31, 39, 491, 659 Trichodesmium sp. strain NIBB 1067 43 Trichodesmium thiebautii 40, 491

Z Zamia integrifolia 8

Gene and Gene Product Index A aadA 588 acpA 78, 79 ApcA 154 apcA 76, 200, 205, 644 apcA1 625 apcA1B1 627, 646 apcA2 200, 625, 627, 646 apcA3 625 apcAB 203, 204, 601, 650, 666, 797, 798 ApcB 154 apcB 76, 200, 205, 644 ApcC 76 apcC 78, 193, 200, 203, 205, 644, 646 ApcD 154, 161, 681 apcD 154, 155, 162, 200, 203–205, 211, 625, 644, 646, 683 apcE 76, 156, 158, 159, 200, 203–205, 602, 625, 644, 646 ApcE See core-membrane linker apcF 76, 155, 200, 203–205, 624, 625, 633, 644, 683 apcL 78 aphII 585 argB 78 argC 509 argD 509 argE 509 argJ 509 atp1 operon 364, 623, 624, 628 atp2 operon 364, 623, 624, 627, 628 atpA 97, 100, 105 atpB 76, 97, 100, 105, 111, 366 atpBE 54, 797 atpC 76, 97 atpD 76, 97, 100 atpE 76, 97, 100, 366 atpF 97, 99, 100 atpG 76, 78, 97, 100 atpH 97, 100 atpI 76, 97, 100

B bch 538 bchB 522, 545 bchD 522, 539 bchE 522, 541, 548 bchG 522, 546 bchH 522, 540, 548 bchI 522, 539 bchJ 522, 542 bchL 522, 545 bchM 522, 541 bchN 522, 545 bchO 539 bchP 522, 546 BifA 631, 635, 802

bifA 631 BifB 631, 634, 635 ble 588 bom 583, 591 brlA 799

C cat 586, 588, 596, 602, 628 cbbM 441 cbpA 473, 565, 745 ccmK 481 ccmL 481 ccmM 481 ccmN 473, 481 ccmO 481

CCM 469, 470, 471, 476, 478, 480, 482 chlB 79, 522, 545 chlL 11, 79, 482, 522, 545 chlL (f'rxC) 95 chlN 79, 482, 522, 545, 591, 593 clpB 78 clpP1 79 clpP2 79 cmp 472 cmpA 472, 473, 565 CmpA 473 cmpB 472 cmpC 472 cmpD 472 cobA 522, 529 cobN 548 cox 427 coxA 427 coxB 427 coxBAC 427 coxC 427 cpc operon 193, 198,. 200, 204 cpc1 operon 201 cpc2 operon 201 cpc3 operon 165, 201 cpcA 37, 76, 164, 198, 200, 644 cpcA1 165 cpcA2 165 cpcA3 165 cpcB 37, 76, 164, 198, 200, 203, 601, 644 cpcBl 165, 625 cpcB1A1 165, 627, 645–647, 650 cpcB2 165, 625 cpcB2A2 165, 603, 627, 645, 646, 648, 650–654 cpcB2A2H212D2 645, 833 cpcB3 165 cpcB3A3 646 cpcBA 203, 599, 665, 797, 798 cpcBACDEF 204, 628, 665 cpcBACDEFG 624

850 cpcCD 644 CpcD 133, 150, 283, 390 cpcD 205 cpcD2 646 cpcD3 646 cpcE 200, 202, 536, 537, 646, 735 cpcF 200, 202, 536, 537, 646, 735 CpcG 76 cpcG 78, 197, 200, 203, 205, 625, 644 cpcG 3' 204 CpcG1 197 cpcG1 195, 204 cpcg1-3 195 CpcG2 197 cpcG2 195, 204 CpcG3 197 cpcG3 195, 196 cpcG4 204 cpcH2 646 cpcH3 646 cpcH1 644 cpcI2 646 cpcI3 646 cpcL 78 cpe operon 200, 201 cpeA 33, 35, 37, 644 cpeB 33, 35, 37, 188, 203, 644 cpeBA 35, 36, 200, 203, 627, 630, 634, 645–654, 833 CpeC 184, 647 cpeC 198, 646, 647 cpeCD 630 cpeCDE 202, 627, 630, 644, 647, 652 CpeD 184, 647 cpeD 198, 646, 647 CpeE 184 cpeE 198, 646, 647 cpn60 See GroEL Crp 510 crp 663 crtB 569, 570, 572 crtE 77, 78, 569, 570 crt1 569, 570 crtL 569, 572, 573 crtP 569, 572, 573 crtQ 569, 572 crtY 569 crtZ 569 cryIVD 601 ctaA 522, 548 ctaB 522, 548 CYC3 522, 523 cyoE 522, 532, 548 CysA 663 cysA 662 cysG 522, 529 cysH 725 cysI 529 CysP 663 cysR 663 CysT 662 cysT 123, 663

Gene and Gene Product Index cysU 663 cysV 663 CysW 662 cysW 123, 663 cyt-2 522, 532

D dab 795 desA 710 devA 793 dnaB 78 DnaJ 752, 754, 755, 759 dnaJ 753, 756 DnaK 752–763 dnaK 77, 753, 756, 757, 759, 761, 763, 764

E erm 588 eve 799

F fabH 78 fbcB 263, 269 fbcC 263 fbcF 263, 275 fbcFBC 290 fbcH 290 fdxH 385, 494, 495, 774, 776, 802 fdxN 495, 598, 774, 775, 776 FecA 741 FepA 741 FhuA 741 FhuE 741 fix 795 fixK 663 fox 795 frxC (chlL) 11, 79, 95, 482, 545, 591, 593 fur 740, 743 Fur 743, 746

G gapA 108 gap A ’ 108 gapB 108 gapC 108, 110 gdhA 509 gidA 545 gidB 545 glbN 496, 775 glnA 506, 508, 510, 597, 601, 622, 624, 627, 631, 632, 797, 802, 835 GlnB 511, 835 glnB 78, 95, 508 glnN 508 gltB 78 gltX 522 gnd 414, 624, 632

Gene and Gene Product Index GroEL (cpn60) 443, 752–764, 775 See also chaperone groEL 77, 78, 441, 753, 761–764, 775 GroES (cpn10) 443, 752–754, 755, 758, 760, 763 groES 77, 441, 753, 763 See also GroEL, chaperone groESL operon 441, 622, 626, 710, 764 GrpE 752, 754, 755, 759 grpE 753, 756 gsa 522 gvpA 828 gvpA1 832 gvpA2 832 gvpABC 627 gvpC 828, 832 gvpD 832

H hemA 79, 522, 525 hemB 522, 528 hemC 522, 528 hemD 522, 529 hemE 522, 530 hemF 522, 547 hemG 547, 548 hemH 522, 529, 531 HemL 522, 526 hemN 522, 547 hemY 522, 530, 531 hen 795 hep 795 hepA 598, 599, 793, 795–797, 799, 801, 802, 804, 810 hepB 801 hetA 39, 598, 801 hetC 800, 805 hetN 800, 806 hetP 800, 805, 810 hetR 597, 599, 635, 770, 793, 796–799, 801, 804, 805, 808–810 hgl 795 hglK 802 hisH 79 hlpA 78 hrdA 616 hrdB 622 hrdC 622 hrdD 622 HtpG 754 HtpI 754 hupB 624, 627

hupL 776

I icfA 479, 481 ilvB 78 iphP 661 irpA 42, 745 isiA 43, 626, 665, 738 IsiA protein 739 isiAB operon 737, 745 isiB 43, 387, 737, 738

851 L lacI 603 603 lacZ 276, 472, 586, 596, 602, 628, 702–705, 773 lcy 572 lpcr 522, 544

luxAB 586, 594, 596, 597, 772, 773, 788, 802, 804

M mbpX 95 mbpY 95 metF 602 mob 583, 588 mpeA 36, 37 mpeB 36, 37 mpeC 186, 202 mpeU 203 mpeV 203

N nadA 71, 77, 79 Nar 499, 502 narB 499

nblA 667 ndhA 100, 101 ndhAIGE operon 418 ndhB 284, 418, 472, 476, 482 ndhC 100, 101, 418, 477 ndhD 100, 101, 418 ndhE 100, 101, 418 NdhF 354, 414 ndhF 284, 477, 625 ndhG 100, 101 NdhH 419 ndhH 100 NdhI 419 ndhI 100, 101 NdhJ 419 ndhJ 100, 101, 418 NdhK 419 ndhK 100, 101, 418, 476 ndhK1 418 ndhK2 418 ndhL 284, 418, 476 nif 20, 29, 622, 631 nifAL 785 nifB 386, 495, 774, 775 nifD 40, 492, 545, 598, 774–776, 802 nifE 496, 774, 787

nifF 387 NifH 789 nifH 11, 40, 492, 545, 591, 596, 631, 772, 774, 775, 796, 802 nifH* 775 nifHDK 385, 495–497, 596, 775, 788, 795, 797 nifJ 495, 496, 599, 777 nifK 40, 492, 545, 774, 775, 802 nifN 496, 774, 787

852 nifS 386, 495, 774, 775 nifU 495, 774, 775 nifV 496 nifW 496, 774 nifX 496, 774 Nir 499, 502 nir 498, 500, 502, 510 nirA 529 npt 585 nptII 414 nrtA 59, 473, 498, 565 nrtABCD 502 NrtB 498 nrtB 473 nrtBCD 498 nrtC 473 NrtD 498 nrtD 473 NtcA 631, 660, 803 ntcA 510 ntr 809 NtrC 631

O oli 522 ompR 78 oriT 583, 584, 589 oriV 589

P PatA 798 patA 635, 805 PatB 798 patB 635 pbsA 78 pecA 198 pecB 198 pecBACEF operon 193, 200, 204 pec-cpc superoperon 200 pecE 202, 204 pecF 202, 204 PepB 631 pet 285, 289, 290 petA 75, 260, 263, 275, 276, 288, 289, 303 petB 52, 54, 75, 100, 101, 260, 263, 285, 288, 289, 303, 422 petC 260, 263, 288, 289, 422 petD 52, 54, 75, 100, 101, 260, 263, 285, 288, 289, 303, 422 petE 279, 289, 392 petF 98, 289, 624, 626, 627, 628, 776 petFI 71, 73, 75, 382, 387, 602, 624, 737 PetG 262, 263, 279 petG 75, 263, 279, 289 PetH See ferredoxin: oxidoreductase petH 283, 289, 390, 416, 417, 625 petI 737 petJ 78, 289, 394, 395 petK 78, 79, 289, 397 petP 290

Gene and Gene Product Index petR 290 PhoA 661 phoA 269, 603 PhoB 40, 41, 45 phoB 37 PhoR 40, 41, 44 phoR 37 pknA 798 ppc 453, 461, 624, 779 preA 77 prk 458, 461 ps2B 696 PsaA 128, 325, 327, 329, 330, 336, 688 psaA 74, 100, 107, 128, 412 psaA1 625 psaA2 625 psaAB 687 PsaB 128, 325, 327, 330, 336, 688 psaB 74, 128, 329 PsaC 128, 130, 325, 328, 333, 343, 345–352 psaC 71, 74, 100, 101, 343, 383, 418, 625 PsaC1 343 PsaC2 343 PsaD 128, 130, 325, 328, 333, 346–350 PsaE 128, 130, 133, 261, 283, 284, 325, 328, 346, 349–352 psaE 78, 107, 284, 625 PsaF 128, 328, 337 psaF 74, 337, 340, 394, 395 PsaG 324 PsaH 324 PsaI 128, 339 psaI 74, 95 PsaJ 128, 328, 339, 340 psaJ 74, 337, 340 PsaK 128, 327, 339, 341 psaK 341 PsaL 128, 327, 339, 342 psaL 78, 342, 683 PsaM 128, 323, 328, 343 psaM 75, 343 PsaN 324 psb1 697 PsbA 130 psbA 51, 52, 54, 75, 103, 105–107, 111, 219, 232, 592, 594, 596, 603, 605, 627, 629, 630, 693–696, 699–710 psbA-1 699 psbA-2 699 psbA-3 699 psbA1 625, 696 psbA2 696 psbA3 696 psbAI 624, 626, 628, 629, 633, 694, 696, 699, 701–705, 707, 708 psbAII 624, 626, 629, 696, 701–705, 707, 708 psbAIII 624, 626, 629, 696, 701–705, 707, 708 psbAIV 624, 696 PsbB 130, 387 psbB 52, 54, 75, 100, 101, 219, 235, 289, 595, 596, 624, 627, 628, 696, 697, 797 PsbC 130, 223 psbC 75, 100, 219, 223, 595, 665, 696, 698, 704, 738 PsbD 130

Gene and Gene Product Index psbD 75, 100, 107, 219, 235, 596, 602, 620, 630, 693, 694, 698, 704,706 psbD1 625, 696 psbD2 625, 696 psbDC operon 698 psbDI 595, 626, 696, 698, 704, 705 psbDII 626, 628, 629, 698, 704, 705 psbE 75, 100, 219, 231, 698 PsbE-PsbF 130 See also cytochrome b559 psbEF 696 PsbF 131 psbF 75, 102, 219, 231, 698 psbG 418 PsbH 131, 228, 696 psbH 52, 54, 75, 100, 101, 228, 229, 289, 696, 698 PsbI 227, 696, 698 psbI 229, 696, 698 PsbJ 697 psbJ 75, 229, 230, 697, 698 PsbK 228, 341 psbK 75, 228, 229, 697,698 PsbL 131 psbL 75, 228, 229, 697, 698 psbM 228, 229 psbN 75, 100, 101, 228, 229, 289, 697, 698 PsbO 124, 219–223 See also woxA psbO 219, 220–223, 695, 699 psbP 219 psbQ 219 psbR 229, 230 PstS 40 pstS 36, 37, 42 psy 572 purE 472 purEK 472 purK 472, 481

R rbc 772 rbcL 52, 54, 76, 95, 103, 106, 441, 449, 457, 460, 624, 627–629, 631, 633, 634, 774 rbcLrbcS 441 rbcLS 469, 472, 473, 479, 480, 482, 496, 597, 603, 797, 803 rbcLS operon 480 rbcS 52, 54, 76, 77, 103, 106, 441, 449, 457, 460, 480, 774 rca 456, 457, 458, 460 RcaA 630, 631, 652, 833, 839 RcaB 630, 631, 652, 833 RcaC 839 rcaC 589, 635, 645, 646, 651, 654 RcaD 833, 839 recA 622, 625, 764 RegA 290 rhdA 603, 663 rotA 482 rpcE 202 rpcF 202 rpeB 102, 203 rpl1 73 rpl2 81

853 rpl3 73 rpl5 74 rpl6 74 rpl18 74 rpl19 73 rpl20 73 rpl21 73 rpl22 81 rpl24 73 rpl33 73, 81 rpl35 73 rpl36 73 rplKAJL 96, 620 rplQ 621 rpmJ 621 rpoA 73, 621 rpoB 76, 620 rpoBC1C2 621 rpoC 620 rpoC1 52, 54, 76, 81, 99, 103, 620 rpoC2 76, 620 RpoD 754 rpoD 621 rpoD1 622 rpoH 752 rps2 73, 76 rps4 71, 73 rps5 74 rps7 81 rps9 73 rps10 73 rps11 73 rps12 81, 98, 102 rps13 73 rps14 73 rps17 74 rps18 73, 81 rps18/rpl33 71 rps19 81 rpsD 621 rpsK 621 rpsM 621

S sacB 584, 586, 593, 773 Sbp 662 sbpA 123, 662 SecA 276, 760 secA 80 SecB 760 secD 760 SecE 760 secE 760 SecE/SecY 760 secF 760 SecY 82, 760 secY 80, 96, 760 SigA 62l sigA 621, 624, 627, 797, 836 SigB 632

Gene and Gene Product Index

854 sigB 621, 622, 797, 836 SigC 632 sigC 621, 797, 836 sigG sigK smtA smtB sodB

799 799 43 43 710

spc operon 96 sphR 40 sphS 40

str operon 98, 102

T tln2 797 tln6 796 TonA 741 TopA 482 tra 584 trnA 70 trnC 71 trnE 524

U unc operon 364 Uncl 364

V vnf 589 vnfD 775 vnfDGK 496 vnfK 775

W woxA 37, 585, 624, 627, 697, 699 See also psbO

X xisA 631, 775, 776, 802 xisC 776 xisF 775

Y y-1 544

trnI 70

Z zds 572 zwf 779 trpA 78 trpG 79 trsA 78 trxA 78 tsf 76, 78 tufA 73, 81, 103, 107, 111

