Advances in Microbial Physiology Volume 52

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Advances in

MICROBIAL PHYSIOLOGY VOLUME 52

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Advances in

MICROBIAL PHYSIOLOGY Edited by

ROBERT K. POOLE West Riding Professor of Microbiology Department of Molecular Biology and Biotechnology The University of Sheffield Firth Court, Western Bank Sheffield S10 2TN, UK

Volume 52

Amsterdam Boston Heidelberg London New York Oxford Paris San Diego San Francisco Singapore Sydney Tokyo ACADEMIC PRESS

Academic Press is an imprint of Elsevier 84 Theobald’s Road, London WC1X 8RR, UK Radarweg 29, PO Box 211, 1000 AE Amsterdam, The Netherlands The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, UK 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA First edition 2007 Copyright r 2007 Elsevier Ltd. All rights reserved No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone (+44) (0) 1865 843830; fax (+44) (0) 1865 853333; email: [email protected]. Alternatively you can submit your request online by visiting the Elsevier web site at http://elsevier.com/locate/permissions, and selecting Obtaining permission to use Elsevier material Notice No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent veriﬁcation of diagnoses and drug dosages should be made ISBN-13: 978-0-12-027752-0 (volume) ISBN-10: 0-12-027752-2 (volume) ISSN: 0065-2911 (series) For information on all Academic Press publications visit our website at books.elsevier.com

Printed in Great Britain 07 08 09 10 11

10 9 8 7 6 5 4 3 2 1

Contents

CONTRIBUTORS TO VOLUME 52 . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

ix

Oxygen, Cyanide and Energy Generation in the Cystic Fibrosis Pathogen Pseudomonas aeruginosa Huw D. Williams, James E.A. Zlosnik, Ben Ryall

1. Introduction to Pseudomonas aeruginosa . . . . . . . . . . . . . . . 2. Oxygen and P. aeruginosa Infection of the Cystic Fibrosis Lung – the Scope of this Review. . . . . . . . . . . . . . . . . . . . . 3. Means of Energy Generation in P. aeruginosa . . . . . . . . . . . 4. Aerobic Respiration in P. aeruginosa. . . . . . . . . . . . . . . . . . 5. Anaerobic Respiration . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6. Fermentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7. Anaerobic Metabolism in the Cystic Fibrosis Lung . . . . . . . 8. Synthesis of the Respiratory Inhibitor Hydrogen Cyanide in P. aeruginosa . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9. Mucoid Conversion of P. aeruginosa in the Cystic Fibrosis Lung: the Role of Oxygen and Energy Metabolism . . . . . . . 10. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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3 4 7 7 31 40 42

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43

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47 50 50

Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Introduction: Enzymic Mechanisms to Combat Oxidative and Peroxidative Stress. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

74

Structure, Mechanism and Physiological Roles of Bacterial Cytochrome c Peroxidases John M. Atack, David J. Kelly

74

vi

CONTENTS

2. Phylogenetic Analysis of Bacterial CCPs Reveals a Novel Sub-Group of Tri-Haem Proteins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79 3. MauG Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83 4. Structure of Bacterial CCPs . . . . . . . . . . . . . . . . . . . . . . . . . . 83 5. Mechanistic Aspects of Catalysis by Bacterial CCPs . . . . . . . . . 88 6. Electron Donors and Electron Transport in Bacterial CCPs . . . 90 7. Roles of CCPs in Bacterial Cells. . . . . . . . . . . . . . . . . . . . . . . 93 8. Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 98 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 98 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 98

Respiratory Transformation of Nitrous Oxide (N2O) to Dinitrogen by Bacteria and Archaea Walter G. Zumft and Peter M.H. Kroneck Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Chemistry of N2O . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Genomic and Organismal Resources . . . . . . . . . . . . . . . 4. Properties of N2O Reductase . . . . . . . . . . . . . . . . . . . . 5. Structure of N2O Reductase . . . . . . . . . . . . . . . . . . . . . 6. Novel Cu Centres in N2O Reductase. . . . . . . . . . . . . . . 7. Organization of nos Genes, Gene Expression, Regulation 8. Evolutionary Aspects and Phylogenetic Relationships. . . 9. Topology and Transport Processes . . . . . . . . . . . . . . . . 10. Cu Centre Assembly . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Electron Donation and Maintenance of Activity in vivo . 12. A Glimpse of History . . . . . . . . . . . . . . . . . . . . . . . . . 13. Conclusions and Perspectives . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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109 110 112 114 127 131 136 152 157 168 175 180 194 196 197 197

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. The Cyanobacterial Circadian Clock: The S. Elongatus PCC 7942 Kai Locus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

231

A Circadian Timing Mechanism in the Cyanobacteria Stanly B. Williams

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CONTENTS

vii

3. Sequence, Structure and Function of Clock Proteins and the Kai-Clock Complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Clock-Controlled Gene Expression . . . . . . . . . . . . . . . . . . . . 5. Clock Input . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6. Other Components: The rpo (Sigma Factor) and cpmA Genes . 7. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

245 266 270 274 276 281 282

AUTHOR INDEX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

297

SUBJECT INDEX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

333

Colour Plate Section to be found in the back of this book

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Contributors to Volume 52

JOHN M. ATACK, Department of Molecular Biology and Biotechnology, The University of Shefﬁeld, Western Bank, Shefﬁeld S10 2TN, UK DAVID J. KELLY, Department of Molecular Biology and Biotechnology, The University of Shefﬁeld, Western Bank, Shefﬁeld S10 2TN, UK PETER M. H. KRONECK, Faculty of Biology, University Konstanz, D-78464 Konstanz, Germany BEN RYALL, Division of Biology, Faculty of Natural Sciences, Imperial College London, Sir Alexander Fleming Building, London SW7 2AZ, UK HUW D. WILLIAMS, Division of Biology, Faculty of Natural Sciences, Imperial College London, Sir Alexander Fleming Building, London SW7 2AZ, UK STANLY B. WILLIAMS, Department of Biology, Life Science Building, University of Utah, Salt Lake City, UT 84112, USA JAMES E. A. ZLOSNIK, Division of Biology, Faculty of Natural Sciences, Imperial College London, Sir Alexander Fleming Building, London SW7 2AZ, UK WALTER G. ZUMFT, Institute of Applied Biosciences, Division of Molecular Microbiology, University Karlsruhe, D-76128 Karlsruhe, Germany

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Oxygen, Cyanide and Energy Generation in the Cystic Fibrosis Pathogen Pseudomonas aeruginosa Huw D. Williams, James E.A. Zlosnik and Ben Ryall Division of Biology, Faculty of Natural Sciences, Imperial College London, Sir Alexander Fleming Building, London SW7 2AZ, UK

ABSTRACT Pseudomonas aeruginosa is a Gram-negative, rod-shaped bacterium that belongs to the g-proteobacteria. This clinically challenging, opportunistic pathogen occupies a wide range of niches from an almost ubiquitous environmental presence to causing infections in a wide range of animals and plants. P. aeruginosa is the single most important pathogen of the cystic ﬁbrosis (CF) lung. It causes serious chronic infections following its colonisation of the dehydrated mucus of the CF lung, leading to it being the most important cause of morbidity and mortality in CF sufferers. The recent ﬁnding that steep O2 gradients exist across the mucus of the CFlung indicates that P. aeruginosa will have to show metabolic adaptability to modify its energy metabolism as it moves from a high O2 to low O2 and on to anaerobic environments within the CF lung. Therefore, the starting point of this review is that an understanding of the diverse modes of energy metabolism available to P. aeruginosa and their regulation is important to understanding both its fundamental physiology and the factors signiﬁcant in its pathogenicity. The main aim of this review is to appraise the current state of knowledge of the energy generating pathways of P. aeruginosa. We ﬁrst look at the organisation of the

Copyright r 2007 by Elsevier Ltd. ADVANCES IN MICROBIAL PHYSIOLOGY VOL. 52 All rights of reproduction in any form reserved ISBN 0-12-027752-2 DOI: 10.1016/S0065-2911(06)52001-6

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aerobic respiratory chains of P. aeruginosa, focusing on the multiple primary dehydrogenases and terminal oxidases that make up the highly branched pathways. Next, we will discuss the denitriﬁcation pathways used during anaerobic respiration as well as considering the ability of P. aeruginosa to carry out aerobic denitriﬁcation. Attention is then directed to the limited fermentative capacity of P. aeruginosa with discussion of the arginine deiminase pathway and the role of pyruvate fermentation. In the ﬁnal part of the review, we consider other aspects of the biology of P. aeruginosa that are linked to energy metabolism or affected by oxygen availability. These include cyanide synthesis, which is oxygen-regulated and can affect the operation of aerobic respiratory pathways, and alginate production leading to a mucoid phenotype, which is regulated by oxygen and energy availability, as well as having a role in the protection of P. aeruginosa against reactive oxygen species. Finally, we consider a possible link between cyanide synthesis and the mucoid switch that operates in P. aeruginosa during chronic CF lung infection.

1. Introduction to Pseudomonas aeruginosa . . . . . . . . . . . . . . . . . . . . . . . . . 3 2. Oxygen and P. aeruginosa Infection of the Cystic Fibrosis Lung – the Scope of this Review. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 3. Means of Energy Generation in P. aeruginosa . . . . . . . . . . . . . . . . . . . . . 7 4. Aerobic respiration in P. aeruginosa . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 4.1. Respiratory Dehydrogenases of P. aeruginosa . . . . . . . . . . . . . . . . . . 9 4.2. Transhydrogenase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19 4.3. Quinones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20 4.4. Cytochrome bc1 Complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20 4.5. Cytochromes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21 4.6. Terminal Oxidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24 4.7. Terminal Oxidases in P. aeruginosa . . . . . . . . . . . . . . . . . . . . . . . . 25 4.8. Cytochrome c Peroxidase. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31 5. Anaerobic Respiration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31 5.1. P. aeruginosa Nitrate Reductases . . . . . . . . . . . . . . . . . . . . . . . . . . 33 5.2. P. aeruginosa Nitrite Reductase . . . . . . . . . . . . . . . . . . . . . . . . . . . 36 5.3. P. aeruginosa Nitric Oxide Reductase . . . . . . . . . . . . . . . . . . . . . . . 37 5.4. P. aeruginosa Nitrous Oxide Reductase. . . . . . . . . . . . . . . . . . . . . . 38 5.5. Regulation of Denitrification Genes . . . . . . . . . . . . . . . . . . . . . . . . . 39 6. Fermentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 40 7. Anaerobic Metabolism in the Cystic Fibrosis Lung . . . . . . . . . . . . . . . . . . 42 8. Synthesis of the Respiratory Inhibitor Hydrogen Cyanide in P. aeruginosa 43 8.1. Physiology of Cyanide Production . . . . . . . . . . . . . . . . . . . . . . . . . . 43 8.2. Genetics of Cyanide Production . . . . . . . . . . . . . . . . . . . . . . . . . . . 44 8.3. Mechanisms of Tolerance to Cyanide . . . . . . . . . . . . . . . . . . . . . . . 44 8.4. Evidence for the Biological Function of Cyanide Produced by P. aeruginosa . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 46

OXYGEN, CYANIDE AND ENERGY GENERATION

9. Mucoid Conversion of P. aeruginosa in the Cystic Fibrosis Lung: the Role of Oxygen and Energy Metabolism . . . . . . . . . . . . . . . . 9.1. Oxygen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.2. Phosphate. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.3. Energy Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.4. Perspective . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.5. A Link between Mucoidy and Cyanide Production. . . . . . . . . 10. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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1. INTRODUCTION TO PSEUDOMONAS AERUGINOSA Pseudomonas aeruginosa is a Gram-negative, rod-shaped bacterium that belongs to the g-proteobacteria (Holt and Kreig, 1984). This clinically challenging, opportunistic pathogen occupies a wide range of niches from an almost ubiquitous environmental presence to causing infections in a wide range of animals and plants (Van Delden and Iglewski, 1998; Tummler and Kiewitz, 1999; Stover et al., 2000). P. aeruginosa has a large genome of around 6.3 million base pairs encoding some 5270 predicted open reading frames on its single chromosome (Stover et al., 2000). Its chromosome possesses signiﬁcantly more distinct gene families (paralogous groups) than Escherichia coli, Bacillus subtilis or Mycobacterium tuberculosis, a factor which may contribute to its broad environmental range (Stover et al., 2000). It is also notable that 9% of the assigned open reading frames of P. aeruginosa encode known or putative transcriptional regulators, which have been hypothesised to enable the bacterium to adapt to a wide range of environments (Juhas et al., 2005). P. aeruginosa is a model organism for a number of key bacterial processes. Considerable research effort has gone into studying its ability to form bioﬁlms (O’Toole et al., 2000), as well as cell-to-cell communication using quorum sensing (Withers et al., 2001; Juhas et al., 2005). Another signiﬁcant characteristic of this bacterium is its intrinsic resistance to antibiotics. This resistance is imparted by a synergy of chromosomally encoded resistance determinants, including an impermeable outer membrane, b-lactamases and efﬂux pumps (Poole, 2001). P. aeruginosa is a classical opportunistic pathogen and is able to cause infections in a range of organisms. Probably reﬂective of its versatility as a bacterium, the range of organisms it is known to infect crosses kingdoms from animals to plants (Walker et al., 2004). P. aeruginosa can cause infections in ﬁsh, reptiles, birds, dogs, sheep and dairy herds. Therefore, although an opportunist, it is clearly a signiﬁcant and versatile pathogen

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capable of not just cross species infections but indeed cross kingdom infections. This range of host species has an advantage for research in permitting the development of numerous infections models. While there are existing infection models in mice (Stieritz and Holder, 1975; Stotland et al., 2000), researchers have taken advantage of P. aeruginosa’s broad host infectivity to develop a range of non-vertebrate eukaryotic infection models. These models include Caenorhabditis elegans, Drosophila melanogaster, Dictyostelium discoideum, Arabidopsis thaliana, Galleria mellonella, silkworm larvae, alfalfa and lettuce (Rahme et al., 1997; Tan et al., 1999; Tan and Ausubel, 2000; D’Argenio et al., 2001; Cosson et al., 2002; Kaito et al., 2002; Silo-Suh et al., 2002; Miyata et al., 2003). In humans, P. aeruginosa is capable of causing a wide range of infections. Many of these are associated with immunosuppression. For example, P. aeruginosa has a high association with AIDS patients (Van Delden and Iglewski, 1998) as well as with neutropenic and mechanically ventilated patients, the latter being associated with high fatality rates (Garau and Gomez, 2003). Indeed, P. aeruginosa is one of the top three causes of nosocomial infections, causing around 10% of all hospital-acquired infections (Van Delden and Iglewski, 1998; Fluit et al., 2000; Hancock and Speert, 2000). Of major signiﬁcance, is the role of P. aeruginosa in causing infections in cystic ﬁbrosis (CF) sufferers, where it is the cause of very high mortality rates (Lyczak et al., 2000). High mortality rates are also seen in infections in burn wound patients (Tredget et al., 1992). Additionally, P. aeruginosa is known to cause severe eye infections in users of soft contact lenses (Lakkis and Fleiszig, 2001). From a researcher’s perspective, these three infections are thought to be especially interesting as they allow examination of infections caused by P. aeruginosa where there is not an underlying susceptibility to infections by a range of organisms as a result of damage to the immune system (Lyczak et al., 2000).

2. OXYGEN AND P. AERUGINOSA INFECTION OF THE CYSTIC FIBROSIS LUNG – THE SCOPE OF THIS REVIEW CF is caused by mutation of the gene encoding the CF transmembrane regulator (CFTR), which functions as a chloride channel in epithelial membranes (Collins, 1992). Several hypotheses link mutations in CFTR to development of chronic P. aeruginosa infections (Ratjen and Doring, 2003).

OXYGEN, CYANIDE AND ENERGY GENERATION

5

A hypothesis for which there is increasing support is the isotonic ﬂuid depletion/hypoxic mucus hypothesis (Fig. 1). This proposes that isotonic salt concentrations resulting from abnormal sodium absorption from the airway lumen coupled with the failure of CFTR to secrete chloride leads to formation of a dehydrated layer above the epithelial cells, which increases mucus viscosity and impairs mucociliary and cough clearance of the lungs (Fig. 1) (Ratjen and Doring, 2003). This allows bacteria to invade and become trapped in this viscous mucus layer. Crucially in CF patients, steep O2 gradients are present within the mucus on CF epithelial surfaces prior to infection, due to abnormal oxygen consumption of the epithelial cells (Fig. 1) (Stutts et al., 1986; Worlitzsch et al., 2002). P. aeruginosa infections are localised within low O2 or hypoxic regions of the mucus and invasion will require P. aeruginosa to move from a region of high to low dissolved oxygen and probably into an anaerobic environment (Fig. 1) (Worlitzsch et al., 2002). CF patients initially become infected with non-mucoid environmental isolates of P. aeruginosa. A switch to a mucoid phenotype occurs in virtually all CF lung infections and is associated with increased inﬂammation, tissue destruction, pulmonary function decline and correlates with poor prognosis for patients (Fig. 2) (Koch and Hoiby, 1993). Mucoidy results from the production of the exopolysaccharide alginate, a linear copolymer of b-D-mannuronic acid and a-L-guluronic acids, which protects the bacterium from phagocytic killing

Cystic Fibrosis Lung Normal Lung

Mucus pO2

Mucus Airway Epithelia

pO2

pO2 P. aeruginosa

Airway Epithelia

Airway Epithelia

Figure 1 Hypoxic mucus hypothesis to explain the events during P. aeruginosa colonisation of the airways of the cystic ﬁbrosis lung. In the normal lung, a usual rate of epithelial O2 consumption produces no O2 gradients within the thin mucus layer. The low viscosity of the mucus combined with cillial beating leads to efﬁcient mucocilliary clearance, denoted by the arrow. In the cystic ﬁbrosis lung, dehydrated viscous mucus sticks to the epithelial layer and mucus transport is reduced, as shown by the bidirectional arrow. Continuous mucus secretion leads to increased thickness of the mucus layer and this combined with increased O2 consumption of the CF epithelial layer generates steep hypoxic gradients in the mucus. P. aeruginosa is not removed by mucus clearance and it moves through the mucus layer from a high O2 to low O2 and then potentially to a hypoxic environment. (Adapted from Worlitzsch et al., 2002.)

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Initial infection

S. aureus & H. influenza

Median age 1 year

Median age 13 years

Initial P. aeruginosa infection NON -MUCOID

Switch to MUCOID P. aeruginosa

By age 30

Respiratory failure & death

Figure 2 The progression of bacterial infections in the cystic ﬁbrois lung Chronic P. aeruginosa infection follows initial infection with a range of bacteria including Haemophilus influenzae and Staphylococcus aureus. Initial P. aeruginosa infection is with non-mucoid environmental isolates but chronic P. aeruginosa infection is associated with switch to alginate overproducing, mucoid phenotype.

mechanisms and prevents phagocytosis of P. aeruginosa by neutrophils and macrophages (Simpson et al., 1989; Pedersen et al., 1992). Alginate also affects leucocyte functions such as the oxidative burst and it has an immunomodulatory role (Pedersen, 1992; Pedersen et al., 1992). High oxygen levels select for a mucoid phenotype and the oxygen-dependent regulation of the alginate biosynthetic genes have been demonstrated (Bayer et al., 1990; Leitao and Sa-Correia, 1993; Leitao and Sa-Correia, 1997). Furthermore, mucoid strains are less sensitive to oxygen. These data suggest that alginate helps protect P. aeruginosa against oxidative damage, a conclusion supported by the ﬁnding that P. aeruginosa converts into a mucoid phenotype when cultures are treated with levels of H2O2 similar to those produced in vivo by polymorphonuclear leucocytes (Mathee et al., 1999). Oxygen is also critical in P. aeruginosa bioﬁlm physiology and recent data indicate that oxygen-limitation contributes to antibiotic tolerance of P. aeruginosa in bioﬁlms (Borriello et al., 2004). Therefore, there is compelling evidence to indicate that oxygen plays an important role in the biology of P. aeruginosa during infection. The colonisation of the single but important pathological niche of the dehydrated mucus of the CF lung will require P. aeruginosa to undergo signiﬁcant metabolic shifts. One can predict that it will have to show metabolic adaptability to modify its energy metabolism as it moves from a high O2 environment, which might favour a respiratory chain terminated by an energetically efﬁcient terminal oxidase, to a region of low O2, favouring the use of a high-afﬁnity terminal oxidase perhaps coupled with aerobic denitriﬁcation if nitrate is available. As anaerobic conditions are encountered, a shift to fermentation or anaerobic denitriﬁcation will take place. Recently, there has been much interest in the anaerobic respiratory capability of P. aeruginosa, in particular, the role anaerobic respiration may play in the sticky dehydrated mucus layer that P. aeruginosa inhabits in the CF lung (Hassett et al., 2002; Worlitzsch et al., 2002). We argue that an

OXYGEN, CYANIDE AND ENERGY GENERATION

7

understanding of the diverse modes of energy metabolism available to P. aeruginosa and their regulation is important to understanding both the fundamental physiology of the bacterium and the factors signiﬁcant in its pathogenicity. Therefore, the aim of this review will be to appraise the current state of knowledge of the energy-generating pathways of P. aeruginosa. We will then consider other aspects of the biology of this bacterium that are linked to energy metabolism or affected by oxygen availability. These will include; cyanide synthesis, which is oxygen-regulated and can affect the operation of aerobic respiratory pathways, and mucoidy and alginate production, which is regulated by oxygen and energy availability, as well as having a role in the protection of P. aeruginosa against reactive oxygen species.

3. MEANS OF ENERGY GENERATION IN P. AERUGINOSA It is in their methods of energy generation that bacteria demonstrate their exceptional metabolic versatility and diversity. Pseudomonads use a remarkably eclectic range of carbon and energy sources (Stanier et al., 1966). Irrespective of the energy source used, energy-generating reactions must accomplish the same metabolic functions, that is, to produce precursor metabolites, reducing power and the energy required by the cell in the form of ATP and Dp (proton and/or sodium electrochemical gradient), in the precise proportions required for biosynthesis and polymerisation (Neidhardt et al., 1990). As different carbon sources will vary in their ability to yield these metabolic products, the cell modiﬁes the proportions produced by changing the relative ﬂow through its various catabolic pathways. P. aeruginosa is capable of both aerobic and anaerobic respiration as well as fermentation of arginine and pyruvate, although it preferentially obtains its energy from aerobic respiration (Van der Wauven et al., 1984; Davies et al., 1989; Zannoni, 1989; Eschbach et al., 2004). We will consider the pathways that P. aeruginosa uses to obtain energy aerobically and anaerobically before looking at its limited fermentative capacity, which is used in the absence of a respiratory electron acceptor.

4. AEROBIC RESPIRATION IN P. AERUGINOSA Bacterial respiratory chains are complex organisations of electron-transfer components, which together can oxidise a broad array of substrates via substrate-speciﬁc dehydrogenases. These initial oxidations of redox couples

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with low negative redox potentials are linked to the four-electron reduction of oxygen to water by a sequence of electron transfer components that are common to all organisms. These include quinones, cytochromes and terminal oxidases and contain as redox centres haems, Fe-S clusters and metals, such as iron and copper. The most striking characteristics of bacterial aerobic respiratory chains are their highly branched nature at both the low potential dehydrogenase end and the high potential terminal oxidase end, and a consequence of this is the apparent redundancy of certain components, particularly the terminal oxidases. A central question is why do bacteria have such an array of respiratory routes? By controlling the expression of particular electron-transfer components, especially terminal oxidases, a bacterium constructs the most appropriate electron transfer chain for the prevailing environmental conditions (Poole and Cook, 2000). The best studied aerobic respiratory chain is that of E. coli, which has two branches terminated by the quinol oxidases cytochrome bo3 and cytochrome bd. A second cytochrome bd oxidase is made but its function is poorly deﬁned (Sturr et al., 1996; Atlung et al., 1997; Lindqvist et al., 2000; Poole and Cook, 2000). Cytochrome bd has a much higher oxygen afﬁnity (Km ¼ 3–8 nM) compared to cytochrome bo3, which has had values measured for the puriﬁed enzyme in the low mM range (Kita et al., 1984), while values of 10–100 fold lower than this were obtained with whole cells using a more reliable methodology (D’Mello et al., 1995). However, a cytochrome bo3-terminated respiratory chain is energetically more efﬁcient as this oxidase can act as a proton pump, while cytochrome bd cannot (Junemann, 1997). Therefore, a simple model explains the function of the branched aerobic respiratory chain of E. coli. Under low oxygen conditions, cytochrome bd levels rise and there is increased electron ﬂux through this pathway, while under high oxygen conditions, particularly at high growth rate, the cytochrome bo3 expression increases and the cells can make use of the increased energetic efﬁciency of electron ﬂux through this branch (Poole and Cook, 2000). However, genome sequencing has indicated that many bacteria have the potential to make far more complex aerobic respiratory chains than E. coli, and so it is difﬁcult to transfer such a straightforward analysis to these bacteria. This is the case for P. aeruginosa whose aerobic respiratory chain is predicted to be a complex highly branched structure, far more complex than the equivalent pathway in E. coli (Fig. 3; Poole and Cook, 2000; Comolli and Donohue, 2002; Cooper et al., 2003). It has at least 17 primary dehydrogenases, ubiquinone and b- and c-type cytochromes, as well as ﬁve terminal oxidases (Fig. 3). We will consider each of these principal components in turn.

OXYGEN, CYANIDE AND ENERGY GENERATION

9

4.1. Respiratory Dehydrogenases of P. aeruginosa A few of the 17 respiratory dehydrogenases of P. aeruginosa have been studied as puriﬁed enzymes or at the whole cell level, but the operation of many can only be inferred from genome sequence analyses and have not been studied experimentally. 4.1.1. NADH Dehydrogenases Three types of NADH dehydrogenases (NADH:quinone oxidoreductase) are recognised in bacteria. The ﬁrst is the proton – translocating NADH dehydrogenase, NDH-1. It is the bacterial counterpart of the mitochondrial Complex I and has been best studied in Paracoccus denitrificans (Yagi et al., 1993, 1998, 2001). It has 13–14 subunits, contains FMN together with two [2Fe-2S] and six to seven [4Fe-4S] clusters. The second enzyme is NDH-2, a simpler enzyme comprising a single polypeptide subunit that has a bound FAD prosthetic group and does not translocate protons (Yagi et al., 2001). The third bacterial NADH:quinone oxidoreductase is the sodium-translocating NADH:quinone oxidoreductase (Na+-NDH or Nqr). Nqr comprises six subunits, FAD and one to two FMN and a single [2Fe-2S] cluster (Pfenninger-Li et al., 1996; Nakayama et al., 1998; Zhou et al., 1999; Hase et al., 2001; Steuber, 2001). The P. aeruginosa genome encodes each of these three NADH quinone oxidoreductases (Fig. 3). The nuoABCDEFGHIJKLMN genes encode a putative type I NADH dehydrogenase. It is predicted that this enzyme will contain two [2Fe-2S] and six to eight [4Fe-4S] clusters and that it would be energy coupling, contributing to the generation of the Dp in P. aeruginosa. Additionally, P. aeruginosa has an ndh gene predicted to encode a type II NADH dehydrogenase, composed of a single FAD-containing subunit. Furthermore, its genome encodes genes annotated nqrABCDEF encoding a predicted energy coupling Na+-translocating NADH-quinone oxidoreductase. However, the biochemistry of these dehydrogenases has not been studied in P. aeruginosa. 4.1.2. Succinate Dehydrogenase (Succinate Quinone Oxidoreductase) These are membrane-bound enzymes that are components of the tricarboxylic acid cycle and catalyse the two-electron oxidation of succinate to fumarate and couple it to the two-electron reduction of a quinone to the corresponding quinol. They can also catalyse the reverse reaction

NADH Dehydrogenase I (nuo)

Cytochrome cbb3 I

10

NADH Dehydrogenase II (ndh)

Cyanide-insensitive oxidase

Succinate dehydrogenase

Cytochrome cbb3 II Cytochrome bo3

L-lactate Dehydrogenase I (lldA) L-lactate Dehydrogenase II (lid?)

Cytochrome caa3

sn-glycerol-3-phospahate dehydrogenase D-amino acid dehydrogenase Proline dehydrogenase

Ubiquinone Pool

Cytochrome bc1

Cyt. c/ azurin Cytochrome c peroxidase

Allohydroxy-D-proline dehydrogenase Glucose dehydrogenase

Nitrite reductase

Gluconate dehydrogenase

Nitrate reductase (Nap)

Electron transfer flavoprotein

Malate Dehydrogenase (MqoA)

Nitrate reductase (Nar)

Formate dehydrogenase Ethanol Dehydrogenase (QEDH)

Cytochrome c551/EDH

?

Nitrous oxide reductase

Figure 3 The proposed respiratory chains of P. aeruginosa. Each of the components and pathways are described in the relevant portions of the text. The scheme is based partly on experimental data and partly on the authors’ analysis of the P. aeruginosa genome sequence. The formate dehydrogenase-nitrate reductase pathway is shown without the participation of quinones, as respiratory chain has been reconstituted in vitro and does not require quinone for its function. However, it has not been ruled out that quinones are required in vivo. Cytochromes c4 and c5 are not shown as it is not clear where they act in the aerobic respiratory chains. The quinoprotein ethanol dehydrogenase (QEDH) donates electrons directly to the cytochrome c551/EDH. However, the electron acceptor from this cytochrome is not known. Azurin is an alternative electron carrier to cytochrome c (Cyt. c) in anaerobic but not aerobic respiratory chains.

HUW D. WILLIAMS ET AL.

Nitric oxide reductase

Malate Dehydrogenase (MqoB)

OXYGEN, CYANIDE AND ENERGY GENERATION

11

acting as a quinol fumarate oxidoreductase (QFR). It is not possible to predict from the protein sequence if an enzyme acts in vivo as a succinate quinone oxidoreductase (SQR) or a QFR. Five types of SQR are recognised (Lancaster, 2002; Lancaster and Simon, 2002). The best studied enzymes are those of E. coli and Pa. denitrificans and comprise four subunits; SdhA, B, C and D. SdhA is a ﬂavoprotein with a covalently bound FAD, while SdhB is an Fe-S protein with three different clusters, a [2Fe-2S], a [4Fe-4S] and a [3Fe-4S] (Cecchini et al., 2002; Hederstedt, 2002). Together these subunits constitute a linear electron transfer pathway from FAD to the membrane anchored subunits SdhC and SdhD which contain a single haem B prosthetic group, although some enzymes such as in the Gram-positive bacteria Paenibacillus macerans and B. subtilus have two haem groups (Hederstedt, 2002). P. aeruginosa has an sdhCDAB operon. As no fumarate reductase activity has been reported for P. aeruginosa we assume at present that this operon encodes a succinate dehydrogenase, and it is predicted to be homologous to the enzymes from E. coli and Pa. denitrificans discussed above. However, a fumarate reductase is postulated as operating in the recently described pyruvate fermentation pathway of P. aeruginosa (see Fig. 8; Eschbach et al., 2004), and so this issue needs an experimental resolution.

4.1.3. Lactate Dehydrogenases Lactate is a signiﬁcant product of the bacterial fermentation of glucose and other carbohydrates. P. aeruginosa has a fermentative lactate dehydrogenase, and this will be considered in Section 6. Bacteria can also possess respiratory chain-linked D- or L-lactate dehydrogenases. The former is typically a two-subunit enzyme with FAD as a prosthetic group, while the latter is a single-subunit enzyme containing FMN. These enzymes allow bacteria to use lactate as a carbon and energy source and lactate oxidation to be coupled to energy transduction (Poole and Ingledew, 1987). Kemp described D- and L-lactate dehydrogenase activities in particulate fractions from P. aeruginosa cells grown aerobically with DL-lactate (Kemp, 1972). The linkage of these activities to the respiratory chain was suggested by their ability to reduce b- and c-type cytochromes. Both activities were inducible by lactate under aerobic conditions. The P. aeruginosa genome has the lldA and lldD genes both of which are predicted to encode single subunit, FMN-containing, membrane-bound L-lactate dehydrogenases with strong similarities to the well studied E. coli enzymes (Poole and Ingledew, 1987). Adjacent to lldD is lldP, a putative lactate permease, and PA4472 which is a

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HUW D. WILLIAMS ET AL.

putative D-lactate dehydrogenase, containing sequence signatures for the binding of a 4Fe-4S clusters and which shows440% similarity to cytochrome type D-lactate dehydrogenases from a number of bacteria. Whether or not these enzymes catalyse the activities described by Kemp has not been tested experimentally (Kemp, 1972). 4.1.4. sn-Glycerol-3-Phosphate Dehydrogenase In P. aeruginosa glycerol is primarily metabolised through the Entner– Doudoroff pathway (EDP) (for review see Lessie and Phibbs, 1984). While there is some confusion over the precise way glycerol is transported (Schweizer and Po, 1996), intracellular glycerol is phosphorylated to sn-glycerol-3-phosphate (G3P) by glycerol kinase (glpK) and subsequently oxidised to dihydroxyacetone phosphate by the membrane-associated enzyme G3P dehydrogenase. This analysis is based on the observed speciﬁcity of the enzyme induction and on the study of mutants defective in speciﬁc glp genes (Wilderman et al., 2004). Mutants defective in G3P dehydrogenase activity fail to grow on either glycerol or G3P (Wilderman et al., 2004). Dihydroxyacetone phosphate is preferentially metabolised to pyruvate by the EDP (Lessie and Phibbs, 1984). The further metabolism of pyruvate by the TCA cycle requires an inducible pyruvate carboxylase (Phibbs et al., 1974), which is probably encoded by ORF PA1400. The annotated P. aeruginosa glpD gene encodes the sn-glycerol-3-phosphate dehydrogenase and the P. aeruginosa glpD is able to complement an E. coli glpD mutant. It encodes a protein of 56 kDa, which is 56% identical to the E. coli GlpD and is also predicted to have an FAD-binding site (Entsch and Ballou, 1989). The E. coli enzyme is involved in a respiratory chain with oxygen as the terminal electron acceptor, it is thought via direct electron transfer to quinones (Poole and Ingledew, 1987). 4.1.5. Amino Acid Dehydrogenases (i) D-Amino acid dehydrogenase. In E. coli, this membrane-bound enzyme is involved in the degradation of D-amino acids, whose oxidation yields imino acids, which are hydrolysed to the corresponding keto acid and NH3. It is a primary dehydrogenase with electrons arising from the oxidation reaction being transferred directly to the respiratory chain. The protein is a heterodimer and contains FAD as well as non-haem iron. It shows a broad speciﬁcity to D-amino acids, of which D-alanine is the best substrate (Stieritz and Holder, 1975; Reitzer and Magasanik, 1987; Elias et al., 2004). The

OXYGEN, CYANIDE AND ENERGY GENERATION

13

dadA gene has been established as a structural gene for D-amino acid dehydrogenase, but only encodes the smaller of the enzyme’s two subunits. The identity of the gene encoding the second subunit remains unknown (Lobocka et al., 1994). However, Marshall and Sokatch (1968) described the presence of a D-amino acid dehydrogenase in P. aeruginosa after growth in the presence of D- or L-forms of either alanine or valine. The enzyme showed high activity with a wide range of amino acids, particularly alanine, valine, norvaline, histidine, methionine, phenylalanine and serine. The enzyme was membrane-bound and its activity was associated with cytochrome reduction. A second amino acid dehydrogenase activity in P. aeruginosa has been described, which also has high activity with D-alanine but differed in its undetectable activity against other amino acids (Pioli et al., 1976). P. aeruginosa can grow well with D- and L-isomers of a-alanine and also with b-alanine as sole sources of carbon and energy. It was shown that the utilisation of D-alanine involved its direct oxidation by an inducible, membrane-associated, cytochrome-linked dehydrogenase that was also required for oxidation of other isomers following their conversion into D-alanine by cytoplasmic enzymes. P. aeruginosa has a dadA gene, which encodes a putative protein that is 75% similar to dadA gene product of E. coli, but there have been no biochemical or genetic studies on this system. Therefore, whether this gene encodes the dehydrogenase responsible for the amino acid oxidising activities described in P. aeruginosa is unclear, although it is clear that P. aeruginosa does have at least one respiratory chain-linked D-amino acid dehydrogenase. (ii) Proline dehydrogenase. Pseudomonads, including P. aeruginosa, can use proline as a sole source of carbon and energy for growth (Stanier et al., 1966; Meile et al., 1982). Following its transport into the cell, proline is catabolised to glutamate in a two-step process catalysed by the multifunctional PutA protein. The PutA protein is a ﬂavoenzyme that in enteric bacteria and in Pseudomonas putida combines catalytic, membrane-binding and DNA-binding activities into a single polypeptide. The E. coli PutA puriﬁes as a dimer of 293 kDa with one covalently bound FAD per monomer (Brown and Wood, 1992). PutA catalyses the two-step oxidation of proline to glutamate using discrete proline dehydrogenase (ProDH) and D1-pyrroline-5-carboxylate dehydrogenase (P5CDH) domains (Menzel and Roth, 1981; Abrahamson et al., 1983; Brown and Wood, 1992). The ProDH domain catalyses the transfer of two electrons from proline to FAD in the ﬁrst step to generate D1-pyrroline-5-carboxylate (P5C) and reduced FAD. In the oxidative half reaction, PutA associates with the membrane where the ProDH domain catalyses the transfer of electrons to the respiratory chain, regenerating oxidised FAD (Abrahamson et al., 1983; Graham

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HUW D. WILLIAMS ET AL.

et al., 1984; Wood, 1987). After hydrolysis of P5C, the P5CDH active site catalyses the NAD-dependent oxidation of g-glutamic acid semialdehyde to glutamate. The P. aeruginosa PutA protein has a similar catalytic activity, but unlike the PutA proteins of enteric bacteria and P. putida it does not act as a repressor of the putAP operon (putP encodes a proline permease) in the absence of proline (Meile et al., 1982; Meile and Leisinger, 1982; MuroPastor and Maloy, 1995; Vilchez et al., 2000). In P. aeruginosa the putAP operon is regulated by a PruR, an AraC/XylS family protein (Nakada et al., 2002). (iii) Allo-hydroxy-D-proline dehydrogenase. P. aeruginosa can grow on minimal medium containing either hydroxyl-L-proline or allohydroxyD-proline as sole sources of carbon and energy and such conditions induced the synthesis of an allohydroxy-D-proline dehydrogenase that is membrane associated and linked to oxygen through the respiratory chain (Bater et al., 1977). The genes encoding this enzyme are not clear from inspection of the genome.

4.1.6. PQQ-Containing Dehydrogenases Pyrroloquinoline quinone (2,7,9,-tricarboxypyrroloquinoline quinone, PQQ) is one of several quinone cofactors used in a class of dehydrogenases, known as quinoproteins, which are distinct from the ﬂavin- and nictotinamidedependent oxidoreductases (Duine, 1999). When grown on glucose or various alcohols P. aeruginosa, like a number of Gram-negative bacteria, synthesises PQQ-dependent dehydrogenases, which are located in the periplasm. (i) The ethanol oxidation system. P. aeruginosa is able to grow with ethanol as its sole carbon and energy source. When growing on ethanol, P. aeruginosa uses a PQQ-dependent ethanol oxidation system (Fig. 4) (Gorisch, 2003). The ethanol-oxidising system is encoded by the exaABC operon and includes a quinoprotein ethanol dehydrogenase (QEDH, exaA), a speciﬁc cytochrome c550 (also known as cytochrome cEDH, exaB) that is an essential component of the respiratory chain and accepts electrons from QEDH (Figs. 3 and 4) (Rupp and Gorisch, 1988; Reichmann and Gorisch, 1993), and an NAD-dependent acetaldehyde dehydrogenase (exaC) (Fig. 4). Ethanol oxidation is inhibited by antimycin A and myxothiazol indicating the involvement of the cytochrome bc1 complex in electron transfer to oxygen. Downstream of the exaABC genes are the pqqABCDE genes, which code for proteins essential for the biosynthesis of the cofactor PQQ. Also essential for growth on ethanol is an acetyl CoA synthase (acsA) and another PQQ-containing dehydrogenase malate:quinone oxidoreductase

PEP-C PEP pyruvate CO2 ATP PC ADP +Pi QH2

Acetaldehyde DH

ACS

acetate

acetyl-CoA

AMP+PPi ATP+CoA

QEDH, Cyt. c550

acetaldehyde

NADH+H+ NAD+

ETHANOL +

-

2H + 2e

PEP-CK oxaloacetate

MQO Q MDH ME malate

NADH+H+ citrate

NAD+ glyoxylate

CO2 NAD(P)H NAD(P)

isocitrate

fumarate

succinate

OXYGEN, CYANIDE AND ENERGY GENERATION

glucose

alpha-ketoglutarate

15

Figure 4 Ethanol oxidation and intermediary metabolism in P. aeruginosa. Acetaldehyde DH, acetaldehyde dehydrogenase; ACS, acetyl-CoA synthetase; MDH, malate dehydrogenase; ME, malic enzyme; MQO, malate:quinone oxidoreductase; PC, pyruvate carboxylase; PEP, phosphoenolpyruvate; PEP-C, PEP carboxylase (PEP+CO2+H2O oxaloacetate+Pi); PEP-CK, PEP carboxykinase (oxaloacetate+GTP PEP+GDP+CO2); QEDH, quinoprotein ethanol dehydrogenase. (Adapted from Kretzschmar et al., 2002.)

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HUW D. WILLIAMS ET AL.

(see below). Analysis of mutants defective in this cytochrome c550 (exaB mutants) shows that it is essential for ethanol oxidation (Schobert and Gorisch, 1999). The role of this cytochrome is thought to be analogous to that of c-type cytochromes in methylotrophic bacteria, which oxidise methanol via a PQQ-containing methanol dehydrogenase, with the primary electron acceptor being a speciﬁc, acidic cytochrome c, known as cytochrome cL (Anthony, 1986, 1990, 1992; Goodwin and Anthony, 1998; Nunn and Anthony, 1988), which is subsequently oxidised by a second cytochrome c, cytochrome cH. QEDH has been co-puriﬁed from P. aeruginosa with two soluble c-type cytochromes (Schrover et al., 1993). One of these (referred to as cytochrome cEDH in this paper) in its oxidised-form was an excellent electron acceptor from the QEDH, and the N-terminal sequence of this cytochrome cEDH indicates that it is the protein predicted to be encoded by exaB. While this cytochrome c550/EDH mediates electron transfer between QEDH and a cytochrome c oxidase, it is unclear whether quinones play a role in this electron transfer route (Gorisch, 2003). (ii) Malate dehydrogenase. P. aeruginosa has two genes encoding putative malate:quinone oxidoreductases, mqoA and mqoB. Malate:quinone oxidoreductase (MQO) was ﬁrst described in P. aeruginosa as a membrane-associated malate dehydrogenase (MDH) activity (Mizuno and Kageyama, 1978). It is a FAD-dependent enzyme activity that catalyses the oxidation of malate to oxaloacetate with evidence, in some bacteria, for the electrons being donated to the quinones of the respiratory chain (Fig. 3) (Kather et al., 2000). This MQO is encoded by the mqoB gene and is involved in the operation of the TCA and glyoxylate cycles (Fig. 4). mqoB mutants are unable to grow on ethanol or acetate, but show no defects when glucose, lactate, succinate or malate are the carbon and energy sources. Under these conditions, an apparent bypass route is used with malic enzymes using malate to generate pyruvate, which is then carboxylated to oxaloacetate by pyruvate kinase (Kretzschmar et al., 2002). The role of the second putative MQO encoded by mqoA, has not been determined. (iii) Glucose and gluconate dehydrogenases. P. aeruginosa readily catabolises glucose, not via the Embden–Meyerhof pathway because it does not possess 6-phosphofructokinase, but through the EDP (Horio et al., 1958). Depending on the physiological conditions, glucose is converted into 6-phosphogluconate for entry into the EDP by one of the two routes (Fig. 5) (Hunt and Phibbs, 1983). The phosphorylative route involves uptake of glucose by an inducible transport system and once inside the organism it is phosphorylated by glucokinase and then converted into 6-phosphogluconate by glucose-6-phosphate dehydrogenase, the product of the zwf gene (Horio et al., 1958; Matsuura et al., 1982; Gavira et al., 2002).

Glucose

Gluconate

2-Ketogluconate

glucose dehydrogenase

gluconate dehydrogenase

Cytoplasmic Membrane Glucose

2-Ketogluconate

Gluconate

ATP

ATP

ATP

glucokinase ADP Glucose 6-P

Glucose-6-P dehydrogenase NAD(P)

2-ketogluconokinase

gluconokinase ADP

ADP reductase

6-Phosphogluconate

2-Ketogluconate 6-P

NAD(P)H

NADP H2O

NADPH

dehydratase

2-Keto-3-deoxy6-phosphogluconate

Cytoplasm

aldolase

Pyruvate

OXYGEN, CYANIDE AND ENERGY GENERATION

Periplasm

Glyceraldehyde 3-P

Figure 5 Multiple peripheral pathways for glucose oxidation in P. aeruginosa. The phosphorylative route is shown by the dashed lines and the oxidative routes by the solid lines. (Adapted from Hunt and Phibbs, 1981.)

17

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HUW D. WILLIAMS ET AL.

The direct oxidative route involves oxidation of glucose to gluconate and gluconate to 2-ketogluconate in the periplasm by membrane-bound glucose and gluconate dehydrogenases, respectively (Gavira et al., 2002). Both gluconate and 2-ketogluconate are transported into the cytosol where they are converted to 6-phosphogluconate. Gluconate is phosphorylated by the cytoplasmic gluconokinase to 6-phosphogluconate. 2-Ketogluconate is also converted into 6-phosphogluconate by the sequential action of a 2-keto-gluconokinase and a reductase (Fig. 5). Both oxidative routes have been shown to be physiologically signiﬁcant through the isolation of mutants blocked in either gluconate or 2-Ketogluconate utilisation (Matsuura et al., 1982). The 6-phosphogluconate is converted into glyceraldehyde3-phosphate and pyruvate through the sequential action of 6-phosphogluconate dehydratase (Edd) and 2-keto-3-deoxy-6-phosphogluconate aldolase (Eda) (Horio et al., 1958). In the context of this review, we are concerned particularly with the glucose and gluconate dehydrogenases. Glucose dehydrogenase is a membrane-bound quinoprotein, which catalyses the oxidation of D-glucose to D-gluconate in the periplasm and transfers the electrons directly to ubiquinone in the respiratory chain (Fig. 3) (Tiwari and Campbell, 1969). It occurs as an apoenzyme in E. coli as this bacterium cannot make PQQ (Matsushita et al., 1997). This is not the case in P. aeruginosa where a functional holoenzyme can be made as this bacterium can synthesise the PQQ cofactor. The E. coli GDH has a tightly bound ubiquinone, which accepts a single electron from PQQH2 to form a semiquinone radical which acts as a single electron mediator of the intramolecular electron transfer in GDH (Elias et al., 2001, 2004). Oxygen availability regulates glucose metabolism in P. aeruginosa. Neither the oxidative nor the phosphorylative pathways are uniquely required for aerobic glucose oxidation (Hunt and Phibbs, 1981). No GDH activity was detectable in P. aeruginosa grown anaerobically with glucose and nitrate, which demonstrated an obligatory role for the phosphorylation route during these growth conditions (Hunt and Phibbs, 1981). However, apoGDH was detected in anaerobically grown P. aeruginosa and by addition of PQQ it could be reconstituted to an active enzyme capable of generating a Dc. Therefore, there would seem to be an uncoupling of the synthesis of the GDH apoprotein from its cofactor synthesis under anaerobic conditions. P. aeruginosa has a ﬁve-gene cluster pqqABCDE and a sixth gene, pqqF, at a separate locus that are homologous to the genes required for PQQ synthesis from a variety of bacteria including P. fluorescens (Toyama et al., 1997; Kim et al., 2003; Magnusson et al., 2004). In the oxidative pathway, the next stage is the oxidation of gluconate to 2-keto-D-gluconate by a membranebound gluconate dehydrogenase (GADH) (Matsushita et al., 1979). It has

OXYGEN, CYANIDE AND ENERGY GENERATION

19

been puriﬁed from P. aeruginosa as a monomer of 138 kDa and contains a cytochrome c1 (c554), which might be a dihaem cytochrome and a covalently bound ﬂavin but not ubiquinone. The P. aeruginosa genome has three adjacent genes, PA2264–2266, which are homologous to the three subunits (dehydrogenase 65 kDa, cytochrome c 45 kDa, and small 20 kDa subunit) of the GADH of Erwinia cypripedii (Yum et al., 1997), and it would seem most probable that these encode the P. aeruginosa GADH, although this has not been experimentally established. 4.1.7. Electron Transfer Flavoprotein The heterodimeric electron transfer ﬂavoproteins (ETF) function as intermediate electron carriers between primary dehydrogenases and terminal respiratory systems. They have been functionally classiﬁed by Weidenhaupt et al. (1996) into two groups, housekeeping ETFs and ETFs that are synthesised under special nutritional conditions. The housekeeping ETFs function in the oxidation of fatty acids and some amino acids and include those enzymes from mammals and Pa. denitrificans (Finocchiaro et al., 1988, 1993; Watmough et al. 1992; Bedzyk et al., 1993; Goodman et al., 1994). The second group functions in the oxidation of secondary metabolites such as trimethylamine, carnitine as well as in nitrogen ﬁxation (Chen and Swenson, 1994; Weidenhaupt et al., 1996). In the latter case, the ETF protein is proposed to link substrate oxidation to the reduction of a cytochrome cbb3type terminal oxidase to provide energy for nitrogen ﬁxation. P. aeruginosa has a putative ETF encoded by the annotated etfA and etfB genes, but there have been no studies on these genes or their putative gene products.

4.2. Transhydrogenase The nictotinamide nucleotide transhydrogenase is an enzyme that couples the transfer of reducing equivalents between NAD(H) and NADP(H) to the translocation of protons across a membrane. It is found in the inner mitochondrial membrane of eukaryotes and in the cytoplasmic membrane of some bacteria. NAD(H) is normally involved in catabolic reactions and NADP(H) in anabolic or biosynthetic reactions. Transhydrogenase is positioned at the boundary between the NAD(H) and NADP(H) pools and under physiological conditions in most bacteria it acts to oxidise NADH and reduce NADP at the expense of the Dp (Jackson, 2003; Jackson et al., 1993). P. aeruginosa contains pntAB genes that are proposed to encode the alpha- and beta subunits, respectively of a transhydrogenase. However, the pntA in P. aeruginosa

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HUW D. WILLIAMS ET AL.

strain PAO1 has a frameshift about two-thirds of the way along its length so whether or not a functional enzyme is made is unknown.

4.3. Quinones Bacterial respiratory quinones are lipid soluble H-carriers that act to transfer electrons from primary dehydrogenases either to the cytochrome bc1 complex or directly to quinol oxidases of the aerobic respiratory chain or reductases of anaerobic respiration (Fig. 3). Bacteria contain two main types of quinone, the benzoquinone known as ubiquinone (or Coenzyme Q) and napthoquinones, the most common of which is menaquinone (MK or vitamin K2). The isoprenoid side chains of these molecules vary in length between species (Soballe and Poole, 1999). Some facultatively anaerobic bacteria such as E. coli can make both types, but reserve UQ primarily for aerobic respiration (although E. coli also uses it for nitrate respiration), and MK for anaerobic respiration with lower potential electron acceptors for which the MK/MKH2 couple has a more suitable midpoint potential (Em,7,–74 mV) than UQ/UQH2 (Em,7,+113 mV). Early studies identiﬁed ubiquinone-9 (UQ-9) as the major quinone extractable from cytoplasmic membranes of aerobically grown P. aeruginosa (Matsushita et al., 1979). Ubiquinone depletion and reconstitution studies showed that UQ-9 was essential for respiration. We are not aware of any reports of quinone analyses of anaerobically respiring P. aeruginosa. Eight genes are recognised as essential for the ubiquinone biosynthesis pathway in E. coli (Soballe and Poole, 1999), and there are clear homologues of six of these (ubiA, ubiB, ubiC, ubiE, ubiG and ubiH) annotated on the P. aeruginosa chromosome. Additionally, the ORFs PA0254 and PA5237 show signiﬁcant similarity to E. coli ubiD, while PA0655 encodes an ORF that shows similarity to the COQ7 protein from yeast, which catalyses the same reaction as UbiF. As expected from the biochemical analyses there are no genes for the biosynthesis of MK annotated on the P. aeruginosa genome.

4.4. Cytochrome bc1 Complex The ubiquinol: cytochrome c oxidoreductase, or cytochrome bc1 complex, comprises intrinsic membrane proteins that catalyse the oxidation of ubiquinol (UQH2) and the reduction of cytochrome c (Fig. 3) (Trumpower, 1990, 1991; Brandt and Trumpower, 1994; Crofts and Berry, 1998; Crofts, 2004). It is a key component of both respiratory and photosynthetic electron

OXYGEN, CYANIDE AND ENERGY GENERATION

21

transfer chains and contributes to the formation of the electrochemical gradient necessary for ATP synthesis. Numerous bacteria harbour a cytochrome bc1 complex comprising three redox-active subunits: a cytochrome b subunit with two B-type haems (bL and bH), a cytochrome c1 subunit with a C-type haem and a subunit with a high potential (2Fe-2S) cluster known as the Rieske cluster. These polypeptides form two catalytic domains, the Qo and Qi sites, located on either side of the cytoplasmic membrane (outside and inside, respectively). The transfer of electrons through the complex leads to the generation of a Dp via the operation of a protonmotive Q-cycle (Trumpower, 1990, 1991; Brandt and Trumpower, 1994; Crofts and Berry, 1998; Crofts, 2004). For every two electrons that pass through the complex, four protons are translocated across the cytoplasmic membrane to the cytoplasm, with the overall reaction catalysed by the cytochrome bc1 being: þ QH2 þ 2Hþ i þ 2cyt:cox ! Q þ 4H0 þ 2cyt:cred

(cyt.cox and cyt.cred are oxidised and reduced cytochrome c, respectively). In bacteria, the polypeptides of the cytochrome bc1 complex are encoded by the petABC genes (or fbcFBC genes). While these genes are not annotated as such on the P. aeruginosa genome there is a three-gene cluster, PA4431, PA4430 and PA4429, which encode ORFs with clear similarity to PetA (Rieske Fe-S protein), PetB (cytochrome b) and PetC (cytochrome c1), respectively. Little experimental work has been carried out on the cytochrome bc1 of P. aeruginosa although inhibitor studies have indicated its operation under aerobic and anaerobic conditions (Matsushita et al., 1983; Ray and Williams, 1996; Hasegawa et al., 2003).

4.5. Cytochromes 4.5.1. c-Type Cytochromes Cytochromes c are distinguished from other cytochromes by the covalent attachment of the haem prosthetic group to the cytochrome polypeptide chain. This is achieved by thioether links between the two protoporphyrin IX vinyl groups and the thiol group of two cysteine residues present in the conserved motif CXXCH. This process requires posttranslational modiﬁcation of the apocytochrome c polypeptide. P. aeruginosa has a ccmABCDEFGHcycH gene cluster, which has been shown in other bacteria to encode proteins with speciﬁc roles in cytochrome c biogenesis

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(Thony-Meyer, 1997; Kranz et al., 1998; Page et al., 1998; O’Brian and Thony-Meyer, 2002). The details of cytochrome c biogenesis will not be discussed further here. A variety of early spectral studies on the cytochrome composition of P. aeruginosa were not particularly informative. They certainly suggested that membranes contained b- and c-type cytochromes (Matsushita et al., 1980, 1983), but the electron transfer step between the cytochrome bc1 complex and the terminal cytochrome c oxidases, that is the speciﬁc cytochrome c involved, was not deﬁned. Anaerobic cultures have c-type cytochromes with a-bands in reduced minus oxidised spectra at 550, 551, 555 and 557 nm (Lemberg and Barrett, 1973; Zannoni, 1989). Cytochromes c produced under anaerobic conditions will be discussed in more detail later. In aerobically grown cultures, cytochromes c with a-bands at 551 and 554 nm were resolved (Gel’man et al., 1975). A cytochrome c554 has been puriﬁed as a component of gluconate dehydrogenase (see above, Matsushita et al., 1979) and this seem likely to be the cytochrome c encoded by PA2266, suggesting that a cytochrome c551 is an electron donor to the cytochrome c oxidase terminated pathways and acts as an acceptor from cytochrome bc1. However, the identity of the gene encoding this cytochrome is unclear. In addition to those c-type cytochromes with clear predicated roles in anaerobic respiration or terminal oxidases (see below), 12 others can be identiﬁed as encoded on the P. aeruginosa genome. ORF PA1600 encodes a cytochrome c and is part of a three-gene cluster encoding a putative aldehyde dehydrogenase. exaB (PA1983) is the cytochrome c component of the ethanol dehydrogenase discussed above and pvdD (PA2257) is a cytochrome c with a role in pyoverdine biosynthesis (Merriman et al., 1995; Ackerley et al,. 2003). The remaining nine have no known roles and presumably include the cytochrome c responsible for electron shuttling between the cytochrome bc1 and the cytochrome c oxidases. 4.5.2. Cytochrome c4 and c5 The cytochrome c5 is a monohaem cytochrome for which there is evidence of a role in the aerobic respiratory chain in certain bacteria (Carter et al., 1985; Rey and Maier, 1997). The cytochrome c4 proteins are dihaem cytochromes c with molecular masses of approximately 20 kDa that are found in a variety of bacteria (Sawyer et al., 1981; Pettigrew and Brown, 1988; Pettigrew and Moore, 1990). Two haem C groups are attached to the polypeptide chain by covalent bonds to two cysteines, and cytochrome c4 has a histidine and a methionine as axial ligands to the haem iron, forming a hexacoordinated low spin complex in the reduced state (Pettigrew and Moore, 1990).

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Cytochromes c4 have been characterised and sequenced from Azotobacter vinelandii, Pseudomonas stutzeri, Thiobacillus ferrooxidans, as well as P. aeruginosa (Sawyer et al., 1981;Ambler et al., 1984; Hunter et al., 1989; Pettigrew and Moore, 1990; Christensen, 1994; Conrad et al., 1995; Kadziola and Larsen, 1997; Rey and Maier, 1997; Abergel et al., 2000). The cytochromes c4 and the associated cytochromes c5 are reported to be predominantly membrane-bound proteins that face the periplasmic space, and it has been proposed that they play a role in aerobic respiration by acting in electron transfer between primary dehydrogenases and terminal oxidases or reductases. However, despite crystal structures being available for a number of these cytochromes, including the P. aeruginosa c4 (albeit at low resolution (Sawyer et al., 1981)), their precise role has not been established convincingly in any bacterium. In the iron-oxidising bacterium T. ferrooxidans, cytochrome c4 could be reduced by Fe2+ in the presence of the blue copper protein rusticyanin (which is the initial acceptor of electrons from Fe2+ during respiratory iron oxidation (Appia-Ayme et al., 1998)). This indicated that rusticyanin catalyses the transfer of electrons from Fe2+ to cytochrome c4 on their way to oxygen via a terminal oxidase. Rusticyanin has been found to form a tight electron transfer complex with cytochrome c4. However, their organisation into an electron transport chain bridging the span from Fe2+ to the membrane-bound, energy-conserving electron transport pathways is still a matter of debate (Giudici-Orticoni et al., 1999). The physiological role of cytochromes c4 and c5 in A. vinelandii is also unclear (see Hunter et al., 1989; Ng et al., 1995; Rey and Maier, 1997). On the basis of data from physiological and spectral analysis of c4 and c5 single mutants and a c4c5 double mutant, it has been proposed that c4 and c5 operate in parallel electron transfer pathways to a terminal oxidase that was described as a cytochrome o. However, this oxidase was poorly deﬁned and recent data indicate that a cytochrome cbb3-type oxidase may oxidise c4 and c5 in A. vinelandii (Bertsova and Bogachev, 2002). The proposed branch point in the pathway is ubiquinone and no cytochrome bc1 complex is incorporated into the model. This model represents a novel electron transfer route but at present it is difﬁcult to reconcile with the organisation of experimentally well-established respiratory pathways and the presence of a cytochrome bc1 complex, which the draft of the A. vinelandii genome sequence tells us is present in these bacteria. Therefore, while a role for cytochrome c4 and c5 in aerobic respiration is likely, its exact nature has not been deduced. In P. aeruginosa the c4 and c5 genes are encoded adjacent to one another on the genome (PA5490 and PA5491), although PA5300 encodes a second c5 gene. The presence of similar amounts of c4 in P. aeruginosa in aerobically grown and denitrifying cells suggests it may also have a role in anaerobic

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respiratory pathways in P. aeruginosa (Pettigrew and Brown, 1988). However, at present, nothing more is known about the roles of these cytochromes in P. aeruginosa.

4.6. Terminal Oxidases The basic role that all terminal oxidases must complete is the efﬁcient fourelectron reduction of oxygen to water, avoiding the production of partial reduction products. Single electron reduction of oxygen yields the superoxide radical anion (Od 2 ), while transfer of two electrons to oxygen or a single electron to superoxide forms peroxide (O2 2 ). Transfer of another electron to peroxide forms the highly reactive hydroxyl radical (OHd), and the fourth forms water. Formation of these reactive intermediates is an inevitable consequence of using oxygen as a terminal electron acceptor, but terminal oxidases operate to try and minimise the production of reactive oxygen species. Bacteria have well described mechanisms for dealing with endogenous and exogenous sources of these reactive oxygen intermediates (van der Oost et al., 1994). The standard redox potential (E00 ) of the O2/H2O couple is +815 mV with the prospect of a much higher yield of free energy than with an alternative electron acceptor such as nitrate/nitrite (E00 ¼ +430 mV). The kinetic inertness of oxygen requires its activation by a metal centre which in an aerobic respiratory chain involves the Fe atoms in two haem prosthetic groups or a Fe-copper centre in the haem-copper oxidases (Garcia-Horsman et al., 1994; van der Oost et al., 1994; Junemann, 1997). Bacterial terminal oxidases can be classiﬁed in a number of different ways. One way is on the basis of whether their physiological electron donor is a quinol or a cytochrome c, (Poole and Cook, 2000). However, they can also be distinguished by their subunit composition, cytochrome content and oxygen afﬁnity, proton translocation ability and the metal centres at their active site (Pereira et al., 2001). Current knowledge of the nature of bacterial terminal oxidases indicates that they fall naturally into two fundamental divisions. Most, including the mitochondrial aa3-type cytochrome c oxidase, are members of the haem–copper oxidase superfamily of proteins (Calhoun et al., 1994; Garcia-Horsman et al., 1994; van der Oost et al., 1994). Irrespective of their cytochrome composition, O2 afﬁnity or proton translocating ability, which vary between individual oxidases, members of this family are characterised by the homology of their major subunit, which contains an iron–copper bimetallic active site where the reduction of molecular oxygen to water takes place. The second group comprises quinol oxidases related to the cytochrome bd of E. coli, which are only found in prokaryotes (Junemann, 1997).

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4.7. Terminal Oxidases in P. aeruginosa Five terminal oxidases are encoded in the P. aeruginosa genome (Comolli and Donohue, 2002; Cooper et al., 2003). Four of these putative terminal oxidases belong to the haem copper oxidase superfamily, three of which are predicted to be cytochrome c oxidases and one a quinol oxidase (Fig. 3). The ﬁfth terminal oxidase is the cyanide-insensitive terminal oxidase (CIO), encoded by the cioAB operon. The CIO is part of the cytochrome bd oxidase family, which is a family of two-subunit quinol oxidases that do not contain Cu and have not been shown to translocate protons. They have no homology to members of the haem-copper oxidase superfamily. We will discuss what is known about each of the ﬁve terminal oxidases in turn. However, our knowledge of the organisation, regulation and function of the terminal oxidases of P. aeruginosa is rudimentary at present, and we only have a limited idea as to why and how these ﬁve pathways operate. 4.7.1. Cytochrome cbb3-Type Oxidases A cytochrome c oxidase, ﬁrst identiﬁed as a cytochrome co-type oxidase, was puriﬁed and biochemically characterised from P. aeruginosa as an enzyme with four subunits of molecular masses of 29, 21, 11.5 and 9.5 kDa (Matsushita et al., 1982). The two largest subunits stained for haem-catalysing peroxidase activity. The puriﬁed enzyme possessed b- and c-type cytochromes, both of which reacted with carbon monoxide leading to the view that the putative B-type haem was actually a cytochrome o. The enzyme had a high TMPD-oxidising activity and it reacted preferentially with a membrane-bound cytochrome c551 leading to the suggestion that this was its natural electron donor in vivo. It is likely that this puriﬁed enzyme was one, or a mixture of both, of the now recognised cytochrome cbb3-type oxidases of P. aeruginosa (Fig. 3) (Comolli and Donohue, 2004). Cytochrome cbb3-type oxidases are exclusive to bacteria and are members of the haem-copper superfamily. They were initially identiﬁed as oxidases with roles in respiration under oxygen-limiting conditions. The ﬁrst was identiﬁed in the symbiotic nitrogen-ﬁxing bacterium Bradyrhizobium japonicum as a product of the fixNOQP operon (Preisig et al., 1993, 1996). Similar enzymes were subsequently found in other bacteria with the genes being named ccoNOQP (for a review see Pitcher and Watmough, 2004). Cytochrome cbb3-type oxidases have been found to have very high O2 afﬁnities. A Km value of 7 nM has been measured for the B. japonicum enzyme (Preisig et al., 1996), which facilitates the growth of this bacterium in the root nodule where the free oxygen concentration is in the range 3–22 nM (Preisig et al.,

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1993, 1996). The oxygen afﬁnities and expression patterns of other cytochrome cbb3 oxidases also supports the view that these enzymes are important in adapting to limited oxygen conditions (Van Spanning et al., 1997; Mouncey and Kaplan, 1998; Myllykallio and Liebl, 2000; Smith et al., 2000; Otten et al., 2001; Swem et al., 2001; Swem and Bauer, 2002; Cosseau and Batut, 2004). The cytochrome cbb3 has four subunits, which are encoded by the ccoNOQP cluster. CcoN is the catalytic subunit with two haem Bs one of which, with the copper ion that is also present, forms the bimetallic active site. The arrangement of prosthetic groups and the bimetallic active site are similar to those of other haem copper oxidases as well as to that of subunit I of the nitric oxide reductases (Castresana et al., 1994; van der Oost et al., 1994, see below). The CcoO and CcoP subunits are mono- and dihaem ctype cytochromes, respectively, which may function to transfer electrons into the active site on CcoN. These cytochromes c replace the CuA-containing subunit II that is found more usually in other members of the haemcopper oxidase superfamily. The role of the fourth subunit, CcoQ, is unclear but it may contribute to the stability of the catalytic centre of the enzyme (Zufferey et al., 1996; Oh and Kaplan, 2002). The presence of a cytochrome cbb3-type oxidase in P. aeruginosa was ﬁrst shown by Thony-Meyer et al., (1994). However, the completion of the genome sequence of P. aeruginosa showed that the bacterium contained two ccoNOQP operons [although the ccoQ genes were not annotated], which are predicated to encode cytochrome cbb3-type oxidases, named cbb3-1 and cbb3-2. In addition, there are two orphan ccoN genes elsewhere on the genome (PA4133 and PA1856). Comolli and Donohue (2004) have investigated the roles of these two oxidases. The cytochrome cbb3-1, unlike other oxidases of this type, has a role at high oxygen tensions. It was highly expressed relative to cbb3-2 at high oxygen tensions and its loss by mutation affected growth rate under these conditions. In contrast, the cytochrome cbb3-2 was more highly expressed under oxygen-limited conditions, and its expression was regulated by the oxygen-sensing transcriptional regulator Anr. While nothing is known at present regarding the biochemistry of these enzymes, it would appear that the two isoforms have specialised roles under different prevailing oxygen levels (Comolli and Donohue, 2004). Cytochrome cbb3 oxidases also have a regulatory role, inﬂuencing bacterial gene expression in response to oxygen availability. Kaplan and colleagues, working on the facultative phototroph Rhodobacter sphaeroides, showed that inactivation of the cbb3 by mutation or the use of speciﬁc inhibitors led to the transcription of genes that are normally only induced by oxygen limitation. This was dependent on a two-component system comprising the sensor kinase PrrB and the response regulator PrrA

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27

(O’Gara et al., 1998; Oh and Kaplan, 1999, 2000, 2001, 2002). They proposed that a change in the redox state of the respiratory chain or the electron ﬂux is sensed by the cbb3 oxidase which in turn generates an inhibitory signal that affects the activity of PrrB and prevents the accumulation of activated PrrA in response to oxygen tension (Oh et al., 2004). RoxRS is the P. aeruginosa homologue of this regulatory system (Comolli and Donohue, 2002) and it has been shown to have a role in the regulation of the cioAB genes encoding the CIO. The activity of cytochrome cbb3-1 not cbb3-2 was found to inﬂuence the expression of the CIO and may act to do this via the RoxRS two component system (Comolli and Donohue, 2004). 4.7.2. Cytochrome aa3 Oxidase A different cytochrome c oxidase was puriﬁed from the P. aeruginosa strain IFO12689 which comprised three subunits with molecular weights of 38, 57 and 82 kDa (Fujiwara et al., 1992). The puriﬁed oxidase contained two molecules of haem A, one of which was CO-reactive, two atoms of copper and one molecule of haem B per protein. However, this cytochrome baa3 accounted for only 1% of the total cytochrome c oxidase activity of membranes of strain IFO12689. This led the authors to suggest that oxidation of cytochrome c may not be of physiological signiﬁcance and that this oxidase was a quinol oxidase. It would seem likely that the oxidase puriﬁed from strain IFO12689 is the enzyme encoded by PA0105–PA0108 (annotated coxB, coxA, PA0107, coIII) on the P. aeruginosa PAO1 chromosome. Although the genome sequence gives the name coIII to the third gene, it will be called coxC here due to its homology to other coxC genes. Another gene lies between coxA and coxC, named PAO107, which has 55% similarity to a probable cytochrome c assembly protein of Pa. denitrificans. The coxA, B and C genes of P. aeruginosa have homology to genes from various bacteria which encode subunits I, II and III, respectively of a cytochrome c oxidase of the haem-copper oxidase family. In Pa. denitrificans and R. sphaeroides, the gene-encoding subunit I (ctaD) is not located with the genes for the other subunits (Cao et al., 1992). However, in B. japonicum, the genes are arranged in a more typical prokaryotic gene order of II-I-III-IV (Surpin et al., 1996), which is closer to that seen in P. aeruginosa. The cytochrome c oxidases encoded by the cox genes of B. subtilis, Pa. denitrificans and R. sphaeroides are all aa3-type oxidases (Garcia-Horsman et al., 1994). The presence of a potential CuA-binding site in the P. aeruginosa coxB subunit (SUII) indicates that the enzyme encoded by these genes is a cytochrome c oxidase and not a quinol oxidase, contradicting the conclusions of Fujiwara et al. (1992). The absence of cytochrome aa3 signals in difference

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spectra of a number of strains of P. aeruginosa suggests that this enzyme is not expressed under normal laboratory growth conditions (Matsushita et al., 1980, 1983; Cunningham and Williams, 1995; Cunningham et al., 1997). Indeed, it is not observed in mutants lacking either the cytochrome cbb3-type oxidases nor the CIO (Cunningham et al., 1997; Comolli and Donohue, 2004).

4.7.3. Cytochrome bo3 Quinol Oxidase The P. aeruginosa genes annotated PA1317–PA1321 share between 72 and 82% homology with the cyoABCDE genes of E. coli. The cyoA, cyoB, and cyoC genes of E. coli encode subunits II, I and III, respectively of a haem-copper oxidase. cyoD is thought to encode a small subunit which assists copper incorporation into subunit I during maturation, while cyoE encodes a farnesyl transferase which is responsible for the production of haem O from haem B (Thony-Meyer, 1997). The mature enzyme is homologous to the aa3-type oxidase of mitochondria but contains haems B and O (bo3-type) and oxidises ubiquinol rather than cytochrome c. All of the cyo genes are required for the catalytic function of the cytochrome bo3 of E. coli (Nakamura et al., 1997). Essential conserved residues within the cyoA and cyoB genes have been identiﬁed as the ligands for the redox centres of the enzyme (Garcia-Horsman et al., 1994). A sequence alignment of the cyoB gene products (subunit I) of P. aeruginosa and E. coli demonstrates that these residues are also present in the P. aeruginosa protein. Histidine residues at positions 106 and 421 are ligands for the low spin haem B, H419 for the high spin haem B and H333 and H334 for the CuB (E. coli numbering) (Garcia-Horsman et al., 1994). Within subunit II of the cytochrome c oxidases, there is also a conserved sequence motif, –HXnCXEXCX3HX2M-, containing the residues responsible for binding the redox centre CuA, which is involved in cytochrome c oxidation. This motif is present in the cytochrome bo3 homologue of Pa. denitrificans, but absent from the E. coli cytochrome bo3 which is a quinol oxidase (Losonczy et al., 1971; Raitio et al., 1987; Surpin et al., 1996). A sequence alignment between these two contrasting bacterial sequences and the P. aeruginosa predicted homologue reveals that the residues required for CuA binding, and therefore necessary for cytochrome c oxidation, are not present in the CyoA of P. aeruginosa, indicating that this enzyme is a quinol oxidase not a cytochrome c oxidase. However, to our knowledge, this putative oxidase has not been studied experimentally and the conditions under which it plays a role in the aerobic respiration of P. aeruginosa are not known.

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4.7.4. The Cyanide-Insensitive Terminal Oxidiase (CIO) The CIO, encoded by the cioAB operon is homologous to the cytochrome bd quinol oxidases of E. coli and A. vinelandii (Cunningham and Williams, 1995; Cunningham et al., 1997; Junemann, 1997), and as such shows no homology to members of the haem-Cu oxidase superfamily (GarciaHorsman et al., 1994; van der Oost et al., 1994). CioA and CioB are homologous to the two subunits of the cytochrome bd quinol oxidase, CydA and CydB, of E. coli. Histidine and methionine residues identiﬁed in E. coli cytochrome bd as being ligands to the low-spin cytochrome b558 (H196 and M329 in CioA) and high-spin b595 (H21 in CioA) are conserved, as is a periplasmic loop – the Q-loop – that contains a putative quinol-oxidising site, although the Q-loop is signiﬁcantly shorter in CioA than in CydA (Cunningham et al., 1997). However, the distinctive absorption bands of a cytochrome bd quinol oxidase are not present in P. aeruginosa membranes (Matsushita et al., 1980, 1982, 1983; Zannoni, 1989; Cunningham and Williams, 1995; Cunningham et al., 1997) and it was proposed that a high-spin haem B replaces the usual haem D in this oxidase. It is likely that there is a family of bacterial quinol oxidases related to the cytochrome bd of E. coli and of other bacteria, which differs in a number of important ways, particularly haem composition, from the E. coli paradigm. Intriguingly, under low O2 conditions, P. aeruginosa synthesises hydrogen cyanide as a metabolic product at concentrations of up to 300 mM. At these concentrations cyanide inhibits the function of members of the haem-copper oxidase superfamily of cytochrome oxidases (Castric, 1975, 1977, 1983; Meganathan and Castric, 1977; Matsushita et al., 1980, 1983; Cunningham et al., 1997; Blumer and Haas, 2000). P. aeruginosa has evolved a respiratory chain that allows its own aerobic respiration to function in the presence of this potent terminal oxidase inhibitor. Therefore, the CIO has been proposed to have a role in allowing aerobic respiration under cyanogenic growth conditions (Cunningham and Williams, 1995; Cunningham et al., 1997). Cyanide has been detected in tissue samples infected with P. aeruginosa (Goldfarb and Margraf, 1967) and recent experiments using mutants that do not make cyanide show that cyanide is a virulence factor in a C. elegans model of infection (Gallagher and Manoil, 2001; and see below). Mutation or overexpression of the cioAB genes has profound effects on the biology of P. aeruginosa. Mutation of cioAB leads to temperature sensitivity for growth, difﬁculty exiting stationary phase, cell division defects and multiple antibiotic sensitivity, probably due to damage to a multidrug efﬂux pump. This may be partly explained by increases in oxidative stress levels in CIOdefective strains as a result of the inability of these strains to synthesise a

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speciﬁc catalase, leading to oxidative protein damage (Tavankar et al., 2003; Mossialos et al,. 2006). The cytochrome bd quinol oxidases are O2-regulated in many bacteria, and normally show increased expression as O2 levels fall (D’mello et al., 1996; Junemann, 1997; Poole and Cook, 2000; Wu et al., 2000; Kana et al., 2001; Larsson et al., 2005). The A. vinelandii enzyme is an exception with its expression increasing at high O2 levels (Moshiri et al., 1991; Wu et al., 2000). Superﬁcially, the P. aeruginosa CIO does not appear to be O2-regulated, at least not in a straightforward way. Varying the O2 transfer coefﬁcient in batch cultures of P. aeruginosa from 11.5 to 87.4 h1 had no effect on CIO expression and there was no correlation observed between CIO induction and the dissolved O2 levels in the growth medium (Cooper et al., 2003). However, a mutant deleted for the O2-sensitive transcriptional regulator Anr de-repressed CIO expression in an O2-sensitive manner, with the highest induction occurring under low O2 conditions (Cooper et al., 2003). Therefore, CIO expression can respond to a signal generated by low O2 levels, but this response is normally kept in check by Anr repression. Anr may play a role in preventing overexpression of the CIO in relation to other terminal oxidases. Cyanide is a potent inducer of the CIO at physiologically relevant concentrations and experiments using spent culture medium from an hcnB mutant, which is unable to synthesise cyanide, showed that cyanide was the inducing factor present in P. aeruginosa spent culture medium. However, the ﬁnding that in an hcnB mutant cioA-lacZ expression is induced normally upon entry into stationary phase indicated that cyanide was not the endogenous inducer of the terminal oxidase. It was suggested that the failure of O2 to have an effect on CIO expression in the wild type can be explained either by the requirement for an additional, stationary phase-speciﬁc, inducing signal or the loss of an exponential phase-speciﬁc repressing signal (Cooper et al., 2003). P. aeruginosa has a two component regulatory system RoxRS, which is related to PrrAB system described previously as regulating electron-transfer components in R. sphaeroides in response to the levels of electron ﬂux through a cytochrome cbb3 terminated respiratory chain (Oh and Kaplan, 2000, 2001; Comolli and Donohue, 2002). It has been shown that mutation of roxR leads to loss of cyanide and azide resistance in P. aeruginosa PAK, and that this is due to CIO deﬁciency. However, the roxR mutant still showed signiﬁcant cyanide-dependent induction of CIO activity and a cioA-lacZ gene fusion, indicating that there are other regulators of the CIO operating that are still to be identiﬁed. Furthermore, these experiments with roxR mutants were performed on exponential phase cultures, that is, conditions under which the CIO expression is at a minimum and during which a putative stationary phase inducing signal is either absent or has its effect repressed

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(Cooper et al., 2003). Therefore, it is premature to suggest that RoxR is the main regulator responsible for the stationary phase induction of the CIO. The role of the CIO in relation to HCN production will be discussed later.