Subject Index A cytochrome c oxidase 410, 426, 428 See also cytochrome c oxidase ABC transporters See ATP-binding cassette transporters acetohydroxyacid isomeroreductase 525 acetolactate synthase 78 acetosyringone 838 acetyl coenzyme A carboxylase 802 N-acetylgalactosamine 191 acetylglutamate kinase 78 N-acetylglutamate semialdehyde dehydrogenase 509 N-acetyl muramic acid 68 acetylornithinase 509 acetylornithine 5-aminotransferase 509 N-acetyl putrescine 69 action spectrum 645, 648, 679, 830 acyl carrier protein (ACP) 78 adaptive responses 43 adenine-nucleotide binding cassette family 371 See ATP-binding cassette transporters adenosine-5'-phosphosulfate (APS) sulfotransferase 661 adenosine 5'-diphosphate (ADP) 368 adenosine 5'-triphosphate See ATP adenosine triphosphopyridoxal 371 adenosylcobalamin 726 adenylate cyclase 809 adenylate kinase 371, 372 adenylylation 508 ADH See alanine dehydrogenase aerobactin 741 See also siderophore affinity chromatography 524, 536 agar, soft 55 akinetes 6, 771–773, 799, 805, 810, 826 ALA See levulinic acid alanine 455 D-alanine 68 alanine dehydrogenase (ADH) 492, 508, 791 algae 92, 93, 95, 96, 100, 102, 105, 106–112 alkaline invertase 779 allophycocyanin (APC) 32, 76, 147, 152–154, 158, 183, 190, 641, 645, 666, 798, 833 See also allophycocyanin B, phycobiliprotein, phycobilisome subunit 152, 205 subunit 152 apc operons 200, 205 subunit 152, 153, 173, 205 See also ApcB amino acid sequences 153 subunit 76, 152, 155, 205, 683 See also ApcF core subcomplexes 160, 193 energy transfer 161 general structure 154 sequence homology 152–153 allophycocyanin B 127, 152,154, 162, 205, 681, 683 See also allophycocyanin, ApcD, phycobiliprotein, phycobilisome subunit 127, 152, 154, 162, 205, 681, 683 mutants 155, 683

sequence homology 152–153 alpha operon 96 alternative nitrogenases See nitrogenase alternative oxidase 427 amino acid assimilation 38 aromatic amino acid biosynthesis 7 sequence 153, 156, 157, 164, 166, 170, 178–180, 183, 188, 194, 195, 198, 366 sequence motifs 197 sequencing 95, 265 sidechains 368 transport 503 L-amino acid oxidase 699 aminoglycoside 3'-phosphotransferase 585 levulinic acid (ALA) 488, 509, 521–527, 535 levulinic acid dehydratase 527 levulinic acid synthase 521, 525 ammonium 488 assimilation 455, 505 inhibition 501 transport 504 -promoted inhibition 502 -promoted repression 497, 502 amphipathic helix 268, 269 anchor polypeptide See ApcE, core membrane linker phycobiliprotein, phycobilisome angiosperms 9, 543–545 anoxygenic photosynthesis 19 anoxygenic phototrophic bacteria 453 antenna protein 53, 56, 142, 560, 679, 680, 689 See also carotenoid-binding protein, light-harvesting chlorophyll protein, phycobiliprotein, phycobilisome anthranilate synthetase 510 antibiotic resistance genes 585 antimycin-A 131, 268, 280, 283–285, 288, 293, 297–299, 422 See also Cyt complex, Cyt complex antisense RNA 832 Apicomplexa 109–111 appressed membranes See membrane, thylakoid APS See adenosine-5'-phosphosulfate AquI See restriction endonuclease arginase 504 arginine 503, 509, 791, 809 transport 504 arginine deiminase 504 aromatic amino acid biosynthesis 7 ascorbate 534 ascorbate peroxidase 123, 129, 132 aspartate 809 aspartate aminotransferase 526, 527 aspartate transcarbamylase 510 astaxanthin See carotenoids ATP 286, 362, 369, 373, 410, 411, 412, 414, 428, 524, 530, 678, 686, 688, 689 ATPase 95, 97, 100, 718, 757 See also ATP hydrolysis, ATP synthase activity induction of activity by organic acids 375

856 ATP-binding cassette (ABC) transporters 371, 805 multicomponent transport systems 498 ATP hydrolysis 368, 371, 375, 757 See also ATPase, ATP synthase activating anions 375 ATP synthase 57, 76, 94, 99, 105, 119–121, 123, 125, 129, 132– 134, 280, 361–376, 623, 686, 761 active state 373–375 activation by or acid-base transition 373 adenine binding 371 adenosine ring binding sites 371, 372 adenosine triphosphopyridoxal modification 371 subunit 362–366, 372, subunit 2, 362–366, 369, 370, 371 binding affinity 369, 371 binding-change mechanism 368, 369 L-bidentate metal-ADP isomer 366 bidentate metal-ATP complex 367 bovine mitochondrial ATPase 367 structure 364 catalytic ATP binding sites 369 model 370 catalytic site 367, 369, 371, 372, 376 competitive inhibition 372 coupling factor ATPase 364–370, 373–375 119, 123, 129, 130, 132, 134 129, 132, 366, 373–375 subunit 362–366 disulfide bond, subunit 373 reduction 374 divalent cation cofactors 366 electrochemical proton gradient 362, 367, 373–375 electron micrograph 364 subunit 369, 373 fluorescent ATP derivative (TNP-ATP) binding 368 F type ATPase 362–376 123 complex 123, 363 ATPase 362, 366, 368, 376 complex 362, 363, 370, 375 stoichiometry 363 subunit 362–366, 368, 373–376 sulfhydryl groups 373, 374 proteolysis 374 induction of ATPase activity by organic acids 375 inhibition by diadenosine tetraphosphate 372 latent state 373–375 metal binding site, subunit 371 metal-nucleotide binding site 369–371 model 373 noncatalytic 369, 372 metal-nucleotide complex 367, 369, 375 structure 367 monodentate metal-ADP complex 367 monodentate metal-ATP complex 369 nucleotide binding sites 363, 370 organization of subunits 363–364 P-loop 371, 373 photoaffinity labeling 371 rat mitochondrial ATPase crystal structure 363, 372 structure 363, 372 reaction mechanism 368

Subject Index reaction mechanism 368 subunit a 363, 364, 366 subunit b 363, 364, 366 subunit b' 363, 364, 366 subunit c 363, 364, 366 subunit I 366 See subunit b subunit II 366 See subunit b' subunit III 366 See subunit c subunit IV 366 See subunit a tight-binding ADP 375 tridentate metal-ATP complex 366 epimer 367 epimer 367 uncomplexed metal inhibition of activity 375 unprotonated amine role in metal-nucleotide binding 369 unisite catalysis 368 vanadyl binding site 369 Walker B consensus region, subunit 372 ATP synthesis binding-change mechanism 368, 369 direct vs. indirect coupling 368 mechanism 366, 367 role of electrochemical proton gradient 362 atrazine 697, 710 autophosphorylase activity DnaK 757 AvaI See restriction endonuclease AvaI methylase 588 AvaII See restriction endonuclease AvaIII See restriction endonuclease axenic strains 55 axial ligands, Cyt b 267, 276 azaserine 504, 505, 508 DL-7-azatryptophan 455, 806 azetidine-3-carboxylic acid 829 azide inhibitor of uptake 426

B b cycle 272, 282 model 272 b heme 262, 266, 281, 293, 294 heme 268 heme 268 b type cytochromes See cytochromc b bacteriochlorophyll (Bchl) 142, 538, 540, 546 bacteriochlorophyll a 546 bacteriochlorophyllide a 546 bacterioferritin 42 baeocyte 5, 6, 20, 826 Bangiophyceae 177 See also red algae barley translational control of PsaA and PsaB synthesis 688 bathophenanthroline 427 inhibition of cyanide-insensitive uptake 427 Bergey’s Manual of Systematic Bacteriology 5 bicarbonate 469, 470, 473, 679, 685, 686 symport 474 chemical potential gradient 475 pump 474

Subject Index uptake 475 role of 474 role of 42-kDa polypeptide 471–473 See also CmpA bilin See phycobilin biliverdin 148, 533–537 biliverdin 15, 16-reductase 522, 536 biosynthesis amino acid 95 aromatic amino acids 7 carotenoid 95, 567–571 chlorophyll 95, 520–548 fatty acid 108 glutamine 95, 504–508 phycobilin 95, 533–536, 735 pigment 108 tetrapyrroles 520–548 biotin carboxyl carrier protein 802 bleaching 664 See also chlorosis bleomycin resistance (ble) 588 blue light 706, 830 blue-green algae 55 bootstrap analysis 14, 104, 106 boron 802 boundary layer role in Ci uptake 476, 479 bovine mitochondrial ATPase 367 structure 364 bryophytes 837

C cI represser 602, 799 cadmium resistance 43 calcium 225 calcium-binding protein 223 calcium-phosphate chromatography 264 calmodulin 809 immunolocalization 809 caloxanthin See carotenoids. Calvin reductive pentose phosphate pathway 438, 457–460, 477 gene arrangement in chemoautotrophic bacteria 458 transcriptional activator for genes of 458 cAMP See cyclic adenosine 3', 5'-monophosphate canavanine 806 canthaxanthin See carotenoids carbamyl phosphate synthase/synthetase (CPS) 439, 455, 510 occurrence in vegetative cells and heterocysts 455 carbon adaptation to ambient levels 471 adaptation mutants 481 assimilation 43 C-deficiency 656 Anabaena variabilis 656 Spirulina platensis 656 Synechococcus 656 Synechococcus lividus 656 inorganic (Ci) 470 accumulation 473 boundary layer, role in uptake 476, 479 -concentrating mechanism 469 efflux (slippage of pump) 476 energy dependence of Ci uptake 470, 476 influx 476

857 transport 473, 474 isotopic fractionation 471 role of symport 474 transporters 125 carbon dioxide 411, 437–462, 469, 473, 474, 792 -dependent promoters 472 diffusion 478 fixation 678, 686 mutants 469, 471, 480, 482 limited chemostat cultures 456 growth 473 metabolism 438–462 scavenging system 476 uptake 475 adaptation to ambient levels 471 mutant SC 477 role of 42 kDa polypeptide 471–473 See also CmpA carbon monoxide (CO) 420, 426, 541 carbonic anhydrase 43, 452, 469, 470, 475, 479, 481, 792 mutant C3P-O 479 role in trans-inhibition 479 carbonylcyanide m-chlorophenyl-hydrazone (CCCP) 683 2–carboxyarabinitol monophosphate (CA1P) 456 carboxylation 472 439, 453 DD-carboxypeptidase 69 LD-carboxypeptidase 69 carboxysomes 43, 51, 52, 58, 66, 438, 469–471, 477–485, 663, 831, 838 cardiolipin 265 carotene 560 See carotenoid hydroxylase 569, 570 See carotenoid See carotenoid desaturase 569, 570, 572 carotenoids 2, 7, 560–575 absorption properties 562, 563 astaxanthin 563 bandshifts 282 biosynthesis 132, 567–574 caloxanthin 563, 565 canthaxanthin 563, 565 128, 559–567, 574, 665 563 563, 569 563 myxoxanthophyll 563, 564 chemical structure 565 caloxanthin 565 canthaxanthin 565 echinenone 565 myxoxanthophyll 565 nostoxanthin 565 oscillaxanthin 565 zeaxanthin 565 composition Anabaena aerulosa 567 Anabaena cylindrica 567 Anabaena flos-aquae 567 Anabaena variabilis 567 Aphanizomenon flos-aquae 567

858 Aphanothece sp. 566 Arthrospira sp. 566 Calothrix parietina 567 Chlorogloea fritschii 567 Coccochloris elabens 566 Cylindrospermum sp. 567 Mastigocladus sp. 567 Merimopedia punctata 566 Microcoleus paludosus 566 Microcoleus vaginatus 566 Microcystis aeruginosa 566 Nostoc commune 567 Nostoc sp. 567 Oscillatoria agardhii 566 Oscillatoria amoena 566 Oscillatoria limosa 566 Oscillatoria princeps 566 Oscillatoria rubescens 566 Oscillatoria tenuis 566 Phormidium foveolarum 566 Phormidium laminosum 566 Phormidium luridum 566 Phormidium persicunum 566 Spirulina geitleri 566 Spirulina maxima 566 Spirulina platensis 566 Synechococcus elongatus 566 Synechococcus sp. 566 Synechococcus sp. PCC 7942 566 Tolypothrix tenuis 567 echinenone 563 inhibitors 572–573 lycopene 563, 569 neurosporene 563, 569 oscillaxanthin 563 phytoene 562, 563 15-cis-phytoene 569 phytofluene 563, 569 specific absorbance coefficients 563 thin-layer chromatography 562 triplet state 561 violaxanthin 561 xanthophyll 130, 142, 560, 561, 562, 564, 565, 567, 571, 573, 574, 575, 579, 665 zeaxanthin 53, 56, 57, 123, 561, 565, 665 zeaxanthin-binding protein 566 carotenoid-binding protein 53, 123, 124, 566, 745 role in blue-light-shielding 53 zeaxanthin-binding protein 566 carotenoprotein See carotenoid-binding protein catalase 788 cell cycle developmental 825, 827, 830 cell division 827, 830 cell membrane 409, 413 cell motility 825 cell wall 58 centrifugation filtering 474, 475 Chamaesiphonales 6 chaotropic agents 223, 279, 348, 351 chaperone 751–765 role in membrane-protein insertion 759–762

Subject Index chaperonin 288, 443, 753, 755, 760, 775 cpn60 14, 443, 753 See also GroEL, chaperone, heat shock electron microscopy 753 immunocytochemistry 762 Charophyceae 96 chemotaxonomy 5 carotenoids 7 lipid composition 5 pigment composition 5, 10, 17 chimaeric genomes 93, 107, 111 chloramphenicol 686, 701, 707, 831, 832 chloramphenicol acetyl transferase Also see cat, CAT 472, 588, 596 chlorin 543 p-chloromercuribenzenesulfonate 538, 540 p-chloromercuribenzoate 540 2–(4-chlorophenylthio)-triethylamine hydrochloride (CPTA) 573 See also lycopene cyclase chlorophyll 50–57, 62, 92, 93, 95, 103, 142, 265, 428, 520, 524, 531, 538, 541, 544–546, 678, 679, 681–688 absorption 735 -binding protein 54, 738, 745 See also Chl a/b protein complex, Chl-protein complex, light-harvesting chlorophyll protein biosynthesis 411, 520–531, 538–548 Chl a 3, 321, 538, 540, 542, 546 dimer 333 Chl a' 333, 546 epimer = Chl a' 546 51 Chl a/b-protein complex 8, 50, 53–59, 60–62, 131, 142, 266, 680, 688, 761 Chl a/b-antenna complexes 54, 56 See also chlorophyllbinding protein, light-harvesting chlorophyll protein trimeric association 53 Chl a/c light-harvesting complex 143 inhibition by o-phenanthroline 547 Chl b 50–57, 61, 62, 70, 538, 540, 546, 547, 695 synthesis 53 Chl 51 Chl c 51 Chl-protein complex 8, 50, 53–59, 60–62, 131, 142, 143, 266, 680, 688, 761 CP1 341 CP43 218, 219, 223, 387, 397, 696, 698, 706, 738, 739 See Photosystem II, PsbC CP47 218, 219, 224, 397, 696, 761 See Photosystem II, PsbC, CP47 chlorophyll a-binding polypeptide 697 divinyl-Chl a 51 donor 218 isocyclic ring 538, 540, 548 triplet state 561 -xanthophyll binding proteins 130 chlorophyll 3-methyl oxygenase 522 chlorophyll synthetase (isoprenyltransferase) 522, 545 chlorophyllide (Chlide) 538, 543, 546 Chlide a 545, 547, 688 Chlide b 545, 547 Chlorophyta 92, 93 chlorophyte 92, 93, 96, 103, 111 chloroplast 66, 261, 264, 418, 680, 686, 706 See also cyanelle, plastid autogenous origin 91

Subject Index endoplasmic reticulum 92, 109 genomes 93–95, 203, 204 See also Cyanophora paradoxa, Porphyra purpurea Astasia longa 95 conifer 545 Epifagus virginiana 94, 95 liverwort 93 tobacco 93–95 rice 93 inverted repeat 94, 96, 105, 106 monophyletic origin 92, 93, 95, 97, 105, 109, 111 phagotrophic host(s) 93, 94, 111 polyphyletic origins 92, 93, 105, 107, 111 proto-chloroplast 53 xenogenous origin 91 chloroplast coupling factor ATPase 364–370, 373–375 119, 123, 129, 130, 132, 134 129, 132, 366, 373–375 chlororespiration 52, 95, 285 chlorosis 546, 664 See also bleaching chlorosome 142, 600, 773 chromatic adaptation See complementary chromatic adaptation chromatography 7 gel exclusion 266 HPLC anion-exchange 265 HPLC-hydroxyapatite 265 Chromophyta 92, 93 pre-chromophyte alga 93 chromosome, map Anabaena sp. strain PCC 7120 600 Synechococcus sp. strain PCC 7002 600 Synechocystis sp. strain PCC 6803 600 Chroococcales 6 circadian rhythm 461, 694 circular dichroism (CD) 53, 448 citric acid cycle 414, 778 See also tricarboxylic acid cycle citrulline 439, 455, 504, 509 cluster analysis 30 coactivator 619 cobaltochelatase 548 codon usage 36–38 Marine Cluster A Synechococcus sp. 36–38 coenzyme 726 cold shock 829 competency 590, 604 See also transformation complementary chromatic adaptation 19, 151, 164, 165, 176, 190, 201, 202, 589, 630, 635, 649, 643–654, 684, 688, 826, 833– 835, 839 action spectrum 645, 648 FdB mutants 650, 653 FdG mutants 650–653 FdR mutant 650–653 model 652, 653 complementation 302, 451, 667 complex III See cytochrome complex conjugation 582–590, 593, 604, 605, 773 RP-4 mediated 773 copper 392–395, 410, 423, 742 coproporphyrinogen III 530 coproporphyrinogen IV 530 coproporphyrinogen oxidase 522, 530–532, 547, 548 HEM13 mutant 522, 530 oxygen-dependent 547

859 oxygen-independent 548 copy-correction mechanism 96, 105 core-membrane linker phycobiliprotein ApcE) See allophycocyanin, ApcE, linker, core-membrane linker phycobiliprotein phycobilisome corrins 520, 529 cosmid 586 coupling factor ATPase 364–370, 373–375 See also ATP synthase 119, 123, 129, 130, 132, 134 129, 132, 366, 373–375 cpn60 See chaperonin CPTA See 2–(4-chlorophenylthio)-triethylamine hydrochloride cross-linking studies 221, 272, 278, 338, 349 Crp family 631 cryptomonad (cryptophyte) 92, 96, 102, 108, 111, 140, 143, 146, 148, 176, 177, 183, 186, 188–190, 203, 204, 210, 211, 534, 536, 537 See also Cryptophyta biliprotein subunits 184, 190 chloroplast 189 Cryptophyta 92, 93, 204 See also cryptomonad cryptoviolin 148. See phycobiliviolin (PXB) See carotenoid crystals 165, 172, 180, 286, 287, 326, 346, 383, 388, structure 171, 173, 176, 181, 185, 186, 286, 287, 326, 346, 363, 365, 372, 383, 384, 388, 389, 391, 393, 395, 441, 442, 493, 507 cucumber 538–542 cyanelle See also Cyanophora paradoxa 15, 16, 19, 66, 92, 95, 97, 98, 100, 102, 103, 104, 105, 107, 111, 263, 279, 460, 539, 545 chromosome map 72 cyanide 283, 414, 420, 424, 426, 427, 541 -insensitive alternative terminal oxidase 427 -insensitive NADPH oxidation 415 -insensitive uptake inhibition by bathophenanthroline 427 -insensitive respiration 411, 427, 428 -insensitive terminal oxidase 284, 413 -sensitive alternative terminal oxidase 413, 427 -sensitive cytochrome c oxidase 426 -sensitive NADPH oxidation 415 -sensitive terminal oxidase 420, 427 cyanoglobin 496 cyanophage 33–35 host resistance 35 Cyanomyovirus 34, 35 Cyanopodovirus 34 Cyanostylovirus 34, 35 Myoviridae 34 Styloviridae 34, 35 N-1 623, 624, 627 cyanophycin 29, 488, 510, 664, 791, 799, 809, 831, 838 granules 838 cyanophycinase 809 cyanophycin synthetase 809 cyanorubrum 479 cycad 8, 9, 837, 838 cyclic adenosine 3', 5'-monophosphate (cAMP) 510, 806, 809 receptor protein 510 cyclic electron transport 226, 280, 283, 284, 323, 342, 343, 347, 354, 398, 495, 678, 682, 683, 689, 776, 781, 782 cyclic phosphorylation See cyclic electron transport

860 cytochrome (Cyt) 284, 732 alternative 284 cytochrome 420, 426, 532 cytochrome oxidase 261 cytochrome b 263, 264, 266–269, 279, 280, 285, 290–294, 296, 297, 299, 300 axial ligands 267, 276 redox centers 267 cytochrome 298, 299 cytochrome 75, 260, 262, 264, 266–268, 270, 275, 279–281, 284, 287–289, 291, 296, 297, 299, 303, 416, 422, 680, 687– 689 amphipathic helix 268, 269 axial ligands 267, 276 b cycle 272, 282 model 272 b heme 262, 266, 281, 293, 294 heme 268, 298, 299 heme 268 bovine mitochondria 267 hydropathy analyses 267, 269 interheme electron transfer 282 model 268 redox centers 267 split Cyt b protein 264 subunit IV 268–270 cytochrome complex 260–264, 266, 272, 279, 281, 282, 285, 288, 289, 291, 294, 296–298, 300, 303, 422 aggregation state 266, 279 cytochrome 298, 299 dimer 266 inhibitors 268, 269, 280, 281 inhibitor resistance mutations 269, 280, 282, 291, 296, 297 interheme electron transfer 282 precursor polyprotein in B. japonicum 290 supercomplex 266 yeast mitochondrial 266 cytochrome complex 75, 94, 119, 120, 121, 123, 125, 129, 130, 131, 133, 134, 259–304, 352, 362, 409, 412, 413, 416, 420–423, 425, 427, 678, 680, 682, 686, 687, 688, 689, 733, 781, 782, 786 aggregation state 266, 284 biogenesis 286, 288, 289 dimers 266, 284 duroquinol-plastocyanin oxidoreductase activity 264 hydropathy analyses 267, 274, 275, 276, 279 inhibitors 268, 269, 280, 281 inhibitor resistance mutations 269, 280, 282, 291, 296, 297 isolation 264 monomer 266 mRNA processing 288 oxidant-induced reduction of Cyt b 281–284 plastoquinol-1-plastocyanin oxidoreductase activity polypeptides 264 Q cycle model 281 site 265, 275, 280, 281, 296, 297, 299, 300 site 275, 280, 281, 298 quinone-depleted complexes 265 redox centers 264 Rieske-depleted complex 280 substrate binding sites 264 subunit IV (SU IV) 260, 262–270, 272, 275, 279–281, 283, 287–289, 291, 296, 297, 299, 422

Subject Index relationship to cytochrome 268–270 subunit topology 264 supercomplex 266 thermophilic Synechococcus sp. 265, 283 tightly-bound chlorophyll in 265 topographical model 263 cytochrome 75, 123, 130, 132, 218, 219, 223, 227, 231, 242, 696, 698, 782 stoichiometry in PS II 231 cytochrome 283, 362, 397, 413, 422 cytochrome c 69, 264, 276, 277, 293, 294, 388, 393, 398, 411, 416–418, 420–423, 425, 426, 532 c-heme 262, 281, 289, 293 horse heart 288 low-potential 396–398, 401 See cytochrome reduction 386 cytochrome c heme lyase 532, 548 cytochrome c oxidase (COX, Cox) 53, 119, 121, 123, 125, 126, 129, 261, 266, 285, 410, 413, 420, 423–428, 531, 532, 688, 732, 786, 787 See also cytochrome cytochrome c oxidase subunit structure 427 cytochrome c reductase 392, 423 cytochrome 279, 659 cytochrome 69, 263, 264, 275, 276, 277, 279, 289, 290 cytochrome 69, 266, 279, 659 cytochrome 11, 289, 325–328, 337, 338, 349, 389, 393–396 See also cytochrome cytochrome 396, 397, 425 cytochrome 78, 218, 225, 226, 229, 285, 289 cytochrome 11, 74, 78, 129, 130, 260, 276–278, 281, 289, 303, 394, 410, 413, 416, 421–426, 659, 786, 787 See also cytochrome cytochrome f 75, 260, 262–264, 269, 275–279, 281, 283, 286– 289, 298, 303, 392–395, 416, 420, 422–425, 532 flash-induced oxidation-reduction 687 hydropathy analyses 276 midpoint potentials 277 plastocyanin affinity-binding studies 278 transmembrane topology 276 cytochrome o oxidase 427 cytochrome P450 775 cytoplasm 413, 470, 475, 686 cytoplasmic membrane (CM) 53, 119–126, 131–133, 261, 288, 411, 413, 418–428, 471, 474, 475 See also plasma membrane, plasmalemma

D tyrosine 218, 237, 239, 240, 244 D1 protein 51, 52, 56, 103, 105, 130, 218, 219, 223, 224, 227, 232, 233, 236–238, 560, 693–696, 706–708, 710, 761 See also PsbA effects of blue light on transcription of psbA genes 706 Form I, D1 (PsbA) protein 707 Form II, D1 (PsbA) protein 705, 707, 708 insertion/deletion at C-terminus 103, 105 light wavelength regulation of expression 706 psbA transcript stability 701–702 pre-protein 695 processing protease 233 signal transduction pathway for psbA expression 706 D2 protein 56, 130, 218, 219, 223, 224, 227, 235–238, 693–696,

Subject Index 698, 705–708, 761 See also PsbD DAPI See 4'-6-diamidino-2–phenylindole DBMIB See 2, 5-dibromo-3-methyl-6-isopropyl-p-benzoquinone DCCD See N, N'-dicyclohexylcarbodiimide DCMU See 3–(3, 4-dichlorophyenyl)-1, 1-dimethylurea DCPIP See 2,6-dichlorophenolindophenol 421 denitrifying bacteria 261 deoxyribonucleic acid See DNA dephosphorylation 60, 680, 682, 826, 833 destacking See state transition, thylakoids detachment model See state transition detergents 56 digitonin 57, 223, 226 dodecyl extraction 56, 59, 129, 223, 227, 265 extraction 264 MEGA-9 264 129, 227, 264, 373, 375 octyl thioglucoside 227 sodium cholate 264 sodium dodecyl sulfate (SDS) 56, 224 Triton X-100 226, 227 Triton X-114 52 Zwittergent 14 56 deuteroporphyrin IX 531, 538 developmental cell cycle 825, 827, 830 developmental control 788 diadenosine tetraphosphate 372 4'-6-diamidino-2–phenylindole (DAPI) staining 70, 109 3,3'-diaminobenzidine (DAB) 126, 795, 800 4,5-diaminovaleric acid 526 diaphorase 416 2, 5-dibromo-3-methyl-6-isopropyl-p-benzoquinone (DBMIB) 265, 266, 271, 272, 280, 284, 285, 417, 418, 420, 421, 422, 683,781, 834, 835 dicarboxylic acid transport 617 2,6-dichlorophenolindophenol (DCPIP) 421, 709 3-(3, 4-dichlorophyenyl)-1, 1-dimethylurea (DCMU) 280, 284, 297, 421, 428, 476, 697, 710, 831, 834, 835 dicistronic message 698, 706 N, N'-dicyclohexylcarbodiimide (DCCD) 371 DiCys-bilin See phycobilin didemnids 50 diethyl sulfate mutagenesis 592 diethylpyrocarbonate 534 differentiation 825, 826, 829, 830 See also akinete, heterocyst, hormogonium digitonin See detergent 15,16-dihydrobiliverdin 148, 188, 189, 190, 533, 536 15, 16-dihydrobiliverdin 2,3-reductase 522 affinity chromatography 536 dihydrogeranylgeranyl 545 dihydroporphyrin 328 dimethylallyl pyrophosphate 569 dimethyl sulfoxide (DMSO) 285 1,3-dimethyluroporphyrinogen dehydrogenase 522 dinitrogen See nitrogen dinitrogenase See nitrogenase dinitrogenase reductase See nitrogenase dinoflagellate 92, 93, 109, 111 Dinophyta 92, 93 See also dinoflagellate dioxygenase 271, 273

861 diphenyl ether herbicide 531, 548 diphenylcarbazide (DPC) 420, 421, 709 dipole-induced dipole (resonance) energy transfer See energy transfer, Förster dipyrrole cofactor 528 dis-equilibrium experiments Ci uptake 474 disulfide bond, subunit ATP synthase 373 reduction 374 dithiothreitol (DTT) 373, 374, 476, 530, 541 diuron See 3-(3, 4-dichlorophyenyl)-1, 1-dimethylurea (DCMU) 280, 297 divinyl-chlorophyll a 51 See also chlorophyll divinylprotochlorophyllide (Mg-2,4-divinylpheoporphyrin 539, 540, 542, 544, 547 divinylchlorophyllide b 547 DNA (deoxyribonucleic acid) amplification fingerprinting (DAF) 10 anti-DNA antibodies 109 base composition 5, 9, 19, 20, 30, 103 binding 95 content 30 -DNA hybridization 3, 4, 10, 12, 17, 20 fingerprinting 10 highly iterated palindromic sequence (HIP1) 43, 44 octameric palindrome 43, 44 mobility-shift 629 modification 31 palindromes 43, 44 588 repetitive sequences 599 short tandemly repeated repetitive (STRR) sequences 4, 10, 648 substitution rate nucleotide 103 substitutional bias 103 topology 482 transfer system 55 DNP-INT See 2–iodo-6-isopropyl-3-methyl-2', 4, 4'-trinitrodiphenyl-ether docking protein, SecA 760 dodecyl See detergents See detergents, dodecyl Dollo parsimony 30 double-crossover 585 See recombination DTT See dithiothreitol duroquinol 264, 424 duroquinol-plastocyanin oxidoreductase 264

E echinenone See carotenoid EDDA See ethylene diamine di-o-hydrophenyl acetic acid electrochemical gradient 683 electrochemical membrane potential 473 hyperpolarization of 474, 475 electrochemical proton gradient 362, 367, 373–375 electrogenic pump 474 electron micrograph 151, 157, 196, 364 electron microscopy 58, 109, 143, 180, 198, 265, 324, 325, 364, 374 freeze-fracture 59, 131, 134 fracture faces 57 electron nuclear double resonance (ENDOR) spectroscopy 239,