4.8. Cytochrome c Peroxidase Cytochrome c peroxidase (CCP) is a periplasmic enzyme that catalyses the reduction of H2O2 to water, using a cytochrome c as an electron donor (Soininen et al., 1970; Millett et al., 1995). It was puriﬁed from P. aeruginosa as long ago as 1970 by Ellfolk’s lab and both this group and a number of others have studied it extensively (Ellfolk and Soininen, 1970). It is a dihaem cytochrome c with the two haems, which have very different redox potentials of +320 and 330 mV, attached to a single 34 kDa polypeptide subunit. The electron donor to CCP is thought to be the periplasmic cytochrome c, reduced by the cytochrome bc1 complex, but azurin can also act as an electron donor for CCP, at least in vitro. The reaction between cytochrome c and CCP has been studied in detail as a fundamental biological electron transfer process (Soininen et al., 1970; Ronnberg and Ellfolk, 1975; Ronnberg et al., 1979, 1981; Greenwood et al., 1984; Foote et al., 1985, 1992; Greenwood and Gibson, 1989; Ellfolk et al., 1991; Ellfolk and Soininen, 1970; Brittain and Greenwood, 1992; Fulop et al., 1993). Despite its structure being known and the extensive studies of its electron transfer processes (Fulop et al., 1993; Millett et al., 1995; Samyn et al., 1995), virtually nothing is known about the physiological role of CCP in P. aeruginosa, although it is thought to be induced by low oxygen concentrations (Ellfolk et al., 1991). There is evidence of a role for Anr in the regulation of CCP in P. aeruginosa, with Anr being an activator of cytochrome c peroxidase under low oxygen conditions (Ray and Williams, 1997). Data support a role for CCP in cellular detoxiﬁcation of H2O2 in Rhodobacter (De Smet et al., 2005) as well as in yeast (Kwon et al., 2003) and the fungal opportunistic pathogen Cryptococcus neoformans (Giles et al., 2005), although such a role has not yet been demonstrated for the P. aeruginosa enzyme.

5. ANAEROBIC RESPIRATION P. aeruginosa is a denitrifying bacterium. It is able to carry out anaerobic respiration with N-oxides as terminal electron acceptors for anaerobic respiration. Denitriﬁcation is the sequential reduction of nitrate to N2 via nitrite, nitric oxide and nitrous oxide, a process that is catalysed by four

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enzymes: nitrate reductase (NAR), nitrite reductase (NIR), nitric oxide reductase (NOR) and nitrous oxide reductase (N2OR) (Figs. 3 and 6; Zumft, 1997). The denitrifying enzymes provide alternative respiratory routes for energy generation and reoxidation of reducing power under anaerobic conditions, using fundamentally the same mechanisms and sharing many of the components of aerobic respiratory chains. Thus, the terminal reductases of an anaerobic respiratory chain receive electrons from quinones or the cytochrome bc1 complex as do the quinol oxidases or cytochrome c oxidases of aerobic respiration (Figs. 3 and 6). However, in general, anaerobic respiratory chains are energetically less efﬁcient than aerobic respiratory pathways (Zumft, 1997; Richardson, 2000). Denitriﬁcation is a process of great ecological importance as a key part of the nitrogen cycle. Nitrogen is introduced into the biosphere as dinitrogen

Figure 6 Proposed arrangement of the denitriﬁcation pathways of P. aeruginosa with NADH as the source of reductant. Nuo, NADH:quinone oxidoreductase I; Ndh, NADH:quinone oxidoreductase II; Nqr, Na+-translocating NADH-quinone oxidoreductase. While these three NADH:quinone oxidoreductases are shown, it is not known whether they all operate under anaerobic conditions in P. aeruginosa. Q, quinone pool; Nar membrane-bound nitrate reductase; Nap, periplasmic nitrate reductase; Nir, nitrite reductase; Nos, nitrous oxide reductase; Nor, nitric oxide reductase; NarK1/2, putative nitrate: nitrite antiporter; Az, azurin; c551, cytochrome c551.

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by biological nitrogen ﬁxation as well as chemical processes and it is removed from it by denitriﬁcation. On one level denitriﬁcation is an undesirable process as it leads to depletion from the soil of an essential plant nutrient, and thereby decreases agricultural productivity. However, on a global scale, it plays the vital role of ensuring that ﬁxed nitrogen does not end up in the oceans, through the constant leaching of the soluble nitrate and nitrite ions, leading to starvation in terrestrial ecosystems of ﬁxed nitrogen. Denitriﬁcation also preserves the potability of freshwaters (Atlas and Bartha, 1997). Anaerobic respiration and denitriﬁcation have been studied in detail in certain bacteria, particularly Pa. denitrificans, P. stutzeri and E. coli. In this section, we will ﬁrst review what is known about these processes from detailed studies of other bacteria and then describe the current state of knowledge for P. aeruginosa.

5.1. P. aeruginosa Nitrate Reductases There are three different nitrate reductases systems in bacteria (Berks et al., 1995; Moreno-Vivian and Ferguson, 1998; Potter and Cole, 1998). All three (Nas, Nap, Nar) belong to the dimethylsulphoxide (DMSO) reductase family of proteins and contain a molybdenum cofactor in the form of a molybdopterin guanine dinucleotide (bis-MGD) prosthetic group. The P. aeruginosa genome encodes all 3 systems. The ﬁrst of these is the assim+ ilatory NAR which is a cytoplasmic enzyme that reduces NO 3 to NH4 allowing it to be used as a source of nitrogen (Moreno-Vivian and Ferguson, 1998). In P. aeruginosa there is a three-gene cluster PA1781-PA1780-PA1779, which is annotated as nirB, nirD and nasC, respectively. These encode for an assimilatory nitrite reductase large subunit (nirB) and small subunit (nirD). The nasC gene encodes the putative assimilatory nitrate reductase (Nas). The P. aeruginosa Nas has the sequence features consistent with the binding of a bis-MGD cofactor as well as an [4Fe-4S] cluster (Stover et al., 2000). 5.1.1. Membrane-Bound, Dissimilatory Nitrate Reductase (Nar) Membrane-bound Nar have been characterised from both nitraterespiring bacteria such as E. coli and from denitrifying bacteria such as Pa. denitrificans, P. stutzeri and P. aeruginosa (Carlson et al., 1982; Carlson and Ingraham, 1983; Berks et al., 1995; Moreno-Vivian and Ferguson, 1998; Potter and Cole, 1998). Nar enzymes typically comprise three subunits; a large subunit (NarG, 118–150 kDa), which contains the active site with the

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bis-MGD cofactor, a smaller subunit (NarH, 55–64 kDa) and an associated cytochrome b subunit (NarI, 19–21 kDa). Nitrate reduction by Nar is an energetically coupled pathway. Nar is a quinol oxidase and QH2 is oxidised at the periplasmic face of the membrane by the cytochrome b subunit (NarI). Two protons are ejected into the periplasm, while electrons ﬂow back across the membrane via the B haems of NarI and then on via the Fe-S centres of NarH to the bis-MGD cofactored cytoplasmic subunit NarG. Here, nitrate is reduced to nitrite and two protons are consumed from the cytoplasm. Therefore, this electron transfer pathway is a classic redox loop and leads to the conservation of energy as a Dp. P. aeruginosa has a narGHJI operon. The puriﬁed Nar from P. aeruginosa is a molybdo-protein and a Fe-S protein and comprises 118 and 64 kDa subunits (Carlson et al., 1982), which presumably correspond to narG and narH-encoded proteins, respectively. A NarI subunit did not co-purify. narJ encodes a subunit that is not predicted to be part of the enzyme, but has been shown to be needed for the assembly of the functional protein (Blasco et al., 1992, 1998; Dubourdieu and DeMoss, 1992; Vergnes et al., 2006). When P. aeruginosa is grown anaerobically with nitrate as the terminal electron acceptor, it synthesises a membrane-bound formate dehydrogenase, which is thought to participate in an anaerobic respiratory chain. The FDH of E. coli is known to contain cytochrome b, selenium, iron sulphur clusters and molybdenum (Enoch and Lester, 1975). The P. aeruginosa enzyme has Mo, a cytochrome b and a 4Fe-4S centre. Spectroscopic studies of the P. aeruginosa enzyme led to a proposal for the organisation of the metal components of the FDH-nitrate reductase chain (Gadsby et al., 1987; Godfrey et al., 1987a,b). The Mo in the FDH takes electrons derived from formate oxidation and passes them to the cytochrome b with the next component being the [4Fe-4S] centre. Electrons are then passed on directly to the [4Fe-4S] clusters of nitrate reductase without the intermediary of quinones. This ﬁnding does not rule out the participation of quinones in this pathway in vivo and their apparent absence contrasts with the situation in E. coli where quinones are electron transfer components for the FDH-nitrate reductases respiratory chain (Poole and Ingledew, 1987). 5.1.2. Periplasmic Nitrate Reductases (Nap) In addition to a membrane-bound nitrate reductase, the P. aeruginosa genome has the napEFDABC genes encoding for the production of a putative periplasmic nitrate reductase (Nap). The Nap enzyme has been studied from a number of bacteria. It is a two-subunit enzyme located in the periplasm and composed of a catalytic 90 kDa protein containing a bis-MGD cofactor

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and one 4Fe-4S centre, together with a 13–19 kDa bihaem cytochrome c, encoded by the napAB genes, respectively (Reyes et al., 1998). Electron transfer to Nap requires a membrane-bound tetrahaem cytochrome c encoded by napC (Berks et al., 1995; Reyes et al., 1996). Like Nar, Nap is a quinol oxidase, but its location and its consequent oxidation of QH2 and transfer of electrons to nitrate for reduction at the periplasmic surface means that it does not form a redox loop and makes no contribution to the Dp. As a result, energy is dissipated. It makes energetic sense for a bacterium to make use of NAR anaerobically when the organism is dependent on nitrate for Dp formation. The predicted physiological roles of Nap are varied and are reﬂected in the ways in which Nap is regulated in bacteria. In some bacteria, it is the sole nitrate reductase and it has a role in supporting anaerobic growth through nitrate respiration. This is the case in Pseudomonas sp. G-179 (Bedzyk et al., 1999). In E. coli, which has two membranebound Nar enzymes in addition to Nap, it is induced by anaerobic conditions but only at low nitrate concentrations which support a role for Nap in anaerobic respiration under nitrate-limiting environments (Potter and Cole, 1998; Potter et al., 1999). In contrast, in Paracoccus pantrophus Nap is made aerobically in the presence and absence of nitrate and it is responsible for the ﬁrst step in aerobic denitriﬁcation (Bell et al., 1990). Indeed, the nap operon is repressed during anaerobiosis and expressed in the presence of oxygen, but only when growth is supported by a highly reduced carbon source such as butyrate (Ellington et al., 2003). This led to the suggestion that its physiological role may be in part a way of maintaining redox balance by removing excess reductant. A similar role has been suggested for Nap in Rhodobacter sp. (Richardson, 2000; Gavira et al., 2002). The P. aeruginosa Nap has not been biochemically characterised nor has its regulation or physiological role been determined in detail. Denitriﬁcation is not a solely anaerobic process and it is well established that P. aeruginosa can carry out aerobic denitriﬁcation (Davies et al., 1989; Chen et al., 2003). Aerobic denitriﬁcation appears to function as a respiratory process that is complementary to, or competitive with, aerobic respiration (Chen et al., 2003). Recent microarray data are consistent with Nap having a role in aerobic denitriﬁcation in P. aeruginosa (Filiatrault et al., 2005). In this study, the changes in gene expression following a switch from aerobic growth in the presence of nitrate to anaerobic growth in the presence of nitrate were examined. It was found that there was no signiﬁcant change in expression levels of some of the P. aeruginosa denitriﬁcation genes nar, nap, nir, nos or nor, suggesting that nitrate but not anaerobiosis may induce their expression. This was supported by the upregulation of the narI gene

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encoding the cytochrome b subunit of Nar, when growth was switched from aerobic without nitrate to aerobic with nitrate. However, the narGH and narJ genes encoding the other components of the P. aeruginosa Nar were not upregulated by this switch. The expression of napB napA and napD was induced by this switch (as was nosRDFYL and nirS) suggesting that nitrate may be sufﬁcient to induce expression of many of the denitriﬁcation genes in P. aeruginosa and suggesting a role for Nap in aerobic denitriﬁcation. This contrasts with the ﬁndings of others that the oxygen-sensitive transcriptional regulator Anr and a homologue Dnr are required for P. aeruginosa to denitrify (Arai et al., 1995b, 1997; Ye et al., 1995). The regulation of denitrifying enzymes in P. aeruginosa needs further study. These data raise the issue of the role Nap plays during colonisation of the CF lung by P. aeruginosa where the presence of nitrate and an oxygen gradient across the mucus layer could favour a role for aerobic denitriﬁcation, particularly if reduced carbon sources are present.

5.2. P. aeruginosa Nitrite Reductase There are two completely distinct types of nitrite reductase in bacteria that differ in their structure and the nature of their prosthetic groups. The ﬁrst type is the copper-containing nitrite reductase (Zumft, 1997). However, in pseudomonads, including P. aeruginosa, the nitrite reductase is of the second type, a cytochrome cd1. Cytochrome cd1 was ﬁrst described by Horio as a cytochrome c oxidase (Horio et al., 1961). Being a soluble enzyme, it was studied for many years as a model for oxygen activation and electron transfer studies, in part because of the difﬁculty at the time in purifying the ‘‘true,’’ membrane-bound cytochrome c oxidases (Horio et al., 1961). However, its physiological role is as a nitrite reductase in denitriﬁcation. It is located in the periplasm of P. aeruginosa (Silvestrini et al., 1994) and is a homodimer with subunit masses of 60 and 264 kDa. It has haem C and D1 prosthetic groups which have redox potentials of +290 and +287 mV, respectively. It catalyses the oxidation of cytochrome c551 (NirM) in reducing nitrite to nitric oxide, but can also accept electrons from the blue copper protein azurin. Cytochrome c551 and azurin are interchangeable in vitro and exist in rapid equilibrium with each other, although cytochrome c551 is the more efﬁcient electron donor in vitro (Zumft, 1997). However, the in vivo role of azurin is uncertain (Vijgenboom et al., 1997), as an azu mutant was not affected in nitrite reduction and growth rate is only affected in an azurin cytochrome c551 double mutant. Furthermore, the expression pattern of the azu gene does not correlate well with the expression of the denitriﬁcation

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system (Vijgenboom et al., 1997). A functional cytochrome bc1 complex is essential for nitrite reduction, and is involved in the transfer of electrons from the quinone pool to nitrite reductase via cytochrome c551 or azurin (Hasegawa et al., 2003). The genes for cytochrome cd1 are located in the nirSMCFDLGHJEN gene cluster. nirS encodes the structural gene for the nitrite reductase, while nirM and nirC encode two monohaem c-type cytochromes (Arai et al., 1991; Kawasaki et al., 1995). nirM encodes the cytochrome c551, which is the physiological electron donor for nitrite reductase and has been extensively studied (Horio et al., 1958, 1961; Wood, 1978; Arai et al., 1991; Kawasaki et al., 1995). It is a small protein (8685 Da), with a C-type haem attached to Cys12 and Cys15, with His16 and Met61 acting as axial ligands to the haem iron (Garau and Gomez, 2003). The biochemistry and biophysics of electron transfer within this cytochrome and its redox partners have been extensively studied. nirC also encodes a monohaem cytochrome c that can act as an electron donor for NIR (Hasegawa et al., 2001). The nirSMC genes are followed by genes for the synthesis of the haem D1 (Kawasaki et al., 1997). The azurin is not encoded by the nir cluster but by PA4922, which is located elsewhere on the chromosome.

5.3. P. aeruginosa Nitric Oxide Reductase While the formation of nitric oxide (NO) by bacteria was demonstrated about 50 years ago, it was a further 30 years before a nitric reductase (Nor) was isolated (see Zumft, 2005). NOR is an integral membrane protein that has been best studied in Pa. denitrificans, P. stutzeri and Ralstonia eutrophus (Heiss et al., 1989; Hoglen and Hollocher, 1989; Carr and Ferguson, 1990; Braun and Zumft, 1991, 1992; Dermastia et al., 1991; Kastrau et al., 1994; Zumft et al., 1994; Fujiwara and Fukumori, 1996; Cramm et al., 1997; Sakurai and Sakurai, 1997; Cheesman et al., 1998; Hendriks et al., 2000). While three different bacterial NORs have been characterised, the best studied is a cytochrome bc, which comprises two subunits NorC (17 kDa) and NorB (53 kDa) (for review see Zumft, 2005). NorC contains a low-spin haem C and acts as an electron carrier from a soluble cytochrome c to the catalytic subunit NorB. The metal centres of NorB consist of a low-spin haem B and a catalytic binuclear centre containing a high-spin haem B (cytochrome b3) and a nonhaem iron (FeB) (Zumft, 2005). The amino acid sequence of NorB is homologous to that of the largest subunit of the cytochrome c oxidases and this has led to the view that NOR belongs to the haem-copper oxidase

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superfamily, and that the two enzymes share a common ancestor (Hendriks et al., 2000; Zumft, 2005). So cytochrome b3 and FeB in Nor are equivalent to the a3-CuB (or b3-CuB or o-CuB) binuclear centre of terminal oxidases of the haem-copper superfamily. It is apparent that a key difference between NORs and haem-copper oxidases is that they have a binuclear centre that contains a FeB rather than CuB. A further difference is that many haem copper oxidases act as an H+ pump. For example, cytochrome aa3 pumps 4H+ across the cytoplasmic membrane of the bacterium for every O2 reduced to H2O. The evidence is that NOR does not have an H+-pumping activity and the proposed periplasmic active site of NO reduction would rule out its function as a redox loop. It catalyses the following reaction: 2NO þ 2e þ 2Hþ ! N2 O þ H2 O If the NorC subunit is present it terminates an electron transfer pathway that includes the cytochrome bc1. In contrast, in some denitriﬁers such as R. eutrophus (Cramm et al., 1997), the NOR lacks the NorC subunit and acts as a quinol oxidase, with a consequent reduction in the energetic efﬁciency of NO reduction. The genes encoding NOR were identiﬁed some time ago in P. aeruginosa (Arai et al., 1995a). In P. aeruginosa, the NOR is encoded by the norCB genes and has been puriﬁed and characterised (Kumita et al., 2004) and it would appear to accept electrons from a pathway including cytochrome bc1. The norCB genes are also adjacent to other genes with putative roles in production of a functional NOR and the nitrite reductase gene clusters.

5.4. P. aeruginosa Nitrous Oxide Reductase Nitrous oxide reductase (N2OR or Nos) catalyses the last step in the denitriﬁcation pathway, the two-electron reduction of N2O to N2. N2OR has been puriﬁed from P. aeruginosa (SooHoo and Hollocher, 1991) and it is very similar to the enzymes isolated from a number of bacteria including those from P. stutzeri and Pa. denitrificans (Snyder et al., 1987; Scott et al., 1989). This type of N2OR has two Cu centres per subunit CuA and CuZ. The former is a two-copper centre similar to the CuA site in haem–copper oxidases and CuZ is at the catalytic centre (Brown et al., 2000a,b). In P. stutzeri the N2OR is encoded in the nosRZDFYL operon, which is clustered with the nir and nor genes. nosZ is the structural gene for the enzyme, nosDFY encodes proteins involved in processing and insertion of the copper and nosL is an outer membrane disulphide isomerase whose role is unknown. nosR encodes a putative membrane protein with a N2O-sensing regulator that is required for N2OR gene expression (Honisch and Zumft,

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2003; Zumft, 2005). In P. aeruginosa, PA3391–PA3396 encode nosRZDFYL genes that are highly similar to the well-studied P. stutzeri system. The percentage identities for corresponding genes being nosR (75%), nosZ (80%), nosD (71%), nosF (70%) nosY (77%) and nosL (60%), provided they are not clustered with nir and nor genes. Unlike most denitriﬁers, P. aeruginosa cannot grow on exogenous N2O as the sole electron acceptor even though it can use endogenous N2O for generation of free energy and for growth during denitriﬁcation (Carlson and Ingraham, 1983; Bryan et al., 1985).

5.5. Regulation of Denitrification Genes The nosR gene encodes a putative membrane protein (Cuypers et al., 1992) Its product is proposed to be an N2O-sensing regulator that is required for the expression of the N2OR genes (Vollack and Zumft, 2001). In P. stutzeri, the nos gene cluster is transcribed from at least three promoters located upstream of nosR, nosZ and nosD, but this has not been conﬁrmed to be the case in P. aeruginosa (Cuypers et al., 1995). P. aeruginosa does not express N2OR in response to N2O, and so the mechanism of regulation of N2OR of P. aeruginosa is expected to differ from that of other denitriﬁers such as P. stutzeri, which are able to grow on N2O (SooHoo and Hollocher, 1990). The expression of the P. aeruginosa nir and nor genes is regulated by Anr and Dnr, which belong to the Fnr regulator family of transcriptional regulators (Arai et al., 1995a, 1997). Fnr regulates genes in response to the availability of oxygen, acting as a direct oxygen sensor. Oxygen depletion is sensed by a 4Fe-4S cluster (Beinert and Kiley, 1999), which is liganded by four cysteine residues. These residues are conserved in Anr, which is a functional analogue of Fnr in P. aeruginosa (Sawers, 1991; Zimmermann et al., 1991). In contrast, Dnr lacks these cysteine residues and is proposed to sense N-oxides, in particular NO and not oxygen (Arai et al., 1995b, 1997). Motifs similar to the Fnr box are located in the promoter regions of nir and nor operons, but these promoters are activated by Dnr, not Anr. However, Dnr itself is regulated by Anr and so Anr indirectly regulates the levels of the nir and nor genes. The effect of the operation of these two regulators is to switch on the expression of nir and nor genes in the presence of N-oxides under anaerobic conditions (Arai et al., 1997). However, this model does not explain the physiological role of these systems under aerobic denitrifying conditions. Dnr is also required for the expression of the nos genes, with an Fnr-binding motif being located upstream of nosR. NO is an inducer of the nos genes as NO and NO-generating agents induced nosR promoter activity but N2O did not (Arai et al., 2003).

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Arginine / ornithine antiporter

Pi

ADI

Arginine (out)

Arginine (in)

Ornithine (out)

Ornithine (in)

Citrulline NH3

ADP cOTC

CK Carbamoylphosphate

ATP CO2 + NH3

CM

Figure 7 The arginine deiminase pathway of P. aeruginosa. ADI, arginine deiminase; CM, cytoplasmic membrane; cOTC, catabolic ornithine carbamoyltransferase; CK, carbamate kinase. (Adapted from Gamper et al., 1991.)

6. FERMENTATION P. aeruginosa has often been described as a non-fermentative bacterium. This is not the case, but it has only limited fermentative capacity. It has been long recognised that in the absence of oxygen and nitrate P. aeruginosa can use arginine as source of energy for growth using the arginine deiminase (ADI) pathway (Shoesmith and Sherris, 1960; Van der Wauven et al., 1984). This pathway catalyses the breakdown of L-arginine to L-ornithine, with the formation of an ATP (Fig. 7). However, anaerobic growth on arginine requires a rich growth medium and is slow, taking 4–5 days for colonies to form under these conditions. Three enzymes are involved in the ADI pathway, arginine deiminase itself, a catabolic ornithine carbamoyltransferase and carbamate kinase (Fig. 7) (Mercenier et al., 1980). The structural genes for the three enzymes (arcABC) are preceded by the arcD gene, which encodes an arginine–ornithine antiporter, and the four genes are organised into the arcDABC operon (Luthi et al., 1986, 1990; Baur et al., 1989). The ADI pathway is induced under anaerobic conditions and its induction is speciﬁcally enhanced by arginine (Mercenier et al., 1980; Park et al., 1997; Lu et al., 1999). Anaerobic induction is mediated by Anr through its binding at the arcD promoter and mutants defective in the anr gene cannot grow anaerobically with arginine as the sole energy source. Arginine induction is mediated by the ArgR regulatory protein, which binds to a conserved sequence motif upstream of the Anr binding site in the arcD promoter. In heterofermentative bacteria, the reduction of pyruvate to lactate (L- (+) or D- (–)) is a major route for the regeneration of NAD+ and it is catalysed by cytoplasmic lactate dehydrogenases. The P. aeruginosa genome was found to have an ldhA gene, encoding a fermentative D-lactate

OXYGEN, CYANIDE AND ENERGY GENERATION

HCO3+ ATP

ADP + Pi Oxalacetate NADH + H+ Mdh NAD+ Malate Fum

41

NADH + H+

NAD

Pyruvate

Lactate LdhA

PycA CoA + NAD+ Pdh CO2 + NADH + H+

Fumarate FADH2 Frd FAD Succinate

Acetyl-CoA Pta

Pi

CoA Acetyl – phosphate

AckA

ADP ATP

Acetate

NADH + H+

Acetaldehyde dehydrogenase

NAD+ Acetaldehyde NADH + H+ NAD+

AdhA

Ethanol

Figure 8 Proposed model for pyruvate fermentation by P. aeruginosa. PycA, pyruvate carboxylase; LdhA, fermentative lactate dehydrogenase; Pdh, pyruvate dehydrogenase; Pta, phosphotransacetylase; AckA, acetate kinase; AdhA, alcohol dehydrogenase. Enzymes of the reductive citric acid cycle: Mdh, malate dehydrogenase; Fum, fumarase; Frd, fumarate reductase. (Adapted from Eschbach et al., 2004.)

dehydrogenase (LDH). A recent report looked at the role of this LDH in P. aeruginosa (Eschbach et al., 2004). In a search for open reading frames with potential roles in fermentative metabolism, these authors also identiﬁed an ackA-pta locus potentially encoding an acetate kinase and a phosphotransacetylase, respectively and the adhA locus encoding a potential alcohol dehydrogenase. It was found that neither pyruvate fermentation nor the fermentation of glucose could allow signiﬁcant anaerobic growth of P. aeruginosa. However, pyruvate fermentation was found to be important for the long-term anaerobic survival of P. aeruginosa under stationary phase conditions and survival was associated with the accumulation of increasing amounts of possible fermentation products, lactate, acetate and succinate, in the growth medium accompanied by stoichiometric decreases in pyruvate levels. These data provide support for a model of mixed acid pyruvate fermentation by P. aeruginosa, which is further strengthened by the ﬁnding that the pts and ldhA genes are essential for pyruvate fermentation. On the basis of the ﬁndings Eschbach et al. (2004) proposed a model for pyruvate fermentation in P. aeruginosa (Fig. 8).

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7. ANAEROBIC METABOLISM IN THE CYSTIC FIBROSIS LUNG Recent research has implicated the formation of anaerobic bioﬁlms and consequently the operation of anaerobic respiratory pathways in the colonisation of and survival in the CF lung by P. aeruginosa. In an elegant study, referred to earlier, Worlitzsch et al. (2002) showed that steep oxygen gradients exist in the mucus lining of the CF lung, whereas no equivalent gradients exist in the mucus of the healthy non-CF lung. There is evidence of raised oxygen consumption by CF epithelial cells (Stutts et al., 1986), which may contribute to oxygen gradients across the mucus. P. aeruginosa forms microcolonies in the mucus of the CF lung, probably in a low oxygen or anaerobic environment (Lam et al., 1980; Worlitzsch et al., 2002). Here P. aeruginosa probably does not grow as a surface-attached bioﬁlm, of the kind used in many model systems, but the bacteria are in a colonial-type structure in a physically heterogeneous environment as opposed to a homogenous planktonic existence. These studies raised the possibility of anaerobic respiration being a major mode of energy generation of P. aeruginosa in the CF lung (Hassett et al., 2002; Yoon et al., 2002). In this context, it is important to realise that CF mucus has signiﬁcant levels of both nitrate and nitrite. Nitrite levels are similar in the sputum from CF and non-CF patients, while nitrate levels are markedly increased in the sputum of CF patients (Linnane et al., 1998; Jones et al., 2000), and it has been reported that the levels are sufﬁcient to support the anaerobic growth of P. aeruginosa (Hassett et al., 2002). P. aeruginosa certainly can form robust bioﬁlms anaerobically in the presence of nitrate (Hassett, 1996; Yoon et al., 2002). Therefore, the operation of anaerobic denitrifying pathways has been proposed in the mucus of the CF lung with the sequential operation of Nar, Nir, Nor and Nos (Hassett et al., 2002). However, as O2 gradients exist within the mucus, the operation of aerobic denitriﬁcation pathways in the CF lung also has to be considered. It has been clear for some years that P. aeruginosa is capable of aerobic denitriﬁcation, as discussed above, and indeed denitriﬁcation can continue at even quite high dissolved oxygen levels (Davies et al., 1989; Chen et al., 2003). For the reasons argued previously, and based on the recent data of Filiatrault et al., 2005, it would seem reasonable to postulate that both aerobic and anaerobic denitriﬁcation occur during P. aeruginosa infection of the CF lung, with the periplasmic nitrate reductases (Nap) operating aerobically and the membrane-bound Nar anaerobically. Consideration should also be given to a role for arginine fermentation and pyruvate fermentation in the survival of P. aeruginosa during chronic lung infections.

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8. SYNTHESIS OF THE RESPIRATORY INHIBITOR HYDROGEN CYANIDE IN P. AERUGINOSA An intriguing aspect of the biology of P. aeruginosa is its ability to synthesise the respiratory inhibitor hydrogen cyanide, which can reach concentrations of 300 mM in laboratory cultures (Blumer and Haas, 2000; Zlosnik and Williams, 2004). The fact that cyanide is synthesised aerobically raises the interesting issue of how can the bacteria respire aerobically while producing cyanide. The fact that cyanide is made only under low oxygen conditions raises the issue of whether it is made in the low oxygen regions of the CF lung and also whether it is made differentially in mucoid and non-mucoid strains. In this section, we will review aspects of HCN production by P. aeruginosa and focus on its interrelationship with respiration, especially the CIO, and the possibility that the CF-lung provides an environment in which the bacterium synthesises HCN. Production of cyanide has been determined in a range of organisms including: bacteria, fungi, algae and in a range of plants (Blumer and Haas, 2000). Currently, bacterial cyanide production is known only in a limited range of bacteria. These include both P. aeruginosa and P. fluorescens as well as Chromobacterium violaceum, P. aureofaciens, P. chlororaphis and Rhizobium leguminosarum (Knowles and Bunch, 1986; Blumer and Haas, 2000). P. aeruginosa itself was probably the ﬁrst bacterium to be described as a cyanide producer, having been reported as cyanogenic in 1913 under its former name Bacillus pyocyaneus (Clawson and Young, 1913).

8.1. Physiology of Cyanide Production In P. aeruginosa, cyanide is produced under microaerophillic conditions on the entry to stationary phase (Castric, 1975, 1983). In ﬂask cultures of P. aeruginosa, levels of cyanide production are known to be around 200–300 mM and it is produced maximally at a temperature between 34 1C and 37 1C (Pessi and Haas, 2000; Zlosnik and Williams, 2004). Also, cyanide production is almost completely abolished (by around 98%) in anaerobic cultures (Castric, 1975). In both P. aeruginosa and C. violaceum, the amino acid glycine is used as the precursor for HCN formation (Michaels and Corpe, 1965; Wissing, 1974, 1975). Indeed, the addition of both glycine and threonine to cultures stimulates cyanide production, with threonine being a precursor to glycine (Castric, 1977). The addition of iron and phosphate, are also known to stimulate cyanide production, although above 10 mM phosphate actually inhibits cyanogensis (Castric, 1975).

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A number of pieces of evidence support the hypothesis that a hydrogen cyanide synthase enzyme catalyses cyanide production by the oxidative decarboxylation of glycine, producing four electrons and four hydrogens per molecule of glycine, in the process (Blumer and Haas, 2000). First, it is known that in C. violaceum the C–N bond is maintained during cyanogenesis from glycine (Brysk et al., 1969; Blumer and Haas, 2000). Second, using radiolabelled carbon experiments, it has been shown that cyanide is formed from the methylene group of glycine, while CO2 is formed from the carboxyl group (Askeland and Morrison, 1983). Finally, alignments of the protein sequences encoded by the hcnABC genes in P. fluorescens with sequences from other organisms indicate that these genes may function as amino acid dehydrogenase/oxidases, which is consistent with the currently proposed mechanism (Blumer and Haas, 2000). An interesting possibility is that the HCN synthase is simply a respiration-linked, amino acid dehydrogenase, which happens to make HCN.

8.2. Genetics of Cyanide Production The regulation of the hcnABC genes is complex. Transcription of the hcnABC genes requires the oxygen-sensitive transcriptional regulator Anr. It is also known that it is regulated via the RhlR quorum sensing system, in response to activation by the GacS/GacA two-component system (Reimmann et al., 1997; Pessi and Haas, 2000; Pessi et al., 2001). Transcription can take place from one of two transcription start sites and a number of potential regulator-binding boxes have been identiﬁed upstream including two lux boxes, similar to those found in other quorum sensing controlled genes. Additionally, it is now clear that a global posttranscriptional regulator, RsmA, can act both to suppress the transcription of the hcnABC genes and, at a posttranscriptional level, the translation of these genes, and thereby the subsequent production of cyanide (Pessi and Haas, 2001). Finally, AlgR, a regulator in alginate biosynthesis, has also been shown to control cyanide synthesis and this is discussed further below.

8.3. Mechanisms of Tolerance to Cyanide The production of cyanide at levels of 200–300 mM is more than sufﬁcient to inhibit most forms of aerobic respiration, catalysed by haem–copper oxidases (Cunningham and Williams, 1995). This, then, strongly implies that P. aeruginosa possesses mechanisms for dealing with the presence of cyanide. In order for aerobic respiration to continue once cyanide production has started, P. aeruginosa will need to be able to use one or both of two

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possible strategies. First, the toxic effects of cyanide could be avoided by the use of a branch of the respiratory chain that is tolerant to cyanide at physiological concentrations or an anaerobic energy-generating pathway could be used. Alternatively, P. aeruginosa could use some mechanism to detoxify the cyanide it has produced. Strong candidates for both such strategies exist. Cyanide-insensitive respiration is well characterised in P. aeruginosa, being mediated by the cyanide insensitive oxidase (CIO) (Cunningham and Williams, 1995; Cunningham et al., 1997; Comolli and Donohue, 2002; Cooper et al., 2003). As a mechanism for cyanide-tolerance, this terminal oxidase is an attractive proposition as, along with the ability to reduce oxygen in the presence of 1 mM cyanide, its synthesis is coincident with cyanide production, both being made on entry to stationary phase (Cooper et al., 2003). Additionally, a role for the CIO has been suggested as a sink for electrons produced during cyanogenesis (Blumer and Haas, 2000). It has been determined recently that the CIO can provide a growing culture with tolerance against exogenous cyanide and can raise the cyanide MIC around four-fold when a stationary phase inoculum is used (Zlosnik et al., 2006). However, this protection appears not to be the absolute mechanism of cyanide tolerance in P. aeruginosa as cio mutants continue to produce cyanide at wild-type levels in early-stationary phase and are not affected in their long-term viability in stationary-phase culture (Zlosnik et al., 2006). This former observation is also signiﬁcant as it rules out the possibility that the CIO is acting as the sole sink for electrons produced in cyanogenesis. Given that cyanide can also inhibit metallo-enzymes (including important protective enzymes such as catalase, peroxidase and superoxide dismutase), it seems unlikely that respiratory protection alone would provide P. aeruginosa with sufﬁcient tolerance to physiological cyanide. Therefore, in addition, it is likely that P. aeruginosa possesses mechanisms to detoxify cyanide. Indeed, over a period of more than a week, cyanide levels in stationary phase cultures of P. aeruginosa, sealed to prevent gas loss, decline (Zlosnik et al., 2006). The recent characterisation of the enzyme rhodanase in P. aeruginosa also lends support to this idea (Cipollone et al., 2004, 2006). Rhodaneses are one of a number of enzymes capable of the catalytic degradation of cyanide, in this case by the transfer of a sulphur group to the cyanide anion to form thiocyanate. Other potential enzymes are known to be produced in other cyanide-producing bacteria. These include g-cyano-aaminobutyric acid synthase and b-cyanoalanine synthase, which are known to be synthesised by the cyanide producer C. violaceum, which also produces a rhodanese (Knowles and Bunch, 1986). P. fluorescens is known to be able

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to scavenge and utilise cyanide as a nitrogenous growth substrate by the action of cyanide oxygenase (CNO). The CNO is a cytosolic enzyme that has been recently characterised as being composed of four separate enzymes working in concert to convert cyanide into ammonia, which is then assimilated (Fernandez and Pizarro, 1996).

8.4. Evidence for the Biological Function of Cyanide Produced by P. aeruginosa The physiological role of bacterial cyanide production in general is far from clear. In P. aeruginosa, it was suggested that cyanide may be produced as a mechanism for detoxifying, via a metabolic shunt to cyanide as a secondary metabolite, excess quantities of glycine and threonine (Castric, 1975, 1977, 1983). This hypothesis, is consistent with the temporal production of cyanide in batch culture. However, given the toxicity of cyanide, it seems perhaps more likely the production of cyanide may provide P. aeruginosa with an advantage in the range of ecological niches it inhabits. Indeed, cyanide is a virulence factor in P. aeruginosa in the paralytic killing model of C. elegans (Gallagher and Manoil, 2001). In human infections, the role of cyanide is far from clear. An initial report of the presence of HCN in tissues infected with P. aeruginosa has not been followed up (Goldfarb and Margraf, 1967). The possibility of cyanide production in CF is discussed further below. Given that P. aeruginosa is an opportunistic human pathogen, it would seem more likely that cyanide production evolved for its signiﬁcance in the environmental niches it inhabits. This is supported by the observation that in P. fluorescens cyanide can suppress black root rot of tobacco in a gnotobiotic system (Voisard et al., 1989). Additionally, it has been reported that P. fluorescens strain CHAO can use cyanide as a biocontrol mechanism against the tomato crown and root rot pathogen Fusarium oxysporum f. sp. radicis-lycoperisci (Duffy et al., 2004). Therefore, further investigation of cyanide production in P. aeruginosa is merited to understand what, if any, signiﬁcance it has in the clinical and environmental settings which it inhabits. Of relevance here is the low O2 environment of the CF mucus, which satisﬁes one of the environmental parameters for cyanide synthesis by P. aeruginosa, another being high population density to facilitate quorum sensing-mediated up-regulation of the hcnABC genes. We have recently assayed sputum samples for cyanide. We found that cyanide was present at concentrations of up to 200 mM in sputum of CF and bronchiectasis patients with P. aeruginosa infections (B. Ryall, J. Davies, R. Wilson and H.D. Williams, unpublished observations). Such cyanide levels may well compromise the function of cells and tissues in the lung and contribute to lung damage

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caused by chronic P. aeruginosa infections. Cyanide may also be an important factor in the ability of P. aeruginosa to exclude other bacterial pathogens from the CF lung, which leads to it being the dominant infecting bacterium. In this last regard, it is interesting that we have recently found that Burkholderia cepacia complex organisms, which are the other major CF pathogens, sythesise HCN (B. Ryall, J. Zlosnik and H.D. Williams, unpublished).