Subject Index

862 241, 243, 244, 247, 272, 369 electron paramagnetic resonance (EPR) spectroscopy ATP synthase 372 cytochrome 267, 282 cytochrome 276 Photosystem I 334, 336, 346, 349, 350 Photosystem II 237–239, 241, 243, 246, 247 g = 4.1 signal 241–243, 245, 247 Rieske protein 265, 270, 271, 279, 280, 282 spectra 385 spin-polarized signal in Photosystem I 334 spin-relaxation studies 275 electron spin-echo envelope modulation (ESEEM) 241, 243, 272, 369 hyperfine couplings, PS II 243 electron transfer 280, 362 electron transport 31, 95, 108, 289, 303, 409–429, 476, 477, 683, 687, 688, 732, 826, 839 alternative pathways 262, 284 chain 683, 835 inter-heme 281 molecules 689 noncyclic 281, 352, 495, 678, 689 pathways 262, 284 respiratory 288, 409, 410, 424, 426, 428, 732, 778 electroporation 33, 589, 591, 604, 605 elongation factor 2, 74, 78, 107 G 74 Ts 78 Tu 2, 74, 371 endopeptidase 69 ENDOR See electron nuclear double resonance spectroscopy endospore formation 615 endosymbiont 92, 93, 94, 95, 107, 108, 109, 110, 111 endosymbiont theory 49, 50, 53 endosymbiosis 66, 67, 92, 93, 95, 97, 107, 108, 109, 111 endosymbiotic event primary 93 secondary 93, 109, 111 endosymbiotic progenitors of chloroplasts 458 energy transfer 139–141, 143, 150, 151, 154, 155, 161–166, 172– 176, 193, 194, 198, 206–215, 680, 681 dipole-induced dipole (resonance) energy transfer 150, 162, 176 Förster energy transfer 150, 162, 176 mechanism 150, 172, 176 kinetic studies 174 rates 176 energy dependence of Ci uptake 470, 476 enterobacteria 505 enterobactin 741 See siderophore entomocidal toxin gene Bacillus sphaericus 601 epifluorescence microscopy 838 equilibrium constant, ATPase 368 EPR spectroscopy See electron paramagnetic resonance spectroscopy erythromycin resistance gene (erm) 588 Escherichia coli divergence from cyanobacteria 15, 364 ESEEM See electron spin-echo envelope modulation ESR See EPR spectroscopy 7

ethine reduction 420 ethionine inhibition of Mg-protoporphyrin methyltransferase 540 ethylene diamine di-o-hydrophenyl acetic acid (EDDA) 742, 743 ethylenediamine 506 ethylidine isomers 533, 535–537 cis-trans isomerization in phycobilin synthesis 535 ethylidine isomerization 535–537 N-ethylmaleimide 373, 541 etiolated barley seedlings 688 eubacteria 120, 123, 134, 366 eucaryote 92, 93, 94, 103, 109, 110, 111 eucaryotic nuclei 366 Euglenophyta 92, 93 euglenophyte 92, 94, 111 evolution 53, 186, 190, 197, 278, 362, 364, 366, 372, 374, 376 convergent 95 EXAFS See Extended X-ray absorption fine structure. excisase 775 See site-specific recombinase XisA 775 XisC 776 XisF 775 excision element See excison excisons 775, 776 exon 97, 102 extracellular polysaccharides 800

F F-type ATPase 362–376 f-type thioredoxin See thioredoxin center See Photosystem I, center centers See Photosystem I, centers center See Photosystem I, center center See Photosystem I, center facultative autotroph 451 FAD See flavin adenine dinucleotide farnesyl pyrophosphate 77 farnesyl pyrophosphate synthase 78 farnesyltransferase 522, 548 fatty acid 2, 5, 53 FBPase See fructose-1,6-bisphosphatase Fd See ferredoxin Fe deficiency See iron deficiency Fe-Mo cofactor See nitrogenase, molybdenum Fe nitrogenase 497 Fe-S center/cluster 129, 270–274, 283, 287, 301, 302, 335, 492, 687, 774 See also Rieske Fe-S protein, ferredoxin, nitrogenase binding domain 273, 302 [2Fe-2S] cluster 270–274, 382, 383 cluster 336, 346 cluster/center 333, 345, 346, 386, 492, 774 See also ferredoxin, Fe-S protein, PsaC cluster 347 mixed-ligand cluster 336 P clusters 492 fermentation 285 ferredoxin (Fd) 11, 43, 270, 280, 283, 284, 289, 320, 323, 346, 349, 352, 382–387, 391, 392, 397, 398, 413–416, 424, 492– 495, 499, 508, 529, 534–536, 545, 654, 678, 716, 722–724, 732, 735–738, 776, 781, 802 ferredoxin H 385

Subject Index ferredoxin I 75, 81, 382–387, 735 ferredoxin II 383–386, 397 ferredoxin III 386 Fe-S center See Fe-S center two-fold rotational symmetry axis, Peptococcus aerogenes 345 ferredoxin-dependent NADPH:cytochrome c reductase 534 ferredoxin-heme oxygenase 387 oxidoreductase (FNR) 79, 80, 82, 108, 133, 149, 150, 192, 261, 283, 289, 382, 384, 388–392, 397, 410, 411, 413–416, 424, 534, 535, 723, 777, 778, 781 ferredoxin-nitrate reductase 133 ferredoxin-nitrite reductase 133 ferredoxin:quinone reductase (FQR) 280, 283 ferredoxin-thioredoxin reductase 722 ferredoxin/thioredoxin system 453 ferrichrome 741 See siderophores ferrochelatase 522, 531, 532 ferrocyanide 426 fibrillogranular body See nucleomorph fimbriae 831, 832 See also pili flagellar synthesis 617 flavin adenine dinucleotide (FAD) 389, 391, 419, 544 FAD-binding 570 flavin mononucleotide (FMN) 387–388, 418 flavodoxin 43 flavodoxin 43, 323, 346, 349, 352, 382, 383, 387–389, 392, 492, 495, 499, 654, 735, 736–738, 745, 776, 777 Nif-specific 387 flavoprotein reductase 722 Florideophyceae 177 flow cytometry 29 fluorescence 60 -assisted sorting 70 decay kinetics 682 delayed 32 emission 56, 60, 150, 153, 154, 180, 199, 680, 682, 683 emission spectra 180, 199 induction I-D dip 421 intensity 679 lifetime 162, 334 low-temperature studies 56, 155, 265, 682 measurements at 77 K 56, 155, 265, 682 microscopy 51 quenching 474, 476 resonance energy transfer (FRET) 370, 373, 374 fluorescent light 685 fluorescing chromophore 163, 174, 175 fluorimetry 474 fluorochloridone 573 fluorodeoxyuridine 285 fluridone 561, 572 footprinting assays 629, 631 Form I, D1 (PsbA) protein 707 Form II, D1 (PsbA) protein 705, 707, 708 Förster dipole-induced dipole energy transfer 162 See also energy transfer mechanism 150, 172, 176 fossil record 5 FNR See oxidoreductase FQR See ferredoxin:quinone reductase fractionation mechanical 57

863 fracture faces 57 freeze-fracture electron microscopy See electron microscopy, freeze-fracture French press 55 FRET See fluorescence resonance energy transfer fructose 412, 414, 421 fructose 1,6-bisphosphate 455 fructose-1,6-bisphosphatase (FBPase) 716, 718, 723 FT-IR spectroscopy See infrared spectrscopy funiculosin 280, 297 Fur represser 743 Fur-binding sequences table 746

G gabaculine 525, 526 GSA aminotransferase inhibition 525, 526 276, 444, 596, 701, 702, 704, 760, 773 subunit See also phycoerythrin ATP synthase 362–365, 369, 373 phycoerythrin 148, 176, 177, 179–181, 184, 185, 190, 202–204 subunit 148, 176, 177, 179–181, 184, 185, 190, 202–204 nuclear-encoded 186 RNA polymerase 620, 632 gas vacuole 832 gas vesicle 10, 11, 17, 51, 52, 825, 827, 830, 832, 834, 835, 838 antisense RNA 832 gas vesicle protein 828 GDH See glutamate dehydrogenase gel exclusion chromatography 265, 266, 526, 535 gel retardation assay 630, 652 gene atp gene organization 364–366 conversion 585, 591, 593 duplication 167 electron transport (pet) 289, 290 families 693 fusions 269, 276 homologous 93, 102, 103 mapping 95 nuclear-encoded 108 overlap 100 Photosystem I components table 324 Photosystem II components table 696 phycobilisome components 199 plastid-encoded 106, 107 replacement 720 sequence 93, 102 transfer 93, 97, 106, 582–592 conjugation 582 genetic analysis reversion 291 site-directed mutagenesis 291 genetic system 582–606 development 604 genome 92–97, 100–102, 105–108, 111 chimaeric 93, 107, 111 copy number 832 inverted repeat 94, 96, 105, 106

864 map Anabaena sp. strain PCC 7120 600, 773 Synechococcus sp. strain PCC 7002 600 Synechocystis sp. strain PCC 6803 600 plasticity 44 size 599–601, 832 geranylgeraniol 545, 546 geranylgeranyl pyrophosphate (GGPP) 568–570, 688 geranylgeranyl pyrophosphate (GGPP) synthase 78, 568–570 geranylgeranyl reductase 522 GGPP synthase See geranylgeranyl pyrophosphate synthase Glaucocystophyceae 66 Glaucophyta 92, 93 gliding motility 837 glucose 191, 412, 414, 471 glucose 6-phosphate 792 glucose 6-phosphate dehydrogenase (G6PDH) 108, 412, 428, 718, 724, 726, 778, 779 glucose-tolerant mutant Synechocystis sp. strain PCC 6803 697, 699 (GUS) 596, 603 D-glutamate 68 L-glutamate 509, 523–526 conversion to 523–527 glutamate dehydrogenase 505, 509 glutamate-oxaloacetate transaminase 791 glutamate-pyruvate transaminase 791 glutamate l-semialdehyde(GSA) 521–526 cyclic ester vs. free 526 glutamate l-semialdehyde (GSA) aminotransferase 108, 522, 525, 526 4,5-diaminovaleric acid 526 glutamate synthase (GOGAT) 78, 455, 492, 501, 504, 505, 508, 790, 806 glutamine 488, 777, 788, 790, 791, 809, 835, 836 transport 504 glutaminase 455, 504 glutamine-amido transferase 504 glutamine oxoglutarate aminotransferase (GOGAT) See glutamate synthase glutamine synthetase 492, 501–508, 632, 660, 724, 725, 790, 796, 826, 829, 833, 835 adenylylation 508 quaternary structure 507 -glutamate synthase pathway 90, 488, 489, 501, 505, 506, 508, 509 glutamyl-tRNA 521 See trnE glutamyl-tRNA reductase 522, 524–526 affinity chromatography 524 glutamyl-tRNA synthetase 522, 524 Chlamydomonas reinhardtii synthetase 525 glutaredoxin 717, 720, 725, 726 glutathione (GSH) 525, 535, 537, 717, 724, 726, 778, 788 glyceraldehyde 3-phosphate dehydrogenase 92, 108, 778 glycine 521 glycocalyx 121, 122 glycogen 412, 421, 663, 734, 789, 838 glycolate 438, 455 glycolic acid 411 glycolipids 5, 772, 783, 800, 801, 802 glycolysis 778 glycoproteins 191 GOGAT See also glutamate synthase

Subject Index Gram-negative bacteria 122 Gram-negative cell wall 58, 122 Prochlorothrix hollandica 58 grana destacking 680 lamellae 53 membranes 680 stacks 53, 680 -type, oxygen-evolving particles 57 green light 165, 190, 191, 198, 642–654, 678, 830, 832–835, 839 green sulfur bacteria 107, 134, 261 growth rate 55 GS See glutamine synthetase GSA See glutamate 1 -semialdehyde GS/GOGAT . See glutamine synthetase/glutamate synthase guanosine triphosphate (GTP) 369 GUS See gymnosperm 95, 544, 837

H See hydrogen See bicarbonate See hydrogen peroxide halobacteria 44 heat shock 95, 634, 752 cyanobacteria 762, 763 relative molecular mass 763 cellular thermometer 756 genes 622 genes 763 proteins 754, 756 hsp60 753 See chaperone, chaperonin hsp70 78, 753, 756, 759 chaperone regulon 753 response 751, 752–765 heliobacteria 134, 261 See also Heliobacterium sp. hematoporphyrin IX 531 heme 148, 520–532, 538, 541 -binding motif 276 binding residues 276 biosynthesis 288, 520–532 ligands 277, 289, 291, 296 axial 267, 276 redox potential 276 -staining reaction 266 thioether linkage 276 heme a 532, 548 heme 266, 281, 282, 295, 299 heme 266, 275, 281, 282, 291, 295, 298, 299 heme c 532 lyase 522, 532 See also CYC3 and cyt-2 heme f 287 heme o 548 heme oxygenase 78, 522, 534, 535, 735 hemoglobin 148 hemoprotein 541 2-n-heptyl-4-hydroxyquinoline-N-oxide (HQNO) 280–283, 297, 414, 420, 422, 427, 687, 782 herbicide 561, 571–573, 710 atrazine 697, 710 DCMU 280, 284, 297, 421, 428, 476, 697, 710, 831, 834, 835 diphenyl ether 531, 548

Subject Index fluridone 561, 572 ioxynil 697 methyl viologen 423, 683 norflurazon 561, 571–573 resistance to 571–573, 695 heterocyst 6, 11, 20, 165, 410, 412, 414, 415, 417, 420, 422–424, 426, 428, 455, 488, 490, 494–497, 510, 532, 631, 634, 660, 663, 665, 666, 724, 725, 736, 770–811, 825, 826, 827, 828, 838 channel 772 dab mutant 795 development 621, 632, 635 differentiation 632, 775, 794, 796, 835, 836, 839 environmental control 788 het mutant 795 time-course 794 electron micrographs 783 envelope 783, 784, 800 fibrous layer 784 glycolipids 801, 802 hen mutant 795 polysaccharide 784, 785, 801 fix mutant 795 fox mutant 588, 589, 596, 599, 795 glycolipids 784, 785, 795 hgl mutant 795 honeycomb membranes 786 10.5-kb insertion element 776 11-kb insertion element 495, 598, 775 55-kb insertion element 495, 598, 775 isolated 773 non-fragmenting mutants 799 pattern formation 803–810 polysaccharide layer 784, 785, 795 hep mutant 795 pores 780 proheterocysts 794, 800, 803–805, 809 -specific ferredoxin 387 See also ferredoxin H terminal 804 heterotrophic growth 55, 451, 590 light-activated heterotrophic growth (LAHG) 285, 345,412 hexahydrogeranylgeranyl 545 See phytol hexaprenylphosphate synthase 78 hexose monophosphate shunt 724 See mercuric chloride high light intensity See light intensity mutants See carbon dioxide high-pressure liquid chromatography (HPLC) 53, 185, 524, 562, 564 anion-exchange chromatography 265 hydroxyapatite chromatography 265 highly iterated palindromic sequence (HIP1) 43, 44 octameric palindrome 43, 44 HIP1 See highly iterated palindromic sequence histidine protein kinase 37, 40 homeostatic regulation PS I: PS II ratio 689 homologous gene 93, 102, 103 honeycomb membranes 786 hormogonium (-ia) 6, 20, 810, 825, 826–840 chemotactic response 838 differentiation 828, 829–840 action spectrum for photoinduction 830

865 effects of light wavelength 830 HPLC See high-pressure liquid chromatography HQNO See 2–n-heptyl-4-hydroxyquinoline-N-oxide hsp60 See heat shock protein, chaperone, chaperonin, GroEL hsp70 See heat shock protein, chaperone, DnaK HUP See hydrogenase, uptake hybridization in situ 93, 109 Southern 109 hydrogen 412, 414, 419, 420, 421, 424, 792 hydrogenase 285, 385, 386, 397, 398–402, 399, 412, 413, 419, 420, 781, 793 uptake 261, 413, 419, 420, 776, 793 reversible 419, 793 hydrogen-cytochrome c reductase 421 hydrogen peroxide 476 hydroquinone 225, 246 See also quinol hydroxyamate siderophores See siderophores hydroxyapatite 280 hydroxylamine 225, 246 hydroxylation in isocylic ring formation in Chl biosynthesis 541 p-hydroxymercuribenzoate 534 hydroxymethylbilane 527 hydroxymethylbilane synthase 527–529 dipyrrole cofactor 528 4-hydroxyphenylpyruvate dioxygenase 421 5'-hydroxyphylloquinone 421 540 hyperfine couplings, PS II 243 hyperpolarization of electrochemical membrane potential 474, 475 hyperscum 400

I ICM See intracytoplasmic membrane, thylakoid immunoelectronmicroscopy 772, 778, 791, 809, 838 immunogold labeling Photosystem I 330 superoxide dismutase 788 immunological comparisons taxonomic studies 8 immunoprecipitation nearest-neighbor analysis for PS II 221 immunostaining analysis of symbiotic cyanobacteria 9 2–iodo-6-isopropyl-3-methyl-2', 4, 4'-trinitrodiphenyl-ether (DNPINT) 271, 272, 280, 284 IncP (conjugative) plasmid See plasmid IncQ plasmid See plasmid infrared spectroscopy FT-IR 240 difference FT-IR 240 inorganic carbon See carbon, inorganic inorganic carbon-concentrating mechanism (CCM) See carbon, inorganic concentrating mechanism inosine 5'-monophosphate 472 inosine 5'-triphosphate (ITP) binding to ATPsynthase 369 insertion element See also nif genes, nitrogenase, heterocyst 10.5-kb 776