9. MUCOID CONVERSION OF P. AERUGINOSA IN THE CYSTIC FIBROSIS LUNG: THE ROLE OF OXYGEN AND ENERGY METABOLISM The production of the exopolysaccharide alginate confers the well-described mucoid phenotype on P. aeruginosa. Alginate is a linear copolymer composed of b-D-mannuronic acid and a-L-guluronic acids and its production in bacteria was ﬁrst described in P. aeruginosa in 1964 (Linker and Jones, 1964). Alginate is also known to be produced as an extracellular polysaccharide by A. vinelandi and other Pseudomonads (Gacesa, 1998). In CF a typical pattern of P. aeruginosa infection has long been established, whereby CF patients become infected by environmental non-mucoid strains, which subsequently undergo a phenotypic switch to the mucoid colony phenotype (for review see Govan and Deretic, 1996). Interestingly, a mucoid isolate of the other major CF pathogen, B. cenocepacia, has been recently described (Conway et al., 2004). While the exopolysaccharide produced in mucoid B. cenocepacia is chemically distinct from alginate, this coupled with the fact that mucoid P. aeruginosa is rarely isolated from other infections or the environment, does suggest the CF lung speciﬁcally provides an environment to select for overproduction of exopolysaccharide. The genetic mechanisms underlying the switch to mucoidy in P. aeruginosa have been well studied (for review see Govan and Deretic, 1996). However, it is still not clear what factors induce this switch or indeed what the precise physiological beneﬁt the mucoid switch confers on P. aeruginosa. While a full consideration of mucoid P. aeruginosa is beyond the scope of this review, there are a number of pertinent observations that link energy generation and oxygen with the mucoid switch that we will consider here.

9.1. Oxygen Several lines of research have demonstrated that exposure to oxygen can result in the mucoid switch. First, Krieg and co-workers showed that the

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mucoid phenotype was retained in aerated batch and continuous cultures, while in a non-aerated batch culture it was progressively lost with nonmucoid revertants dominating the culture after 33 h (Krieg et al., 1986). The beneﬁt of mucoidy under aeration conditions was further established by the observation in the same study that mucoid strains would eventually predominate in a continuous culture started as an equal mix of mucoid and nonmucoid strains. The possibility that oxygen itself could serve as a trigger for conversion into mucoidy has been demonstrated by the observation that high levels of oxygen, reﬂective of the levels in the left-hand cardiac circuit, can select for mucoid P. aeruginosa (Bayer et al., 1989, 1990). Additionally, Sabra et al. (2002) have reported that non-mucoid PAO1 can produce a mucoid exopolysaccharide under conditions of oxygen stress. These data suggest that alginate might help to protect P. aeruginosa against oxidative damage, a conclusion supported by the ﬁnding that P. aeruginosa converts into a mucoid phenotype when cultures are treated with levels of H2O2 equivalent to those produced by polymorphonuclear leucocytes (Mathee et al., 1999). Oxygen has also been shown to regulate the transcription of the alginate biosynthetic genes algA, algC and algD (Leitao and Sa-Correia, 1997). However, this upregulation of transcription was not followed by a concomitant increase in either speciﬁc enzyme activity or alginate production. It is thus conceivable that alginate could act as a barrier to oxidative stress for P. aeruginosa in the CF lung. Alginate can be produced during anaerobic respiration and it can act to limit the diffusion of oxygen (Hassett, 1996). Interestingly, in A. vinelandii, alginate has been shown to function to provide protection to the highly oxygen-sensitive enzyme nitrogenase (Sabra et al., 2000).

9.2. Phosphate Phosphate-limited cultures of non-mucoid P. aeruginosa can give rise to mucoid sub-populations (Terry et al., 1991, 1992). Further research has shown a link between phosphate starvation, central metabolism and algR2 (also known as algQ) – one of the genes involved in the regulation of alginate biosynthesis. Speciﬁcally, algR2 regulates nucleoside diphosphate kinase (ndk) and the TCA cycle enzyme succinyl-CoA-synthetase (scs) (Schlictman et al., 1994). This would appear to suggest that algR2 could modify central metabolism, perhaps to facilitate alginate production. The promoter region of algR2 has been shown to respond to phosphate starvation and experimental evidence has been presented to support a model where phosphate starvation leads to low NTP levels, which results in increased expression of algR2. This in turn up-regulates ndk and scs resulting

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in increased levels of NTPs, along with an upregulation of alginate production (Sundin et al., 1996; Kim et al., 1998). The production of alginate itself requires both phosphorylated sugars and GTP; therefore, this might function to ensure sufﬁcient GTP is available for alginate biosynthesis.

9.3. Energy Inhibitors The direct inhibition of energy generation has also been shown to result in the conversion of non-mucoid to mucoid P. aeruginosa. Using strain PAO1, Terry et al. (1992) showed that treatment with gramicidin (a proton translocation inhibitor) and N0 ,N00 -dicyclohexylcarbodiimide (DCCD, inhibits ATP synthase) resulted in conversion of a small percentage (0.1–1.8%) of the population into mucoidy. However, this effect was much more marked when DCCD was combined with phosphate limitation, where more than 50% of a culture converted to the mucoid phenotype.

9.4. Perspective While it is not clear what causes the mucoid switch in the CF lung, substantial evidence exists that this can be affected in the laboratory under a range of metabolic stress conditions, in addition to those listed above. It is also known that mucoidy can arise from non-mucoid strains through nutritional stress, such as growth on acetamide, nitrogen limitation, iron limitation and antibiotic treatment (Govan and Fyfe, 1978; Speert et al., 1990; Terry et al., 1991, 1992). The CF lung itself is likely to be an environment of both nutrient limitation and oxidative stress for P. aeruginosa. However, paradoxically, it is also known that the production of alginate is an energetically costly process to P. aeruginosa, using over 50% of the glucose supplied in a continuous culture (Mian et al., 1978). Therefore, it would seem clear that mucoidy must offer a considerable beneﬁt to an energetically challenged cell.

9.5. A Link between Mucoidy and Cyanide Production Recently, it has been shown that cyanide synthesis is also controlled by the AlgR transcriptional regulator, part of the alginate biosynthesis regulatory network (Carterson et al., 2004; Lizewski et al., 2004). These two studies have shown that AlgR can also modulate transcription of the hcn genes by

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binding to a site between 410 and 402 bp upstream of the hcnA gene and that the effect of AlgR on transcription is different depending on whether the strain is producing alginate or not. Carterson et al. (2004) showed, using an assay that traps cyanide given off by plate cultures, that in mucoid strains (alginate over producing strains) AlgR acts as an activator of the hcnABC genes, whilst in non-mucoid strains it acts as a repressor. While this work implies that alginate biosynthesis and cyanide production are co-ordinately regulated, these studies have not attempted to integrate the role of this regulator with the other previously well-characterised factors involved in regulation of the hcnABC genes, nor have they provided experimental evidence for the mechanism by which this switch in regulatory function takes place. The signiﬁcance of cyanide in CF is not currently understood. However, in a survey of over 167 isolates of P. aeruginosa from 103 CF patients, 74% were cyanogenic (Carterson et al., 2004). Quantiﬁcation of cyanide from 41 CF isolates from 24 patients was also reported and it was shown, using the plate trapping assay, that mucoid strains produced markedly more cyanide than non-mucoid strains, raising the possibility that cyanide could be signiﬁcant in these infections (Carterson et al., 2004).

10. CONCLUSION P. aeruginosa has broad-ranging energy-generating pathways which will be important in the adaptation of P. aeruginosa to the niche of the CF lung. The recent demonstration of oxygen gradients across and anaerobic conditions in the depths of the mucus layer of the CF lung suggest that a full range of energy generating pathways will function in P. aeruginosa during colonisation and chronic infection of the CF lung. This information, coupled with the HCN production in the mucus layer and the potential role of oxygen and energy metabolism in the phenotypic switch to mucoidy associated with the CF lung, emphasises the need for more in-depth studies of the operation and regulation of the respiratory pathways of P. aeruginosa.

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Structure, Mechanism and Physiological Roles of Bacterial Cytochrome c Peroxidases John M. Atack and David J. Kelly Department of Molecular Biology and Biotechnology, The University of Sheffield, Western Bank, Sheffield S10 2TN, UK

ABSTRACT Cytochrome-c peroxidases (CCPs) are a widespread family of enzymes that catalyse the conversion of hydrogen peroxide (H2O2) to water using haem co-factors. CCPs are found in both eukaryotes and prokaryotes, but the enzymes in each group use a distinct mechanism for catalysis. Eukaryotic CCPs contain a single b-type haem co-factor. Conventional bacterial CCPs (bCCPs) are periplasmic enzymes that contain two covalently bound c-type haems. However, we have identiﬁed a sub-group of bCCPs by phylogenetic analysis that contains three haem-binding motifs. Although the structure and mechanism of several bacterial dihaem CCPs has been studied in detail and is well understood, the physiological role of these enzymes is often much less clear, especially in comparison to other peroxidatic enzymes such as catalase and alkylhydroperoxide reductase. In this review, the structure, mechanism and possible roles of bCCPs are examined in the context of their periplasmic location, the regulation of their synthesis by oxygen and their particular function in pathogens.

Copyright r 2007 by Elsevier Ltd. ADVANCES IN MICROBIAL PHYSIOLOGY VOL. 52 All rights of reproduction in any form reserved ISBN 0-12-027752-2 DOI: 10.1016/S0065-2911(06)52002-8

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Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Introduction: enzymic mechanisms to combat oxidative and peroxidative stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1. Catalase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2. Glutathione, Thioredoxin and NADH-linked Peroxidases . . . . . . 1.3. Cytochrome-c Peroxidases . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Phylogenetic analysis of bacterial CCPS reveals a novel sub-group of tri-haem proteins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. MauG PROTEINS. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Structure of bacterial CCPS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Dimer Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Calcium-binding Sites. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. Haem Sites in Bacterial CCPs . . . . . . . . . . . . . . . . . . . . . . . . 5. Mechanistic aspects of catalysis by bacterial CCPs . . . . . . . . . . . . . 6. Electron donors and electron transport in bacterial CCPS . . . . . . . . . 6.1. Nature of Interactions with Electron Donors . . . . . . . . . . . . . . . 6.2. Electron Transport between Haems. . . . . . . . . . . . . . . . . . . . . 7. Roles of CCPS in bacterial cells . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1. Detoxification of Periplasmic Hydrogen Peroxide . . . . . . . . . . . 7.2. Hydrogen Peroxide as an Electron Acceptor . . . . . . . . . . . . . . 7.3. Alternative Substrates and Multiple Enzymes . . . . . . . . . . . . . . 7.4. A Paradox in the Regulation of Expression of Bacterial CCPs? . 8. Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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ABBREVIATIONS bCCP CCP FDH hp lp MADH TTQ

bacterial cytochrome-c peroxidase cytochrome-c peroxidase formate dehydrogenase high potential low potential methylamine dehydrogenase tryptophan tryptophylquinone

1. INTRODUCTION: ENZYMIC MECHANISMS TO COMBAT OXIDATIVE AND PEROXIDATIVE STRESS Any organism, and particularly one that uses oxygen as a terminal electron acceptor, is subject to oxidative stress. Oxidative stress results from the formation of toxic oxygen intermediates formed by incomplete reduction of

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oxygen. Reduction of oxygen to water is a four electron reaction, but toxic oxygen intermediates formed by incomplete reduction include the superox ide anion (O 2 ), hydrogen peroxide (H2O2), and the hydroxyl radical (HO ) (Storz and Imlay, 1999). Superoxide and H2O2 interact to form the highly toxic hydroxyl radical. Build up of these highly reactive species can lead to damage to proteins, nucleic acids and membranes. Reactive oxygen species are also produced by the immune system to kill invading microorganisms. Therefore, the ability to combat these toxic compounds is key to survival in the environment and the host (Storz and Imlay, 1999), and microbes have evolved an impressive array of enzyme systems to detoxify reactive oxygen species. H2O2 is not an abundant molecule in many environments. It is formed naturally mainly by the action of sunlight on water and is thus found in traces in natural water bodies, rain and snow. H2O2 as a molecule is fairly weakly reactive, but the single bond between the two oxygen atoms is easily broken, so that it readily fragments into a hydrogen and a hydroperoxyl radical or into two hydroxyl radicals. H2O2 is generated both endogenously within cells that use oxygen as a terminal electron acceptor and exogenously by, for example, certain types of host cells when pathogens elicit an immune response (Storz and Imlay, 1999; Imlay, 2003). H2O2 can damage proteins through oxidation of co-factors such as iron–sulphur clusters (Imlay, 2002), membranes and DNA, and can react with superoxide (O 2 ) to form the hydroxyl radical (HO ) using iron via the Fenton reaction (Imlay et al., 1988). H2O2 is also produced as an intermediate in superoxide breakdown by superoxide dismutase (SOD), which is a major defence enzyme against damage caused by superoxide. Using two protons, SOD converts two superoxide anions to a molecule of H2O2 and a molecule of oxygen. SOD is widely distributed in nature – nearly all aerobes possess the enzyme, but it is present in only some anaerobes (Imlay, 2002, 2003). The major sources of endogenously produced H2O2 have traditionally been thought to be respiratory chain redox enzymes, especially ﬂavoproteins, that react with molecular oxygen to produce H2O2, but recent studies using mutants of Eschericia coli suggest that this may not be the case. Seaver and Imlay (2004) constructed an E. coli strain lacking both catalases and alkyl-hydroperoxide reductase. This strain released accumulated H2O2 into the medium, where it could be measured. Further mutations were made in this background and, surprisingly, mutants lacking either or both NADH dehydrogenase I or II and/or fumarate reductase continued to produce H2O2 at normal rates. The conclusion is that, in E. coli at least, most peroxide is generated outside the respiratory chain, but as yet, the sources of

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endogenous H2O2 remain to be elucidated (Seaver and Imlay, 2004). Nevertheless, irrespective of the source, bacteria possess three major types of enzymes to remove H2O2, of which the cytochrome-c peroxidases are the particular focus of this review.

1.1. Catalase Catalase removes H2O2 from cells by splitting it to water and oxygen. Catalases can be divided into three types; mono-functional catalases, bi-functional catalase–peroxidases and manganese-containing (non-haem) catalases (Chelikani et al., 2004). Many micro-organisms contain more than one type of catalase, these often being induced under different stress conditions by the action of redox-sensing regulatory systems, e.g. OxyR in E. coli (Zheng and Storz, 2000). Although catalase is the most-widely studied H2O2 detoxiﬁcation mechanism in the cytoplasm (Storz and Imlay, 1999; Chelikani et al., 2004), it may not represent the major way in which H2O2 is removed by bacteria at low concentrations (Seaver and Imlay, 2001).

1.2. Glutathione, Thioredoxin and NADH-linked Peroxidases Glutathione peroxidases (Gpx) are selenoenzymes, which catalyse the reduction of hydroperoxides (H2O2 or ROOH) in the presence of glutathione (GSH). Although glutathione peroxidases are widespread among eukaryotes where they function as a major peroxide defence, they have not been well studied among prokaryotes. However, evidence from mutant studies in Neisseria meningitidis (Moore and Sparling, 1996) and Streptococcus pyogenes (Brenot et al., 2004) indicate a role in the protection against H2O2, and in the latter case a role in virulence. Alkyl-hydroperoxide reductase (Ahp) and the related proteins thiolperoxidase (Tpx) and ‘‘bacterioferritin-comigratory protein’’ (Bcp) are non-haem peroxidases, and are members of the peroxiredoxin family (Poole et al., 2000; Poole, 2005). These enzymes reduce reactive hydroperoxides to their corresponding alcohols using thioredoxin or NADH as an electron donor, and can also reduce H2O2 (Poole et al., 2000; Baker et al., 2001; Guimaraes et al., 2005). In fact, mutant studies have shown that in E. coli, Ahp is more important than catalase in removing low concentrations of H2O2, due to a higher afﬁnity (Seaver and Imlay, 2001). AhpC has also been shown to be a peroxynitrite reductase (Bryk et al., 2000). Hydroperoxides are a problem in cells as they are able to initiate and propagate free radical

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chain reactions that can cause signiﬁcant damage to DNA and membranes (Baillon et al., 1999). In E. coli, Ahp consists of two subunits. AhpC contains the catalytic site, and the ﬂavoprotein AhpF is the electron donor to AhpC (Poole et al., 2000). AhpC homologues are found in many organisms, and all contain highly conserved cys residues at their N- and C-termini (Baillon et al., 1999). Activity was shown to be dependent on the N-terminal cysteine (Ellis and Poole, 1997; Bryk et al., 2000) with the C-terminal cysteine stabilising the oxidised protein via an intersubunit interaction (Ellis and Poole, 1997). In catalysis, the N-terminal cysteine is oxidised to a cys-sulphenic acid upon addition of the hydroperoxide substrate, with the second cysteine stabilising this via disulphide bond formation (Wood et al., 2003). Like catalase, AhpC is cytoplasmic. In many organisms, AhpC exists as an oligomer, usually as a pentamer or hexamer of dimers (Guimaraes et al., 2005; Parsonage et al., 2005). In Helicobacter pylori, mutation of ahpC led to a signiﬁcant decrease in catalase activity. This is due to the disruption of the haem environment of catalase by an increase in organic peroxides in the cell. Thus, AhpC prevents catalase inactivation by removing these organic peroxides (Wang et al., 2004). It is not clear if AhpC is an important virulence factor. Cells of Porphyromonas gingivalis, an aetiological agent of periodontitis, lacking AhpC were more sensitive to organic peroxides, but were just as virulent as wild-type cells in the mouse model of infection (Johnson et al., 2004). Campylobacter jejuni mutants lacking ahpC were hypersensitive to killing by cumene hydroperoxide and signiﬁcantly less aerotolerant than wild-type cells (Baillon et al., 1999). Cells in which ahpC was overexpressed were hyperresistant to killing by cumene hydroperoxide. Tpx and bacterioferritin co-migratory protein (Bcp) are members of a novel class of peroxiredoxins catalysing the removal of peroxides. In E. coli, Tpx is dependent on two cysteine residues for antioxidant activity – Cys61 and Cys94 (Cha et al., 1995; Zhou et al., 1997). Like AhpC, catalytic activity is dependent on an N-terminal cysteine residue (Choi et al., 2003). This cysteine is oxidised to a cys-sulphenic acid intermediate upon addition of peroxides. Catalytic activity is highest with alkyl-hydroperoxides (Choi et al., 2003). Thioredoxin was shown to be the electron donor to the E. coli Tpx (Cha et al., 1995; Zhou et al., 1997) with enzymatic activity apparently localised mainly in the periplasm (Cha et al., 1995). However, Tpx does not have any signal peptide and the direct use of cytoplasmic thioredoxin as a direct electron donor is not consistent with a periplasmic location. Therefore, there is still some doubt as to the true sub-cellular location of this enzyme. Mutants lacking Tpx in H. pylori were signiﬁcantly more sensitive to organic peroxide stress, but loss of Bcp gave only a weak oxidative stress phenotype

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(Comtois et al., 2003; Olczak et al., 2003; Wang et al., 2005a). However, in E. coli, bcp mutants were more sensitive to both H2O2 and organic peroxides (Jeong et al., 2000), indicating differential importance in different bacteria. In many organisms, catalase and AhpC are regulated by the peroxide stress regulator OxyR (Herbig and Helmann 2001; Jakubovics and Jenkinson, 2001; Charoenlap et al., 2005) or by the Fur homologue PerR (Baillon et al., 1999; van Vliet et al., 1999, 2002; Horsburgh et al., 2001).

1.3. Cytochrome-c Peroxidases Cytochrome-c peroxidases (CCPs) are extracytoplasmic haem-containing enzymes that have a role in peroxide stress resistance, and are found in both eukaryotes and prokaryotes. In eukaryotes, such as Saccharomyces cerevisiae, CCPs contain a single b-type haem as a co-factor and are located in the inter-membrane space of mitochondria (Erman and Vitello, 2002). On the other hand, in prokaryotes, CCPs are di-haem proteins, containing two c-type haems, and are found in the periplasm. Both eukaryotic and prokaryotic CCPs play a role in resistance to H2O2 (Minard and McAlister-Henn, 2001; Seib et al., 2004). However, in S. cerevisiae mutants lacking CCP, cell viability and growth were not signiﬁcantly affected, indicating some redundancy with other peroxidatic enzymes (Kwon et al., 2003). Bacterial CCPs (bCCPs) use mono-haem cytochrome c as an electron donor that has been reduced by the respiratory chain. Therefore, CCPs also allow the use of H2O2 as a terminal electron acceptor. bCCP are thought to defend cells against exogenous sources of peroxide stress, protecting membrane lipids and proteins (Minard and McAlisterHenn, 2001). As described above, cells have many other enzymes to defend against exogenous and endogenous peroxides, such as catalase, AhpC, Gpx, Tpx and Bcp. Not all of these enzymes necessarily occur in the same bacterium, but this plethora of peroxidatic enzymes might explain why cells lacking CCPs are not totally killed by addition of peroxide. However, it also poses the question of why cells need to detoxify peroxides in both the periplasm and cytoplasm. CCPs are not universally present in bacteria, can be dispensed under certain conditions, and are not always required for resistance to either H2O2 or organic peroxides. These features suggest some special reasons why CCPs may be a feature of bacterial physiology.

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2. PHYLOGENETIC ANALYSIS OF BACTERIAL CCPS REVEALS A NOVEL SUB-GROUP OF TRI-HAEM PROTEINS Bacteria where CCPs have been studied biochemically or physiologically include Pseudomonas aeruginosa (Ellfolk and Soininen, 1970; Fulop et al., 2001), Rhodobacter capsulatus (Hanlon et al., 1992), Paracoccus pantotrophus (Goodhew et al., 1990), Nitrosomonas europaea (Arciero and Hooper, 1994), Methylococcus capsulatus (Zahn et al., 1997), Pseudomonas stutzeri (Villalain et al., 1984), Pseudomonas nautica (Alves et al., 1999), Paracoccus denitrificans (Gilmour et al., 1993) and Neisseria gonorrhoea (Turner et al., 2003). Genome sequencing shows CCPs to be present in many (but not all) Gram-negative bacteria, but they appear to be generally absent from those Gram-positive bacteria and archaea that have been sequenced to date. This may relate to the absence of a periplasm in the latter organisms, but there are several examples of other periplasmic redox proteins, which do have counterparts across bacterial and archaeal groups. Among Gram-negative bacteria, there is no particular correlation with metabolic lifestyle or type of organism, and a wide variety of both free-living and pathogenic bacteria contain ccp genes. Figure 1 shows a phylogenetic analysis of a selection of biochemically characterised and putative CCPs, after multiple sequence alignment in CLUSTAL X (Thompson et al., 1997) and analysis by the PHYLIP suite of programmes (Felsenstein, 1989). Of those CCPs characterised to date, the P. aeruginosa, R. capsulatus, P. pantotrophus and P. denitrificans enzymes are known to share mechanistic similarities. In the archetypal P. aeruginosa enzyme, which has been studied in much detail at the biochemical and structural level (Fulop et al., 1995, 2001), the protein must undergo a conformational rearrangement to become catalytically active. This occurs only after reduction by cytochrome c, and involves movement of a ﬂexible loop from the peroxidatic site, changing the co-ordination state of the haem from six ligands to ﬁve ligands, which allows entry of peroxide (see Fulop et al., 2001; Section 4.3.2. for a detailed explanation). The P. aeruginosa and mechanistically similar enzymes form a closely related phylogenetic group (Fig. 1). All CCPs in this group are di-haem enzymes. On the other hand, the CCP from N. europaea is active even in the fully oxidised state – the ﬂexible loop that needs to reorientate in P. aeruginosa type CCPs is always in the ‘out’ conformation, the haem is always penta-coordinate, and hence the enzyme is always able to accept its substrate (Arciero and Hooper, 1994). This enzyme is part of a small but distinct phylogenetic group, including the enzymes from Vibrio cholerae and V. vulnificus, and, interestingly, from Pseudomonas fluorescens. Whether the latter CCP is mechanistically similar

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Figure 1 Phylogenetic tree showing the relationship between the bacterial CCPs and the MauG proteins. Trees were produced by aligning sequences in CLUSTAL X (Thompson et al., 1997), with the output ﬁle used in PHYLIP (Felsenstein, 1989) to produce a distance matrix tree, which was viewed in TREEVIEW (Page, 1996). The CCPs that have been biochemically characterised are underlined.

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to the N. europaea enzyme, or it is like the CCP from its close relative P. aeruginosa, is yet to be elucidated. It would be interesting if two bacteria of the same genus have evolved two mechanistically distinct CCPs. Again, the N. europaea-type enzymes all contain two c-type haems. A third major group is also present in Fig. 1, containing enzymes from E. coli and related enterobacteria, as well as bacteria from a wide variety of other genera. These enzymes are all distinct from the well-studied CCPs in that they contain three haem-binding motifs, not two as in the other CCP families. The extra haem-binding site is located in a domain at the N-terminus of the protein which is absent in the di-haem CCPs. This is illustrated by the sequence alignment in Fig. 2. The exact role of this third haem domain is unclear, although it can be postulated that the extra haem is acting as an electron donor to the conventional CCP part of the enzyme, taking the place of the soluble mono-haem protein, which is the usual type of electron donor to bCCPs. It is possible that this extra portion of protein evolved through fusion of a mono-haem cytochrome c gene to the ccp gene in these organisms, although BLAST sequence analysis has shown that the N-terminus of these proteins is unique to the tri-haem-containing CCPs, and not signiﬁcantly similar to current mono-haem cytochrome cs. The remainder of the sequence of tri-haem CCPs has homology to the full-length di-haem CCPs, which lends weight to the possibility that this extra haem domain has been added on at some point during the evolution of this subfamily of CCPs. It should be noted that the tri-haem CCP from Haemophilus ducreyi contains the unusual XXXCH haem-binding motif (Fig. 2) in its N-terminal haem-binding domain (all the others contain the conventional CXXCH). This XXXCH motif has been found in c-type cytochromes from eukaryotic protozoa (Allen et al., 2005), but has also recently been identiﬁed in some prokaryotic c-type cytochromes (Hartshorne et al., 2006), where a dedicated haem lyase is required to attach the haem. Although most bacteria possess a single CCP, C. jejuni, a major foodborne pathogen (Butzler and Skirrow 1979; Kelly, 2001) contains two putative ccp genes (Parkhill et al., 2000; Myers and Kelly, 2005). These are encoded by Cj0358 and Cj0020c. Cj0358 appears to be a typical CCP, which clusters with other similar enzymes from the bacteria in the epsilon group of the proteobacteria (e.g. H. pylori). These enzymes form a distinct cluster but are distantly related to the P. aeruginosa type of CCP. However, Cj0020 appears not to be a closely related paralogue of Cj0358, and is not contained within any of the four major groups apparent in Fig. 1, although it does appear to be distantly related to the tri-haem enzymes from Zymomonas mobilis and Gluconobacter oxydans, even though Cj0020c contains only two haem-binding motifs.

82 JOHN M. ATACK AND DAVID J. KELLY

Figure 2 Sequence alignment of the N-terminus of the tri-haem-containing CCPs. The E. coli sequence was used in a BLAST search to extract other tri-haem CCPs from the databases, and a multiple sequence alignment generated with CLUSTAL X. The additional haem-binding motif is highlighted. Areas of homology are shaded. The aligned N-termini of the di-haem CCPs are shown on the right of the ﬁgure.

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3. MauG PROTEINS A fourth group is also present in Fig. 1; the MauG proteins. These are related to bCCPs in sequence and structure, but not in role. MauG proteins are required for methylotrophy and have around 30% sequence similarity to bCCPs (Chistoserdov et al., 1994a,b; Gak et al., 1995). Like bCCPs they are di-haem proteins, but the haem-spectral proﬁle of reduced MauG proteins is more typical of haems that bind oxygen than of typical c-type haems (Wang et al., 2005b). MauG is one of the four proteins needed for biosynthesis of methylamine dehydrogenase (MADH), the enzyme needed to utilise methylamine. Mutants lacking MauG are unable to form functional MADH (van der Palen et al., 1995; Wang et al., 2005b). MauG itself is involved in the generation of the tryptophan tryptophylquinone (TTQ) co-factor used by MADH. This consists of two covalently linked tryptophan residues, one of which contains two extra oxygen atoms (Wang et al., 2003). MauG forms TTQ by binding and activating oxygen, and linking two tryptophan residues of MADH to form TTQ (Wang et al., 2003). Speciﬁcally, MauG is needed for incorporation of a second oxygen atom onto one of the tryptophan residues to be modiﬁed in MADH, and the joining of the two tryptophan residues to form TTQ (Pearson et al., 2004; Wang et al., 2005b). In mutants lacking MauG, a biosynthetic intermediate accumulates that contains tryptophan with only one incorporated oxygen atom and no covalent link to the second tryptophan (Wang et al., 2005b). The ability of c-type haems to bind oxygen is unusual, and when taken together with the low peroxidase activity of the MauG proteins (Wang et al., 2003), suggests that they perform an unrelated function to CCPs. It is likely that MauG proteins and CCPs diverged from a common ancestral gene (Hu et al., 1997) if located in the same species. This divergence is evident in Fig. 1. This ﬁgure also shows that the MauG group of proteins are themselves highly divergent from one another, and from the CCPs. The reason for this is unclear. Interestingly, the analysis showed one MauG, that from Mannheimia succiniciproducens, to be much more similar to true CCPs than other MauG proteins. Perhaps this should be re-classed as a putative CCP rather than a MauG protein. Due to their divergence from bCCPs and their low peroxidase activity, MauG proteins can be viewed as oxygenases rather than peroxidases.

4. STRUCTURE OF BACTERIAL CCPS Bacterial CCPs are two-domain proteins, encoded by a single polypeptide. Each domain contains a single c-type haem – the high potential (hp) haem at

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the C-terminal domain and the low potential (lp) haem at the N-terminal domain. Each domain has a structure similar to the classical c-type cytochromes, but contains extra secondary structure elements (Moore and Pettigrew, 1990; Fulop et al., 1995). The P. aeruginosa enzyme is the best studied of bCCPs with a 2.4 A˚ crystal structure currently available (Fulop et al., 1995). The N. europaea enzyme was crystallised with diffraction to 1.8 A˚ (Shimizu et al., 2001). The P. denitrificans enzyme has also been crystallised (Echalier et al.., 2004), as has the CCP from P. pantotrophus (Echalier et al., 2006). The molecular weight of bCCPs ranges from 35–40 kDa depending on species. The two domains are related by a pseudo two-fold axis, with the interface between the two being hydrophobic. Interestingly, the domain interface holds a calcium ion (Ca2+) binding site (Fulop et al., 1995; Shimizu et al., 2001). The CCP from P. nautica has been crystallised in two conformations (Dias et al., 2004), although this was done at low non-physiological pH, and the protein cannot be physiologically reduced. This implies that these conformations are not relevant forms of this enzyme. The only enzyme where the inactive and active forms of the enzyme have been crystallised under physiological conditions is that from P. pantotrophus (Echalier et al., 2006). The inactive, fully oxidised form is most similar to the structure of the CCP from P. aeruginosa. The half-reduced active form of the enzyme resembles the enzyme from N. europaea, which is always active, even in the oxidised form (Arciero and Hooper, 1994; Echalier et al., 2006). This is consistent with structural rearrangements expected to occur upon the switch from oxidised inactive, to half-reduced active states. This is discussed in detail later in the review.

4.1. Dimer Formation bCCPs exist as homodimers in solution (Arciero and Hooper, 1994; Gilmour et al., 1994; Fulop et al., 1995; Zahn et al., 1997; Alves et al., 1999; De Smet et al., 2001). Like the domain interface in the monomer, the dimer interface is also hydrophobic (Fulop et al., 1995). Additional to this in the N. europaea enzyme, ﬂexible loops containing Histidine 59 (His59 loops) from each subunit form inter-subunit interactions at the dimer interface (Shimizu et al., 2001). The equivalent loops in the P. aeruginosa CCP dimer (His71 loops) occur only after they have reorientated from the ‘in’ to the ‘out’ conformation, giving an interface similar to the N. europaea dimer (Shimizu et al., 2001). Dimer formation may be essential for activity as the CCP from P. denitrificans loses activity upon dilution – an equilibrium may exist between

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the monomer and the dimer, with the dimer being the active form (Hu et al., 1997).

4.2. Calcium-binding Sites Bacterial CCPs have two different Ca2+-binding sites (Gilmour et al., 1995). Site I, which exists in the monomer at the domain interface, is high afﬁnity and is always occupied by Ca2+ (Fulop et al., 1995; Gilmour et al., 1995). Site II is a low-afﬁnity Ca2+-binding site, formed by dimerisation – each monomer contributes half the residues needed to form this site. In the CCP from P. pantotrophus this involves the motif GLGGVDGL (residues 61–68) from each monomer, which form a complete Ca2+-binding site at the dimer interface (Gilmour et al., 1995). This is also proposed for the P. denitrificans CCP (Hu et al., 1997). It is currently unclear as to whether binding of Ca2+ at site II is needed for dimer formation, as the R. capsulatus enzyme is able to form dimers in the presence of the metal ion chelater EDTA (De Smet et al., 2001). However, in P. pantotrophus, enzyme treated with metal ion chelater was monomeric and inactive (Gilmour et al., 1994; Pettigrew et al., 2003b) indicating Ca2+ is essential for dimerisation and activity in this enzyme at least. Thus, whether Ca2+ induces dimerisation or binds after dimerisation is uncertain. Ca2+ binding at site II is needed to switch the lp haem from the inactive low-spin state to the active high-spin state in the P. denitrificans CCP (Gilmour et al., 1993, 1994, 1995), but is not needed to induce this low- to high-spin state switch in the P. stutzeri enzyme (Timoteo et al., 2003). Ca2+ is essential for the reduction and hence activation of the CCP from P. pantotrophus (Prazeres et al., 1993). It is unclear why some CCPs require Ca2+ for activation whereas others do not. In P. nautica, Ca2+ is reported to be needed to induce the ‘out’ conformation around the lp haem (Dias et al., 2004), although disruption of the calcium-binding site by the acidic conditions used to crystallise the enzyme leaves this conclusion open to debate. In the P. aeruginosa, P. nautica and R. capsulatus enzymes, the positive charge of the calcium ion is not balanced by any negatively charged amino acids, signifying that it may modulate electron transport between the hp and lp haems (Fulop et al., 1995; De Smet et al., 2001, 2006; Dias et al., 2004).

4.3. Haem Sites in Bacterial CCPs As already mentioned, bacterial CCPs possess two haem sites – the hp site, where electrons from cytochrome c are donated, and the lp site, the so-called

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peroxidatic site, where H2O2 is reduced to water. Both haems are bound in hydrophobic environments. 4.3.1. High Potential Haem Site The hp haem site accepts electrons from cytochrome c, and therefore has a high positive midpoint redox potential. The exact value varies between bacterial species; for example, P. aeruginosa CCP hp site is +320 mV (Ellfolk et al., 1983), R. capsulatus +270 mV (De Smet et al., 2001), and N. europaea +130 mV (Shimizu et al., 2001). After accepting electrons from cytochrome c, the hp haem donates them to the lp haem. In many CCPs, the hp haem must be in the reduced Fe2+ form for the enzyme to be functional, such as in P. aeruginosa and R. capsulatus (Fulop et al., 1995; De Smet et al., 2001). However, this is not the case in N. europaea CCP, which is active even when fully oxidised (Shimizu et al., 2001). In the enzymes studied, the hp haem is present in the C-terminal domain – in P. aeruginosa residues 165–302 (Fulop et al., 1995). As in the majority of c-type cytochromes, the haem group is histidine and methionine ligated, and is covalently bound to two cysteine residues by thioether bonds. In P. aeruginosa, the His and Met ligands are His201 and Met275, respectively (Fulop et al., 1995). The equivalent residues in N. europaea are His187 and Met258 (Shimizu et al., 2001). In bacterial CCPs, the hp haem replaces the need to generate the tryptophan free radical that is used as an electron store in eukaryotic CCPs (Sivaraja et al., 1989). 4.3.2. Low Potential Haem Site As the lp haem is the site of peroxide reduction, it is known as the peroxidatic site. Like the hp site, the exact redox potential of the lp haem varies between species. For example, the P. aeruginosa CCP lp haem has a redox potential of 330 mV (Ellfolk et al., 1983) – a difference of 650 mV compared to the hp haem. The R. capsulatus enzyme lp haem redox potential varies from 190 to 310 mV (De Smet et al., 2001) – again, a large potential difference. However, the N. europaea CCP lp haem has a redox potential of +70 mV – a difference of just 60 mV (Shimizu et al., 2001). Like the hp haem, the lp haem is anchored to the protein via covalent thioether links to two conserved cysteine residues, but is coordinated to two histidine residues, not a histidine and a methionine. In the P. aeruginosa CCP, these two histidines are His55 and His71 (Fulop et al., 1995). The lp haem is coordinated to the N-terminal domain, which corresponds to residues 17–164 in the P. aeruginosa CCP (Fulop et al., 1995). In the P. aeruginosa,

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R. capsulatus and all other enzymes studied, with the exception of the N. europaea enzyme, the lp haem site must undergo a conformational change to allow H2O2 binding. A ﬂexible loop containing His71 (P. aeruginosa numbering) moves from the ‘in’ conformation to the ‘out’ conformation allowing the enzyme to bind to H2O2 and become catalytically competent (Fulop et al., 1995). His55 was initially thought to reorientate, but cannot as it is buried next to one of the cysteines that is covalently bound to the haem (Fulop et al., 1995). This reorientation of the lp haem site probably also involves movement of other residues in the active site as well (Fulop et al., 1995). Rebinding of the shedded His71 ligand – the distal histidine – is hypothesised to be slow (Fulop et al., 1995; De Smet et al., 2001). However, the N. europaea enzyme does not need to undergo this conformational change as its lp haem is pentacoordinate – it is only associated with a single histidine ligand. The second histidine (His59, equivalent to His71 in P. aeruginosa) is always in the ‘out’ conformation, so there is no need to undergo this structural rearrangement to become active. Thus, in the N. europaea CCP, the lp haem site is always open to H2O2, and therefore the enzyme is always active. The conformational change needed to activate the P. aeruginosa and R. capsulatus CCPs is redox linked – that is, the His71 loop can move to the ‘out’ conformation only after the hp haem site has been reduced. Hence, the P. aeruginosa and R. capsulatus enzymes are active only when then hp site is in the Fe2+ state (Shimizu et al., 2001). Re-orientation of this loop causes a low- to high-spin state switch in the lp haem that is necessary for activity. It is thought that the signal is transmitted between the haems via a b-sheet motif that is present in the P. aeruginosa enzyme, but not the N. europaea CCP (Shimizu et al., 2001). The recent publication of the structure of the P. pantotrophus enzyme in the fully oxidised and halfreduced states by Echalier et al. (2006) has shed some light on how the signal may be transmitted between the hp and lp haem domains. Reduction of the hp haem leads to the reorientation of a proprionate group and rotation of a histidine (His275 in P. pantotrophus CCP) that results in reorientation of a ﬂexible loop (residues 225–257) from the hp haem site to the interface between the hp and lp haem domains. The arrival of the 225–257 loop at the domain interface causes movement of another ﬂexible loop (residues 105–132) away from the domain interface to the lp haem site. This loop contains an arginine residue (Arg111) that is conserved in all bCCPs, with the exception of N. europaea, where the residue is an isoleucine. This arginine interacts with a proprionate at the lp haem. The arrival of the 105–132 loop is thought to trigger the shedding of the distal histidine around the lp haem, and the reorientation of the His loop to the ‘out’ conformation, and activation of the enzyme (Echalier et al., 2006).

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5. MECHANISTIC ASPECTS OF CATALYSIS BY BACTERIAL CCPS Eukaryotic mono-haem CCPs and di-haem bCCPs differ in the mechanism by which they reduce H2O2 to water, by the way in which oxidising equivalents are stored (Shimizu et al., 2001), and in the type of haem they utilise. Eukaryotic CCPs contain a single non-covalently bound b-type haem and carry out their chemistry in three steps (Sivaraja et al., 1989), generating a tryptophan free radical. In the ﬁrst step, H2O2 reacts with the enzyme to form compound I, which contains a tryptophan free radical and an oxyferryl center. Then, compound I is reduced to compound II by an electron from cytochrome c that is passed to the trp free radical. Finally, a second cytochrome c passes its electron to compound II, reducing the enzyme back to the ground Fe3+ state and forming water (Shimizu et al., 2001): (i)

Fe3þ Trp þ H2 O2 ! Fe4þ O Trpdþ þH2 O

(ii)

Fe4þ O Trpdþ þreduced cyt c ðFe2þ Þ ! Fe4þ O Trp

ðcompound IÞ ðcompound IÞ

(iii)

ðcompound IIÞ

þoxidised cyt c ðFe3þ Þ 4þ 2þ Fe O Trp þreduced cyt c ðFe Þ ! Fe3þ Trp ðcompound IIÞ

þoxidised cyt c ðFe3þ Þ þ H2 O

The bacterial CCPs all contain two covalently bound c-type haem groups, as opposed to a single b-type haem in the eukaryotic enzymes. As already mentioned, these two haem groups differ in their chemistry allowing reduction of H2O2 to water without the need to generate a free radical intermediate. Figure 3 shows a generalised bCCP mechanism (adapted from Greenwood et al., 1988; Fulop et al., 1995; Shimizu et al., 2001; Bradley et al., 2004). First, the hp haem must be reduced to the Fe2+ state by cytochrome c. The active form of the enzyme then reacts with H2O2 to generate compound I, which contains an [Fe4+QO] group at the lp haem, and produces water. Unlike the eukaryotic CCP, no free radical is formed. Reduction of compound I to II yields an [Fe3+–OH] group at the lp haem, which is subsequently used to form water. At this point both haems are in the Fe3+ state, meaning the hp site must be reduced back to Fe2+ for the enzyme to be active again. Thus, in most bacterial CCPs, the hp haem serves to store electrons before donating them to the lp haem for use in H2O2 reduction. In the N. europaea enzyme, the initial reduction of the hp haem is not needed for enzyme activity (Arciero and Hooper, 1994; Shimizu et al., 2001;

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hp cyt-c

3+

Fe

89

lp Fe3+

(inactive)

Fe2+ cyt-c Fe3+

hp

lp

Fe2+

Fe3+

(active)

H2O2 hp

lp

Fe3+

Fe4+=O

compound I

+

H2O H+ + e-

hp

lp

Fe3+

Fe3+-OH

compound II

H+ H2O

hp

lp

Fe3+

Fe3+

(inactive)

Next cycle Figure 3 Generalised reaction mechanism for the ‘‘classical’’ (Ps. aeruginosa type) bacterial CCP. First, the hp haem must be reduced to the Fe2+ state by cytochrome c. The active form of the enzyme then reacts with H2O2 to generate compound I, which contains an [Fe4+QO] group at the lp haem, and produces water. Unlike the eukaryotic CCP, no free radical is formed. Reduction of the compound I to compound II yields an [Fe3+–OH] group at the lp haem, which is subsequently used to form water. At this point both haems are in the oxidised Fe3+ state, meaning the hp site must be reduced back to Fe2+ for the enzyme to be active again.

Bradley et al., 2004). Also, an extra group, possibly a porphyrin radical (termed R in Fig. 4), is involved in compound I formation (Bradley et al., 2004). Compound II in the N. europaea CCP contains the ferryl [Fe4+ ¼ O] at the lp haem and not the di-ferric Fe3+ at both hp and lp haems as seen in the conventional CCP cycle (Bradley et al., 2004). Figure 4 shows the scheme proposed for the N. europaea CCP by Bradley et al. (2004).

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hp 3+

Fe

lp Fe3+ H2O2

hp

lp

Fe3+ [Fe4+=O] R•+ compound I

+

H2O e-

hp 3+

Fe

lp [Fe4+=O]

R

compound II H+ + ehp

lp

Fe3+

Fe3+-OH H+ H2O

hp

lp

Fe3+

Fe3+

Next cycle

Figure 4 Reaction mechanism for the CCP from N. europaea. The enzyme is active even when the it is fully oxidised, unlike the majority of CCPs studied, where the hp haem must be reduced for the enzyme to be active.

6. ELECTRON DONORS AND ELECTRON TRANSPORT IN BACTERIAL CCPS The physiological electron donors to bCCPs are often small monohaem cytochrome cs. Other physiological electron donors can include azurin and pseudoazurin – small copper proteins found in some bacteria (Ronnberg et al., 1981; Pauleta et al., 2004a,b). In vitro, horse heart cytochrome c can also act as an artiﬁcial electron donor (De Smet et al., 2001; Pauleta et al., 2004b).

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6.1. Nature of Interactions with Electron Donors In P. denitrificans and P. aeruginosa, cytochrome c is thought to interact electrostatically with the CCP at a small binding patch on the enzyme’s surface (Crowley and Ubbink, 2003). This patch has a hydrophobic centre and a charged periphery. The small size of the interface site ensures that interactions are transient (high dissociation rate constant) yet desolvation (the dissociation of water and binding of the electron donor) of this patch favours complex formation (high association rate constant). Therefore, binding is transient but highly speciﬁc. The charged periphery of the binding patch is thought to enhance the association rate through electrostatic steering, but this does not contribute to the speciﬁcity of the interactions or improve electron transfer rates (Hart et al., 2003). In the R. capsulatus CCP, activity is highest at low ionic strength – activity drops off steeply as ionic strength increases (Koh et al., 2003). Mutational analysis on the R. capsulatus CCP electron donor, cytochrome c2, indicated that the interaction surface of the donor with the CCP involves both non-speciﬁc charge–charge interactions and salt bridges (Koh et al., 2003) – both electrostatic in nature and therefore dependent on ionic strength. The P. nautica enzyme, on the other hand, interacts with its electron donor in a hydrophobic manner (Alves et al., 1999). This was concluded from studies showing that the enzyme was active across a wide range of ionic strengths, indicating a hydrophobic and not an electrostatic interaction with cytochrome c. The interaction surface of the P. nautica CCP with cytochrome c is much more hydrophobic in character when compared to that in the P. aeruginosa enzyme (Alves et al., 1999). In those CCPs that interact with pseudoazurin, for example the P. pantotrophus CCP, the nature of the interaction is electrostatic (Pauleta et al., 2004a). The site of electron transfer from pseudoazurin is a His residue, surrounded by a hydrophobic patch, which is itself surrounded by a ring of Lys residues (Pauleta et al., 2004a). These lysines interact with acidic residues on the surface of the CCP, which is thought to prevent charge interactions forming with the CCP. The interaction surface of the R. capsulatus CCP electron donor, cytochrome c2, also contains several lysine residues, the loss of which disrupts interaction with the CCP (Koh et al., 2003). The studies by Pauleta et al., (2004b) also revealed that the CCP from P. pantotrophus is able to accommodate more than one electron donor at the same time, indicating a large capture surface for electron donors. Pseudoazurin binds as a dimer, and horse heart cytochrome c binds at two sites of differing afﬁnities. Studies on the CCP from P. denitrificans showed

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that it was able to bind two horse heart cytochrome cs (Pettigrew et al., 2003a, b). The binding of an ‘active’ and ‘waiting’ cyt-c in a so-called ternary complex may serve as a rate enhancement mechanism in vivo. It was also shown that pseudoazurin and cytochrome c bind at the same site (Pauleta et al., 2004b) – this could represent different binding sites for different electron donors, or that the binding sites overlap, again indicating a large capture surface for the electron donors. It is possible that the binding of a second ‘waiting’ electron donor promotes dissociation of the ‘active’ one (Mei et al., 2002). Structural studies on a complex of the P. denitrificans CCP and a single physiological electron donor, monohaem cytochrome c, showed that the haem of the donor lies above the hp haem site of the enzyme (Pettigrew et al., 1999). In comparison to this, two molecules of horse heart cytochrome c are able to bind between the two haems of the enzyme (Pettigrew et al., 1999). This again indicates that CCPs contain a large capture surface for electron donors.

6.2. Electron Transport between Haems The haem sites in bCCPs are over 10 A˚ apart, but electron transfer is possible between them through electron tunnelling (Fulop et al., 1995). Four possible routes for electron transport between the hp an lp haems of P. aeruginosa CCP have been proposed (Fulop et al., 1995), where electrons jump between groups: (i) (ii) (iii) (iv)

From His201 adjacent to the hp haem via the protein backbone (residues 246–248) to the lp haem propionate. From the hp haem propionate, via the calcium ion at site I and the protein backbone (residues 79–81) to reach His55 of the lp haem. His 261 connects the hp haem propionate with Asn79 to join the second path. From the hp haem propionate via Trp94 to the lp haem propionate.

It appears that His261 is critical for activity of the enzyme, as removal of the equivalent histidine, His275, in the P. denitrificans enzyme completely abolishes activity (McGinnity et al., 1996). Thus, route (iii) appears to be the most likely route of electron transfer, although multiple pathways should not be ruled out. Trp94 is found in the hydrophobic cavity between the hp and lp haem domains, and may also be a possible electron transport route as it lies in the same plane as hp haem and perpendicular to the lp haem and is able to interact with both (Fulop et al., 1995).

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7. ROLES OF CCPS IN BACTERIAL CELLS Although structurally and biochemically well characterised, the exact role of CCPs in bacterial physiology is often not obvious. The fact that these enzymes are not present in all bacterial cells, but rather are restricted to certain Gram-negative species, indicates that the known alternative enzymic strategies for the removal of H2O2 are sufﬁcient for viability in many cases. Nevertheless, in some bacteria a convincing case can be made for a distinct function.

7.1. Detoxification of Periplasmic Hydrogen Peroxide When cells possess catalase (and in many cases AhpC, Tpx and Bcp), which can remove H2O2 from the cytoplasm, the obvious conclusion to make is that CCPs have evolved to remove H2O2 from the periplasm. But in what situations is periplasmic H2O2 a particular problem? Cells are commonly exposed to exogenous peroxides, which can damage periplasmic or outer membrane components as, for example, would be the case in pathogens encountering the oxidative burst in macrophages or during tissue inﬂammation. The relative importance of CCP versus catalase has been investigated in pathogenic Neisseria species. In N. gonorrhoea, single mutants that lacked CCP were not signiﬁcantly more sensitive to H2O2 than the wild-type parent (Turner et al., 2003), whereas single-catalase mutants showed a very signiﬁcant increase in growth sensitivity to H2O2 in disc diffusion assays. However, a double mutant lacking both CCP and catalase was more sensitive than the catalase single mutant alone (Turner et al., 2003). Although this indicates a speciﬁc physiological role for CCP in peroxide detoxiﬁcation, the results suggest that H2O2 is more damaging in the cytoplasm than in the periplasm, and that catalase is able to compensate for the loss of the CCP to some extent. Indeed, cells that lack catalase are known to show large-scale DNA damage after exposure to H2O2 (Johnson et al., 1993). Interestingly, N. meningitidis lacks a CCP, but it is still an effective pathogen, indicating that CCPs are not always an important part of a pathogens defence against H2O2, and it is clear that these two Neisseria species have evolved different strategies for oxidative stress defence related to their individual tissue tropism and niches (Seib et al., 2004). The N. gonorrhoea CCP is also unusual in that it is the only bacterial CCP studied to date that has been convincingly demonstrated to be a membrane-anchored lipoprotein (Turner et al., 2003) – all the others are free periplasmic enzymes. It is

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unclear whether the enzyme is associated with the outer or inner membrane, or why it is membrane associated, but one possibility is that being a lipoprotein keeps the enzyme in closer proximity with its membrane-associated c-type cytochrome electron donor or possibly nearer the bacterial surface where it may be important in pathogenicity-related peroxide detoxiﬁcation (Turner et al., 2003). The analysis carried out by Turner et al. (2003) showed possible cleavage sites for the lipoprotein-speciﬁc signal peptidase II in several other known and putative CCPs, including the H. pylori, E. coli and Pseudomonas enzymes, with no particular correlation as to pathogenic status of the organism or mechanistic class of the enzyme. Thus, the general signiﬁcance of the lipid modiﬁcation and membrane anchoring is not clear. Another explanation for the existence of periplasmic CCPs in some bacteria is to remove H2O2 generated within the periplasm itself. This latter situation was postulated to be the case in the veterinary pathogen C. mucosalis, which was proposed to generate H2O2 in the periplasm as a side reaction of the formate dehydrogenase (FDH) when using formate as an electron donor to its respiratory chain (Goodhew et al., 1988). However, C. mucosalis lacks catalase, meaning that CCP may be the only or major way of removing H2O2 from the cell in this organism. Nevertheless, the use of CCP to remove FDHgenerated peroxide in other formate-oxidising Campylobacter species, which are catalase positive, may prove to be physiologically important.

7.2. Hydrogen Peroxide as an Electron Acceptor An additional aspect of the mechanism of bacterial CCPs that is distinct from, for example, cytoplasmic thiol peroxidases is that they can act as terminal reductases of an electron transport chain, where H2O2 would be the terminal electron acceptor. Proton translocation and energy conservation may thus accompany electron transport from a primary electron donor to H2O2, although the periplasmic location of the peroxidase means that the enzyme itself is not energy conserving. Rather, the reduction of H2O2 would allow proton translocation to occur at the level of the primary dehydrogenase and also through the operation of the cytochrome bc1 complex (if present), which is often the immediate electron donor to soluble periplasmic cytochrome c, from which electrons ﬂow to the CCP, as described above. The importance of the energy conserving function of CCPs has not been widely investigated, but Goodhew et al. (1988), working with the catalase-negative microaerophile C. mucosalis, were able to show that addition of H2O2 resulted in uncoupler-sensitive proton extrusion in classical proton pulse-type experiments, albeit with a low stoichiometry (about 0.6 H+/H2O2). It was

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suggested that this may be important in the bioenergetics of the bacterium during growth on formate where, as noted above, periplasmic H2O2 may be generated. The role of CCP as a functional component of the bacterial respiratory chain in intact cells was ﬁrst demonstrated by Richardson and Ferguson (1995) in studies to explain the almost complete inhibition of nitrate reduction by H2O2 in cells of Thiosphaera pantotropha (now P. pantotrophus) and R. capsulatus. The inhibition had previously been ascribed to oxygen, produced as a result of the action of catalase, inhibiting nitrate reduction, but it was shown that oxygen itself did not signiﬁcantly affect this process and that the inhibition arose because H2O2 competed with nitrate for electrons via the operation of a CCP. These bacteria are found in a variety of aerobic to anaerobic niches in soils, water and other environmental situations where their capacity for nitrate reduction or denitriﬁcation is a key element in the nitrogen cycle. It is therefore interesting that H2O2 producing bacteria have been identiﬁed (Arcobacter sp.) which may co-exist in the same environments (Kontchou and Blondeau, 1990), and which could potentially lead to inhibition of nitrate respiration but also provide a source of an alternative electron acceptor.

7.3. Alternative Substrates and Multiple Enzymes It has been generally assumed that all CCPs are speciﬁc for H2O2, but there is some limited indirect evidence that some of these enzymes may be able to handle other substrates, and it is possible that this may be physiologically important. This evidence chieﬂy comes from Bacteroides fragilis, where mutants lacking CCP were signiﬁcantly more sensitive to organic peroxide stress generated by cumene hydroperoxide or t-butyl hydroperoxide, but not to that generated by H2O2 (Herren et al., 2003). However, nothing is known about the substrate speciﬁcity of the CCP enzyme in this bacterium so it is not yet possible to correlate this with the mutant data. Most bacteria appear to contain a single gene encoding a CCP, but there are some clear examples where two or more ccp genes appear to be present in a given genome. As noted above, analysis of the C. jejuni genome reveals two separate genes encoding putative CCPs homologues (Parkhill et al., 2000; Myers and Kelly, 2005). Cj0020 and Cj0358 are 34 and 37 kDa proteins, respectively, with both proteins containing two cytochrome c haembinding site signatures. However, as can be seen in the phylogenetic tree in Fig. 1, the two proteins are distinct in that Cj0020 appears to form a separate clade, while Cj0358 is more like the P. aeruginosa enzyme which has an

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activation step as described previously in Section 4. Mutants in the Cj0020c gene are unable to effectively colonise chicks (Hendrixson and DiRita, 2004), indicating that Cj0020 plays an important role in colonisation. As there is no good animal model for disease for C. jejuni, it is not yet known if this putative CCP also plays a role in virulence. The role of the second CCP in C. jejuni, Cj0358, has not yet been studied, although some data on its regulation are available (see below). Methylococcus capsulatus strain Bath is a methanotrophic bacterium, which appears to contain multiple ccp-like genes. One CCP has been the subject of a detailed biochemical characterisation (Zahn et al.., 1997) and shown to be a typical CCP with a subunit mass of 35.8 kDa. Recently, an unusual CCP-like protein which is much larger (78 kDa) has been discovered in the same bacterium (Karlsen et al., 2005). This protein is copper repressible and appears to be localised to the cell surface, facing the external environment, rather than located in the periplasm. The deduced sequence shows the presence of two typical haem-binding motifs, albeit spaced much further apart due to the longer length of the protein, and also conservation of the amino acids that are involved in calcium binding in bona fide CCPs. Haem staining on gels conﬁrmed the presence of c-type haem. Sequence comparisons revealed several other similar hypothetical proteins in genome databases for Photobacterium profundum, P. fluorescens and Nostoc punctiforme. Structural modelling strongly suggested that this novel protein and its homologues are related to the bCCP family as a distinct sub-group (Karlsen et al., 2005). However, peroxidase activity has not yet been conﬁrmed for this protein and it is difﬁcult to suggest what its physiological role might be. It is also not clear how electrons would be transferred across the outer membrane to reach the enzyme on the cell surface.

7.4. A Paradox in the Regulation of Expression of Bacterial CCPs? It is well known that hyperoxic conditions result in an increase in oxidative stress and it has been shown that exposure of bacterial cells to excess oxygen increases the rate of H2O2 production (Seaver and Imlay, 2004). Paradoxically however, bacterial CCPs appear to be upregulated preferentially under low oxygen (microaerobic) or anaerobic conditions. In a number of bacteria, including N. gonorrhoea, P. denitrificans and P. stutzeri, the global regulator FNR positively up-regulates the ccp gene at low oxygen tensions (Van Spanning et al., 1997; Vollack et al., 1999; Turner et al., 2003). FNR is a well-characterised global regulator of anaerobic gene expression in E. coli

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(Iuchi and Lin, 1991; Spiro and Guest, 1991). In P. aeruginosa, ccp expression is regulated by Anr, a homologue of FNR (Zimmermann et al., 1991). Like FNR, Anr also up-regulates CCP expression under low oxygen conditions. In B. fragilis, CCP is up-regulated by H2O2, but in a manner independent of its peroxide stress regulator OxyR (Herren et al., 2003). The method of CCP control in B. fragilis is yet to be determined. The CCP in E. coli appears to be regulated by both FNR and OxyR (J. Partridge and J. Green, personal communication). This is the ﬁrst time a CCP has been shown to be controlled by both anaerobic and peroxide stress response systems. How is this apparent paradox in ccp gene expression to be explained? In one sense, the pattern of regulation makes sense if H2O2 is considered purely as an alternative terminal electron acceptor. Many other alternative reductases, e.g. fumarate and nitrate reductases, are also positively up-regulated by FNR under anaerobic conditions, allowing respiration to continue with a variety of alternatives to oxygen. Nevertheless, H2O2 seems to be a special case as it is much more unstable than other potential electron acceptors and it seems unlikely that in many environments it would be present at higher concentrations under anaerobic conditions compared to aerobic conditions. As noted above, there may be some circumstances in free-living bacteria (during formate respiration or in the presence of peroxide-producing bacteria) where electron transport to H2O2 might be quantitatively important. However, the increased expression of CCPs under low oxygen conditions may be much more relevant in pathogenic bacteria, as a drop in oxygen levels can be a signal of entry into the host. Pathogens living in the gut or on mucosal surfaces may well be subject to microaerobic or severely oxygenlimited conditions, in addition to having to contend with the host immune response, which will invariably involve the attack of H2O2-producing phagocytes. Under these conditions, the upregulation of ccp genes would allow the deployment of a periplasmic ‘‘ﬁrst line of defence’’ against exogenous peroxide. If this were overwhelmed, then other cytoplasmic peroxidatic defence enzymes described above could be brought into play. Currently, there is little direct experimental evidence to support this scenario. Careful colonisation and pathogenicity studies are needed with single ccp mutants, and also double or triple mutants lacking catalase and/or thiol peroxidases, to dissect out the contribution made by each system. In fact more work has been done on pathogenic eukaryotes with regard to the role of CCPs in vivo. For example, fungal pathogens (such as Cryptococcus neoformans, the leading cause of fungal meningitis) which lack CCP are just as virulent as wild-type strains, but are more vulnerable to exogenous peroxide stress (Giles et al., 2005). The CCP from the parasitic trematode worm Fasciola hepatica has been shown to protect deoxyribose from damage by

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H2O2 in vitro (Campos et al., 1999) by preventing formation of the highly damaging hydroxyl radical (HO ). F. hepatica lacks catalase (Barrett, 1980), making CCP a major defence against H2O2 and, therefore, playing a key role in defending the organism from the host immune system. In addition to oxygen, ccp genes are also regulated by iron. In C. jejuni, Cj0358, one of the two CCPs in this organism, is negatively regulated under iron-limited conditions by the iron response regulator Fur, and its homologue PerR, the peroxide-response regulator (Holmes et al., 2005). Cj0358 is also regulated by a temperature-responsive regulator, RacR (Bras et al., 1999), indicating a complex regulatory pattern for the CCP in this organism, which may well be related to pathogenicity.

8. CONCLUDING REMARKS Despite the availability of a thorough understanding of the structure and mechanisms of bacterial CCPs, the true physiological roles of these enzymes are clearly much less apparent. A key common feature is their upregulation under microaerobic or anaerobic conditions, suggesting a role in peroxide detoxiﬁcation under conditions where electron transport to oxygen is limiting. CCPs may be especially important in some pathogenic bacteria but few in vivo studies with mutants have yet been carried out to suggest a function distinct from the many other possible peroxidatic enzymes often present in such bacteria. The roles of paralogous enzymes and newly identiﬁed novel types of CCP, for example the tri-haem CCPs, also require more detailed investigation.

ACKNOWLEDGEMENTS JMA is supported by a UK Biotechnology and Biological Sciences Research Council (BBSRC) doctoral training award. DJK gratefully acknowledges the BBSRC for ﬁnancial support of research in his laboratory.

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Respiratory Transformation of Nitrous Oxide (N2O) to Dinitrogen by Bacteria and Archaea Walter G. Zumft1 and Peter M.H. Kroneck2 1

Institute of Applied Biosciences, Division of Molecular Microbiology, University of Karlsruhe, D-76128 Karlsruhe, Germany 2 Faculty of Biology, University of Konstanz, D-78464 Konstanz, Germany

ABSTRACT N2O is a potent greenhouse gas and stratospheric reactant that has been steadily on the rise since the beginning of industrialization. It is an obligatory inorganic metabolite of denitrifying bacteria, and some production of N2O is also found in nitrifying and methanotrophic bacteria. We focus this review on the respiratory aspect of N2O transformation catalysed by the multicopper enzyme nitrous oxide reductase (N2OR) that provides the bacterial cell with an electron sink for anaerobic growth. Two types of Cu centres discovered in N2OR were both novel structures among the Cu proteins: the mixed-valent dinuclear CuA species at the electron entry site of the enzyme, and the tetranuclear CuZ centre as the ﬁrst catalytically active Cu–sulfur complex known. Several accessory proteins function as Cu chaperone and ABC transporter systems for the biogenesis of the catalytic centre. We describe here the paradigm of Z-type N2OR, whose characteristics have been studied in most detail in the genera Pseudomonas and Paracoccus. Sequenced bacterial genomes now provide an invaluable additional source of information. New strains harbouring nos genes and capability of N2O utilization are being uncovered. This reveals previously unknown

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relationships and allows pattern recognition and predictions. The core nos genes, nosZDFYL, share a common phylogeny. Most principal taxonomic lineages follow the same biochemical and genetic pattern and share the Z-type enzyme. A modiﬁed N2OR is found in Wolinella succinogenes, and circumstantial evidence also indicates for certain Archaea another type of N2OR. The current picture supports the view of evolution of N2O respiration prior to the separation of the domains Bacteria and Archaea. Lateral nos gene transfer from an e-proteobacterium as donor is suggested for Magnetospirillum magnetotacticum and Dechloromonas aromatica. In a few cases, nos gene clusters are plasmid borne. Inorganic N2O metabolism is associated with a diversity of physiological traits and biochemically challenging metabolic modes or habitats, including halorespiration, diazotrophy, symbiosis, pathogenicity, psychrophily, thermophily, extreme halophily and the marine habitat down to the greatest depth. Components for N2O respiration cover topologically the periplasm and the inner and outer membranes. The Sec and Tat translocons share the task of exporting Nos components to their functional sites. Electron donation to N2OR follows pathways with modiﬁcations depending on the host organism. A short chronology of the ﬁeld is also presented. Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Chemistry of N2O . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Chemical Reduction of N2O . . . . . . . . . . . . . . . . . . . . . . . . 2.2. N2O–Transition Metal Complexes. . . . . . . . . . . . . . . . . . . . 3. Genomic and Organismal Resources . . . . . . . . . . . . . . . . . . . . . 3.1. Formation and Consumption of N2O by Bacteria . . . . . . . . . 3.2. Genomics and N2O Respiration . . . . . . . . . . . . . . . . . . . . . 4. Properties of N2O Reductase. . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Finding the Enzyme by a Classical Microbiological Strategy . 4.2. Z-type N2O Reductase . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Structure of N2O Reductase . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1. Crystal Structure and Subunit Interactions . . . . . . . . . . . . . 5.2. Surface Charge . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6. Novel Cu Centres in N2O Reductase . . . . . . . . . . . . . . . . . . . . . 6.1. CuA Centre. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2. CuZ Centre. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7. Organization of nos Genes, Gene Expression, Regulation . . . . . . 7.1. Patterns in nos Gene Clusters . . . . . . . . . . . . . . . . . . . . . . 7.2. Gene Expression and Regulation . . . . . . . . . . . . . . . . . . . . 8. Evolutionary Aspects and Phylogenetic Relationships . . . . . . . . . 8.1. Phylogenetic Relationships among Nos Proteins . . . . . . . . . 8.2. Relationship of N2O Reductase to Cytochrome Oxidase . . . 8.3. Inorganic Metabolism of N2O by Archaea . . . . . . . . . . . . . .

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109 110 112 112 114 114 114 120 127 127 128 131 133 135 136 136 147 152 152 154 157 157 163 163

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8.4. Gram-Positive Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.5. Plasmid-Encoded nos Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9. Topology and Transport Processes . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.1. The Tat System Exports NosZ and NosX to the Periplasm. . . . . . . 9.2. Sec Export-Dependent Components NosR, NosD, NosY and NosL 9.3. Lipoprotein Targeting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10. CU Centre Assembly . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.1. Supplying Cu: NosA, NosL and ScoP . . . . . . . . . . . . . . . . . . . . . . 10.2. ABC Transporter. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Electron Donation and Maintenance of Activity in vivo . . . . . . . . . . . . . . 11.1. Electron Transfer Components. . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2. Role of Flavoproteins NosR and NosX . . . . . . . . . . . . . . . . . . . . . 12. A Glimpse of History . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13. Conclusions and Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

167 167 168 168 173 174 175 175 177 180 181 184 194 196 197 197

ABBREVIATIONS ABC CD COX Dnr EPR EXAFS MCD NGC(s) N2OR(s) Nar Nir Nor Nos Tat

ATP binding cassette Circular dichroism Cytochrome c oxidase Denitriﬁcation regulator Electron paramagnetic resonance Extended X-ray absorption ﬁne structure Magnetic circular dichroism nos gene cluster(s) Nitrous oxide reductase(s) Gene designation for nitrate respiration Gene designation for nitrite respiration Gene designation for respiratory nitric oxide reduction Gene designation for nitrous oxide respiration Twin arginine translocation

When a thing is new, people say ‘It is not true’. Later, when its truth becomes obvious, they say ‘It’s not important’. Finally, when its importance cannot be denied, they say ‘Anyway, it’s not new’. William James (1842–1910) psychologist–writer known as the ‘nitrous oxide philosopher’, brother of Henry James

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1. INTRODUCTION In reviewing the geochemical fate of N2O, Kaplan and Wofsy (1985) summarized two decades ago that an increase in the concentration of N2O is expected to deplete stratospheric ozone levels and raise the temperature of the lower atmosphere. The radiative forcing of N2O is about 200 times that of CO2. Anthropogenic perturbations from agriculture, forest clearing and combustion of fossil fuel, causing a steady rise in the atmospheric N2O concentration to 314 ppbv currently, have intensiﬁed concerns since (Stein and Yung, 2003). The atmospheric concentration of N2O is growing exponentially in substantial part due to the increased use of fertilizers (Mosier et al., 1998). High input of nitrogen is stimulating the microbial communities underlying the biogeochemical N transformations. The rise in atmospheric N2O concentration shows that the prokaryotic N cycle is off balance, particularly in the terminal step that generates N2 from N2O (Fig. 1). N2O is an obligatory inorganic metabolite of the bacterial cell during denitriﬁca tion, a respiratory process which converts an ionic N oxide, NO 3 or NO2 , to the gaseous forms, NO, N2O or N2 (Zumft, 1997). Denitrifying bacteria constitute the principal group of N2O producers, with secondary roles by nitrifying and methanotrophic bacteria and by fungi. Among these groups, only the denitrifying prokaryotes have the ability to respire N2O to N2. We will address here the microbiological aspect of N2O metabolism connected to denitriﬁcation. The process is found among the three domains of life, Archaea, Bacteria and Eukarya, but is limited in the latter to certain fungi. There is no evidence that fungi would possess the capability of N2O reduction and therefore terminate nitrate denitriﬁcation with N2O. Conversion of N2O into N2 is the last step of an unabridged denitriﬁcation pathway, and represents a respiratory process in its own right. Many denitrifying bacteria grow at the expense of N2O as the sole electron acceptor. We consider complete denitriﬁcation of modular structure of four respiratory processes that utilize nitrate, nitrite, nitric oxide or N2O, and are tied by regulatory means. N2O respiration sustains the entire bioenergetic needs of a bacterium when O2 is absent. No wonder, we ﬁnd the process in organisms living in a broad spectrum of habitats, ranging from subzero temperatures to the boiling point, from saturated salt solutions to a hydrostatic pressure of 110 MPa. Extreme habitats and N2O utilization are compatible and demonstrate that the underlying enzymatic apparatus has adapted to environmentally challenging conditions. N2O is formed in the denitrifying cell by respiratory NO metabolism which utilizes a distinct type of NO reductase (Zumft, 2005a).

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Figure 1 The biogeochemical nitrogen cycle drawn to emphasize the role of nitrous oxide as a greenhouse gas. Roman numerals give the formal oxidation state of the nitrogen atom.

Reduction of N2O is a strongly exergonic reaction: N2O+2H++2eN2+H2O [Eo0 (pH 7.0) ¼ +1.35 V; DGo0 ¼ 339.5 kJ mol1]. A dimeric multicopper enzyme that has been intensively studied for Pseudomonas spp. and Paracoccus spp. catalyses the reaction. Its ﬁrst representative was isolated from the ZoBell strain of Pseudomonas stutzeri. It will be referred to as the Z-type N2O reductase (N2OR) because of its CuZ catalytic site and overall head-to-tail structure that is symbolized by ‘Z’. The activity of N2OR serves as the electron sink for an anaerobic electron transport chain. Charge separation and energy conservation depend on coupling sites of the electron pathway towards N2O. From microbiological and biochemical viewpoints we wish to know which groups of bacteria metabolize N2O, what is the necessary genetic information and the biochemistry to handle it, how is N2O activated and reduced at the catalytic site, how is the process regulated and what are the factors that determine the N2O-transforming capability of a prokaryote? Inorganic N2O metabolism has been treated in the past as part of reviews on denitriﬁcation (Berks et al., 1995; Zumft, 1997). We will cover here the process as an independent respiratory mode and give a comprehensive description (in part including unpublished data analyses) of the current understanding of the underlying chemistry, biochemistry, molecular

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genetics and organismal variety that in all their diversity make up N2O respiration.

2. CHEMISTRY OF N2O N2O is a remarkably inert compound at room temperature and under pressure of one atmosphere (E0.101 MPa) as expected for a 16-electron triatomic species. Among the simpler N compounds it is second only to N2 in inertness, despite its high positive free energy of formation (DG1 ¼ +104.2 kJ mol1) and potent oxidizing power. When we disregard highenergy radicals and molecules formed by pulse radiolysis, only few compounds react homogeneously with N2O at room temperature, such as metal alkyls and amides (LiPh and NaNH2), or of transition metals the chlorides TiCl3 and ammoniacal CrCl2. TiCl3 reduces N2O to ammonia, whereas CrCl2 converts it to N2; both reactions proceed at very slow rates (Banks et al., 1967). Since Pratt and colleagues reported these data almost 40 years ago the chemistry and reactivity of dinitrogen complexes has advanced remarkably. By comparison, in spite of the importance of N2O as a constituent of the atmosphere, little progress has been made in the ﬁeld of transition–metal chemistry of N2O. N2O is a linear, asymmetrical molecule that is isoelectronic to CO2 or the cyanate anion, OCN. Its electronic and geometrical structure can be described by the valence bond resonance forms N N O

N N O

The NN (1.1282 A˚) and NO (1.1842 A˚) bonds are shorter than the respective average double bond values of 1.25 and 1.21 A˚ in agreement with the calculated bond orders of 2.73 (NN) and 1.61 (NO) (Jug, 1978). For a discussion of the structural and spectroscopic properties of N2O in the context of chemical reactivity, the reader is directed to the review by Trogler (1999).

2.1. Chemical Reduction of N2O N2O is thermodynamically unstable; above 600 1C it decomposes to N2 and O2 by ﬁssion of the weaker NO bond. The activation energy of this process

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is around 250 kJ mol1 (Jones, 1975). With dihydrogen, N2O reacts to yield N2 and H2O in a process that is more exothermic than the H2/O2 reaction (Melville, 1933). Cu catalyses the reaction between N2O and CO, which produces N2 and CO2 at 200–250 1C (Schwab and Drikos, 1940). Cu and cuprous oxide are effective catalysts of the cleavage of the NO bond of N2O (Dell et al., 1953a,b; Scholten and Kovalinka, 1969; Sankar et al., 1992). N2O reacts rapidly with a number of low-valent transition metal complexes under mild conditions; cobalt(I) complexes catalyse the reduction of N2O to N2 by borohydride, BH 4 (Banks et al., 1967, 1968; Pu et al., 1969): 2Co(I)vitaminB12+N2O-2 Co(II)vitaminB12+N2. Co(I), Ni(I), and Cu(I) complexes with unsaturated nitrogen macrocycles, which were produced by pulse radiolysis, react very rapidly with N2O to form N2 (Buxton et al., 1976; Tait et al., 1976a,b; Jubran et al., 1985). The Ni(I) complex of 1,4,8,11-tetraazacyclotetradecane (L) is oxidized to the corresponding Ni(III) species according to the stoichiometry: [Ni(I)L]++N2O+2 H3O+[Ni(III)L]3++N2+3 H2O. No evidence was obtained in those studies for the addition of N2O into the coordination sphere of the transition metal, but oxygen atom transfer from N2O into the coordination sphere of hafnium and zirconium compounds was reported (Vaughan et al., 1987, 1988). Thus, it appears that when N2O reacts with transition–metal complexes N2 is extruded and oxo species are formed, M+N2O-O¼M2++N2. These O-atom transfer reactions should proceed in a very clean fashion since the sole byproduct is N2. N2O is a substrate of the Mo-Fe enzyme nitrogenase which reduces it to N2 (Liang and Burris, 1988, 1989). Actually, N2O was the ﬁrst reported example of a substrate reduced by this enzyme other than N2 (Mozen and Burris, 1954; Jensen and Burris, 1986). N2 is competitive with N2O, and N2O is competitive with acetylene, HCCH. On the other hand, acetylene is noncompetitive with N2O (Liang and Burris, 1988). N2O also reacts with the multihaem enzyme cytochrome c nitrite reductase. This enzyme converts nitrite to ammonia in a 6-electron transfer reaction without the release of potential reaction intermediates, such as NO or NH2OH (Stach et al., 2000; Einsle et al., 2002). Whereas NO and NH2OH are reduced also to ammonia, the reaction product of N2O remains unknown. N2O was reported to reversibly inhibit O2 utilization by bovine heart and bean mitochondria (Sowa et al., 1987). In view of its use as a general anaesthetic, the interaction of N2O with cytochrome c oxidase (COX) from bovine heart was investigated (Einarsdo´ttir and Caughey, 1988; Chervin and Thibaud, 1992). N2O molecules were shown by infrared spectroscopy to occupy sites within the oxidase, but no evidence was found for N2O serving as ligand to a metal. Furthermore, N2O interacts with CO dehydrogenase (Lu and Ragsdale, 1991) and methionine synthase (Drummond and Matthews, 1994a,b).