11-kb 495, 598, 775

Subject Index

866 55-kb 495, 598, 775 insertion sequences (IS) 10, 598 IS701 598 IS702 598 IS703 598 IS891 598 IS892 598 IS893 599 IS894 599 IS895 598 IS895B 599 IS895C 599 IS897 599 IS898 599 integration host factor (IHF) 36, 619 integration platforms 597, 601 integrative vectors 584 internal transcribed spacer (ITS) 21 interposon mutagenesis 340, 342, 345, 349, 352, 390, 472, 480 intracellular inclusions 663 See also carboxysomes, cyanophycin, glycogen, phycobilisomes, polyphosphate granules intracytoplasmic membrane (ICM) 409, 411, 413–421, 423–429 See also thylakoid chromatophore 540 intrathylakoid space 413, 424 introgression, of gene clusters 97 intron 96, 102, 203, 366 A + T content 102 group I 102 group II 96, 102 twintron 102 inverted repeat, in plastid genomes 94, 96, 105, 106 ioxynil 697 iron 42, 387, 388, 396, 732, 829 See also Fe -regulated genes 733 stress 54, 734, 738 transport 740 -transport systems 740 iron deficiency 387, 388, 546, 655, 664 Anabaena cylindrica 655 Anabaena flos-aquae 655 Aphanocapsa 6714 655 Synechococcus 655 Synechococcus 7002 655 Synechococcus cedrorum 655 Synechococcus sp. (marine) 655 iron-sulfur center/cluster See Fe-S center/clusters centers iron-sulfur proteins 280, 345 See also ferredoxin, Fe-S center/ cluster, PsaC, Photosystem I, NADH dehydrogenase, nitrogenase, Rieske protein iso-Cyt 279 isocitrate dehydrogenase 724, 778, 779, 792, 796 isocyclic ring 538, 540, 548 hydroxylation during formation 541 inhibition by azide 541 sensitivity to mercurial sulfhydryl reagents 540 isomerase role in phycobilin synthesis 536 isomerization phycobilins 535–536 proteins 758 isopentenyl pyrophosphate (IPP) 77, 569 isoprenoid biosynthesis 545–546, 567–573

isopropyl isotopic fractionation carbon 470–471 isotopic labeling 240 isotopic substitution 240 isozyme chloroplast and cytosolic 108 patterns 7

(IPTG) 603

K K-edge See X-ray absorption spectroscopy 3-ketoacyl-ACP synthase 78 522, 835, 836 dehydrogenase 414, 778 540 kinase LHC 60, 61, 680 transcription factor 839 Knallgas reaction 792

L See linker polypeptide, core See linker polypeptide, core-membrane linker phycobiliprotein See linker polypeptide, rod See linker polypeptide, rod-core lactose 760 lacZ translational fusions 702 lambda promoter See promoter, phage lambda lateral gene transfer 93, 103, 107 lateral heterogeneity 52, 53, 57–59, 680 lateral separation 50, 61 leader peptidase 276, 287 leghemoglobin 531 legume 96, 97 leucine zipper 330 LHC II See light-harvesting chlorophyll protein lichen 831, 838 light -activated heterotrophic growth (LAHG) 285, 345, 412 blue 706, 830 blue-green 678 chromatic 684 -dark transitions 456 -driven electron transfer 373 fluorescent 685 gradient 700 -induced shift 682 intensity 55, 56, 60, 61, 699–704, 707, 826, 829 effect on prochlorophytes 61 high 700, 702–706, 709 quality 706, 826, 830, 834, 836, 839 red light 164, 165, 706, 642–654, 830–834, 839 regulation 290 -responsive elements 699, 702, 703 -responsive regulation 694, 699, 702–706 -sensitive mutants 724 shielding 53 light-harvesting antenna 560, 680, 679, 689 See also antenna, phycobilisome, light-harvesting chlorophyll protein light-harvesting chlorophyll protein (LHC II) 50, 53, 57, 59, 60–

Subject Index 62, 142, 266, 680, 688, 761 apoproteins 56 fluorescence spectroscopy at 77 K 56 trimeric association 53 kinase 61, 286 phosphorylation 60, 61, 680, 681 regulatory role 60, 61 polypeptides 56 regulation of light harvesting 51, 56 light-harvesting pigment 678, 681 See also antenna, chlorophyll, light-harvesting chlorophyll protein, phycobiliprotein, phycobilisome linker polypeptide 139–205, 642, 645, 833 See also phycobilisome core 152, 191, 193, 205 overproduction of 154 reconstitution studies with allophycocyanin 154 core-membrane linkerphycobiliprotein 148, 152, 155, 156, 159–162, 170, 172, 191–194, 198, 200, 205 681 See also ApcE, phycobilisome Anabaena sp. strain PCC 7120 159 Aglaothamion neglectum 156 amino acid sequence 157 Calothrix sp. strain PCC 7601 155, 159 chromophore 156 Cyanophora paradoxa 155 domain structure 156 Mastigocladus laminosus 156, 159 phylogenetic development 168–170, 197 repeat (REP) domains 152, 156, 158–160 spacing ARM 156, 158, 159 structure 152, 156, 158–160 Synechococcus sp. strain PCC 6301 155 Synechococcus sp. strain PCC 7002 155 electrostatic interactions with phycobiliproteins 196 functional domains 196 isoelectric points 196 -phycobiliprotein complexes 152–160, 191–198 rod polypeptide 150, 165, 191, 193, 194, 195, 198, 645, amino acid sequence 195 rod-core polypeptide 8, 152, 165, 166, 173, 191-198 amino acid sequence 195 red-shift in phycocyanin absorption 194 lipid 5, 7, 710 bodies 831, 838 di-unsaturated 710 importance in resistance to photoinhibition 710 tri-unsaturated 710 lipopolysaccharide 51, 122 3-O-methyl-xylose 51 liverwort See Marchantia polymorpha Lon protease 754 long-term adaptation 679, 683–689 model 690 low-potential cytochrome c See cytochrome c LPP See Lyngbya-Plectonema Phormidium LSU See ribulose 1, 5-bisphosphate carboxylase/oxygenase, large subunit luciferase 596, 598, 603, 773 luminescence 772 lumen See intrathylakoid space luteolin 838 lycopene See carotenoid

867 lycopene cyclase 133, 569, 570, 572 inhibition by 2–(4-chlorophenylthio)-triethylamine hydrochloride 573 inhibition by 2–(4-methylphenoxy)-triethylamine hydrochloride 572, 573 lysyl-tRNA synthetase 754

M macromolecule synthesis 31 magnesium, See Mg magnetic circular dichroism (MCD) spectroscopy 267, 276 magnetic resonance 241 See also electron paramagnetic resonance, electron-nuclear double resonance, electron spin echo envelope modulation, nuclear magnetic resonance maize 8 malate dehydrogenase 7, 717, 723 manganese 218, 222, 244, 246, 366 cluster 232, 235, 238, 244, 247 complex 240 K-edge 245 ligands 222, 232, 244 carboxylate groups 232, 244 multiline signal 241, 242, 245 site in the S2 state 244 stabilizing protein (MSP) 218–223, 699, 709 See also PsbO, WoxA mapping 594, 599–601 maps of chromosomes 600 Anabaena sp. strain PCC 7120 600, 773 Synechococcus sp. strain PCC 7002 600 Synechocystis sp. strain PCC 6803 600 marine cyanobacteria 8, 9, 27–45, 461 adaptive responses 43 marine environment 461 mass spectrometry 474, 475, 526 mastigamoeba 92 MEGA-9 See detergent megaplasmid 774 membrane See also cytoplasmic membrane, intracytoplasmic membrane, outer membrane, thylakoid appression 53, 57, 58, 62 -bound NADPH dehydrogenase 417 chromatophore 540 honeycomb 786 -intrinsic cytochrome 425 organization 119–120 -parallel helix 268 potential 474 protein insertion, role of chaperones 759–762 localization 121–134 positive-inside rule 287 targeting 288 structure 120 topology 287 vesicles 474 menadione 414, 420 menaquinone-Cyt c oxidoreductase 261 mercaptoethanol 541 mercurial sulfhydryl reagents 540 mercuric chloride 284 merodiploids 482, 584, 591, 601

868 mesobiliverdin 148, 188, 534 See also phycobilin mesoheme 534 mesoporphyrin IX 531, 538 metal binding site 371 See also ATP synthase metal-nucleotide binding site 369, 371 See also ATP synthase model 373 metal-nucleotide complex 367, 369, 375 See also ATP synthase structure 367 metallothionein 43 Metaphyta 92, 93 methanolysis 533, 535, 537 methionine 526, 530 methionine sulfoxide 717 reduction 716 methionine sulfoximine 497, 505, 506, 508, 826, 829, 836 inhibitor 281 See also MOA-stilbene, myxothiazol 5-methylaminomethyl-2-thiouridine 524 N-methylmesoporphyrin IX 538 methyl oxidase 548 See CtaA 2-(4-methylphenoxy)-triethylamine hydrochloride (MPTA) 572, 573 See also lycopene cyclase N-methylprotoporphyrin IX 538 methyl viologen 423, 683 3-O-methyl-xylose occurrence in lipopolysaccharide of P. hollandica 51 methylation of DNA 591 173, 174, 178, 179, 181, 187, 189 methyltransferase See Mg-protoporphyrin methyltransferase mevalonic acid 569 366, 375, 524, 528, 538 Mg chelatase 522, 538–540, 548 Mg-2,4-divinylpheoporphyrin 540, 541 Mg-protoporphyrin IX 538–542 Mg-protoporphyrin IX methyltransferase 522, 540 inhibition by ethionine 540 Mg-protoporphyrin IX 6-monomethyl ester 539, 541, 542 6-acrylic derivative 542 cyclase 522 vinyl reduction in 542 microbial loop 29 microplasmodesmata 780 microscopy electron 109 epifluorescence 838 immunoelectronmicroscopy 772, 778, 791, 809 immunoelectronmicrosopy 838 midpoint potential 335, 347, 680 mithramycin nuclear DNA staining, C. paradoxa 70 mitochondria 261, 264, 366 cytochrome complex 260–263 membrane 760 ATPase 368, 372 NADH dehydrogenase 414, 419 transfer event 366 mitomycin 806 mixed-ligand cluster 336 Mn See manganese Mo See molybdenum MOA-stilbene 281, 282, 287 See methoxyacrylate mobile antenna 682 mobility-shift experiment 630, 652 See also gel retardation assay

Subject Index modified Q cycle 281 molecular chaperone 441, 752–754, 758, 759, 762–764 mutational studies 764 role in maintaining unfolded conformation of proteins 760 molecular evolution 2 molecular modeling 720, 723 molecular phylogeny 27 molten-globule state 758 molybdenum 496, 499, 775 -dependent nitrogenase 589 Mo-Fe cofactor 495 See also nitrogenase monophyletic origin of chloroplasts 92, 93, 95, 97, 105, 109, 111 monovinylprotochlorophyllide 542, 544 Mössbauer spectroscopy 42 motility 827 m-type thioredoxin See thioredoxin mucidin 280, 292, 294 multicomponent transport systems See transport multiline signal 241, 242, 245 muramidase 69 murein 92 mutagenesis 592–596 chemical 471, 592 diethyl sulfate 592 interposon 340, 342, 345, 349, 352, 390, 472, 480, 584, 591, 592, 606 nitrosoguanidine 592 random 451, , 594, 595, 635 cartridge 594 integrational 595 site-directed mutagenesis 156, 176, 231, 232, 237, 241, 267, 273, 277, 278, 291, 296, 297, 388, 394, 447, 593, 594, 757 targeted 337, 340, 342, 345, 349, 352, 390, 472, 480, 584, 591, 592, 606 ultraviolet (UV) light 592 mutant 194, 450, 649, 651 C3P-O 479 carbon-concentration mechanism 474, 476-482 D4 472 dnaK 757 El 478 JR12 472 M3 478, 482 O221 472, 473 psaD mutant 349, 350 psaE mutant 328, 354 psaF mutant 339 psaF psaJ double mutant 341 psaK mutant 327, 342 psaL mutant 326, 327, 342 RI4 472 segregation 595 SF33 (short filament mutant) 839 site-directed 176, 336, 346 mutational analyses 194, 202 mutations 293, 451 lethal 302 positive selection 587 reversion 291 second-site (suppressor) 291, 452 site-directed 156, 162, 174, 176, 268, 294, 299, 301, 302, 303, 371 temperature-sensitive Rieske 300, 301

Subject Index myoglobin 148, 167, 197 myxothiazol 272, 280, 281, 292, 293, 294, 300 myxoxanthophyll See carotenoid

N N See nitrogen N-1 See cyanophage N-l 414, 416 dehydrogenase 492 NADH 288, 412, 413, 418, 420, 423 NADH:cytochrome c oxidoreductase 421 NADH dehydrogenase 94, 119, 121, 129, 133, 261, 284, 285, 354, 414–421, 472, 476, 477, 481, 683 bacterial-type 419 inhibition by o-phenanthroline 419 inhibition by rotenone 131, 418, 419 ndhF/psaE double mutant 354 soluble NADH dehydrogenase 413, 419 Type-1 enzyme 418, 419 Type-2 enzyme 419 NADH:ubiquinone oxidoreductase 125, 133, 415 See also NADH dehydrogenase 283, 411–417, 420–424, 678 photoreduction 338, 349, 352 malate dehydrogenase 717, 723 dehydrogenase 492, 509 dehydrogenase 718 NADPH 286, 409, 412, 413, 416, 420–423, 428, 456, 524, 534, 535, 541–545, 678 NADPH-DBMIB oxidoreductase 417 NADPH dehydrogenase 125, 413, 415, 421, 476 NADPH-dependent cyt c reductase activity 392 NADPH-dependent oxidoreductases 806 NADPH ferrihemoprotein reductase 534 NADPH-menadione oxidoreductase 415, 420 NADPH/NADH transhydrogenase 284 NADPH plastoquinone oxidoreductase 415, 419, 781 NADPH protochlorophyllide oxidoreductase 522, 544, 545 presence in etiolated oat seedlings 544 inhibition by quinacrine 544 1, 4-naphthoquinone 335, 415, 421 nearest-neighbor analyses 349 Nernst equation 680 neurosporene See carotenoid neutral site 597, 601 vector 602 nickel 399 nickel-iron center 398 nitrate 400, 402, 488, 498, 500, 502, 679 assimilation 39, 497–502 transport 498, 501, 502 transporter 501 nitrate reductase 492, 499 nitric oxide reductase 284 nitrite 400, 402, 500 assimilation 497–502 transport 501, 502 nitrite reductase 383, 492, 499, 500, 529 nitrogen 37, 488–511 acquisition 659 assimilation 492, 497–511, 617 deficiency