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2.2. N2O–Transition Metal Complexes N2O is expected to act as a unidentate ligand, bonding either through nitrogen to form an essentially linear moiety MeNNO, or through oxygen with a bent MeONN group. As a further possibility, N2O could bind in a bidentate manner through N and O. Surprisingly, only one metal complex with N2O has been prepared to date. The Ru(II) complex, [Ru(NH3)5 (N2O)]2+X2 was ﬁrst reported as an unstable species in solution (Armor and Taube, 1969, 1970, 1971), but was later isolated with different counterions (Diamantis and Sparrow, 1970; Diamantis et al., 1975): [Ru(NH3)5 (OH2)]2++N2O-[Ru(NH3)5(N2O)]2++H2O. End-on coordination of N2O has been proposed for both N and O, based on vibrational and mechanistic data or nuclear magnetic resonance spectra for a related system (Pamplin et al., 2001). No crystal structure of the [Ru(NH3)5(N2O)]2+ moiety is available. Yet recently, the spectroscopic properties and electronic structure of the complex were investigated by infrared and resonance Raman spectroscopy. Both types of spectra are fully assignable and allow an unambiguous deﬁnition of the binding mode of N2O (Paulat et al., 2004). The data are only compatible with N2O being coordinated end-on via its terminal N atom in agreement with calculations on the binding mode of CNO and N2O (Tuan and Hoffmann, 1985). The optimized structure of [Ru(NH3)5(N2O)]2+ shows a linear coordination geometry with RuNN and NNO angles close to 1801. N2O coordinated to the terminal oxygen atom would produce a strongly bent structure with a calculated angle for RuONN of 1381. The calculated relative stability of the N- versus O-coordinated isomer is approximately 33.47 kJ mol1 in favour of the N isomer. In solution the complex loses easily N2O, which supports the view of N2O being a weak ligand. Thus, the synthesis of more stable complexes of N2O will be difﬁcult. Since it is a weak s and p donor, a more stable metalNNO bond must be achieved by increasing p back bonding.

3. GENOMIC AND ORGANISMAL RESOURCES 3.1. Formation and Consumption of N2O by Bacteria 3.1.1. The Complete Denitrification Pathway with N2O as Intermediate N2O as an intermediate of the denitriﬁcation process may be liberated transiently. Its relative amount is inﬂuenced by the speciﬁc growth conditions,

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including nitrate and nitrite concentrations, pH, and O2 supply. Table 1 lists examples of bacteria where N2O in various ways is product or substrate of inorganic N metabolism. For example, Thauera mechernichensis ﬁrst forms mostly N2O from nitrate, which subsequently disappears as it is being used as an electron acceptor (Scholten et al., 1999). The dominant controlling factor is O2. N2O production by Alcaligenes faecalis has been studied in dependence of transient aerobic conditions (Otte et al., 1996). In two systematic studies the dynamics of denitriﬁcation with respect to O2 were followed in P. stutzeri and Paracoccus denitrificans (Ko¨rner and Zumft, 1989; Baumann et al., 1996). The amount of N2O liberated can also be manipulated in P. stutzeri by the heterologous expression of ﬂavohaemoglobin (Takaya and Shoun, 2002). Flavohaemoglobin inﬂuences NO homeostasis and thus indirectly affects cellular N2O formation and consumption. The biochemistry of ﬂavohaemoglobin has been reviewed recently (Wu et al., 2003). A fast and simple test to identify N2O is lacking. The gas produced by a denitrifying bacterium has often not been identiﬁed and in the taxonomic description only gas production from nitrate or nitrite is stated, if at all. The best method to identify N2O is by gas chromatography. The demonstration of N2O as a denitriﬁcation intermediate is facilitated by the acetylene blockage technique (Balderston et al., 1976; Yoshinari and Knowles, 1976). The breakdown of cyclic aliphatic and aromatic compounds, particularly those with toxic properties to the environment, is possible under anaerobic, denitrifying conditions. For example, the denitrifying Azoarcus sp., strain 22Lin, grows on cyclohexane-1,2-diol as the sole electron donor and carbon source, with nitrate as electron acceptor (Harder, 1997). Azoarcus sp., strain EbN1, is a versatile bacterium with 10 anaerobic and four aerobic pathways directed at various aromatic compounds and hydrocarbons (Rabus et al., 2005). However, toluene and ethylbenzene are degraded only under anaerobic (denitrifying) conditions. Most pathways converge at the level of benzoyl-CoA, which then undergoes reductive dearomatization and hydrolytic ring cleavage. Deviating from the genus traits, the strain EbN1 lacks diazotrophy. It has been suggested that it may belong to a genus other than Azoarcus. The combination of the denitriﬁcation process with mineralizing capabilities directed at aromatic compounds and aromatic pesticides is an important aspect in maintaining the environment. In a recent study, acetylene blockage was applied to show that the strain M91-3 of Cupriavidus (formerly Ralstonia) basilensis is capable of utilizing the herbicide atrazine and other s-triazines as N source under conditions of complete denitriﬁcation (Stamper et al., 2002). Aquifer microorganisms mineralize benzene and

Reaction

116

Table 1

Bacterial N-oxide transformations involving nitrous oxide 1

2 NO 3 -[N2O] -N2

Observations or distinctive mark

Reference

Azoarcus anaerobius Microvirgula aerodenitrificans Roseobacter denitrificans

Strict anaerobe, sewage sludge Aerobic denitriﬁcation, sewage sludge Marine, aerobic anoxygenic photosynthesis Marine, 120–130 m depth, Baltic Sea Aerobic denitriﬁcation, landﬁll leachate

Springer et al. (1998) Patureau et al. (1998) Shiba (1991)

Shewanella denitrificans Thauera mechernichensis

Brettar et al. (2002) Scholten et al. (1999)

2 3 NO 2 -[N2O] -N2

Alcaligenes odorans Neisseria spp.

Clinical isolate Human commensals or pathogens

Pichinoty and Chatelain (1973) Berger (1961)

4 NO 3 -N2O

Hydrogenivirga caldilitoralis

Marine hot spring, 75 1C, chemolithotroph Thiosulfate-utilized, waste-water plant Earthworm gut Terrestrial oil well, thermophilic, 55 1C Soil

Nakagawa et al. (2004)

Finkmann et al. (2000)

Pseudoxanthomonas taiwanensis

Waste-gas bioﬁlter, animal rendering plant Thermophilic, hot spring 50–75 1C

Wolinella succinogenes Azotobacter vinelandii6

Symbiotic, rumen Diazotrophic, nondenitrifying

Yoshinari (1980) Liang and Burris (1988)

Ottowia thiooxydans Paenibacillus anaericanus Petrobacter succinatimendens Streptomyces thioluteus 3,4 NO 2 -N2O

N2O-N25 1

Luteimonas mephitis

Reaction may be strain-speciﬁc. N2O is obligate intermediate and may equilibrate with free N2O. 3 Nitrate is not reduced. 4 N2O is endproduct. 5 Nitrite is not reduced. 6 Reaction due to nitrogenase. 2

Spring et al. (2004) Horn et al. (2005) Salinas et al. (2004) Shoun et al. (1998)

Chen et al. (2002)

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Representative organisms

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alkylbenzenes at the expense of N2O, which indicates that the process is of signiﬁcance in natural habitats (Hutchins, 1991). N2O can be used in enrichment cultures as sole electron acceptor. Growth on N2O had been used as a screening method for the enrichment of N2Outilizing P. stutzeri from Senegal soil (Pichinoty et al., 1977), to isolate Alcaligenes spp. (Pichinoty et al., 1978b), and Bacillus azotoformans (Pichinoty et al., 1978a), or to isolate mutants affected in N2O respiration (Zumft et al., 1985b). In quite a special setting, N2O has been used as an alternative electron acceptor to select under anaerobic conditions for trimethylbenzenedegrading bacteria and to circumvent potentially toxic effects because of nitrite accumulation (Ha¨ner et al., 1997). The potential of using N2O in enrichment culture does not seem to have been exploited to its full advantage, and many more N2O respirers may exist. Sometimes a complete denitriﬁer, that has all four sets of enzymes, does not reduce exogenous N2O (Bryan et al., 1985; Snyder et al., 1987; Gokce et al., 1989). For Pseudomonas aeruginosa it has been shown that this is due to a regulatory effect caused from the lack of NO as the inducer for nos gene expression (Arai et al., 2003). Consequently, no N2OR is produced under N2O (SooHoo and Hollocher, 1990). We also know of bacteria that respire N2O without being denitriﬁers (Yoshinari, 1980; McEwan et al., 1985), which is conﬁrmed by genomic data (Haselkorn et al., 2001; Baar et al., 2004). Wolinella succinogenes is capable of respiring nitrate to nitrite and N2O to N2. It oxidizes formate by reducing nitrate to nitrite, nitrite to ammonia and N2O to N2. NO is not utilized by whole cells and no N2 is formed from nitrate reduction. Since N2O respiration is an autonomous energy-conserving module, there is no reason why such Wolinella-type N2O respirers should not be more numerous. 3.1.2. Truncated Denitrification: Lack of N2O Reduction The denitriﬁcation process may terminate prematurely at the level of N2O, either in nitrate or nitrite denitriﬁcation. The phenomenon has been known for some time (Greenberg and Becker, 1977; Kaspar, 1982). The truncated denitriﬁcation variant was initially described for ‘Corynebacterium nephridii’ (Hart et al., 1965; Renner and Becker, 1970), Pseudomonas chlororaphis and a strain of Pseudomonas fluorescens (Greenberg and Becker, 1977), and on subsequent search found as the end product in most carboxidotrophic bacteria (Frunzke and Meyer, 1990), in Roseobacter denitrificans (Doi and Shioi, 1991; Shiba, 1991), Thauera aromatica (Anders et al., 1995), Thauera selenatis (Macy et al., 1993), Azoarcus evansii (Anders et al., 1995), or in Bacillus (now Virgibacillus) halodenitrificans (Denariaz et al., 1989). Table 1

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lists a number of recent isolates. For example, Hydrogenivirga caldilitoris from a coastal hot spring in Japan grows optimally at 75 1C. It is an obligately chemolithotrophic, sulfur- and hydrogen-oxidizing bacterium of the phylum Aquificae. Autotrophic growth occurs with H2 by nitrate reduction to N2O as the product. We now see via genomics the underlying causes for the termination of denitriﬁcation with N2O in the absence of nos genes or their fragmentation. The loss of N2O reduction can be mutationally induced (Zumft et al., 1985b). This was important during the discovery of N2OR: comparing N2Orespiring bacteria with those lacking this capability provided genetic evidence to identify the underlying Cu enzyme. The genus Streptomyces, important for its trait of antibiotic production and as dominant soil inhabitant, has members that denitrify nitrate or nitrite to N2O (Shoun et al., 1998). Frequently, organisms found in soil, sludges or scum of sewage, terminate denitriﬁcation with N2O and contribute to an increase of N2O in the atmosphere and enhanced greenhouse effect fuelled by the high anthropogenic N input into the environment. Certain fungi that constitutively have an incomplete denitriﬁcation system and lack N2O reduction, will add to this effect (Shoun et al., 1992). Oil wells are distinct ecosystems because of their physicochemical and geochemical conditions. A moderately thermophilic bacterium, Petrobacter succinatimandens, which uses organic acids as a main carbon source was isolated from an Australian terrestrial oil reservoir. It is an obligate aerobe, but reduces nitrate to N2O (Table 1). A signiﬁcant habitat for both complete denitriﬁers and those that terminate the process with N2O is the gut of the earthworm, which seems to stimulate ingested denitrifying bacteria to produce N2O. The contribution of this specialized soil ecosystem to the global N2O budget appears to be of importance and is estimated at 3 105 t annually (Horn et al., 2005). Cases where a bacterium performs only a two-step denitriﬁcation, that limits the process to the reduction of nitrite to N2O, are relatively rare (Table 1). Such organisms qualify as true denitriﬁers since they convert an N-oxide anion into gas. Luteimonas mephitis, a yellow-pigmented bacterium, was isolated together with other N2O-producing representatives from a bioﬁlter designed to purify odours from an animal-rendering plant (Finkmann et al., 2000). Neisseria meningitidis harbours a two-step denitriﬁcation by lacking both nitrate and N2O respiration. When oxygen is limiting for growth the pathogen supports growth by supplementing energy needs by nitrite denitriﬁcation to N2O (Rock et al., 2005). Pseudomonas taiwanensis was isolated from a hot spring in Taiwan. N2O reduction is absent from this denitriﬁer, as is nitrate reduction.

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119

3.1.3. Oxygen and N2O Utilization Investigations of how bacteria degrade aromatic compounds under anaerobic conditions led to the isolation of the strictly anaerobic, denitriﬁer Azoarcus anaerobicus from enrichment cultures with resorcinol. This bacterium reduces nitrate by the respiratory pathway to N2, but also assimilates it as the nitrogen source, and grows with N2O as the sole electron acceptor (Gorny et al., 1992; Springer et al., 1998). At the other side of the extreme, one ﬁnds aerobically denitrifying strains like Microvirgula aerodenitrificans, an isolate from sewage sludge. In the presence of O2 and nitrate it co-respires both substrates. Nitrite and N2O are transiently formed and the ﬁnal product is N2 (Patureau et al., 1998). Aerobic denitriﬁcation is a desired trait for biotechnological applications to remove N from wastewater. Managing the traditional process of the sequential operation of aerobic nitriﬁcation and anaerobic denitriﬁcation is difﬁcult and results in the release of considerable amounts of N2O (Takaya et al., 2003). Oxygen exerts a regulatory effect on N2O utilization (Zumft et al., 1997). A cell suspension of P. stutzeri switches off N2O utilization nearly instantaneously when O2 is being added. When O2 has been consumed, N2O uptake resumes. The kinetics of the transitions from N2O respiration to O2 respiration and back are such that a fast regulatory response at the enzyme level must occur. The mechanism behind this is unknown. 3.1.4. N2O Formation by Ammonium-Oxidizing Bacteria The obligate aerobic, chemolithoautotrophic nitriﬁers, produce the gas species of denitriﬁcation. It is well established by now, that nitrifying bacteria have also denitrifying capability, and key enzymes and genes have been identiﬁed. Their Cu-containing nitrite reductase, NirK, must be a main source of NO formation (DiSpirito et al., 1985; Miller and Nicholas, 1985), whereas the further reduction of NO to N2O is accomplished by an NO reductase of the cytochrome bc or NorB type (Zumft, 2005a). Nevertheless, when the nirK gene is inactivated, formation of NO and N2O does not cease and additional mechanisms for gas production are suggested by this observation (Beaumont et al., 2002). Nitrosomonas europaea denitriﬁes nitrite to N2 even though the information to encode a Z-type N2OR is not present in its genome. The question of another type of reductase has been raised for this bacterium (Schmidt et al., 2004). In heterotrophic nitriﬁers such as A. faecalis N2O production is associated with hydroxylamine oxidation (Otte et al., 1999). Another notable group of N2O producers are

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methanotrophic bacteria. N2O production by nitrifying and by methanotrophic bacteria has been reviewed elsewhere (Arp and Stein, 2003). The differentiation of N2O produced by nitriﬁcation from that of denitriﬁcation is an important aspect to gauge the relative contributions of these processes to changes in the composition of the atmosphere. Stable isotope labelling has been used for this purpose (for instance see Wrage et al., 2004). A more recent technique relies on the analysis of the intramolecular distribution of 15N in N2O (Sutka et al., 2006, and references therein). N2O is ionized for that and fragmented in the mass spectrometer to N2O+ and NO+, to determine the relative abundance of 15N in the a (14N15N16O) and b (15N14N16O) isotopomers. An a-15N site preference is taken as indicative for N2O produced by nitriﬁcation.

3.2. Genomics and N2O Respiration A new source of information comes from genomics. By October 2005 we have identiﬁed, or newly annotated in this review, a total of 48 species or strains that carry a nos gene cluster (NGC) (Table 2). Sequenced bacterial genomes subsume known denitriﬁers and organisms with previously undisclosed capability of N2O utilization. It is estimated that the portion of N2O-respiring taxa is 10–15%. We now see astonishing examples where the system for N2O utilization is associated with extreme environments, ranging from the greatest depth and highest pressure to salt-saturated environments, and from low to high extremes of temperature. It underlines that N2O respiration is a basic process that may allow extremophiles to survive in harsh environments. The biochemical adaptations must be signiﬁcant to accommodate such an ample spectrum of physical conditions, generally adverse to the common prokaryote. In the following, we brieﬂy discuss bacteria with remarkable metabolic traits and exhibiting N2O-respiring capability with such inference retrieved from genomes (Table 2). 3.2.1. Halorespirers Halorespiration is an anaerobic respiratory process that couples the reductive dehalogenation of chlorinated hydrocarbons to growth. Genomic evidence shows that this trait is found in association with N2O respiration in Anaeromyxobacter dehalogenans, Dechloromonas aromatica and Desulfitobacterium hafniense. An. dehalogenans, a pigmented bacterium with gliding motility, grows anaerobically on acetate as the electron donor using 2-chlorophenol as the electron acceptor (Sanford et al., 2002). Its closest

Code

Taxon1

Achromobacter cycloclastes IAM1013 Anaeromyxobacter dehalogenans 2CP-C Azoarcus sp. EbN1 Bradyrhizobium japonicum USDA110 Brucella abortus 9-941 Brucella melitensis 16M Brucella suis 1330 Burkholderia mallei ATCC23344 Burkholderia pseudomallei K96243 Burkholderia thailandensis E264 Campylobacter fetus 23D Colwellia psychrerythraea 34H Cupriavidus necator H16 (plasmid) Cupriavidus metallidurans CH34 Dechloromonas aromatica RCB Desulfitobacterium hafniense DCB-2 Haloarcula marismortui ATCC43049 Haloferax volcanii DS2 Lyngbya majuscula 19L Magnetospirillum magnetotacticum MS-1 Marinobacter aquaeolei VT8 Neisseria gonorrhoeae FA1090 Neisseria meningitidis Z2491(serogroup A) Neisseria meningitidis MC58 (serogroup B) Neisseria meningitidis FAM18 (serogroup C) Paracoccus denitrificans PD1222 Photobacterium profundum SS9 Pseudomonas aeruginosa DSM50071 Pseudomonas aeruginosa PAO1 Pseudomonas aeruginosa UCBPP-PA14 Pseudomonas fluorescens C7R12 Pseudomonas sp. MT1

Acyc Adeh Azoar Bjap Babo Bmel Bsui Bmal Bpse Btha Cfet Cpsy Cnec Cmet Daro Dhaf Hmar Hvol Lmaj Mmag Maqu Ngon NmenA NmenB NmenC Pden Ppro PaerD PaerP PaerU Pﬂu PseuMT1

? d b a a a a b b b E g b b b g+ A A cy a g b b b b a g g g g g g

Gene arrangement within nos gene cluster Rx

Zx

Zx Zx R|R x D x Rx Zx Rx Zx Rx Zx Rx Zx vX Zx vX Zx vX Zx 3

Rx vX vC vX vC Lxy Lxy vC

Zx Zx Zx Zx Zx Zx

Dx c Fx Dx Dx Dx Dx Dx Dx Dx Dx Dx Rx Rx C1 x c c Dx

Z5 x Z|Z3 x D x Rx Zx Dx R|R3 x Z x D|D3 x R6 x R6 x R6 x Rx Zx Dx Rx Zx Dx R5 x Zx D5 x Rx Zx Dx Rx Zx Dx Rx Zx D Rx Zx Dx

Fx Dx Yx Fx Fx Fx Fx Fx Fx Fx Gx Fx Dx Dx C2 x Dx D2 x Fx

Yx Fx Lx Yx Yx Yx Yx Yx Yx Yx C1 x Yx Fx Fx c Lx F1 x Yx

Lx c c Lx Lx Lx Lx Lx Lx Lx C2 x Lx Yx Yx Dx Yx Y1 x Lx

Xx Yx b Xx Xx Xx Xx c c c Hx

Gx Fx Fx Dx Dx Dx Fx Fx

Hx Yx Yx Fx Fx Fx Yx Yx

Fx Lx Lx Y6 x Y6 x Y6 x Lx Lx

Lx

Fx Fx Fx Fx

Yx Yx Yx Yx

Lx Lx L5 x Lx

vC

c c c Fx

Cx

Cx Cx Cx Yx

Rx Rx Rx Lx

Lx Lx Lx Gx Hx Fx c Yx Fx [L x y v D1]4 y Lx

Rx Xxy

Source

Lx Lx Lx Xx

Yx

Cluster Genome2 Genome Genome Genome Genome Genome Genome Genome Genome2 Cluster Genome2 Genome Genome Genome2 Genome2 Genome Genome2 Gene Genome2 Genome2 Genome Genome Genome Genome2 Cluster Genome Cluster Genome Genome2 Cluster Cluster

(Continued )

121

Species and strain

RESPIRATORY TRANSFORMATION OF N2O

Table 2 Organizational patterns of nos genes in bacteria and archaea

122

Table 2 (continued ) Code

Taxon1

Pseudomonas stutzeri ATCC14505 Pyrobaculum aerophilum IM2 Ralstonia solanacearum GMI1000 Rhodobacter capsulatus SB1003 Rhodoferax ferrireducens DSM15236 Rhodopseudomonas palustris HaA2 Rhodopseudomonas palustris CGA009 Rhodobacter sphaeroides IL106 (plasmid) Salinibacter ruber DSM13855 Shewanella denitrificans OS-217 Shewanella oneidensis MR1 Silicibacter pomeroyi DSS-3 (plasmid) Sinorhizobium meliloti 1021 (plasmid)) Thermomicrobium roseum DSM5159 Thiobacillus denitrificans ATCC 25259 Thiomicrospira denitrificans ATCC33889 Wolinella succinogenes DSMZ1740

Pstu Pbaer Rsol Rcap Rfer RpalH RpalC Rsph Srub Sden Sone Spom Smel Tros Tden Tmden Wsuc

g A b a b a a a cfb g g a a gns b e e

Gene arrangement within nos gene cluster Rx vX RX x y vX vC

vC Rx Rx Rx Rx

Rx Rx Rx

Zx Zx Zx Zx Zx Zx Zx Zx Zx Zx Zx Zx Zx Zx Zx

Dx Rx Dx Dx Dx Dx Dx Dx Dx Lx Dx Dx D7 Rx c c

Fx sco1 x Dx Fx Fx Fx Fx Fx Fx Fx Dx Fx Fx Fx Dx Dx Dx

Yx c Fx Yx Yx Yx Yx Yx Yx Yx Fx Yx Yx Yx Fx Gx Gx

Lx Lx Yx Lx Lx Lx Lx Lx Lx Yx Lx Lx Lx Yx C1 x C1 x

c Lx Xx c Xx Xx

Source

Dx Fx

c

Lx

Xx Lx c c Lx C2 x H x F x L x C2 x H x F x L x

Cluster Genome Genome Genome2 Genome2 Genome2 Genome Cluster Genome2 Genome2 Genome Genome Genome Genome2 Genome2 Y x L x L x Genome2 Y x L x L x Genome

Note: x , transcriptional direction of gene; c, ORF encoding a hypothetical protein. 1 Greek letters denote the corresponding group of the proteobacteria. A, archaeon; cy, cyanobacterium; cfb, cytophaga-ﬂavobacter-bacteroides group; gns, green sulfur bacteria group; g+, gram-positive. 2 Unﬁnished by time of writing. 3 Consists of two adjacent ORFs. 4 Plasmid-encoded. 5 Incomplete sequence. 6 Atypically short. 7 Fused with downstream nosF.

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Species and strain

RESPIRATORY TRANSFORMATION OF N2O

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relatives are members of the Myxococcales, which were considered as obligately aerobic bacteria until recently. De. aromatica belongs to the b-proteobacteria but its taxonomic position among the Rhodocyclaceae has not yet been validly described. Halorespiring bacteria have not only been isolated from stream sediment, soil and municipal sludge, but also from the gut of the earthworm, where Dechloromonas denitrificans was shown to be an active nitrate denitriﬁer with N2O as an intermediate. N2O production is particularly enhanced by nitrite denitriﬁcation (Horn et al., 2005). Dechloromonas strains RCB and JJ produce N2 from nitrate. They oxidize under anaerobic conditions benzene and seem to be the ﬁrst-known examples of their kind (Coates et al., 2001). D. hafniense represents the ﬁrst instance where information about an N2OR can be deduced for a grampositive bacterium (see Section 8.4). 3.2.2. Piezophiles: N2O Utilization at Extreme Depth Not only are nos genes associated with common marine bacteria but also with deep-sea inhabitants. Denitriﬁcation is found over the entire vertical extent of the oceans, stretching from coastal and surface waters to the most profound depths. The sequenced strain SS9 of Photobacterium profundum (Vezzi et al., 2005) was isolated at 2550 m depth in the Sulu Trough, off Mindanao Island in the Philippines. The strain was found there in association with amphipods. It grows optimally at 20 MPa and is also a moderate psychrophile. Only reduction of nitrate to nitrite was reported in the description of this new species (Nogi et al., 1998). The genome of Ph. profundum has a tripartite organization with two circular chromosomes of 4.1 and 2.2 Mb, and an 80-kb plasmid. The NGC is found on chromosome II, the smaller one of the two principal genetic elements (Table 2). The most remarkable habitat of a bacterium that carries nos genes is the Mariana Trench. Pseudomonas sp. MT1 was isolated from mud at the ocean ﬂoor at 11,000 m depth (Tamegai et al., 2002). The bacterium is only a facultative and not a strict anaerobe. It is held that even at these extreme depths the water is ﬂuctuating with respect to its O2 content. Pseudomonas sp. MT1 is by 16S rRNA analysis closely related to P. stutzeri. It is of interest in this context that the ZoBell strain of P. stutzeri is viable under high pressure up to E60 MPa (ZoBell and Oppenheimer, 1950). Thus, we see several examples where denitriﬁcation is associated with piezophilic (high-pressure adapted) or piezotolerant bacteria. Silicibacter pomeroyi, not a piezophile and living in coastal areas, is a marine bacterium of interest for its role in carbon cycling (Moran et al.,

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2004). It has a mixed litho-heterotrophic metabolism that utilizes CO for bioenergetic purposes but not as the carbon source. Si. pomeroyi participates together with members of the Roseobacter group in the cycling of dimethylsulfoniopropionate, (CH3)2S+(CH2)2COO, in the oceans, a process that affects cloud formation and climate. Although the genus description of Silicibacter includes reduction of nitrate to nitrite, lack of nitrite reduction was speciﬁcally reported for Si. pomeroyi (Gonza´lez et al., 2003). Its genome sequence, however, reveals sets of nir, nor and nos genes on a 492-kb megaplasmid, but no nar genes for respiratory nitrate utilization (Moran et al., 2004), thus presenting quite the opposite picture of the genus traits.

3.2.3. Halophiles: nos Genes in Bacteria and Archaea Salinibacter ruber, a bright-red pigmented bacterium, was isolated from saltern crystallizer ponds in Alicante and Mallorca, Spain (Anto´n et al., 2002). It has an extreme salt requirement of 20–30% saturation for optimal growth, and is the most halophilic representative among the Bacteria known to date. Until the isolation of this species such salt requirement was held to be an exclusive trait of Archaea. Sa. ruber was described as a strict aerobic chemoorganotroph that would not reduce nitrate to nitrite or to gas. With the caveat of the still unﬁnished sequence, no nar or nir genes appear in the genome in agreement with the physiological data. However, the genome has revealed the presence of the core nosZDFY cluster, suggesting the possibility of an anaerobic growth mode (Table 2). ‘Marinobacter aquaeolei’ was isolated from an oil-ﬁeld brine from an offshore platform in southern Vietnam. The bacterium has a complete NGC, but was reported to grow only anaerobically on nitrate with various organic acids (Huu et al., 1999). The strain is mesophilic and moderately halophilic. It degrades n-hexadecane and several components from crude oil. ‘Halobacterium marismortui’, the halobacterium of the Dead Sea and redescribed as Haloarcula marismortui, was the ﬁrst archaeon to be shown capable of gas evolution from nitrate (Elazari-Volcani, 1940). The gases were identiﬁed as N2O, NO and N2 by mass spectrometry (Werber and Mevarech, 1978). The genome carries the entire complement of denitriﬁcation genes, including an NGC, in agreement with the report of complete denitriﬁcation for this archaeon. Nitrate reduction to nitrite, but no growth on nitrate was initially stated for Haloferax volcanii (Mullakhanbhai and Larsen, 1975; Grant et al., 2001). The genome reveals genes encoding a

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NirK nitrite reductase, NorZ NO reductase and an NGC, however without evidence for nosZ (Table 2). N2O metabolism among archaea is discussed in more detail in Section 8.3.

3.2.4. Psychrophiles and Thermophiles Colwellia psychrerythraea lives at the lower end of the biological temperature scale (Methe´ et al., 2005). The strain 34H, whose genome sequence was determined, was isolated from Arctic marine sediments at 305 m depth and 0.7 1C. It grows well in heterotrophic media over a temperature range of 1 to 10 1C. Its development is delimited by extremes of 10 and 19 1C; at 10 1C it is still mobile. The bacterium serves as a biochemical model of cold adaptation and properties of cold-active enzymes. The interest in psychrophilic bacteria extends to bioremediation and biotechnological applications. The taxonomic description of Co. psychrerythraea states that nitrate is reduced to nitrite without gas production (D’Aoust and Kushner, 1972). No information was given for strain 34H of its nitrate respiratory or denitrifying capabilities (Huston et al., 2000); however, its genome reveals a full set of nos genes (Table 2). Rhodoferax ferrireducens is only a moderate psychrophile. Its closest relatives are purple nonsulfur bacteria. Unlike them, the bacterium does not grow phototrophically or by fermentation, but respires nitrate, Fe(III) and O2 among several electron acceptors. The products of nitrate reduction were not reported (Finneran et al., 2003). At the extreme end of thermophiles with complete denitriﬁcation is the archaeon Pyrobaculum aerophilum, which grows by respiration of nitrate (Vo¨lkl et al., 1993). A static culture supplied with nitrite does not grow, but under continuous sparging with an N2–CO2 mixture, the bacterium oxidizes organic substrates at the expense of nitrite reduction to N2, with trace production of N2O and NO. N2O alone does not serve as an electron acceptor. The bacterium grows between 75 and 104 1C. Several nos genes are present in the genome of strain IM2 but the situation of a gene encoding an N2OR is unclear (see Section 8.3). Thermomicrobium roseum was isolated in Yellowstone National Park from a hot spring, at 74 1C and pH 8–9. In the systematic description the bacterium is noted as strictly respiratory, obligate aerobic bacterium, but not as a denitriﬁer. Nitrate is used as a source of inorganic N (Garrity and Holt, 2001). The fact that Te. roseum has a set of nosZDFYL genes qualiﬁes it as an N2O-respiring bacterium. Thermomicrobium had been

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considered as a separate phylum, but has now been moved to Chloroflexi (Hugenholtz and Stackebrandt, 2004). The taxon is gram-negative and is characterized by the absence of peptidoglycan in the cell wall. Instead of glycerolipids the cell membrane of Te. roseum is composed of long-chain diols, which are thought to be a factor for membrane stabilization at high temperature (Pond et al., 1986). 3.2.5. Mutualistic and Pathogenic Relationships The diazotrophic genera Azoarcus, Azospirillum, Bradyrhizobium, Sinorhizobium and Rhodobacter harbour denitrifying species. The genera Bradyrhizobium and Sinorhizobium are important symbionts of legumes. They serve as a model organism to study denitriﬁcation and N2O utilization in the context of the root nodule. W. succinogenes is part of the complex rumen symbiosis. Previous surveys of denitrifying bacteria (Zumft, 1992, 1997) have identiﬁed a number of pathogens. NGCs are present in P. aeruginosa, Brucella spp., Burkholderia spp., Neisseria spp. and Campylobacter fetus (Table 2). Nitrate-denitrifying conditions have been shown to be important for bioﬁlm formation by P. aeruginosa in cystic ﬁbrosis (Yoon et al., 2002). 3.2.6. Taxonomic Transfers Changes in the taxonomic position of several bacteria relevant to this article are the following: Alcaligenes xylosoxidans transferred to (-) Achromobacter xylosoxidans (Yabuuchi et al., 1998; Coenye et al., 2003), Bacillus halodenitrificans - Virgibacillus halodenitrificans (Yoon et al., 2004); Bacillus thermodenitrificans-Geobacillus thermodenitrificans (Nazina et al., 2001); Hydrogenomonas eutropha-Alcaligenes eutropha-Ralstonia eutrophaWautersia eutropha-Cupriavidus necator (Vandamme and Coenye, 2004); Marinobacter aquaeolei-Marinobacter hydrocarbonoclasticus (Ma´rquez and Ventosa, 2005); Paracoccus denitrificans GB-17-Thiosphaera pantotropha-Paracoccus pantotrophus (Rainey et al., 1999); Pseudomonas nautica-Marinobacter hydrocarbonoclasticus (Spro¨er et al., 1998); Pseudomonas perfectomarinus-Pseudomonas stutzeri ZoBell (Do¨hler et al., 1987); Ralstonia metallidurans-Cupriavidus metallidurans (Vandamme and Coenye, 2004); Archaea: Halobacterium denitrificans-Haloferax denitrificans (Tindall et al., 1989) Halobacterium marismortui-Haloarcula marismortui (Oren et al., 1990); Halobacterium mediterranei-Haloferax mediterranei (Torreblanca et al., 1986); Halobacterium vallismortis-Haloarcula vallismortis

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(Torreblanca et al., 1986); Halobacterium volcanii-Haloferax volcanii (Torreblanca et al., 1986).

4. PROPERTIES OF N2O REDUCTASE 4.1. Finding the Enzyme by a Classical Microbiological Strategy N2O can serve as the sole electron acceptor for growth by denitrifying bacteria. Nevertheless, early attempts to isolate the underlying enzyme failed. An indirect strategy to reach this objective was based on the nutritional requirements of N2O-respiring cells. Classical examples to identify the essential trace element of a biological process by following the physiological response towards a metal are dinitrogen ﬁxation (Mo), photosynthetic O2 evolution (Mn) or methanogenesis (Ni). The same strategy worked successfully for N2OR (Cu). It was shown that Cu speciﬁcally stimulates anaerobic growth under N2O, whereas Cu deﬁciency impairs the reaction (Iwasaki et al., 1980; Iwasaki and Terai, 1982; Matsubara et al., 1982). Since a transition metal can be expected in N2O activation and reduction, the search had found an object and was directed towards Cu-containing proteins (Matsubara and Zumft, 1982). This was greatly helped by the fact that taxonomically closely related pseudomonads provided backgrounds with and without N2O reduction to compare. The combined strategy resulted in the discovery of N2OR from the ZoBell strain of P. stutzeri (Zumft and Matsubara, 1982). A third element for the successful outcome came from a coupled assay based on the reduction of benzyl viologen by H2 and hydrogenase, which was available because of the research on N2 ﬁxation in the laboratory of one of the authors (WGZ). The ﬁnding was initially received with skepticism (Snyder and Hollocher, 1984). But the alternative view that N2O reduction would not be associated with a Cu protein could not be upheld in light of further biochemical and corroborating genetic evidence (Zumft et al., 1985a,b). In hindsight, one can reconstruct from spectral evidence that 10 years before its enzymatic identiﬁcation, N2OR had been isolated as an unusual Cu protein from A. faecalis IAM 1015 (Matsubara and Iwasaki, 1972) that, having no biological function, faded into oblivion among the Cu proteins. The same protein was independently found a second time as a side product in the puriﬁcation of cytochrome cd1 nitrite reductase from P. stutzeri, where preparative isoelectric focusing is the most effective means to separate haem and Cu proteins, and N2OR becomes visible as a bright magenta band

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in the ﬁnal puriﬁcation step (Zumft and Vega, 1979; see also Zumft, 1997). When T. Matsubara from the Mori-Iwasaki laboratory at Nagoya University, Japan, spent a sabbatical in one of the authors’ laboratories, common interests and the above strategy ﬁnally untangled the problem.

4.2. Z-type N2O Reductase Biochemical studies have been directed nearly exclusively at Z-type N2ORs, the exception being the enzyme from W. succinogenes, which is modiﬁed by carrying a covalently attached c-type cytochrome (Teraguchi and Hollocher, 1989; Zhang et al., 1991; Simon et al., 2004). Depending on the source and the puriﬁcation procedure, distinct forms of the Z-type enzyme are obtained which differ considerably with regard to their activity and spectroscopic properties (Fig. 2; Table 3). In spite of this, all N2ORs isolated are homodimers that contain two multinuclear Cu centres per subunit: (i) the CuA mixed-valent electron transfer site with two bridging cysteine ligands forming a highly spin-delocalized Cu2S2 rhomb, and (ii) the catalytic tetranuclear CuZ site, which represents the ﬁrst biologically active CuS cluster known. In CuZ, the Cu atoms are coordinated solely to imidazole nitrogen from seven histidine residues (Fig. 3). Cu adopts in biological systems usually the

(A)

(B)

Figure 2 (A) UV–vis absorption and (B) EPR spectra of the various forms of N2OR from P. stutzeri. N2OR I, enzyme as isolated in the absence of O2; N2OR III, enzyme form I, reduced with dithionite; N2OR IV, apoprotein reconstituted with Cu(en)2SO4; N2OR V, enzyme form isolated from a nosD mutant defective in CuZ centre assembly. EPR spectra recorded at 9.5 GHz (X-band), temperature 15 K.