869 Anabaena 657 Calothrix parietina 657 Mastigocladus laminosus 657 Oscillatoria 657 Phormidium 657 Plectonema boryanum 657 Pseudanabaena catenata 658 Spirulina platensis 658 Synechococcus 658 Synechococcus 7002 658 Synechococcus sp. (marine) 658 Synechocystis 6308 658 Synechocystis 6803 658 fixation 39, 414, 428, 617, 621, 632, 770, 772, 774–782 gene cluster (nif) 20, 29, 385, 494–497, 596, 774, 775, 788, 795, 797 See also nif insertion element (10.5 kb) 776 insertion element (11 kb) 495, 598, 775 insertion element (55 kb) 495, 598, 775 nif elements 495, 598, 775, 776 nif region 774 nif transcription 461, 785 -fixing species 724 pathways 489 control 461, 489, 510 metabolism 488–511, 659 reserve 32, 38 sources 488–511, 839 starvation 621, 632, 829 nitrogenase 5, 30, 40, 385, 386, 399, 400, 410, 428, 461, 488–499, 545, 770–772, 774, 775, 778, 781, 786–792, 796, 802 alternative nitrogenases 775 dinitrogenase 492, 774, 787 dinitrogenase reductase 492, 493, 497, 591, 774, 787, 789, 793 labeling 772 Fe-Mo cofactors 492 Fe nitrogenase 497 Fe protein 40, 493 See also NifH genes 495–496, 774–776 See also nif genes insertion element (10.5 kb) 776 insertion element (11 kb) 495, 598, 775 insertion element (55 kb) 495, 598, 775 intervening sequence elements 495–496, 774–776 immunolocalization in heterocysts 772 MoFe-protein 545 P clusters 492 vanadium-dependent 496, 589 X-ray structures 493 nitrosoguanidine 592 NMR See nuclear magnetic resonance noncyclic electron transport See electron transport, noncyclic non-fragmenting mutants 799 non-heme iron 264, 322, 329 norflurazon 561, 571–573 Nostocales 6 nostoxanthin 563 See carotenoid See NtrC nuclear magnetic resonance (NMR) 148, 165, 245, 246, 276, 366, 393, 526 nuclear Overhauser enhancement studies 366 NMR spectroscopy 265 solution structure, PsaE 351 spectroscopic analysis 383

Subject Index

870 nuclease 605 nonspecific 605 sugar-nonspecific 605 nuclease protection techniques 623 nucleic acids 2, 7 See deoxyribonucleic acid and ribonucleic acid nucleomorph 92, 93, 94, 109, 110, 204 fibrillogranular body 109 nucleophilic 368 nucleotide binding sites 363, 370 nucleus 92, 93, 94, 95, 96, 97, 107, 109, 110 nutrients deficiency 654–668 C 655 Fe 655 N 655 P 655 S 655 ions 689 reserves 663 nutrition 37

O ocean biased light regime in 678 cyanobacteria in 28 (OGP) See detergent, pyranoside octyl thioglucoside See detergent oligomeric proteins role of molecular chaperones in assembly 441 oligotrophic waters 462 aminotransferases 525 gabaculine inhibition of 525 cassette 588, 605, 702, 703 streptomycin/spectinomycin resistance gene 605 operator 618 ORF (open reading frame) ORF31 100 ORF173. See rpcF ORF283 100 ORF290 100 ornithine 504, 509 ornithine acetyltransferase 509 ornithine carbamyl phosphate transferase 439 ornithine transcarbamylase (OTC) 455, 509, 510, 791 immunolocalization 791 vegetative cells and heterocysts, occurrence in 455 Oscillatoriales 6 oscillaxanthin See carotenoid osmotic pressure 679 high 679 outer membrane 119, 121, 122, 123, 124, 134, 288, 411, 488 oxidative pentose phosphate pathway 412, 778 oxidative phosphorylation 375, 414, 415, 417, 420 complexes 362 oxychlorobacteria 49, 50, 55, 62 See also prochlorophyte oxygen compensation point 474 singlet-state 561 oxygen evolution 219–247, 679, 680, 707 action spectrum for quantum yield 679 oxygen-evolving complex (OEC) 51, 52, 126, 219–247

oxygenic photosynthesis 3 oxyhydrogen reaction 792 oxyphotobacteria 50, 55, 59

P P clusters 492 P loop 371, 373 protein 508, 511, 826, 835, 836, 839 P680 See Photosystem II P700 See Photosystem I PAGE See polyacrylamide gel electrophoresis palindromes 43, 44 588 See also highly iterated palindromic sequences (HIP1) PAPS See 3'-phosphoadenosine-5'-phosphate parsimony analysis 12, 14, 106 particle bombardment Chlamydomonas reinhardtii transformation 302 passive diffusion 474 channels 122 Pasteur Culture Collection 5, 697 pattern formation 803–810 PBP See phycobiliprotein PBS See phycobilisome PC See phycocyanin PCB See phycocyanobilin, phycobilin PE See phycoerythrin PEB See phycoerythrobilin, phycobilin Pchlide a 688 See protochlorophyllide a PCR See polymerase chain reaction pea 538, 686, 687 chloroplasts 203 PEC See phycoerythrocyanin penicillin 68 enrichment 592 -binding proteins 69 pentose phosphate cycle (reductive, Calvin) 457–460 PEP carboxylase See phosphoenolpyruvate carboxylase peptide elongation 688 peptidoglycan 19, 51, 68, 69, 93, 120, 121, 122, 123, 800, 831 peptidyl-prolyl cis-trans isomerase 482 peripheral rods 144, 146, 149, 150, 151, 152, 158, 160, 162, 163, 164, 176, 177, 194, 199, 689 See also phycobilisome periplasmic phosphate protein 37, 40 periplasmic space 121, 413, 424 periplastidal space 94, 109 permeabilized cells 456 phage See cyanophage phagotrophic host(s) 93, 94, 111 1,10-phenanthroline 538 m-phenanthroline 547 o-phenanthroline inhibition of chlorophyll b synthesis 547 inhibition of NADH dehydrogenase activity 419 phenolic compounds 838 phenylglyoxal 450 pheophytin a 546 phosphatase 60, 603, 661, 680, 829, 839 inhibitor NaF 60 phosphate 679 -anhydride bond 368 -binding protein, periplasmic 37, 40 oxygen binding sites 371

Subject Index P deficiency Anabaena variabilis 656 Calothrix 656 Gleocapsa alpicola 656 Nostoc 656 Oscillatoria 656 Phormidium luridum 656 Plectonema boryanum 656 Pseudanabaena catenata 656 Synechococcus 656 Synechococcus 7002 656 3'-phosphoadenosine-5'-phosphate (PAPS) 725 3'-phosphoadenosine-5'-phosphate (PAPS)reductase 725 3'-phosphoadenosine-5'-phosphate (PAPS)sulfotransferase 661 phospholipids phosphatidylcholine 265 phosphatidylglycerol 265 phosphoenolpyruvate (PEP) 452 phosphoenolpyruvate (PEP)carboxylase 439, 453–455, 461 6-phosphofructokinase 414 6-phosphogluconate 446, 456, 792 6-phosphogluconate dehydrogenase (6PGD) 412, 428, 632, 778 3-phosphoglyceraldehyde (3PGA) dehydrogenase 477 3-phosphoglyceric acid 438 2–phosphoglycolate 438 phosphoglycolate phosphatase (PGPase) 438, 455, 471 phosphoribosyl aminoimidazole carboxylase 472 phosphoribulokinase 438, 452, 452–453, 477, 718, 723, 724 chemolithoautotrophic bacteria 453 reactions 439 phosphorus 40, 829 See also phosphate phosphorylation 51, 56, 60, 61, 228, 367, 471, 680, 682, 716, 826, 833, 835 cascade 652 polypeptides 60, 61 seryl-phosphorylation 835 photoacoustic measurements 682 photoaffinity analog 369 photoaffinity labeling 264, 269, 371 photoconversion 320 photodynamic sensitizer 531 photoheterotroph 93, 590 photoinactivation 708, 709 See also photoinhibition photoinhibition 235, 237, 412, 694, 699, 706–709, 764 See also photoinactivation photooxidation 561 photophosphorylation 362, 781 photoreceptor 649, 653, 826, 834 See also photoreversible pigment, phytochrome photorespiration 411, 438, 471, 480 photoreversible pigment 826 See also photoreceptor, phytochrome photosynthetic membrane See thylakoid, intracytoplasmic membrane photosynthetic photon flux density (PPFD) 701 Photosystem I 49, 50–62, 74, 100, 107, 119, 120, 127–132, 134, 140, 142, 155, 261, 265, 266, 278, 280, 283, 284, 286, 320– 354, 373, 383, 388, 390, 392, 395, 397, 400, 412, 413, 415, 419–425, 429, 476, 534, 546, 560, 665, 678–685, 688, 733, 776, 781, 782, 786, 788, 789, 834 321–323, 328, 328, 333, 334 335 321, 329, 333, 335

871 335 abundance 684 apoprotein synthesis 687 back reactions 335, 336 biochemical resolution and reconstitution 326 catalytic domain 326 charge recombination 321 charge separation 321, 333 charge stabilization 333 cofactors 324, 328, 329 composition 324 connecting domain 326 CP 1 341 de novo synthesis during light-intensity acclimation 686 3-dimensional structure 326 2–fold rotation axis 327 3-fold rotation axis 327 4-membered bundle 333 domain-specific antibodies 330 electron density map 326 electron micrograph 128, 325 electron microscopy 128–130, 324–328 EPR analysis of oriented complexes 346 center 329, 336, 342–344, 349 center 329, 336, 342–344, 349 centers 329, 336, 342–344, 349, 352 322, 327, 328, 330, 336, 346 336 See Photosystem I, clusters 336, 346 [4Fe-4S] cluster/center 333, 345, 346, See also Fe-S protein, PsaC ferredoxin docking 349 gene/protein nomenclature 323 heterodimer 327 immunological determination during light adaptation 686 mixed-ligand cluster 336 monomers 324, 327 mutants 326, 327, 337, 339, 341–343, 346, 347, 349, 350, 352– 354 See also psa genes P700 321, 326, 328, 333, 342, 343, 394, 395, 425, 546, 682 P700+ cation 334, 335, 337, 476, 678 P700 triplet 334 formation 334 photoreduction of 346 plastocyanin docking 338 plastocyanin: ferredoxin oxidoreductase 320 PS I:PS II stoichiometry 54, 61, 679, 682, 684–690 homeostatic regulation 683–689 PsaA/PsaB heterodimer 349, 352 PsaC 343–347 C14D mutant protein 347 C51D mutant protein 347 PsaD 347–350 chemical modification 348 PsaE 350–353 Src homology 3 (SH3) domains 351 quantum yield, 320 reconstitution studies 347, 349 separation of PS I and PS II in P. hollandica 56 spin-polarized EPR signal 334 topological studies 330 translational control of PsaA and PsaB synthesis 688 trimers 52, 119, 128–130, 324, 327

872 image analysis, Prochlorothrix hollandica 59 trimeric complex 325 trimeric core 325, 333, 335, 346, 349 cluster 347 Photosystem II 54, 75, 100, 103, 105, 107, 119, 120, 123, 126– 134, 140, 142, 155, 156, 162, 218–247, 286, 289, 290, 387, 395, 397, 400, 412, 413, 420, 422, 423, 428, 476, 546, 560, 665, 678–688, 693–711, 733, 735, 694–711, 733, 738, 761, 764, 782, 834 See also Psb/psb abundance 685 carotenoids 560 complex 629 core complexes 226, 227, 245 CP43 218, 219, 223, 387, 397, 696, 698, 706, 738, 739 See Photosystem II, PsbC CP47 218, 219, 224, 397, 696, 761 See Photosystem II, PsbC, CP47 chlorophyll a-binding polypeptide 697 cytochrome 75, 123, 130, 132, 218, 219, 223, 227–231, 242, 696, 698, 782 D1 protein 51, 52, 56, 103, 105, 130, 218, 219, 223, 224, 227, 232, 233, 236–238, 560, 693–696, 706–708, 710, 761 See also PsbA D2 protein 56, 130, 218, 219, 223, 224, 227, 235–238, 693– 696, 698, 705, 706, 708, 761 See also PsbD g = 4.1 EPR signal 241 –243, 245, 247 genes See also psb genes table 219, 229, 230, 696 5-kDa hydrophilic protein 228, 229 6.1-kDa protein 230 9-kDa protein 218, 225, 226, 228, 229 See also PsbH 10-kDa protein (PsbR) 230 12–kDa protein 218, 225, 226 See also PetK, cytochrome 17-18-kDa protein (PsbQ) 218, 219, 224, 225, 243 24-kDa protein (PsbP) 218, 219, 224, 225, 243 43-kDa protein (CP43) 218, 224, 236 See also PsbC 47-kDa protein (CP47) 218, 221, 223, 235, 236 See also PsbB ligands to manganese 222, 232, 244 manganese stabilizing protein (MSP, 33-kDa) 218–223, 699, 709 membranes 226 multiline signal 241, 242, 245 multigene families 695 nearest neighbor analysis by immunoprecipitation 221 particles 227 polypeptides 219, 229, 230, 696 P680 218 P680 dimer 130 PS I:PS II stoichiometry 54, 61, 679, 682, 684–690 homeostatic regulation 683–689 PsbK 228–230, 341 218, 223, 224, 228, 238, 244, 698 218, 297, 477, 698, 709, 710 site 697 separation of PS I and PS II in P. hollandica 56 tyrosine radicals tyrosine 218, 237–240, 244 tyrosine 238 tyrosine 218, 224, 233, 237–240 phototrophic bacteria 458 phthalate dioxygenase 272 phycobilin 143, 148, 149, 520, 531, 533–537 absorption properties 148 phycocyanobilin lyase 202, 204

Subject Index biliverdin 148, 533–537 biosynthesis 533–536 Cys-bilin 584 148 Cys-bilin 618 148 DiCys-bilin 584 148 15, 16-dihydrobiliverdin 148, 188–190, 533, 536 doubly linked 149 ethylidine isomer 533, 535–537 ethylidine isomerase 522 ligation 148, 202, 204, 536, 537 mesobiliverdin 148, 188, 534 phycobiliviolin (PXB) 148, 149, 163, 165, 173 phycocyanobilin (PCB) 148, 149, 163, 166, 167, 174, 179, 189, 190, 202, 533–535, 537 (3E)-phycocyanobilin 533 (3Z)-phycocyanobilin 533, 536 phycoerythrobilin (PEB) 148, 149, 163, 166, 176–181, 186, 187, 189, 202, 536 See also phycobilin (3E)-phycoerythrobilin 533 (3Z)-phycoerythrobilin 533, 536 phycourobilin (PUB chromophores) 29, 148, 149, 163, 166, 176–178, 181, 186, 189 thioether linkage 148, 533, 536, 537 3-vinyl-2,3-dihydrobilin 535 (3Z)-phycobilins 535–537 phycobiliprotein (PBP) 8, 31, 93, 100, 139–205, 395, 531, 533, 536, 630, 641, 642, 645, 665, 678, 679, 681, 684, 734, 782, 798, 809, 833, 834, 835, 836 absorption properties 146–148, 153 absorption spectra 147, 180, 199 absorption maxima 146, 153 phycocyanobilin lyase 202, 204 amino acid sequences 168 subunit 147, 173 chromopeptides 186 chromophore 164, 174, 177, 178, 186, 189 composition 163 -protein interaction 173 structure 174 chromophorylation 203 cryptophytan subunits 177, 184 amino acid sequence comparison 184 crystals 167 domain 158 families 155, 163, 168–170 fluorescence spectra 180, 199 (gamma, subunit 148, 176, 177, 179–181, 184, 185, 190, 202–204 nuclear-encoded 186 gene organization 200–205, 644 hexamers 147, 161 immunological properties 163, 190 -linker polypeptide complexes 152–160, 191–198 phylogenetic tree 168, 170 primary structure 153, 166–170, 182–184 sensitizing chromophore 163, 174, 175 spectroscopic diversity 148 subunit structure 147 group 537 trimer 147 phycobilisome (PBS) 44, 50–54, 59, 75, 120, 125–129, 133, 134, 140–205, 343, 390, 642–645, 649, 664–666, 678, 679, 682– 684, 686, 688, 694, 734, 782, 838 See also linker polypep-

Subject Index tide, phycobiliprotein absorption spectra 147 abundance 684 architecture 159, 196 assembly 143–161 block-shaped 143 bundle-shaped 143 components 145, 146, 192 composition 145, 150 regulation 150 See complementary chromatic adaptation, light-intensity adaptation core 144, 152–161, 191, 194, 196, 681, 683 cylinders 159, 196 electron micrograph, M. laminosus 157 energy transfer 161 four-cylinder core 159 subcomplexes 160 core-membrane linker phycobiliprotein ( ApcE) 148, 152, 155, 156, 161, 162, 191–193, 198, 200, 681 See also ApcE, linker polypeptide degradation 668 domain structure 156 immunological properties 155 ‘loop’ domain 156 degradation 666 electron microscopy 143, 151, 157, 196 energy transfer 150 genes 54, 202–205, 644 ‘half core’ subcomplex 160 hemidiscoidal PBS 143–145, 150–152, 160, 161, 686 hemiellipsoidal 143 isolation 144 light intensity adaptation 152 PBS-less mutant 682 peripheral rod 144, 161–191, 193–199 proteins 54, 145, 146 polypeptide composition 145, 146 -PS II complexes 127 reconstitution 156 regulatory mutants 653 rod complex/element See peripheral rod rod-core complex 199 SDS-PAGE 160 phycobiliviolin (PXB) 148, 149, 163, 165, 173 phycocyanin (PC) 8, 10, 31, 76, 146, 153, 162–166, 176, 177, 190, 265, 276–279, 283, 532, 535, 537, 630, 641, 643, 649, 653, 666, 678, 735, 798, 826, 836 subunit 164 apoprotein 202, 537 subunit 164 C-phycocyanin 153, 163, 167, 171, 172, 599 crystal structure 167, 171, 189 secondary structure elements 167 turners 171, 175 constitutive 165, 645 crystal structure 176 hexamer 171 inducible 164, 165, 645, 649, 653 R-phycocyanin 165 amino acid sequences 166 subunit 173 R-phycocyanin-I (R-PC-I) 163, 165–167 R-phycocyanin-II (R-PC-II) 163, 166, 167, 202

873 R-phycocyanin-III (R-PC-III) 163, 166 red-light inducible PC 164, 165 spectroscopic properties 176 structure 164 transient absorption spectroscopy 176 phycocyanin 1 165, 522, 544, 833 See also constitutive PC phycocyanin 2 165, 833, 835, 839 See also inducible phycocyanin phycocyanin 3 165 phycocyanin-645 (PC-645) 163, 183, 188, 190 and subunits 148, 188, 190 subunit 189, 189 crystals 189 phycocyanin phycocyanobilin lyase 202, 204, 536, 537 phycocyanobilin (PCB) 148, 149, 163, 166, 167, 174, 179, 189, 190, 202, 533, 534, 535, 537 See also phycobilin attachment 202 binding pocket 156 (3E)-phycocyanobilin 533 (3Z)-phycocyanobilin 533, 536 replacement by PEBs 166 phycoerythrin (PE) 8, 18, 19, 27, 28, 29, 30, 32, 76, 101, 146, 167, 176–179, 188, 190, 198, 536, 537, 630, 641, 643, 645, 649, 653, 678, 826, 833–836, 838, 839 subunit 204 amino acid sequence 177 b-phycoerythrin (b-PE) 186 See also phycoerythrin, Bphycoerythin B-phycoerythrin (B-PE) 167, 173, 179, 181, 185–189, 190 See also phycoerythrin, b-phycoerythrin, R-phycoerythrin subunit 173, 189, 190 crystal structure 173, 185, 186 crystals 179 subunit 203 C-phycoerythrin (C-PE) 167, 177, 188 C-phycoerythrin I (C-PE-I) 181, 183, 189 amino acid sequences 183 subunit 189 C-phycoerythrin II (C-PE-II) 178, 183, 189 subunit 189 Cryptomonas sp. strain CS-24 204 evolution 190 subunit 148, 176, 177, 179–181, 184, 185, 190, 202– 204 nuclear-encoded 186 immunolocalization 838 picosecond time-resolved fluorescence 33 R-phycoerythrin (R-PE) 186, 203 structure 172, 179 phycoerythrin I 166 phycoerythrin II 166, 189, 190, 202 phycoerythrin-545 (PE-545) 183, 188, 190 subunit 189 phycoerythrin-566 (PE-566) 148, 183, 189 subunit 189 phycoerythrobilin (PEB) 148, 149, 163, 166, 176–181, 186, 187, 189, 202, 536 See also phycobilin (3E)-phycoerythrobilin 533 (3Z)-phycoerythrobilin 533, 536 doubly linked 149 phycoerythrobilin:phycocyanobilin isomerase 522, 536 phycoerythrocyanin (PEC) 8, 19, 146, 153, 165, 167, 173, 174, 537 light intensity regulation 165

874 photochemistry 165 phycourobilin (PUB chromophores) 29, 148, 149, 163, 166, 176– 178, 181, 186, 189 doubly linked 149 See also phycocyanin, phycoerythrin, phycobilin phylloquinone (vitamin 321, 328, 335, 420, 421 5'-hydroxyphylloquinone 421 phylogenetic trees 11, 14–21, 168, 170 phylogeny 14–21, 50–55, 81, 92, 93, 95, 96, 102–111 ‘physiological specialization’ Trichodesmium sp. bundles 39 phytochrome 645, 834, 839 phytoene 562, 563 See carotenoid 15-cis-phytoene 569 phytoene desaturase 108, 123, 132, 133, 568, 569, 570, 571 phytoene synthase 108, 133, 569, 570, 572 phytoflavin See flavodoxin phytofluene 563, 569 See carotenoid phytol 545, 546, 688 biosynthesis 545 phytoplankton 27–30, 462 phytylation 545, 546 phytyl pyrophosphate 546 picoplanktonic cyanobacteria 27–30, 462 pigment 92, 93, 104, 108, 142 See also carotenoid, chlorophyll, phycobiliprotein pili 831, 834 See also fimbriae pilot protein 760 See also SecB plankton 28 plant 91–97, 100–109, 111, 681 plasma-desorption mass spectrometry muropeptides from peptidoglycan of C. paradoxa 69 plasmalemma 470, 780 See also plasma membrane, cytoplasmic membrane plasma membrane 53, 119–126, 131–133, 261, 288, 411, 413, 418–428, 471, 474, 475, 480 See also cytoplasmic membrane, plasmalemma -thylakoid membrane contacts 125 plasmid 33, 107, 418 cosmid 586 helper 587, 588, 773 RP-4 mediated conjugation 773 IncP (conjugative) 583, 584 IncQ 584, 586, 590 neutral-site 602 integrative vectors 584 pDU1 773 platform vector 602 self-transmissible conjugative 587 shuttle vectors 584 table, Chapter 19 586 plastid 11 autogenous origin 91 complex 93 DNA 186 evolution 82, 83 genome 66–84, 91–111, 203, 204, 545 origins 81–84, 91–111 phagotrophic host(s) 93, 94, 111 Marchantia polymorpha (liverwort) 15, 16 polyphyletic origins 92, 93, 105, 107, 111 procaryotic progenitors 93 xenogenous origin 91

Subject Index plastocyanin 11, 129, 130, 260, 264, 277, 281, 287, 289, 303, 325, 326, 328, 337, 338, 339, 389, 392–396, 410, 411, 413, 416, 420–425, 602, 659 crosslinking to PsaF 338 kinetic studies of donation to 337 poplar 393 plastocyanin:ferredoxin oxidoreductase See Photosystem I plastohydroquinone See plastoquinol plastoquinone/plastoquinol 125, 218, 223, 224, 260, 264, 269, 281, 283, 295, 296, 298, 322, 397, 409, 410, 420, 678, 680, 682, 687, 689, 781, 826, 835 oxidation 277, 352 plastoquinol-1 264 -plastocyanin oxidoreductase 264 plastoquinone-9 288, 420–423 pool 60, 261, 283, 284, 286, 680 plastoquinol-cytochrome oxidoreductase 259 piastoquinol-plastocyanin oxidoreductase 259, 264, 266, 422 platform vector 602 See plasmid Pleurocapsales 6 polyacrylamide gel electrophoresis (PAGE) discontinuous 8 ‘green’ gels 56 nondenaturing 444 sodium dodecyl sulfate 56, 61, 160, 191, 192, 198, 477 polyamines 7 polymerase chain reaction (PCR) 10, 12, 21, 95, 279, 461 ‘polyphasic’ taxonomy 3 polyphosphate granules 664 polyprotein 290 polysaccharide 772, 784, 785, 800 See also glycogen granules 412, 421, 663, 734, 789, 831, 838 polysomes 688, 760 thylakoid-bound polysomes 688 pores 780 porin 121–124, 135, 488 porphobilinogen (PBG) 527, 529 porphobilinogen (PBG) deaminase 522 porphobilinogen (PBG) synthase 521, 522, 527, 528 positive selection 587 positive-inside rule 287 post-transcriptional regulation 694, 701–704, 798 post-transcriptional responses 702 post-translational modification 173, 290, 789, 798 potassium thiocyanate use in Photosystem II core preparation 223 See plastoquinone/plastoquinol pre-chromophyte alga 93 See also Chromophyta precorrin 2 529 prenyl transferases 77, 78 prephytoene pyrophosphate 569 presequence 276, 288, 337, 339, 341 preuroporphyrinogen 528 primary productivity 27, 29, 42 limitation by iron in oceans 42 primer extension mapping 623 See also promoter, transcription PRK See phosphoribulokinase procaryote 92, 93, 96 Prochlorales 50, 55 prochlorophyte 28, 29, 49–64, 67, 92, 93, 103–105, 120, 134, 220, 538, 546, 695 phylogenetic relationships 10, 14–17, 19, 54, 55 transmission electron micrograph Prochlorothrix hollandica 58

Subject Index ultrastructure 56–59 See also oxychlorobacteria unstacked membranes (stroma thylakoids) 57 proheterocysts 794, 800, 803–805, 809 promoter 101, 441, 614–634, 702–704 activity analyses in vivo 628 assay vector 702 Anabaena sp. strain PCC 7120 624 basal 617, 618, 623, 627–629, 632 Calothrix sp. strain 7601 627 cis elements 629, 634, 635, 694 eucaryotic 619 gene expression for cyanobacteria 602 phage lambda promoter 587 602, 603 602, 603 phoA 667 primer extension mapping 623 psbA 585, 587, 605 untranslated leader regions 702 psbO 585 recA 603 regulated 618 promoter 290, 617, 618 promoter 619 switching 632 Synechococcus sp. strain 7002 625 Synechococcus sp. strain PCC 7942 626 T4 627 T7 598 tac 587 propionate 530 protease 798 inhibitors 145 protein 2, 7, 11, 12, 142 crystallography 363 disulfide reductase 716, 718, 720, 723, 726 electrophoresis 7 export 95 See also protein secretion bacterial 760 docking protein, SecA 760 pilot protein, SecB 760 folding 716, 757–762, 760 intermediates 758 kinase 798, 839 localization 288 phosphorylation 44, 716, 798 secretion 95 sequencing 276 synthesis inhibitors 203 translocation 80 protein A-colloidal gold complex 124 proteobacteria (‘purple bacteria’) proteobacteria 366 proteobacteria 106 proteobacteria 106 proteoliposomes 280, 362 proto-chloroplast 53 protochlorophyll 544 protochlorophyllide a 524, 538, 540–547, 688 reduction 95, 522 protochlorophyllide reductase 11, 95, 387, 482, 542–545, 591 See also NADPH:protochlorophyllide oxidoreductase

875 light-dependent 522, 543, 544 light-independent 522, 543–545 protoheme 266, 534 protoheme 8-methyl oxidase 522 proton pump 376 See also ATP synthase mechanism 369 transfer 280 translocation 281 protoplasmic fracture faces 59 protoporphyrin IX 531, 538–540 photodynamic sensitizer 531 protoporphyrinogen IX 530,531 protoporphyrinogen IX oxidase 522, 530, 531, 532, 547 inhibition by diphenyl ether herbicide 531, 548 PS I See Photosystem I PS II See Photosystem II psaD mutant 349, 350 psaE mutant 328, 354 psaF mutant 339 psaF psaJ double mutant 341 psaK mutant 327, 342 psaL mutant 342 pteridophyte 95, 837 PUB See phycourobilin pulse-labeling 686, 688 pulse-chase 686 pulsed-field gel electrophoresis 109, 773 purine 471, 472, 481 purple bacteria 134, 261 group 366 See also proteobacteria nonsulfur photosynthetic bacteria 451 reaction center 697 PXB See phycobiliviolin pyridine nucleotide dehydrogenase 781 pyridoxal-phosphate 526 pyridoxamine 526 pyrrole 2,3-reductase 535 pyruvate 439, 440, 462, 496, 777, 778 :flavodoxin oxidoreductase 496, 777, 778 oxidase 383

Q Q cycle 281, 282, 283, 284, 298, 422 model 272, 281 modified 281 See Photosystem II See Photosystem II See Cytochrome Qred site See Cytochrome site See Cytochrome quantum yield 320, 679, 683 quinacrine 544 quinol 416, 420 -binding region, cytochrome 295 oxidoreductase 427 oxidase 284, 303 oxidation 268–270, 279–282, 291–296, 297, 299 inhibitors 271 inhibitor-resistance mutations 292 site, Cytochrome site quinolinate synthetase 77

876 quinone 2, 419, 420, 687, 697 See also plastoquinone/plastoquinol, ubiquinone/ubiquinol, napthoquinone/napthoquinol analogs 280 -binding sites 264, 269, 279, 280, 291, 297 photoaffinity labeling 264, 269 cycle 281 site See Cytochrome -reduction 268, 269, 281, 282, 296, 299 inhibitors 297 inhibitor resistance mutations 296, 297 quinone pool 412, 413, 414, 415, 416, 418, 420, 421

R Raman spectroscopy 272, 276 resonance Raman spectroscopy 272 random cartridge mutagenesis 594 random integrational mutagenesis 595 random mutagenesis 451, 635 reaction center 266, 689, 694, 697 See also Photosystem I, Photosystem II bacterial 267, 697 complexes 226, 227 iron-sulfur type 321 L subunit 107 M subunit 107 Photosystem II 57 quinone-type 321 recombination double-crossover 585 non-reciprocal 591 single-crossover 585 reconstitution studies phycobiliprotein-linkerpolypeptides 154, 158, 197–199 Rieske protein 275, 279, 280, Photosystem I 335, 346, 352, 362, 373 red algae 146, 166, 167, 177, 178, 179, 181, 186, 190, 191,203, 679, 681, 682, 684, 686 phycoerythrins 178 red light 164, 165, 706, 642–654, 830–834, 839 redox -active tyrosine residues 237, 240 centers 263 difference spectroscopy 277 titration 266, 680 trap 336, 347 -sensing 286 -state dependence 60 reductive pentose phosphate cycle 457–460 REP domains See linker polypeptide, core-membrane, ApcE repetitive DNA sequences 599 replicon 584 reporter fusions/systems 269, 594, 596–598, 702, 773 See transcription cat (chloramphenicol acetyl transferase) 596 GUS 596, 603 lacZ 596 psbA-lacZ fusions 701 luxAB (luciferase) 596,597 repression 618, 630 repressors 628, 631 repressor proteins 618 binding sites 634

Subject Index resonance Raman spectroscopy 272 respiration 284, 354, 409, 410–429, 495, 772, 789 dark 53 respiratory electron transport chain 288, 409, 410, 424, 426, 428, 732, 778 See also electron transport chain respiratory-defective mutations 301 respiratory enzymes 125 response regulators 37, 40, 290 See also two-component sensory systems restriction endonuclease 31, 35, 588, 604, 773 AquI AvaI 587, 588, 604 AvaII 587, 588, 591, 604 AvaIII 587, 588 Type I restriction systems 605 restriction fragment length polymorphism (RFLP) 8–10, 21, 30, 42 cluster analysis 30 RFLP analysis 30 restriction modification enzymes AvaI methylase 588 Eco47II methylase 588 EcoT22I methylase 588 reverse genetics 302 reverse transcriptase 107 rhodanese 122, 663 Rhodophyta 92, 93 See also red algae and rhodophyte rhodophyte 92–96, 100–111, 534, 539, 450 ribonucleotide reduction 239, 716–718, 720, 726 ribonucleotide reductase 239, 718, 720, 726 ribosomal RNA See rRNA ribosome 70S ribosomes 203 80S ribosomes 203 polysomes 688, 760 thylakoid-bound polysomes 688 ribosomal protein 73, 74, 94, 96, 98 alpha operon 73, 74, 96, 98 S10 operon 73, 74, 96–98 spc operon 73, 74, 96–98 str operon 73, 74, 96–98, 102 ribosomal RNA See also rRNA 5S rRNA 81, 103 16S rRNA 2, 7, 10, 12, 14, 15, 16, 17, 19, 20, 21, 54, 70, 81 23S rRNA 2, 21, 70 ribonucleic acid See RNA ribulose 1,5-bisphosphate (RuBP) 438, 480 ribulose 1,5-bisphosphate carboxylase/oxygenase (RubisCO) 76, 81, 92, 94, 100, 103, 105–107, 111, 203, 411, 438–462, 470, 471, 477–480, 482, 484, 761, 792, 833, 838 activase 447, 456–458 activator 440 carbamate formation 440 active site 449 conformational changes induced by S subunit 449 assembly 443 binding protein 441 biological selection of mutant enzymes 451, 452 isotope discrimination 470 carbon isotopic fractionation 471 2-carboxyarabinitol monophosphate (CA1P) 456 chromophytic enzyme 450 electrophoretic mobility of the recombinant holoenzyme 444