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Table 3 Properties of the different forms of N2O reductase from Pseudomonas stutzeri Form of N2OR1

Preparation or enzyme state

Cu/Mr2

N2Oreducing activity3

UV-vis absorption4

EPR features5

I6

Anoxic; CuZ* low7

E8

70

540 (14); 480/620 sh; 780 (4.0)

II

Oxic; CuZ* high

E7

30

480(5.4); 530 (6.7); 620 (4.2); 780 (4.0)

III

Reduced8

E8

Inactive

650 (6.1)

IV

Regenerated9

E4

Inactive

Similar to II; 620 absent

V

Mutant MK40210

E4

Inactive

Similar to IV

CuA, S ¼ 1/2; 7 lines at gJ ¼ 2.180; AJE3.8 mT; g? E2.03 CuA, S ¼ 1/2; gJ-region less resolved because of underlying CuZ* signal, g?E2.046 CuZ, S ¼ 1/2; gJ-region not resolved; gJ ¼ 2.160; g? ¼ 2.055 Similar to II but CuZ*signal is absent Similar to IV

1

Data from Coyle et al. (1985) and Riester et al. (1989). Average value from preparations obtained with [Cu] in growth medium of 1 mM; increase of [Cu] in growth medium to 5 mM yields to up to 12 Cu atoms/Mr in highly active N2OR I. 3 nkat mg protein1, pH 6.0, electron donor benzyl viologen; maximal value of 1000 nkat mg 2 protein1 at pH 9.8 for N2OR I; CO activates whereas CN, N 3 , NO, S2O4 , and C2H2 inhibit the enzyme. 4 lmax nm (e in mM1cm1); sh, shoulder; spectroelectrochemical titration at 540 nm, pH 7.5, 25 1C, gives E0 0 ¼ +260 mV versus SHE for CuA. 5 X-band (E9.5 GHz); temperature 15 K. 6 No freely accessible –SH groups are present in N2OR I; four disulﬁde groups are determined in the holoenzyme dimer which may play a role in intermonomeric contacts and proper positioning of CuA and CuZ (Dreusch et al., 1996); 1.670.2 inorganic sulfurs/dimer, Cu/S ratio 6.2 (Rasmussen et al., 2000). 7 CuZ* contributes to the absorbance around 620–640 nm, and leads to a broadening of the EPR spectrum in the gJ-region at X-band. 8 Two to 10-fold excess dithionite in the absence of O2. 9 Cu removed by dialysis against KCN followed by incubation of apoN2OR with Cu(en)2SO4; all operations in the absence of O2. 10 Tn5-induced mutant with defective chromophore biosynthesis. 2

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CuA

Figure 3

Cuz

Structures and oxidation states of CuA and CuZ of N2OR.

oxidation states +1 and +2. Hence, in principle three different oxidation and resulting spin states can arise of CuA, and ﬁve different states of CuZ (Fig. 3). For the CuA centre only the S ¼ 1/2 and S ¼ 0 spin states have been observed, whereas for CuZ the states [Cu4S]4+, S ¼ 0, 1 and [Cu4S]3+, S ¼ 1/2, and the colourless super-reduced state [Cu4S]2+, S ¼ 0, have been characterized by spectroscopic techniques including multifrequency electron paramagnetic resonance (EPR), circular dichroism (CD), magnetic circular dichroism (MCD), and extended X-ray absorption ﬁne structure (EXAFS) spectroscopy (Coyle et al., 1985; Riester et al., 1989; Rasmussen et al., 2002; Chen et al., 2004; Oganesyan et al., 2004; Wunsch et al., 2005). The enzyme from P. stutzeri has been studied extensively with regard to its biochemical and spectroscopic properties, metal centre assembly, analysis of inorganic sulfur and putative role of sulfhydryl and disulﬁde groups, and recombinant protein variants (Zumft and Kroneck, 1996; Zumft, 1997, 2005b). Thus, even though its X-ray crystal structure has not been solved, the N2OR from P. stutzeri serves as the paradigm of the Z-type N2OR family. The properties of its different spectral forms are summarized in Table 3. The principal biochemical properties of N2OR from various bacterial sources have been compiled (Zumft, 1997); the recombinant derivatives are described in Sections 5 and 6. The X-ray structures of the enzymes from Marinobacter hydrocarbonoclasticus (formerly Pseudomonas nautica) and Pa. denitrificans (Brown et al., 2000b; Haltia et al., 2003) were obtained from material that was isolated and puriﬁed in the presence of O2. In solution these enzymes show the principal features of the UV–vis and EPR spectra of N2OR from P. stutzeri puriﬁed in air resulting in N2OR form II (Prudeˆncio et al., 2000; Haltia et al., 2003). Spectrally unequivocal forms of

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N2OR I have been isolated from Alcaligenes sp. (Matsubara and Sano, 1985) and Achromobacter (formerly Alcaligenes) xylosoxidans (Ferretti et al., 1999). Of other important biometals, such as Mo, Mn, Co, Ni or Zn, none was required for N2OR activity. However, calcium appears to play an important structural role in dimer formation, which is essential for catalysis (see Section 5). Most enzyme preparations have an average Cu content of 7–9 Cu atoms per N2OR dimer, which is below the value deduced from the X-ray structure. In P. stutzeri it was found that an increase of the Cu concentration from 1 mM in the growth medium to 5 mM will yield up to 10–12 Cu atoms per N2OR dimer in highly active N2OR I (Charnock et al., 2000; Rasmussen et al., 2000). A change in metal centre occupancy by the exogenous Cu concentration indicates the possibility of a noncatalysed process for Cu delivery. Inhibition of N2O reduction by acetylene was discovered long before the enzyme was identiﬁed (Fedorova et al., 1973). Acetylene is still the most selective inhibitor of N2OR. The inhibition is noncompetitive with a Ki of 28–45 mM (Balderston et al., 1976; Kristjansson and Hollocher, 1980). The inhibition is reversible; the mechanism of action is unknown. In the absence of O2 under reducing conditions nearly all of the Cu is released from N2OR by dialysis of the protein against cyanide (Coyle et al., 1985; Riester et al., 1989). The colourless apo-protein is catalytically inactive. Soaking of crystalline N2OR from Pa. denitrificans with cyanide suggests that an average of one Cu atom of the tetranuclear CuZ cluster is removed (Haltia et al., 2003). In solution, approximately 50% of Cu originally present in N2OR from P. stutzeri can be reconstituted using Cu(en)2SO4 as the source for Cu. The resulting regenerated enzyme form, N2OR IV, is catalytically inactive but shows the optical and EPR features of the mixed-valent CuA centre (Table 3). Attempts to regenerate the catalytic CuZ site of N2OR using either Cu(II) or Cu(I) compounds, or mixtures of both, have failed so far, which is not surprising in view of the complex machinery responsible for the biogenesis of the CuZ centre (Zumft and Kroneck, 1996; Zumft, 2005b), and the difﬁculty to provide apt Cu and sulﬁde sources simultaneously.

5. STRUCTURE OF N2O REDUCTASE Almost two decades after the identiﬁcation of N2OR, the X-ray structure was reported (Brown et al., 2000b). The enzyme for crystallization was

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puriﬁed from Ma. hydrocarbonoclasticus in the presence of O2, whereby two forms appeared: a blue form (lmaxE635 nm) that crystallized, and a pink form with lmax ¼ 480, 540, E800 nm (Prudeˆncio et al., 2000). The difference between the two forms was seen as a difference in the oxidation state of the CuA centre, and the authors noted from the 4-line EPR spectrum, S ¼ 1/2, that the CuZ centre in both forms remained reduced. N2ORs characterized thus far have been described as homodimers carrying four Cu ions per monomer on the average (Zumft and Kroneck, 1996). The prevailing opinion held that four Cu ions assemble in two dinuclear centres: one constitutes the CuA site, the other, termed CuZ, represents the catalytic site. Whereas the structure of the CuA centre was accurately predicted from complementary spectroscopic studies, that of the CuZ site was unexpected. It comprises four Cu ions arranged in a novel ‘butterﬂy’ cluster and solely coordinated by histidine residues. The N2OR structure marks the discovery of a new type of biological metal centre (Rosenzweig, 2000). Initially, based on the analysis of optical, EPR, and MCD data, CuZ was proposed to be coordinated by sulfur from cysteine similar to CuA. Sequence comparisons of N2OR indicated that there are not enough conserved cysteine residues to build two thiolate-bridged dinuclear centres. However, a number of histidine residues are highly conserved and led to the notion that CuZ might be coordinated by several histidines (Holz et al., 1999; Charnock et al., 2000). This type of coordination is found in the trinuclear Cu centres in the multicopper oxidases, ascorbate oxidase, ceruloplasmin or laccase (Solomon et al., 1996). By a difference-spectrum approach, using the Cu K-edge X-ray absorption spectrum of the native enzyme and that of the CuA-only form, histidine was identiﬁed as a major contributor to the back scattering with contributions from Cu and/or S atoms being observed also (Charnock et al., 2000). In the ﬁrst crystal structure of N2OR, in addition to the seven histidine residues, three exogenous ligands were assigned as being either water or a hydroxide anion. A sulfur-containing ligand was not considered, which made it difﬁcult to reconcile the structure of the CuZ centre with the spectroscopic data that strongly suggested sulfur ligation. However, this picture changed rapidly. Elemental analysis and resonance Raman spectroscopy of isotopically labelled N2OR conclusively demonstrated the presence of one acid-labile sulfur ligand (Rasmussen et al., 2000; Alvarez et al., 2001). Almost at the same time, the structure of N2OR from Pa. denitrificans at 1.6-A˚ resolution became available (Brown et al., 2000a) and conﬁrmed the existence of one sulﬁde at the CuZ site. The importance of sulfur and its astounding chemical versatility among the 12 nonmetallic elements essential for life has been elaborated (Beinert, 2000). Unusual reaction pathways involving sulfur apparently become

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possible by the speciﬁc properties of this element. CuZ and its intriguing catalytic properties represent an important example among the metal-sulfur sites in biology. Whether the current structure also represents that of the active enzyme puriﬁed under anaerobic conditions still remains to be seen.

5.1. Crystal Structure and Subunit Interactions The crystal structures of N2OR from Ma. hydrocarbonoclasticus at 2.4-A˚ resolution (Brown et al., 2000a,b) and Pa. denitrificans at 1.6-A˚ resolution (Haltia et al., 2003), have been solved. In both cases, the protein used for crystallization was puriﬁed in the presence of O2 producing a ‘blue’ form of N2OR. The UV–vis spectrum of the blue crystal of the Ma. hydrocarbonoclasticus enzyme exhibits a major absorption maximum around 635 nm as described for dithionite-reduced N2OR, or form III (Fig. 2; Coyle et al., 1985; Riester et al., 1989). These crystals have been obtained from ‘form B’ of the Ma. hydrocarbonoclasticus enzyme, whose solution spectrum showed maxima around 540 and 635 nm (Prudeˆncio et al., 2000). In contrast, the solution spectrum of the blue N2OR from Pa. denitrificans has distinct absorption maxima at 480, 540 and 640 nm, similar to aerobically puriﬁed N2OR from P. stutzeri or the so-called N2OR II (Riester et al., 1989). The N2OR from Pa. denitrificans has 61% positional amino sequence identity with that of Ma. hydrocarbonoclasticus, which is clearly reﬂected in a high structural similarity between the two enzymes (Haltia et al., 2003). The enzyme is a head-to-tail homodimer with a large dimerization interface stabilized by a multitude of highly speciﬁc polar and nonpolar as well as chelating interactions (Fig. 4). Each N2OR monomer is composed of two distinct domains: the C-terminal domain carrying the mixed-valent [CuA(1.5+) y CuA(1.5+)] electron transfer centre and the N-terminal domain, which hosts the catalytic, m4-sulﬁde-bridged CuZ site. The latter domain adopts a seven-bladed b-propeller homologous to other enzymes, such as methylamine dehydrogenase or galactose oxidase. The CuZ centre is located at one end of the sevenfold pseudoaxis of the propeller at a position similar to that of other b-propeller structures (Fu¨lo¨p and Jones, 1999). b-Propellers are involved in diverse functions such as cell-cycle control, signalling and catalysis. In N2OR, the propeller domain seems to be engaged both in the formation of the active site and in protein–protein interactions that are essential for dimerization. The C-terminal domain consists of the canonical cupredoxin fold observed in type-1 Cu proteins and in subunit II of COX (Murphy et al., 1997).

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Figure 4 (A) Stereoview of the structure of the N2OR homodimer from Pa. denitrificans. (B) Views of the CuA electron transfer centre and the catalytic centre CuZ. (C) The N2OR monomer seen from the contact side with the other subunit. In this orientation the outermost b-sheets run counterclockwise (Protein Data Bank ﬁle 1FWX; drawn with MolScript and Raster3D softwares). (See Colour Plate Section in back of this volume.)

The shortest CuACuZ distance within a monomer is E40 A˚, which is too long for an efﬁcient electron transfer between the two sites. On the other hand, the corresponding intermonomer distance of 10 A˚ is well suited for

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135

fast electron transfer within a protein (Fig. 4) (Page et al., 1999). Obviously, the dimeric architecture is essential for catalysis and the residues at the dimer interface tend to be highly conserved. In addition to Cu, N2OR binds Ca2+ and chloride ions. The two tightly bound Ca2+ ions in each monomer seem to have a role in dimer formation. In the Pa. denitrificans N2OR structure a third Ca2+ resides on the surface of the protein, and several CuA and CuZ ligands participate in intermonomeric contacts. These ligands located at the dimer interface may provide an efﬁcient electron transfer route from CuA to the substrate molecule bound at the CuZ site. The CuA domain is important for the structural integrity of N2OR. Deletion of the domain renders N2OR unstable and subjects it to rapid degradation. Mutations of the CuA ligands His583, Cys618 and Met629 affect protein export and result, to various degrees, in the accumulation of a cytoplasmic N2OR (Heikkila¨ et al., 2001). The ﬁrst three-dimensional model for an N2OR at about 20-A˚ resolution, deduced from X-ray scattering of the enzyme from A. xylosoxidans (Ferretti et al., 1999), had the two subunits arranged in a twofold symmetry unit around a central axis. A possible interpretation of such an arrangement was that CuA and CuZ are juxtaposed across the subunit interface, which was later conﬁrmed by the X-ray structure. A perturbation in one type of the Cu centre may, thus, also be propagated to the other. Note that a distinct property of the CuA centre in COX and quinol oxidases is to function in the electron transfer pathway across the subunit interface.

5.2. Surface Charge Carboxylic acid residues have been identiﬁed to be required for the binding of cytochrome c to COX (Lappalainen et al., 1993, 1995), where several of them form a negatively charged patch surrounding the CuA site on subunit II (Witt et al., 1998). Asp178 has been shown in Pa. denitrificans to be important for binding of cytochrome c (Witt et al., 1998). The homologous residue of N2OR, Asp580 was mutated (Charnock et al., 2000). A charge loss affected the activity (Asp580Asn substitution) and so did a modiﬁcation of the side chain that maintained the charge (Asp580Glu substitution). The largest decrease in activity was observed when both features were combined (Asp580Ser substitution). These results indicate an indirect role of this conserved acidic residue rather than that of a carboxylate ligand of the CuA core. Since more than one charged residue is expected to form a docking site for the electron donor, the observed response towards the affected amino acid substitutions is compatible with such a role. Positioned N-terminally to Asp580 are the notably conserved acidic residues Asp566, Glu567, Asp576,

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Glu579 and Asp540, several of which may be part of a charged surface patch on the protein serving as the docking site for an electron donor.

6. NOVEL Cu CENTRES IN N2O REDUCTASE 6.1. CuA Centre 6.1.1. Development of the Field In his recollections ‘Crystals and structures of cytochrome c oxidases – the end of an arduous road’, and ‘Copper A of cytochrome c oxidase, a novel long-embattled biological electron transfer site’ Beinert (1995, 1997) traces the history of unravelling the CuA site in COX from the beginning, when few believed that there was any signiﬁcant amount of Cu in this enzyme, followed by the veriﬁcation of three Cu atoms per monomer, and the ﬁnal identiﬁcation of the site as a dinuclear, cysteine-bridged mixed-valence centre. The detection of a binuclear CuA centre in N2OR, mainly by spectroscopic techniques, was central for recognizing the same structure in COX (Beinert, 1997). The EPR spectra of CuA were ﬁrst studied in detail by Beinert et al. (1962) who established that CuA is different from any other Cu centre studied at that time. The physicist in the team, R. H. Sands, realized that the signal observed with COX was quite unusual for Cu. In view of the low g-values and the lack of any hyperﬁne structure (the spectra were recorded at 77 K, 9.5 GHz), he suggested that the signal might come from an interacting pair of Cu atoms in the cupric and cuprous states (Beinert et al., 1962). Another rejected proposal, invoking interaction between two paramagnetic species, was brought forward 17 years later by Froncisz et al. (1979). They interpreted the Cu hyperﬁne structure in the EPR signal of CuA (observed for the ﬁrst time at low frequency and 10 K) to originate from the interacting paramagnets CuA and cytochrome a. The third time this proposal was made by Kroneck et al. (1988, 1989), N2OR was fortunately available to show what a Cu pair might look like. An absorption band around 830 nm, which has been known for a long time to be due to CuA (Wharton and Gibson, 1966), can be used to monitor COX activity in vivo (Jo¨bsis-VanderVliet et al., 1988). Because of the dominance of absorption from haem groups in the UV–vis spectra of COX, it was not until the experiments of Thomson, Greenwood and colleagues (Greenwood et al., 1983, 1988; Thomson et al., 1986) that the complete MCD spectrum of CuA was obtained. It reveals the presence of two intense,

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oppositely polarized transitions in the visible region with the polarization direction perpendicular to the gmax direction, which distinguishes CuA from any other Cu compound studied until then. The two Cu ions in CuA, each one coordinated to a histidine, are bridged by the thiolate sulfurs of two cysteine residues. Weak axial ligands, such as the thioether sulfur of methionine and carbonyl oxygens, are also present. These structural elements are responsible for the unique optical and EPR features and the functional properties including the reduction potential, the low energy of reorganization and the fast rate of electron transfer (Kroneck, 2001). The similarity of numerous spectroscopic features between the CuA-carrying enzymes N2OR and COX is reﬂected at the level of the amino acid sequence in a short region constituting the CuA-binding domain (Zumft et al., 1992). More recently, the NO reductase of B. azotoformans was identiﬁed as the third example of a CuA-containing enzyme (Suharti et al., 2001). Early attempts to model the spectroscopic properties by biomimetic CuS2N2 complexes (Toftlund et al., 1985) failed to produce compounds with EPR, optical or MCD spectra similar to those of the CuA site of COX. Harding et al. (1991) prepared a mixed-valence complex [Cu2L]3+ that was later structurally and spectroscopically characterized (Barr et al., 1993; Farrar et al., 1995b). The complex [Cu2(LiPrdacoS)2]+ is the ﬁrst example of a structurally characterized dithiolate-bridged mixed-valence Cu dimer, speciﬁcally designed to mimic the physical properties of the CuA site (Houser and Tolman, 1995; Houser et al., 1996). The existence of the [Cu(m-RS)2Cu]+ core had been discussed already by Hemmerich et al. (1966), and was proposed as a transient species in Cu thiolate systems (Hemmerich, 1966). More recently, a series of fully delocalized [Cu(1.5+) y Cu(1.5+)] mixed-valence complexes has been assembled, having oxygen donor bonds (LeCloux et al., 1998), a 1,8-naphthyridine-based dinucleating ligand (He and Lippard, 2000), two ureate bridging ligands (Gupta et al., 2002) or the novel Cu2N2 diamond core (Harkins and Peters, 2004). 6.1.2. Cupredoxin Fold and Loop-Directed Mutagenesis The structure of the CuA centre is homologous to the cupredoxin fold that includes both the blue Cu proteins and the CuA electron transfer centres (Adman, 1991; Dennison and Canters, 1996; Kroneck, 2001; Vila and Fernandez, 2001; Lu, 2003). CD spectra of the CuA domain in COX document the presence of the cupredoxin fold, a Greek key b-barrel and common structural motif in small, type-1 Cu proteins of bacteria and plants (Williams, et al., 1999) (Fig. 5). This structural relationship is crucial for engineering by loop-directed mutagenesis the CuA dinuclear centre into the type-1 Cu proteins amicyanin

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Figure 5 Ribbon representation of the cupredoxin fold of CuA azurin from P. aeruginosa (PDB 1CC3).

(Dennison et al., 1995) and azurin (Hay et al., 1996; Robinson et al., 1999) (Fig. 6). A tetragonal mononuclear Cu(II)S(Cys) intermediate was proposed for the incorporation of the Cu2+ ion into apo-CuA azurin (Wang et al., 1999). Furthermore, the CuA redox site of bacterial COX of Pa. denitrificans was transformed into a mononuclear type-1 Cu centre by changing one bridging cysteine ligand to serine (Zickermann et al., 1997). At the same time it became possible to express the CuA-carrying subunit II of the membrane-bound enzyme COX as a water-soluble fragment in a host organism. Since the haem groups and CuB are located in the membrane part, optical and EPR spectra unimpaired by other chromophores were thus obtained (von Wachenfeldt et al., 1994; Larsson et al., 1995; Wilmanns et al., 1995; Farrar et al., 1996; Slutter et al., 1996). 6.1.3. Spectroscopic Properties and Electronic Structure Interest in CuA is directly related to its unique optical and EPR properties (Fig. 2). Multifrequency EPR spectroscopy of N2OR (Fig. 7) provides

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Figure 6 Schematic diagram of the engineered CuA site in azurin from P. aeruginosa. (A) View of the Cu2S2 rhomb; (B) side view of the CuA site (numbering as of PDB 1CC3). (See Colour Plate Section in back of this volume.)

evidence that two Cu atoms per enzyme subunit are organized in a binuclear mixed-valent centre, properties that are also intrinsic to the CuA centre of COX (Kroneck et al., 1988, 1990; Antholine et al., 1992). The crystal structures of COX and of N2OR conﬁrm the binuclear nature of CuA (Iwata et al., 1995; Tsukihara et al., 1995; Brown et al., 2000b; Svensson-Ek et al., 2002; Haltia et al., 2003). Spectroscopic studies by EPR, electron nuclear double resonance (ENDOR), electron spin echo envelope modulation (ESEEM), MCD, resonance Raman and X-ray absorption (Dooley et al., 1987, 1991; Jin et al., 1989; Andrew et al., 1994; Farrar et al., 1996, 1998; Neese et al., 1996, 1998; Charnock et al., 2000; Epel et al., 2002) have provided a clear picture of the electronic properties of the CuA site of N2OR, and the interactions of the

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Figure 7 EPR spectra of N2OR I recorded at 15 K and frequencies (GHz): S, 2.9; C, 4.5; X, 9.4; and Q, 34.5 (Neese, 1997).

metal atoms with neighbouring amino acids. A comparison of the deduced primary structures from bacterial nosZ genes and the CuA-binding consensus of the COX subunit II resulted in a set of candidate CuA ligands. In the absence of a crystal structure of N2OR, site-directed mutagenesis was crucial to identify the ligands of the CuA centre in the enzyme from P. stutzeri (Charnock et al., 2000). EXAFS associated with the Cu K-edge spectrum was used to deduce the local environment of the Cu atoms. An earlier Cu K-edge EXAFS study of the soluble CuA fragment from COX of Bacillus subtilis indicated a directly bonded CuCu unit with a CuCu distance of ca. 2.5 A˚ (Blackburn et al., 1995). Each Cu was coordinated to one histidine and two cysteine residues, and a CuCu metal bond was suggested. Based on calculations of the electronic structure and spectroscopic properties of the CuA centre the model structure consisted of a dimer of two type-1 Cu centres, with no bridging ligands but a direct CuCu bond (Larsson et al., 1995; Ramirez et al., 1995). Cu K-edge EXAFS studies on mixed-valent [Cu(1.5+) y Cu(1.5+)] and reduced [Cu(1+) y Cu(1+)] species reﬁned the structure of the Cu2S2 core to a CuCu distance of 2.43 A˚ for the mixed-valence, and 2.51 A˚ for the reduced centre in CuA fragments of COX from B. subtilis and Thermus thermophilus (Blackburn et al., 1997). The Cu K-edge X-ray absorption

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spectrum of bovine COX also showed evidence for a binuclear Cu centre (Henkel et al., 1995). N2OR has an EPR spectrum with a 7-line hyperﬁne pattern in the g|| region when recorded at 9.30 GHz and 10 K (Coyle et al., 1985) (Figs. 2 and 7). Multifrequency EPR spectroscopy performed at 2.5–35 GHz, g-factor analysis, and computer simulation of the EPR spectra led to the interpretation of the cogent features to arise from a binuclear mixed-valence S ¼ 1/2 site, i.e. a 2Cu(1.5+)2SCys,2NHis core (Figs. 3, 4 and 6), as opposed to the mononuclear type-1 Cu(II)SCys,2NHis,SMet centre (Antholine et al., 1992; Malmstro¨m and Aasa, 1993). The suggestion of a binuclear mixed-valent nature for CuA in COX, based on EPR spectroscopic investigations, initially generated controversy (Li et al., 1989; Malmstro¨m and Aasa, 1993), which was settled with the crystallographic structures of COX from Pa. denitrificans and bovine heart (Iwata et al., 1995; Tsukihara et al., 1995), and an engineered binuclear Cu centre from the membrane-exposed domain of a quinol oxidase complex (Wilmanns et al., 1995). Notice that the X-band EPR spectra of bacterial and mammalian COX, as well as some of the engineered soluble CuA fragments, do not show a well-resolved 7-line pattern in the gJ region around 2.18 as seen with N2OR (Beinert et al., 1962; Coyle et al., 1985; Riester et al., 1989; Antholine et al., 1992; von Wachenfeldt et al., 1994; Farrar et al., 1995a; Neese et al., 1996). Enhanced resolution is accessible by the second harmonic display, or by pseudomodulation treatment of EPR spectra (Antholine et al., 1992). EPR spectra of either 63Cu- or 65Cu-enriched N2OR established the mixed-valence delocalized conﬁguration. The EPR spectrum after incorporation of both 65Cu and 15N-histidine into N2OR improved the resolution of the Cu hyperﬁne lines, but no superhyperﬁne lines from nitrogen and proton couplings were resolved. Based on high-resolution X-ray structural data of the CuA centre (Wilmanns et al., 1995), molecular orbital calculations on a sulfur-bridged [(NH3)Cu(1.5+)(SCH3)2Cu(1.5+)(NH3)]+ core were used to interpret the EPR data. The calculated spin distribution shows an excellent agreement with the values derived from the analysis of the Cu hyperﬁne structure (Neese et al., 1996). ENDOR spectra for the [Cu(1.5+) y Cu(1.5+)], S ¼ 1/2 site were obtained of the 63Cu-, 65Cu- or 65 Cu- and 15N-histidine-enriched enzyme (Neese et al., 1998; Epel et al., 2002). The 14N, 15N isotopic substitution allows for an unambiguous deconvolution of proton and nitrogen hyperﬁne couplings in the spectra. A single-nitrogen coupling with an average value of 12.970.2 MHz was detected. The anisotropy in the nitrogen hyperﬁne values is characteristic for imidazole bound to Cu. The [(NH3)Cu(1.5+)(SCH3)2Cu(1.5+)(NH3)]+ core structure was predicted to be similar to the crystallographically

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determined CuA structure (Wilmanns et al., 1995), and distinct from the CuA structure of Pa. denitrificans (Iwata et al., 1995). The analysis of the g-values and Cu hyperﬁne couplings suggests 15–20% spin density on each Cu ion in CuA, and reveals that the interpretation of the gmax shift is only possible under the assumption of a low-lying excited state. The small value of the Cu-hyperﬁne splitting indicates a high degree of covalent delocalization of the unpaired spin in direction of the ligands, a situation which is reminiscent of that found in type-1 Cu centres (George et al., 1993; Shadle et al., 1993; Solomon and Lowery, 1993). The position of CuA in the Peisach–Blumberg plot of gJ versus AJ (Va¨nngard, 1972; Peisach and Blumberg, 1974) is primarily caused by the apparent small value of AJ. However, a correct comparison between the delocalized mixed-valence site and mononuclear Cu centres in this plot requires multiplication of the AJ value by two, which is directly related to the fact that the unpaired electron resides equal time on each half of the dimer. The position of CuA in the Peisach–Blumberg diagram comes closer then to the domain of type-1 Cu, being displaced only slightly because of its low gJ. Considering the magnitude of gJ in terms of subsite g-tensor noncolinearities, a gmax value of 2.20 (for a hypothetical monomer) is more appropriate. The remaining differences may result from a more pronounced covalency of the purple CuA centre compared to the blue type-1 Cu centre. It appears that, even though each Cu atom is coordinated by two thiolate sulfurs from cysteine, the reduction of the g and ACu values is approximately equivalent to a situation where each Cu atom has only one sulfur in its coordination sphere. In the CuA centre the unpaired spin is almost equally distributed over the Cu and S atoms in the electronic ground state, and there is only 3–5% spin density on the nitrogen donors of CuA. Thus, CuA in its electronic ground state, is viewed best as a completely delocalized radical in which the delocalization occurs via the in-plane s/p framework rather than the out-of-plane p framework as for delocalized organic radicals. This situation may be described by the resonance structures [CuIIRSRSCuI2CuIRSRSCuI2 CuIRSRSCuI2CuIRSRSCuII] (Neese, 1997). Based on the spectroscopic studies and the X-ray structures of CuA proteins and model complexes, theoretical calculations have been carried out ﬁrst by Neese et al. (1996) and Farrar et al. (1996) to develop a molecular orbital picture of the dinuclear mixed-valent centre, which was followed by similar calculations from other groups (Gamelin et al., 1998; Randall et al., 2000; DeBeer George et al., 2001; Olsson and Ryde, 2001). An unusual physical property of the CuA site is the extremely fast relaxation for which possible mechanisms have been advanced. Pure absorption spectra were obtained for CuA by multiquantum EPR spectroscopy (Mchaourab

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et al., 1993). Saturation recovery data recorded at 6–25 K conﬁrm that the intrinsic electron spin-lattice relaxation time, T1, for N2OR is unusually short for Cu centres. The short T1 is attributed to the vibrational modes of type-1 and/or the metal–metal interaction in [Cu(1.5+) y Cu(1.5+)] (Pfenninger et al., 1995). Extensive assignments have been achieved of the 1H-NMR spectra of soluble CuA-containing COX fragments from Thermus thermophilus (Bertini et al., 1996), Pa. denitrificans (Luchinat et al., 1997), and Paracoccus versutus (Salgado et al., 1998), from CuA-amicyanin (Dennison et al., 1997) and also of N2OR from P. stutzeri (Holz et al., 1999). UV–vis, CD and MCD spectra of the mixed-valence CuA site in N2OR have been measured in a range of 33,000–5000 cm1. These spectra differ signiﬁcantly from the spectra of the well-characterized blue type-1, nonblue type-2 and EPR-silent type-3 Cu sites. The purple colour of the CuA site is the result of predominant S(Cys)-Cu charge transfer bands at E480 nm (E21,000 cm1), 530 nm (E19,000 cm1) and a class III mixed-valence charge transfer band in the near infrared region around 800 nm (E13,400 cm1). These features dominate absorption spectra recorded at 5 K, but several weaker bands at both lower and higher energies have also been detected. The MCD spectra at 4.2 K are dominated by intense, oppositely signed bands and a negatively signed band in this spectral region. The MCD spectra are in good agreement with those of COX from bovine heart. An assignment of the electronic spectrum has been made in terms of a covalent planar core [Cu(RS)2Cu]+, with a CuS distance of 2.2 A˚, a CuCu distance of 2.5 A˚ and a CuSCu angle of 701. A delocalization energy of E4500 cm1 estimated from the data is at least one order of magnitude larger than the vibrational energies of the core, as expected for a stable class III mixed-valence site (Farrar et al., 1996, 1998). 6.1.4. Cu– Cu Interaction and Metallic Bond The CuCu distances among the CuA proteins are close to 2.4–2.5 A˚ (Iwata et al., 1995; Tsukihara et al., 1995; Wilmanns et al., 1995). The mean value of E2.26 A˚ for the CuS bond is in the range found in Cu–thiolate complexes (Fox et al., 1993) and longer than the short CuS bond of E2.11 A˚ observed in the oxidized type-1 Cu sites (Collyer et al., 1990; Chapman, 1991; Nar et al., 1991; Durley et al., 1993; Redinbo et al., 1993; Vakoufari et al., 1994). In all CuA centres the CuSCu angle falls in the range of 65–701. The CuN distances show a rather large scatter of 70.22 A˚ with the values reported for CyoA below and that for COX of Pa. denitrificans above the 1.95–2.05-A˚ distance commonly found in copper–nitrogen complexes (Hathaway, 1983). The CSSC dihedral angle differs by more than 401 in the structures reported for CuA in CyoA and Pa. denitrificans. Relative to

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the CuA centres found in proteins, the model complex [Cu2(LiPrdacoS)2]+ (Houser et al., 1996) shows with 2.93 A˚ a much larger CuCu distance and also a larger CuSCu angle of 801. More recently, the X-ray structure of the engineered CuA site in azurin from P. aeruginosa was reﬁned to 1.65-A˚ resolution (Fig. 6). Two azurin molecules in the asymmetric unit slightly differ in their CuCu distances (2.42 versus 2.35 A˚), the angular position of the two histidine imidazole rings with respect to the [Cu(m-RS)2Cu]+core plane, and the distance between Cu and the axial ligand. It appears, thus, that the variation in ﬁne structure of different CuA centres is correlated with the angular positions of the two histidine rings. From these positions one can predict the relative axial ligand interactions which are responsible for modulating the CuCu distance and the electron transfer properties of the various CuA sites (Robinson et al., 1999). Some discussion centres on whether a CuCu bond exists in CuA (Randall et al., 2000; DeBeer George et al., 2001; Olsson and Ryde, 2001). Convenient, albeit not objective, theoretical measures for bond strength are indices such as the Mulliken overlap population (Mulliken, 1955) or the Wiberg bond strength (Wiberg, 1968). Both parameters indicate that the CuCu bond strength is about one-ﬁfth that of the CuS bond. The CuCu stretching frequencies and Cu quadrupole interaction are other indicators of the CuCu bond strength. Of experiments that have been performed towards this objective (Wallace-Williams et al., 1996), none points to a particularly strong CuCu bond (Neese, 1997). In view of the fact that all antibonding molecular orbitals, except one, are completely ﬁlled, the stability of this electron-rich cluster is surprisingly high. Bonding contributions from Cu 4s, 4p and sulfur 3d orbitals (Mehrotra and Hoffmann, 1978; Merz and Hoffmann, 1988) are considered important. Donation of electrons from the sulfur into the empty Cu orbitals as well as back bonding of the ﬁlled Cu orbitals to the empty sulfur 3d orbitals may contribute to the stabilization of the cluster. The mixed-valence state of CuA is energetically very favourable. The oxidation to the Cu(II)Cu(II) form has not been achieved for any CuA centre to date. Remarkably, the CuA centre is formed from apoprotein and Cu(II) with N2OR and other CuA-containing proteins (Coyle et al., 1985; Riester et al., 1989; van der Oost et al., 1992; Dennison et al., 1995; Slutter et al., 1996). Besides, the mixed-valence anion [Cu2S2] is formed in the gas phase by laser ablation from CuS (Fisher et al., 1996). 6.1.5. Recombinant CuA Centres of N2O Reductase Recombinant N2OR derivatives with His494Cys, Asp580Ser or Met629Cys substitutions exhibit the CuA-type electronic spectrum with absorbance

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features at 480–530 nm, and a broad maximum around 790 nm (Fig. 2). Only the Asp580Ser recombinant protein shows a weak contribution to the overall spectrum of a species absorbing at 650 nm. The spectra of the mutant proteins show slight shifts in the positions of absorption bands when compared to the wild-type protein (Charnock et al., 2000). In agreement with the electronic spectra, the EPR parameters at X-band for the His494Cys, Asp580Ser and Met629Cys recombinant enzymes clearly document the presence of the mixed-valent, S ¼ 1/2 CuA site as the dominant paramagnetic species. In the recombinant CyoA fragment CuA does not form when Met118 is mutated to glycine (Kelly et al., 1993), while a Met227Ile substitution in CuA of Pa. denitrificans results in a trapped-valence state and shows reduced electron transfer activity (Zickermann et al., 1995). Thus, this residue must inﬂuence the structural integrity and function of CuA centres signiﬁcantly, similar to the methionine ligand in type-1 Cu. Apparently, it affects the reduction potential of the site (Guckert et al., 1995), but has only a minor inﬂuence on the spectroscopic properties (Lowery and Solomon, 1992; Solomon et al., 1992). The substitutions Met629Cys and Asp580Ser of N2OR are not expected to affect the CuZ catalytic site. However, the conversion of the electronic absorption characteristics of the recombinant enzymes to those of a CuAtype protein indicate that alterations in these amino acid positions also affect the structure of the protein outside the CuA domain. This becomes even more evident on substituting the cysteine and histidine ligands of the CuA centre. Histidine substituted by glycine to remove a binding site from Cu, and cysteine substituted by aspartate to provide a potential alternative ligand as a bridging oxygen function (Charnock et al., 2000), result in the complete (His626Gly, Cys618Asp and Cys622Asp) or nearly complete (His583Gly) loss of enzyme activity. The metal is bound only weakly in the His626Gly and Cys618Asp derivatives; the His626Gly substitution results in a colourless protein with E2 Cu/homodimer. It is unexpected that the collapse of the CuA centre, by removing one cysteine bridge or by introducing the His626Gly exchange, would affect the Cu occupancy of the CuZ catalytic site. However, we now know from the crystal structure that several CuA and CuZ ligands participate in intermonomer contacts (Fig. 4). Thus, perturbation of the CuA site, through intersubunit or interdomain interactions, can lead to a metal-depleted protein. The His583Gly and Cys622Asp mutant proteins, in contrast, retain their Cu, but show signiﬁcant changes in electronic and EPR spectra (Charnock et al., 2000). Both derivatives exhibit a pronounced absorption maximum around 650 nm, similar to that described for dithionite-reduced N2OR and the less active form of the enzyme (Coyle et al., 1985). For the Cys622Asp

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derivative the extinction coefﬁcient at 654 nm (7400 M1 cm1) is in good agreement with the value calculated for reduced N2OR (form III) at this wavelength (Riester et al., 1989). In the EPR spectra of the two protein variants the multiline feature of CuA at g ¼ 2.18 is no longer resolved at X-band. Such perturbations in the EPR spectra are also observed for the enzyme treated with azide, CO or NO (Riester et al., 1989). The His583Gly substitution in N2OR resembles the azurin mutant His46Gly of P. aeruginosa (van Pouderoyen et al., 1996). Even though His46 is buried in azurin, the cavity created in the protein by substituting histidine with glycine can be ﬁlled with an external ligand. Although the structure of this recombinant azurin is not changed, its spectroscopic properties are strongly affected forming a mixture of type-1 and type-2 Cu sites. According to the structural model of the P. stutzeri enzyme (Charnock et al., 2000), His626 is close to the surface, whereas His583 is buried deeper in the CuA fold. A similar arrangement is seen in the crystal structure of Pa. denitrificans N2OR (Haltia et al., 2003). 6.1.6. Electron Transfer Electron transfer mediated by CuA is very efﬁcient. The reason for utilizing a binuclear electron transfer centre in COX and N2OR, instead of mononuclear type-1 Cu, is thought to be the unidirectional electron transfer through the site, or the lower energy of reorganization. The rate constant, kET, of long-range intramolecular electron transfer in an engineered azurin mutant with purple CuA is almost threefold higher than for the same process in the wild-type protein (650760 versus 250720 s1, 298 K, pH 5.1), in spite of a smaller driving force (0.69 versus 0.76 eV) (Farver et al., 1999). For the electron transfer pathway in COX from CuA to haem a, His204 (by bovine enzyme numbering) has been identiﬁed (Ramirez et al., 1995). Williams et al. (1997), on the other hand, have suggested one of the bridging cysteine sulfurs, Cys196, because of the large amount of S(Cys) character in the singly occupied molecular orbital (SOMO) of CuA, analogous to the situation in blue Cu proteins (Regan et al., 1998). Electronic structure descriptions show that CuCu compression and removal of axial ligands are critical determinants of the orbital ground state in the mixed-valent CuA dimer. The weakened axial interaction in CuA appears to parallel the mechanism for protein control of electron transfer function observed in the blue type-1 Cu proteins (Gamelin et al., 1998). A pH-dependent transition between delocalized and trapped mixedvalence states of an engineered CuA centre in azurin has been described more recently. Protonation of one of the histidine ligands, at pH 4.0, results

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not only in a trapped-valence state but also in the increase of reduction potential from 160 to 340 mV (Hwang and Lu, 2004). Exchange of the histidine ligand by asparagine or glycine changes the EPR hyperﬁne signature from a 7- to a 4-line pattern, consistent with a trapped-valence site, although ENDOR data indicate that the electronic structure of the CuA core remains unchanged (Lukoyanov et al., 2002). On the other hand, by substituting in engineered CuA of azurin the highly conserved axial methionine by glutamate, asparate, leucine or glutamine, it was found that the axial ligand substitution affects much less the reduction potential of CuA than that of the mononuclear type-1 Cu (Hwang et al., 2005).