877

Subject Index enediol intermediate 438, 440, 447 fallover products 456 flanking gene organization unicellular strains 457 heterocystous strains 457–460 hybrid enzyme 450 for 446 440 dimers 440 structure 440, 445, 446 enzyme 440 large (L) subunit 81, 92, 106, 440, 443, 697 barrel domain structure 442, 450 loop 6 450 monooxygenase reaction 438 open reading frames near rbcLS genes 457 oxygenase 455 partition coefficient 440 rbcLS operons 457, 458 antisense and sense RNA probes 461 spacer region 366, 457 reactions 439 recombinant protein 441 rhodophytic enzyme 450 small subunit 81, 106, 440–450 hairpin loop, role in expression 447 nuclear-encoded 449 specific activity 445 specificity factor 440 structure 442 ternary complex in activation 440, 455 transcriptional control 460 turnover number 440 Type I enzyme 441 Type II enzyme 446 vector-promoter system for expression 451 whole-cell assays for activity 456 X-ray structure 441 ribulose 5-phosphate 438 rice plastid genome 93 Rieske Fe-S protein 75, 260, 262–264, 269–276, 279, 281–283, 287, 288, 290, 298–303, 413, 422 [2Fe-2S] center 75, 262–264, 270–276, 277, 281, 282, 287, 288, 298–302 EPR 271 pH dependence 271 redox potential 270, 271 hydropathy analysis 274, 275 model 273 resonance Raman spectroscopy 272 role as assembly protein 288 temperature-sensitive mutations 300, 301 rifampicin (rifamycin) 701, 707, 806 RNA See ribonucleic acid RNA polymerase 94, 95, 100, 101, 614, 615, 617, 618, 619, 620, 622, 623, 627, 628, 630, 633, 634, 797 subunit 620, 621 See also RpoA subunit 620 See also RpoB subunit 620, 632 See also RpoC2 subunit (plastid) 620 See also RpoC2 subunit 620, 632 See also RpoC1 Anabaena sp. strain PCC 7120 626

core structure 615, 620 footprinting studies 632 sigma factor/subunit 290, 614–619, 621–628, 630–634, 702, 704, 705, 797, 826, 833, 835, 836, 839 RNaseA inhibition of formation 523 RNasin 523 rotenone 131,418, 419 rRNA 5S rRNA 81,103 16S rRNA 2, 7, 10, 12, 14, 15, 16, 17, 19, 20, 21, 54, 70, 81, 105 secondary structure model, 16S rRNA 13 23S rRNA 2, 21, 70 internal transcribed spacer (ITS) 21 LSU rRNA 105 See also 23S rRNA SSU rRNA 93, 103 See also 16S rRNA

S S states 218, 235 state 245, 246 state 223, 245, 246 state 222, 223, 241–245, 247 multiline signals 244 state 222, 245 S-adenosyl-L-methionine 540, 541 S-adenosyl-L-methionine:Mg-protoporphyrin IX methyltransferase 540, 548 schizokinen 741,742 SDS See detergent SDS-PAGE 56, 61, 160, 191, 192, 198, 477 second-site (suppressor) mutations 291, 452 secondary endosymbiotic events 109, 111 secondary metabolite 806 secondary structure elements 167 prediction 167 secretory pathway (protein) 759 See also protein export bacterial 760 sedoheptulose-1,7-bisphosphatase 718 segregation of mutants 595 self-transmissible conjugative plasmids 587 semiquinone 270, 272, 281, 298 See also plastoquinone, quinone anion 282 anion radical 335 cycle 282 serine protease 798 seryl-phosphorylation 835 short-term adaptation 679, 679–683, 687, 689 model 690 shuttle vectors 584 siderophore 659, 732, 740 aerobactin 741 catechol-type 742 enterobactin 741 ferrichrome 741 hydroxyamate-type 742 production 42 table 744 factor/subunit 290, 614–619, 621–628, 630–634, 702, 704, 705, 797, 826, 833, 835, 836, 839 See also RNA polymerase, transcription

878 alternative 615, 616, 618, 622, 634 Group 1 616 Group 2 616 Group 3 616 (RpoH) 622, 752 (RpoN) 616–619, 622 Bacillus subtilis (RpoD) 621, 797 domains 616 Escherichia coli (RpoD) 621 principal/primary 615, 616, 618, 622, 633 290, 615–619, 622–628, 630, 632, 702, 704, 705, 754 Streptomyces coelicolor Hrd proteins 616, 622 signal sequence targeting 204 processing site 339 signal transduction pathway psbA expression 706 singlet-state oxygen 561 siroheme 529, 532 biosynthesis 529 sirodihydrochlorin 529 siroheme ferrochelatase 522 site-directed mutagenesis See mutagenesis, site-directed site-directed mutant See mutant site-directed mutations See mutations, site-directed site-specific recombinase 775, 776 See also excisase XisA 775 XisC 776 XisF 775 Sn-protoporphyrin IX 534 sodium 474, 792 -dependent ATP synthesis 369 antiporter 474 pump 369, 474 role of in bicarbonate uptake 474 symport 474 sodium cholate See detergents sodium dodecyl sulfate See detergents spacing ARM See linker, core-membrane linker phycobiliprotein species 2 specific absorbance coefficients carotenoids 563 spectinomycin resistance 588 spectral light quality 826, 830, 834, 836 See also light spectroscopy See also fluorescence fluorescence spectroscopy at 77 K 56 FT-IR 240 magnetic circular dichroism (MCD) 267 optical kinetic 336 optical redox-difference 266 NMR 265 redox difference 277 resonance Raman 272, 276 two-color femtosecond transient absorption 176 X-ray absorption 244 spheroplasts 415,426,429 spillover 680, 682 spinach 227, 245, 265, 267, 364, 528, 534, 536 chloroplasts 245 PS II membranes 245 thylakoids 536 spores 771

Subject Index SQR See sulfide-quinone reductase Src homology 3 (SH3) domains 351 state transition 52, 56, 60, 61, 266, 286, 343, 162, 679–683, 689 detachment model 682 light state 1 60, 155, 162, 679 light state 2 60, 155, 162, 679 mechanism in phycobilisome-Chl a system 681 model 681 preferential excitation 680 PS I light 684 PS II light 684 signal monitoring system 687 thylakoid destacking 680 stereochemistry 366, 367 stigmatellin 271, 272, 280, 292–294 Stigonematales 6 streptomycin resistance 588 See also omega cartridge stringent response 798 strobilurin A 280, 292, 294 See also mucidin stromal thylakoids See thylakoids, stromal Styloviridae 34, 35 See also cyanophage substitution rate nucleotide 103 substitutional bias 103 subunit IV (SU IV) 260, 262–270, 272, 275, 279–281, 283, 287– 289, 291, 296, 297, 299, 422 See also cytochrome complex relationship to cytochrome 266–270 succinate 414, 422 succinate -cytochrome c oxidoreductase 414 succinate -cytochrome c reductase 300 succinate dehydrogenase 778 oxidoreductase 414 succinyl coenzyme A 521 sucrose 780 density-gradient centrifugation 56, 224, 264, 266 sucrose synthase 779, 796 sulfate 123, 402, 679, 725 metabolism 716, 725 permease 121, 123, 662 transporter 123 uptake 95, 661, 662 sulfate reductase 383, 725 sulfhydryl group 373 sulfide 725 sulfide-quinone reductase (SQR) 261 sulfite 725 sulfite reductase 529 heme protein 500 sulfur See also sulfate, sulfide, sulfite, thiosulfate availability 201 -limited growth conditions 165, 201 S-deficiency Anabaena variabilis 657 Gleocapsa alpicola 657 Nostoc muscorum 657 Oscillatoria prolifera 657 Phormidium luridum 657 Plectonema boryanum 657 Synechococcus 657 Synechococcus cedrorum 657 starvation 151, 657 sun-shade adaptation 689

Subject Index supercomplex 266 See cytochrome complex, cytochrome complex superoxide dismutase 132, 134, 222, 710 immunolocalization 788 superoxide radical 132 suppressor mutations 291, 452 symbiosis 589, 825, 826, 837 hormogonia role in infection process 837 Macrozamia sp.-Nostoc sp. 455 symbiotic cyanobacteria 8–10 analysis by immunostaining 9 systematics 2 See also taxonomy

T T7 DNA polymerase complex with thioredoxin 716, 717 T7 RNA polymerase gene 598 targeted mutagenesis See mutagenesis targeting process 203 targeting sequence See signal sequence 204 taxonomy 4 bacteriological approach 4 botanical approach 4 ‘polyphasic’ 3 terminal emitter See ApcD, ApcE, linker polypeptide, coremembrane linker phycobiliprotein, phycobilisome core terminal oxidase 278, 284, 423–425, 428 See also cytochrome oxidase 423 tetrahydrogeranylgeranyl See phytol biosynthesis tetrahydroporphyrin (sirodihydrochlorin) 529 N, N, N', N'-tetramethyl-p-phenylenediamine (TMPD) 285 tetrapyrrole 140, 142, 143, 146, 148, 520–548 See also chlorophyll, heme phycobilin biosynthesis 520–548 enzymes (table) 522 thermal regulation role of DnaK 756 thermal tolerance 756 See also heat shock thermophilic Synechococcus sp. 265, 283, 421–424 thin-layer chromatography carotenoids 562 thioether linkage 148, 276, 533 heme 276 phycobilin 148, 533 thioredoxin 78, 373, 716–726,778 complex with T7 DNA polymerase716, 717 denaturation 722 f-type (thioredoxin-f) 718 genes 718 immunolocalization in heterocysts 778 m-type (thioredoxin-m) 717, 718, 723 tertiary structure 721 thioredoxin-f 718 thioredoxin-m 717,723 three-dimensional structures 720 thioredoxin reductase 383, 721, 722 thiosulfate 663 thylakoid membrane 50, 51, 53, 56, 57, 58, 59, 60, 61, 62, 119, 120, 121, 125, 126, 127, 130, 132, 133, 134, 261, 288, 373, 409, 411, 418, 532, 678–690, 762 abundance, Synechocystis sp. strain PCC 6714 686

879 appressed 57, 62 assembly 761 -bound polysomes 688 destacking 680 centers 411 lumen 204 non-stacked stromal 680 phycobilisome-covered 681 spinach 245, 536 stacking 53, 57, 59, 680 stromal thylakoids 53, 57, 266, 680 topographical mapping 134 topography 133 ‘unit thylakoid’ 686 TMAO See trimethylamine-N-oxide TMPD See N, N, N', N'-tetramethyl-p-phenylenediamine tobacco mosaic virus movement protein 603 topoisomerase I (TopA) 482 toxins See Microcystis sp. trans-acting factors roles in transcription 618, 628, 634 trans-splicing 97, 102 transaminases 510 transamination 509, 510 transcription 614–635, 701 accessory factors 630 activation 618, 630, 633 activator 39, 628, 631, 634, 635 activator binding sites 618, 619, 634 activator gene for Calvin cycle enzymes 458 activator protein 618, 619 assays performed in vitro 633 cis elements 629, 634, 635, 694 coactivator 619 control in heterocyst development 796 in rbcLS expression 460 in heat shock response 764 eucaryotic 634, 635 footprinting assays in vitro 631 in vivo 631 induction 701 initiation 101, 632 initiation sites 617, 628, 633 integration host factor (IHF) 619 in vitro assays 623, 626, 627, 633 in vivo assays 623, 629 negative element in psbA regulation 703 primer extension mapping 623 regulation 95, 102 regulators 510 Crp family 631 reporter fusions 472, 594, 596, 597, 702 cat 472 luxAB 594, 597 psbA-lacZ 594, 596, 702 start points 627 termination 36 transcript stability, psbA 701–702 units 201 transduction 774

Subject Index

880 transfer RNA See tRNA transformation 54, 302, 590, 604, 606 competency 590, 604 natural 54 particle bombardment, Chlamydomonas reinhardtii transformation 302 transhydrogenase 414, 416 transhydrogenation 782 transient absorption spectroscopy 176 two-color femtosecond 176 transit sequence/peptide (gamma) subunit of phycoerythin of Aglaothamnion neglectum 186 glyceraldehyde 3-phosphate dehydrogenase 108 PsaF 337 translation 94–97, 108 peptide elongation 688 translational control of PsaA and PsaB synthesis 688 translocation See also protein export, protein secretion of genes 99 of proteins into mitochondria 759, 760 transport 474 dicarboxylic acid 617 multicomponent systems 498 transport channels 121, 129 transposon 589, 598, 801 mutagenesis 33, 593, 594, 598, 774 Tn5 594, 773 tricarboxylic acid cycle 414, 778 See also citric acid cycle tricylindrical (APC) core 159–161 See also phycobilisome, core trimethylamine-N-oxide (TMAO) 285 triose-phosphate 438 triose-phosphate isomerase 108 triparental matings 584 triplet state chlorophyll and carotenoid 561 Triton X-100 See detergent Triton X-l14 See detergent tRNA 94, 95, 96, 97, 102, 524 -Ala 21 -Glu 521, 523 -Ile 21 trypsin 60, 275, 287, 373, 374 tryptophan 544 fluorescence 391 tryptophan synthase 78 turbidostat 700, 707 turgor pressure 832 two-component sensory systems 44, 798 See also transcription regulator tyrosine radical See also tyrosine, tyrosine, tyrosine 218, 223, 238

U ubiquinol-cytochrome c oxidoreductase See cytochrome complex

ubiquinone 261, 419 UDP-N-acetylmuramyl pentapeptide 69 UHDBT See 5-n-undecyl-6-hydroxy-4,7 dioxobenzothiazol ultraviolet (UV) light absorption studies 245, 246 mutagenesis 592 uncoupled respiration 364 phenotype of unc (atp) mutants 364 undecaprenyl phosphate role in peptidoglycan biosynthesis 69 5-n-undecyl-6-hydroxy-4,7 dioxobenzothiazol 271, 272, 280 University of Texas (UTEX) Culture Collection 697 uptake hydrogenase See hydrogenase urea 503, 504 N-source 503 urease 492, 503 uridylylation 835 uroporphyrinogen decarboxylase 522, 530, 532 uroporphyrinogen I 528, 530 uroporphyrinogen III 527, 528, 530 uroporphyrinogen III synthase 522, 527, 528, 529 spiro intermediate 528 uroporphyrinogen methyltransferase 522

V See BifA valinomycin diffusion potential in thylakoids 373 vanadium 496, 775 vanadium-dependent nitrogenase See nitrogenase, vanadiumdependent 496, 589 vegetative trichomes 825–828, 831, 833, 838, 839 violaxanthin See carotenoids 3-vinyl-2,3-dihydrobilin 535 4-vinyl reductase 522, 542, 543, 547 See also BchJ ‘viral loop, the’ 34 vitamin 521, 529, 548 vitamin E 546

W wheat 220, 542 etioplasts 541, 542

X XANES See X-ray absorption spectroscopy xanthophyll See carotenoids xenogenous origin hypothesis for plasitids 91, 92 X-ray absorption spectroscopy 223, 241, 244–247 K-edge shifts 246 X-ray absorption extended fine structure (EXAFS) 244–247, 367 X-ray absorption near-edge structure (XANES) 244,246 X-ray crystallography 267, 276, 286, 528 crystal structure 277

Subject Index Y yeast 267, 524, 528, 530–532, 716, 717

Z tyrosine 218, 224, 233, 237, 238, 240 zeaxanthin See carotenoids (zeta) carotene See carotenoids Zwittergent 14 See detergent

881