6.2. CuZ Centre 6.2.1. Development of the Field The existence of a second Cu site in N2OR can be traced back to the ﬁrst isolation of the enzyme (Zumft and Matsubara, 1982). Addition of dithionite to the purple enzyme from P. stutzeri produced a blue species (lmaxE640 nm) with an EPR signal clearly different from the signal observed for N2OR as isolated. The Cu content of this ﬁrst preparation accounted for E8 Cu/120,000 Mr, an amount that has been determined for numerous preparations of the enzyme from various sources and by different laboratories since (Zumft and Kroneck, 1996; Wunsch et al., 2005, and references therein). The second Cu site was termed as CuZ and turned out to be the ﬁrst biological example of an inorganic sulﬁde ion as part of a multicopper cluster (Farrar et al., 1991; Rasmussen et al., 2000). From the X-ray structure it became clear that N2OR carries 12 Cu atoms per dimer: four atoms in two dinuclear CuA centres, and eight atoms in two tetranuclear CuZ centres (Brown et al., 2000b). In spite of the differences in Cu content, the analytical data on the various preparations of N2OR, including UV–vis and EPR parameters, correlate well. Because of dimer and Cu-site interactions in N2OR it seems that metal centre heterogeneity is low, even in partially Cu-depleted enzyme species. An unusual EPR signal, with g-values at 2.18 and 2.06 clearly distinct from both CuA and type-1 Cu EPR signals, remains even in the presence of excess reductant (Coyle et al., 1985). This state of the catalytic centre CuZ was lately called ‘resting CuZ’, with the CuA sites of N2OR in the spectroscopically silent Cu(I)Cu(I) state, and CuZ in the [Cu4S]3+, S ¼ 1/2 state (Chen et al., 2004). The resonance Raman spectrum of the blue form of the enzyme is essentially identical with that of a type-1 Cu(II) centre

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(Dooley et al., 1987). The conversion of the purple enzyme to the blue species is fully reversible in the absence of O2, using dithionite as the reductant and ferricyanide as the oxidant (Riester et al., 1989). Two kinetically distinct phases of the reaction with stoichiometric amounts of dithionite are observed: in the fast phase the absorbance at 540 nm almost disappears within seconds, mainly due to the reduction of CuA, whereas in the slower phase a blue species is generated in the course of minutes. Interestingly, the position of the absorption maximum of the blue species shifts signiﬁcantly depending on the reductant employed: maxima are positioned at 650, 648 and 665 nm with dithionite, sulﬁde, and 2-mercaptoethanol or glutathione, respectively. These properties are not seen in a catalytically inactive form of N2OR obtained from strain MK402 with defective Cu centre biosynthesis, which carries a transposon Tn5 insertion in the nosD gene. This mutant protein proved invaluable for unraveling the spectroscopic features of the CuA and CuZ centres. Thus, through a comparison of the UV–vis, EPR and EXAFS spectra of the wild-type and mutant forms of the enzyme, the individual contributions of CuA and CuZ to the spectral features could be determined. The presence of a second paramagnetic species, in addition to the CuA centre, became apparent from the Q-band EPR spectra of N2OR as isolated. It was absent in the mutant form (Antholine et al., 1998). The amount of this CuZ* species (Farrar et al., 1991; Alvarez et al., 2001; Rasmussen et al., 2002) depends on the preparation method of N2OR reductase and is minimal in the enzyme from P. stutzeri when isolated under the exclusion of O2. This second paramagnetic species in N2OR as isolated correlates with a band in the UV–vis spectrum centred around 650 nm (Neese, 1997) (Fig. 8). It was also noticed that the persistence of the blue form of N2OR in the presence of strong reductants points towards a catalytic Cu site in a ‘reduced’ state, stabilized by thiol or disulﬁde sulfur with substantial spin density delocalized onto sulfur (Riester et al., 1989). Based on CD spectra, a highly covalent [Cu(II)-S(cys)2Cu(I)-S(cys)] site was proposed (Dooley et al., 1991). In a ﬁrst description, the catalytic site was seen as a binuclear centre which could exist in an active form, CuZ, which also had a cysteine ligand, and an inactive form, CuZ* (Farrar et al., 1991). From sequence alignments mainly histidine coordination was assumed for CuZ, but the binding of one cysteine residue, from outside of the CuA domain, was also seen feasible. This candidate cysteine was later shown not to be a ligand of CuZ (Zumft et al., 1992; Dreusch et al., 1996). In the very ﬁrst crystal structure of N2OR, a sulfur-containing ligand was not yet assigned to the CuZ centre (Brown et al., 2000b). However, elemental analysis and resonance Raman spectroscopy of isotopically labelled N2OR demonstrated the

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Figure 8 Deconvolution of the room temperature UV–vis spectrum of N2OR. (A) N2OR I, native enzyme as isolated; (B) N2OR V, nosD mutant protein; (C) difference spectrum of A and B. Insert, resolution of spectrum C into two Gaussian bands centred at 650 nm (15,300 cm1) and 540 nm (18,300 cm1) (Neese, 1997).

presence of one acid-labile sulfur ligand (Rasmussen et al., 2000; Alvarez et al., 2001) and led to the current view of the catalytic site. 6.2.2. Spectroscopic Properties and Structure The CuZ site comprises four copper ions, CuI–CuIV, arranged in a tetranuclear m4-sulﬁde-bridged Cu cluster solely coordinated by histidine residues (Figs. 3 and 4). CuZ is located in the bulk part N-terminal domain of N2OR, which adopts a seven-bladed b-propeller (Brown et al., 2000a,b; Haltia et al., 2003). It appears that CuI is the predominantly oxidized Cu atom, which is consistent with CuI having a four-coordinate structure and the other three Cu atoms lower coordination numbers. Thus, the CuI centre sits roughly above a plane built by the three Cu centres CuII, CuIII and CuIV, and the inorganic sulfur (Oganesyan et al., 2004). Biomimetic complexes, featuring this structural unit with Cu in higher oxidation levels, have not been reported (Helton et al., 2003; Brown et al., 2004). Most recently, however, the synthesis of a trinuclear [Cu3(m-S)2]3+ cluster coordinated to N-donor ligands was achieved. Remarkably, this complex shows a low-energy band with lmax at 605 nm (e ¼ 3000–4000 M1 cm1), which resembles the spectral feature at 640 nm observed for the resting CuZ state (Brown et al., 2005).

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The spectroscopic properties of the CuZ centre in N2OR from Ma. hydrocarbonoclasticus (puriﬁed in the presence of O2 and with an X-ray structure available) and from Paracoccus pantotrophus (puriﬁed in the presence and absence of O2, no X-ray structures available of both forms) have been studied in greater detail (Chen et al., 2004; Oganesyan et al., 2004). The Cu4 core has approximate Cs symmetry with CuISCuII deﬁning the mirror plane (Figs. 3 and 4). The average CuS distance is E2.3 A˚, and the CuCu distances vary from E3.4 to 2.6 A˚. The CuISCuII angle is E1601, all other CuSCu angles are close to 901. The high-resolution structure of the enzyme from Pa. denitrificans shows a weakly bound ligand, most likely water derived (Haltia et al., 2003). Its nature has not been assigned. Whether the two X-ray structures of ‘blue’ N2OR represent the structure of the active ‘purple’ enzyme, puriﬁed in the absence of O2, remains to be seen in view of the signiﬁcant spectral and catalytic differences between the various forms of N2OR (Zumft and Kroneck, 1996; Oganesyan et al., 2004; Wunsch et al., 2005). The resting state of CuZ is assigned to a [Cu4S]3+, S ¼ 1/2 centre based on MCD, EXAFS and EPR results (Chen et al., 2004; Oganesyan et al., 2004). The gJ value of resting CuZ is rather small, which could be due to extensive covalent character in the metal–ligand bonding or high d–d transition energies as pointed out earlier (Riester et al., 1989). X-ray absorption spectroscopy at the S and Cu K-edges was used to determine the Cu oxidation states and the bridging-sulﬁde covalency in the ground state of resting CuZ in Ma. hydrocarbonoclasticus. Compared to CuA with approximately 46% S character in the ground state, the corresponding sulfur character value for resting CuZ is with 15–22% much smaller (Chen et al., 2004). 6.2.3. Electron Transfer and Activation of N2O N2OR from P. stutzeri is activated by base treatment (Coyle et al., 1985), or incubation with CO (Riester et al., 1989). The mechanism of activation is unknown, although one may speculate that under those conditions the CuICuIV edge of the cluster gets altered. This edge has an additionally bound ligand ‘X’ – most likely an oxygen ligand in N2OR II – and is thought to bind the substrate (Fig. 3). Furthermore, the enzyme from different sources is activated in vitro by reduction with the redox dyes methyl viologen or benzyl viologen (Kristjansson and Hollocher, 1980; Prudeˆncio et al., 2000; Ghosh et al., 2003; Chan et al., 2004). From these studies it appears that the catalytically relevant form of CuZ is the fully reduced [Cu4S]2+, S ¼ 0 state. The UV-vis spectrum of this form is similar to the spectrum of the dithionite-reduced enzyme, but the intensity of the absorption at

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lE650 nm is signiﬁcantly lower. Incubation of the reductively activated N2OR from Achromobacter cycloclastes with N2O-saturated buffer leads to a partially oxidized enzyme as shown by optical and EPR spectroscopy. The activated form of N2OR converts N2O to product and thus represents the ﬁrst report of substrate reduction in vitro followed spectroscopically (Chan et al., 2004). The outcome of computational methods suggests that in the lowest energy structure of the CuZ(4CuI)-N2O complex N2O binds at the CuICuIV edge in a bent, m-1,3 bridging mode with the terminal N atom coordinating to CuI (Fig. 3) (Chen et al., 2004). By Cu-N2O p* backbonding interaction the NO bond will be weakened signiﬁcantly, which might facilitate the direct NO bond cleavage via simultaneous transfer of two electrons from CuZ to the m-1,3 bridged N2O. Remember that in the [Ru(NH3)5(N2O)]2+ complex N2O is preferentially coordinated end-on by its terminal N atom (Paulat et al., 2004). Since N2O is a weak s and p donor, a more stable metal–NNO bond is achieved by an increase in p back bonding, as suggested for the tetranuclear m4-sulﬁde-bridged CuZ cluster. Unsolved still is the structure of CuZ* and its potential involvement in catalysis. When O2 is present during the preparation of N2OR, CuZ* is formed. CuZ* gives an EPR signal but does not show redox activity with either dithionite or ferricyanide. In N2OR, prepared under the exclusion of O2, the catalytic Cu site is capable of undergoing a one-electron reversible redox cycle at Em ¼ +60 mV (Riester et al., 1989; Rasmussen et al., 2002; Oganesyan et al., 2004). The EPR-active redox state of CuZ* must be structurally similar to that of CuZ in the [Cu4S]3+, S ¼ 1/2 state, since spectroscopic properties, including EPR (X- and Q-bands), MCD and CD spectra, are closely similar (Neese, 1997; Alvarez et al., 2001; Rasmussen et al., 2002). An interesting development with regard to the role of the CuZ* centre comes from the study of an N2O-respiration negative, double mutant of the paralogous genes nosX and nirX of Pa. denitrificans. In spite of the absence of whole-cell N2O-reducing activity, puriﬁed N2OR from this mutant is active in the enzyme assay in vitro, and exhibits the spectroscopic features of CuA and the redox-inert CuZ* form of the catalytic centre (Wunsch et al., 2005). While CuZ* is usually generated in N2OR when puriﬁed in the presence of air (Zumft and Kroneck, 1996), or in the case of Pa. pantotrophus by exposure of frozen cell extract to air (Rasmussen et al., 2002), it was now detected in a cellular context. Surprisingly, in spite of the exclusion of air, the double mutant yields a blue enzyme with the UV–vis spectrum featuring a band representative of CuZ* (lmaxE635 nm, eE4.8 mM1 cm1). Since the absence of the NosX function enhances CuZ* formation, this centre will have to be considered for the catalytic cycle. It also suggests that the assay

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in vitro, based on reduced viologens, might not probe the internal electron ﬂow between CuA and CuZ. The formation of a cytochrome c–N2OR complex was described most recently as obligatory for N2O reduction by the enzyme from Pa. pantotrophus. Remarkably, in the absence of cytochrome c, reduced N2OR ([Cu4S]3+, S ¼ 1/2 state and with dithionite present) is inert towards its substrate, even though sufﬁcient electron equivalents are stored to initiate a single turnover. In the presence of cytochrome c and N2O, however, the single turnover occurs after a lag phase. This ﬁnding is interpreted as a conformational change in N2OR speciﬁcally induced by the interaction with cytochrome c (Rasmussen et al., 2005). The source of cytochrome c in these experiments was horse heart, which points to a low selectivity of N2OR towards the electron donor (see also Section 11.1).

7. ORGANIZATION OF nos GENES, GENE EXPRESSION, REGULATION 7.1. Patterns in nos Gene Clusters The positional clustering of nos genes is largely conserved in the genomes of denitrifying bacteria. The strongest conserved pattern in the NGCs is a tricistronic nosDFY arrangement downstream of nosZ (Table 2). It is found in this arrangement even in the archaeon H. marismortui. In the singular case of D. hafniense one ﬁnds a nosDLYF arrangement. An equally strongly conserved pattern in N2O-respiring bacteria is the location of nosL downstream of nosY. Its constant association with other nos genes (missing only in the unﬁnished genomes of An. dehalogenans and Sa. ruber) is suggestive of an essential function provided by NosL. Rarely is an individual gene such as nosX or nosR transposed to a different site as seen in Cupriavidus spp., Burkholderia spp. and Ralstonia solanacearum. Sometimes more copies of nosF, nosY or nosL are present (Table 2). The functionality of these putatively paralogous genes needs investigation. The most conspicuous example of gene duplication is seen with nosL. In a few cases, the dominant nosDFYL gene pattern is interrupted by intercalations of nap homologues, nosC genes, or short open-reading frames (ORFs) encoding hypothetical proteins. Analysis of W. succinogenes resulted in the discovery of two homologues of nap genes, nosH and nosG, intercalated in the NGC. NosG proteins show similarity to the C-terminal part of NosR. They are thought to be involved in electron donation to

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N2OR (Simon et al., 2004). Homologous genes are present in Ca. fetus, De. aromatica, Magnetospirillum magnetotacticum and Thiomicrospira denitrificans. W. succinogenes has a complex nos gene arrangement. It is exactly duplicated in Tm. denitrificans, and is similar in Ca. fetus, all three bacteria being e-proteobacteria (Table 2). As Ca. fetus was reported to reduce N2O (Payne et al., 1982) its NGC, currently incompletely sequenced, is anticipated to carry also a nosZ gene. The NGCs of Azoarcus sp. EbN1, Burkholderia spp., Cupriavidus spp., Pa. denitrificans and several other denitrifying bacteria display one or two nosC genes. The deduced proteins exhibit a conserved CysXaaXaaCysHis sequence suggestive of a c-type cytochrome (see Section 11.1). When nosR is associated with the core nosZDFYL genes it is mostly located immediately upstream of nosZ. The order is turned around in the genera Azoarcus, Cupriavidus, Ralstonia and Thiobacillus. Deduced from its frequency in the NGCs, nosX is another important gene for denitriﬁers belonging to the a- and b-proteobacteria. The gene arrangements are identical in the genomes of different strains of a species, such as of P. aeruginosa PAO1 and UCBPP-PA14, Rhodopseudomonas palustris HaA2 and CGA009 and the serogroups A, B and C of N. meningitidis, represented by strains Z2491, MC58 and FAM18, respectively. On the other hand, the plasmid-borne NGC of Cupriavidus necator H16 (formerly R. eutropha) is absent from the 635-kb megaplasmid of R. eutropha strain JMP134. A BLAST search of the genome of strain JMP134 did not reveal any chromosomally located nos genes, although the strain should be able to denitrify due to the presence of nar, nir and nor genes. It is legitimate to ask whether the complete set of nos genes has been identiﬁed and whether another type of N2OR might exist, other than the modiﬁcation observed with W. succinogenes. The entire denitriﬁcation system consists of over 50 identiﬁed and classiﬁed genes, of which 10 are speciﬁc for N2O utilization (Zumft, 1997). The cluster nosDFYL is found in every N2O-reducing prokaryote, whereas NosR, nosX, nosC, nosG and nosH are distributed mostly according to taxonomic patterns and are not ubiquitous (Table 2). Strong autonomy for a plasmid-borne NGC does not support further candidate nos genes. The possibility of encoding catalytically active holoN2OR from heterologous nosRZDFY in Pseudomonas putida (though nonfunctional in vivo) also limits the number of essential nos genes. This conclusion is further supported from mutational evidence, but clearly is limited to the investigated model organisms. Beyond the Z-type enzyme and its assembly requirements, we do have, for principal reasons, the drawback that a different type N2OR, assuming there is one, cannot be recognized by comparative analysis. It can come only from

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the traditional approach of searching in an interesting organism for the enzyme activity and characterizing it in the same way that had been done for the current model organisms. Genomics provides us with the implication of another type of N2OR for certain archaea where N2O-metabolizing physiology lacks the information of the Z-type enzyme (see Section 8.3).

7.2. Gene Expression and Regulation 7.2.1. Promoter Studies and Anaerobic Gene Expression In the currently known gene clusters the nos genes are transcribed nearly exclusively in a single direction. Only nosX or nosCX are transcribed upstream of nosZ in the opposite direction in the NGCs of Burkholderia spp., Cupriavidus spp., R. solanacearum and Rf. ferrireducens. Transcriptional organizations within NGCs are shown in Fig. 9. The nos genes are arranged

Figure 9 Transcriptional organization of nos gene clusters. Mapped promoters (!) are indicated above the respective gene box; arrow, experimentally identiﬁed transcript; broken arrow, complementation group or putative transcriptional unit. Triangles represent sequence motifs for binding of Crp-Fnr type regulators DnrD, Dnr or Nnr; open triangle, half-site of Crp-Fnr motif.

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in P. stutzeri in three units of monocistronic nosR and nosZ, and the nosDFYLtatE operon (Cuypers et al., 1992; Vollack and Zumft, 2001; Honisch and Zumft, 2003). Three complementation groups, nosR, nosZ and nosDF(Y) are found in S. meliloti and indicate a transcriptional arrangement like that of P. stutzeri (Holloway et al., 1996). The NGC of Bradyrhizobium japonicum has an individual nosZ promoter responsive towards microaerobiosis and nitrate (Velasco et al., 2004). A single nosZ transcript was identiﬁed in P. fluorescens (Philippot et al., 2001). Combining these ﬁndings, the given gene arrangement, and the existence of binding motifs for a CrpFnr-type regulator in the promoter regions of nosZ and nosR, at least three transcriptional units are suggested for these bacteria. The transcriptional organization of the nos cluster of Pa. denitrificans comprises transcripts of nosCR, nosZ and nosDFYLX (R. J. M. van Spanning, personal. communication). Cotranscription of nosC and nosR may indicate a Nos-related function for the nosC product. In P. aeruginosa the nos genes are arranged in a single hexacistronic nosRZDFYL operon (Arai et al., 2003). The stoichiometry required for the different functions encoded in the polycistronic nos messages needs a control mechanism. The nos operons may be subjected to regulation of translational efﬁciency or the stability of distinct mRNA segments. The transcription of nosZ in P. stutzeri depends on the multidomain transmembrane protein NosR (see also Section 11.2). Tn5 insertions in the 50 region of the nosR gene cause the loss of the nosZ transcript, which led to the consideration of NosR as a regulatory component. NosR is also active in the transcription of the nosD operon (Honisch and Zumft, 2003). A nosR mutant of B. japonicum is ineffective in activating the nosZ promoter, which is manifested in the termination of nitrate denitriﬁcation with N2O instead of N2 (Velasco et al., 2004). In P. aeruginosa, NosR is not obligatorily required for nos gene expression, but disruption of nosR approximately halves transcriptional activity of the nosR promoter (Arai et al., 2003). In Pa. denitrificans a homologue of nosR, nirI, representing thus far a singular case among the denitrifying bacteria, has been studied for its role in the transcription of nirS, encoding the cytochrome cd1 nitrite reductase (Saunders et al., 1999). The nirI gene is under the control of the Crp-Fnr factor Nnr (a homologue of Dnr or DnrD; see below), and shows a distinct positional effect with respect to nirS expression, in so far as it cannot be complemented in trans. Against this experimental background the absence of features in NosR of a transcription factor is unexpected. The C-terminally located metal centres of NosR might have DNA-binding properties, but mutating or removing these regions hardly affects the cellular amount of NosZ (Wunsch and Zumft, 2005). NosR does not exhibit a winged helix-turn-helix domain

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like the membrane-bound transcription factor ToxR (Crawford et al., 2003); neither is it part of a two-component regulatory system, either of the classical type or the extracytoplasmic function sigma factors (Helmann, 2002). A mechanistic understanding of these observations will require further studies of how NosR-type proteins exert their apparent transcriptional effects. 7.2.2. NO Signalling via Crp-Fnr Transcription Factors The nos genes are under the control of transcription factors that belong to the Crp-Fnr family of regulators. They carry different mnemonics, such as Dnr, DnrD, Nnr or FixK2 but designate the same structural type of regulator (Ko¨rner et al., 2003). In P. stutzeri dnrD is part of an operon that also encodes DnrN, a haemerythrin-like protein. In response to denitrifying conditions, DnrD of P. stutzeri is also active in the expression of the nnrSorf247 operon, which is located downstream of the nosD operon (Honisch and Zumft, 2003). NnrS of the purple bacterium Rhodobacter sphaeroides, strain 2.3.4, is a haem–Cu oxidase that affects chemotactic behaviour towards nitrite (Bartnikas et al., 2002). nnrS orthologues are often associated with nos genes, but they also exist in nondenitrifying bacteria, which suggests for NnrS a more generalized function, not limited to denitriﬁcation. Aerobic cells of P. stutzeri transcribe their nos genes at a low level, which results in a small constitutive amount of N2OR. Under anaerobic, N2Orespiring or nitrate-denitrifying conditions nos gene expression is activated and leads to a manifold increase in cellular N2OR content. N2O is only a weak inducer of nosZ in several bacteria (Ko¨rner and Zumft, 1989; Richardson et al., 1991; Sabaty et al., 1999). In contrast, NO as the signal molecule strongly upregulates the nosR, nosZ and nosD promoters. The NO signal is probably processed via Dnr/DnrD/Nnr (van Spanning et al., 1999; Vollack and Zumft, 2001; Arai et al., 2003). nosZ expression in Rb. sphaeroides IL106 depends on one of the reduction products of nitrate, pointing here to NO as the candidate signal, too (Sabaty et al., 1999). nosZ of P. stutzeri has a complex promoter structure of six transcriptional start sites. The anaerobically active promoter, P3, has a potential recognition motif for DnrD binding. The nosR promoter, in turn, has DnrD recognition sites spaced in multiples of about 60 nucleotides at 137.5, +67.5 and +127.6 from the start of transcription (Cuypers et al., 1995). A signal transduction pathway may be operative in P. stutzeri, where NO derived from nitrate denitriﬁcation activates DnrD, which results in nosR expression and subsequent activation of nosZ and the nosD operon: NO-DnrD-nosR/ NosR-nosZ, nosD. In P. aeruginosa this is shortened to NO-Dnr-nosR.

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The mechanism of Dnr/DnrD/Nnr activation by NO is not resolved (Zumft, 2002). A signiﬁcant observation is that NO activates heterologous Nnr in Escherichia coli, which indicates a generalized mechanism not speciﬁc to the denitriﬁcation process (Hutchings et al., 2000). Mutational evidence for B. japonicum suggests a dependence of nos gene expression on the haembased O2-sensing two-component system FixLJ in cooperation with the CrpFnr regulator FixK2 (Velasco et al., 2004). FixK2 has been proposed to function solely in a concentration-dependent manner that is controlled by the hierarchically positioned FixLJ. Puriﬁed FixK2 itself does not carry a cofactor and is active at low O2 tension without requiring a further physiological signal (Mesa et al., 2005). Nevertheless, microaerobic conditions alone are not sufﬁcient to activate the nosZ promoter of B. japonicum; also needed are nitrate or an N oxide derived from it (Velasco et al., 2004). In P. aeruginosa, Dnr is under the control of the Fnr homologue and FeS protein Anr, functioning as the O2 sensor (Arai et al., 1997). The Fnr homologue, FnrP, is the O2 sensor in Pa. denitrificans (van Spanning et al., 1997). In P. stutzeri mutational inactivation of FnrA, a further homologue of Fnr, does not affect nos transcription and the question of an O2 signal for nos gene activation remains unanswered here.

8. EVOLUTIONARY ASPECTS AND PHYLOGENETIC RELATIONSHIPS 8.1. Phylogenetic Relationships among Nos Proteins Until very recently, structural information about N2OR came only from individual genetic analysis of N2O respiration and was known for a-, b- and g-proteobacteria. This picture is rapidly changing because of the expanding genome database to which currently six more principal taxonomic lineages, exhibiting inorganic N2O metabolism, have been added. Remarkably, all taxonomic groups, although phylogenetically distant and quite diverse, exhibit the Z-type folding pattern in their deduced N2ORs. Figure 10 shows the phylogenetic relationship among NosZ proteins from those bacteria for which a complete NosZ sequence is known. The protein tree shows three clades consisting of a-proteobacteria (represented by the Rhizobium–Paracoccus lineage), b-proteobacteria (represented by the Burkholderia–Ralstonia lineage), and the equally densely populated Pseudomonas– Colwellia clade formed by various members of the g-proteobacteria. The other groups, identiﬁed as nos genes-carrying species from their

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Figure 10 Unrooted phylogenetic tree of NosZ sequences. The sequences were retrieved from the NCBI database by identifying respective entries in the annotations and by BLAST searches (Altschul et al., 1997) of the core nucleic acid, protein and genome databases. Ongoing genome projects were screened by FASTA or BLAST algorithms depending on the hosting database server of TIGR Microbial Database, The Joint Genome Institute Microbial Genomics Database, The Wellcome Trust Sanger Institute, and ERGO Database of Integrated Genomics Inc. The data set excludes environmental N2OR fragments obtained from nosZ genes ampliﬁed by PCR. Sequences were processed by the multiple-alignment program ClustalX 1.83 (Thompson et al., 1994), using the Gonnet series as protein weight matrix and parameters set to 10 for gap opening, 0.2 for gap extension and divergent sequences delay at 30%. Subsequently, a protein distance matrix was calculated by the use of ProtDist component of the PHYLIP v. 3.65 package (Felsenstein, 1989). Phylogenetic afﬁliation was calculated by the neighbour-joining method (Neighbor program of PHYLIP), and displayed by the TreeView 1.6.6 software (Page, 1996). Numbers on branchings indicate percentages of partitions in an extended majority rule consensus tree based on a bootstrap with 1000 replicates, carried out with PHYLIP’s SeqBoot and Consense applications. The scheme of colours for the clusters of branches indicates the taxonomic group on the class or phylum level based on 16S rRNA. For strain speciﬁcations and abbreviations see Table 2; Mhyd, Ma. hydrocarbonoclasticus. (See Colour Plate Section in back of this volume.)

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genomes, form deeply branching isolated lineages in the NosZ tree. Most consist currently of only a single representative such as the archaeon H. marismortui, the gram-positive bacterium D. hafniense, Sa. ruber from the cytophaga–ﬂavobacter–bacteroides superphylum, Te. roseum from the green nonsulfur bacteria phylum, and the d-proteobacterium An. dehalogenans. The e-proteobacteria are represented by W. succinogenes and Tm. denitrificans. The NosZ sequence of W. succinogenes DSM1740 was corrected for the insertion element IS1302 (Simon et al., 2004); NosZ of Brucella melitensis 16 M, which is segmented by a stop codon, was joined into a single ORF as suggested in the databank annotation (accession number gi 37537970). Other fragments of NosZ sequences present in databanks were excluded from consideration. Two species of Azospirillum, represented by fragments of E200 amino acids, do not cluster in their expected taxonomic position (Rich et al., 2003) for reasons that are unclear. A NosZ fragment of 512 amino acids from Pa. pantotrophus is more similar to Pseudomonas spp. (Scala and Kerkhof, 1998) than to another short NosZ fragment from Pa. pantotrophus (Nogales et al., 2002), indicating that the former sequence is not genuine of that species. NosZ of ‘P. denitrificans’ clusters with the P. aeruginosa PAO1 sequence, but since organisms carrying this name comprise two genera and several species (Doudoroff et al., 1974; Judicial Commission of the International Committee on Systematic Bacteriology, 1982), this sequence fragment also was not considered. We were interested in how well the tree of NosZ proteins would follow the phylogenetic relationship of the same bacteria in a 16S rRNA tree. For this purpose we have extracted the nucleotide sequences of 16S rRNA genes from the genomes of strains listed in Table 2 (when unavailable we resorted to the type species) and reconstructed the corresponding 16S rRNA phylogeny. The NosZ protein tree agrees remarkably well with the 16S rRNA tree (data not shown). The Nos trait seems to be an early evolutionary event that has subsequently followed the phylogenetic trajectory of speciation. With the information from genomes it is also possible to investigate the phylogeny of the other Nos proteins in support of the above conclusion. The argument of an autonomous respiratory mode implies that an NGC evolved along a comparable trajectory for the components making up the system. The congruous phylogenetic relationship, illustrated in Fig. 11, indicates coevolution of the accessory nos genes with nosZ. The grouping of the deduced gene products in the proteobacteria classes a, b and g and the other lineages is remarkably consistent. This is particularly evident for NosD, -F and -Y, which are tied by a common functional element. In Section 10, we will discuss the obligatory dependence of the biosynthesis of N2OR on these accessory gene products. The NosL tree follows in its basic pattern that of

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Figure 11 Phylogenetic relationships of NosD, NosF, NosY and NosL proteins. The trees were constructed from species that harbour a nos gene cluster. For discussion of the positions of Neisseria spp. and A. cycloclastes see the text. For abbreviations see Table 2; Halosp, Halobacterium sp. NRC1; for methods and colour code see Fig. 10. (See Colour Plate Section in back of this volume.)

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the other Nos protein trees, however, in several lineages it is broadened by NosL duplications, generating a pool of presumably paralogous entities. Three gene duplications are also found for nosF and nosY in the archaeon H. marismortui (data not shown). The exceptions, where the NosZ tree and the 16S rRNA tree do not coincide, may represent events of lateral gene transfer. Expectation is that any lateral gene transfer should also be manifest in the phylogeny of the other nos genes, because of the functional coupling of the nos gene products. The best cases of such an event can be made for the a-proteobacterium M. magnetotacticum and the b-proteobacterium De. aromatica. Both bacteria cluster in the NosZ tree with the e-proteobacteria Wolinella and Thiomicrospira. To this lineage Ca. fetus is added in the NosDFYL trees. The NGCs of the e-proteobacteria are distinguished from the rest of noscarrying strains by the acquisition of the nap-like genes nosG and nosH. The same feature holds for M. magnetotacticum and De. aromatica, adding thereby another distinctive element. Both cases are consolidated by the phylogeny of the other Nos proteins (Fig. 11), which are grouped in each case with the homologous proteins of e-proteobacteria. Thiobacillus denitrificans pairs for all nos gene products closely with Azoarcus sp., but phylogenetically it resides in the 16S rRNA tree close to Neisseria spp. and more distant from the genus Azoarcus. It is feasible that acquisition of denitriﬁcation by this aerobic bacterium has been a late event. The nosZ genes of A. cycloclastes and N. gonorrhoeae were suggested previously to have been derived from lateral gene transfer (Philippot, 2002). Indeed, the nos genes of Neisseria spp. have a tendency to associate with the a-proteobacteria rather than the cognate taxon, b-proteobacteria. The genes are on a deep lineage and a certain plasticity of the NGCs of Neisseria spp. is evident. This does not seem to be exclusive to pathogenicity, since the NGCs of Brucella sp. and Burholderia sp. exhibit a consistent pattern of tight phylogenetic relation for all nos genes. The case of ‘A. cycloclastes’ was resolved recently. NosZ of this bacterium lines up in the a clade, whereas the genus Achromobacter belongs to the b-proteobacteria. However, A. cycloclastes was not considered when the genus Achromobacter was newly formed (Yabuuchi et al., 1998) on rejection of the original genus ‘Achromobacter Bergey et al. 1923’ (Holt, 1979), whereby the former is not identical with the latter. To evaluate whether this is a case of putative lateral gene transfer or only a taxonomically misplaced a-proteobacterium, we determined the 16S rRNA sequence (accession no. AM292630), which shows that A. cycloclastes is an a-proteobacterium belonging to the genus Ensifer (Sinorhizobium).

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The three strains of N. meningitidis, which represent serogroups A, B and C, have incomplete NGCs that lack nosZ. The nosR gene is atypically short and translates into an N-terminal fragment of 173 (serogroup A) to 190 (serogroups B and C) amino acids only. The nosY genes encode protein fragments of 39, 151 and 173 amino acids in serogroups A, B and C, respectively, which for about 40 amino acids are highly similar to the C-terminal part of other NosY proteins. N. gonorrhoeae does have a complete nosZ gene, but its nosD and nosR genes are fragmented and each consists of two separate reading frames. Jointly they translate into the respective complete gene product. Absence of selective pressure for N2O reduction in Neisseria spp. seems to have resulted in the loss or fragmentation of nos genes. Shewanella oneidensis represents another interesting case. This bacterium, differing from its close relative Shewanella denitrificans (Brettar et al., 2002) is not known to denitrify yet carries an isolated nosLDFY sequence. The nos genes of Sh. oneidensis could be remnants of a once-complete N2O respiratory system. Whereas Sh. denitrificans shows for each Nos protein constant tree positions next to Ph. profundum and Co. psychrerythraea within its cognate group of the g-proteobacteria, Sh. oneidensis does not do so. It is only loosely associated with the g-proteobacteria group and its NosF protein pairs with the gram-positive bacterium D. hafniense. In all cases a deep branching is indicative of an independent trajectory. Thus, the Nos proteins of Sh. oneidensis, because of an early separation, may have undergone a functional shift away from its homologous genes involved in N2O reduction; the more so particularly since the key gene, nosZ, is not present. The deduced NosZ sequence of M. magnetotacticum consists of two overlapping fragments; a short N-terminal one of 262 amino acids that carries three of the seven histidine ligands of CuZ, and the C-terminal fragment, making up the bulk of NosZ. A two-subunit situation seems unlikely considering the highly ordered b-propeller structure of the CuZ domain. NosD and NosF of Te. roseum DSM5159 are encoded from a single ORF, contrasting with their location at either side of the membrane. These features require reinvestigation on completion of the sequencing projects. A large NosZ pool in the databank of about 850 NosZ fragments of unknown bacterial origins stems from environmental samples. Most belong to two groups of incomplete primary structures covering different parts of the CuZ domain as the result of the PCR primer design. In ecological studies, dissecting the denitrifying bacterial community of a habitat with nosZ gene probes, it is often stated that the diversity of N2OR is much larger than what is represented by the preferred model strains in the laboratory (Scala and Kerkhof, 1999; Ro¨sch et al., 2002; Rich et al., 2003; Stres et al., 2004). Environmental samples yield in phylogenetic analyses clusters of

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well-populated NosZ clades, often harbouring a ‘laboratory strain’ as a lead member. The lack of taxonomic identiﬁcation, however, in a large number of unassignable clusters does not mean principal structural differences within N2OR, or another type of enzyme in these cases. Considering the entire ‘landscape’ of partial NosZ sequences present in the databank, we ﬁnd in the overall alignment only the structural features of the Z-type N2OR (data not shown). However, ecological studies clearly emphasize that many more taxa, waiting to be identiﬁed, do have a Z-type N2OR.

8.2. Relationship of N2O Reductase to Cytochrome Oxidase The seminal ﬁnding of a sequence motif shared between the CuA centre of N2OR and the subunit II of COX (Viebrock and Zumft, 1988) led to the issue of a putative evolutionary relationship of the two enzymes (Zumft, 1992). The hypothesis of a common evolutionary origin of O2 and N-oxide respiratory enzymes was formulated explicitly on recognizing that NO reductase has structurally conserved metal-binding centres and extended sequence homology with the haem–Cu-type oxidase family (Saraste and Castresana, 1994; van der Oost et al., 1994). It is now possible to unite NO reductases and O2 reductases in a phylogenetic relationship (Zumft, 2005a). Elucidation of the evolutionary origin of respiration will be achieved from genomes. The sequenced genomes already include a representative number of denitrifying bacteria from different taxons and have revealed valuable new information about N oxide-metabolizing capabilities.

8.3. Inorganic Metabolism of N2O by Archaea Nitrate denitriﬁcation to N2 is a sequence of four reaction steps, each one requiring a distinct metalloenzyme in the Bacteria. Physiological, biochemical and genetic evidence have provided a robust picture for this. When the process of denitriﬁcation in the Archaea was ﬁrst reviewed (Zumft, 1997), it was cautiously suggested, given the paucity of information then, that Archaea might follow the bacterial pattern. Fresh evidence puts us in a position to view the nature of the pathway in the Archaea again and discuss the conclusion of a biochemistry homologous to the bacterial situation. The evidence for archaeal dissimilatory N-oxide metabolism can be summarized by the same reaction sequence, homologous genes and biochemistry as found in Bacteria: NarG

NirK

NorZ

NosZ

NO 3 ! NO2 ! NO ! N2 O ! N2 NirS

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Informational gaps can be ﬁlled now by viewing several species and by considering complementary biochemical and genomic evidence, even though there is still no single archaeon whose denitriﬁcation pathway would have been uncovered at both the enzyme and genetic levels. Denitriﬁcation genes encoding the necessary enzymes have been detected in genomes of Archaea by homology search with the bacterial counterparts. Surveying this evidence, we see no difference in pathway or biochemistry. Archaea and Bacteria exhibit the same denitriﬁcation pathway with the same metalloenzymes to let us assume that the process existed in a common ancestor before both domains separated. 8.3.1. N2O Formation and Reduction Gas formation from nitrate and/or nitrite was reported for a number of halophilic archaea. The gas is usually assumed to be N2, but N2O has also been identiﬁed. Within the genus Haloarcula the denitrifying species are H. vallismortis, H. hispanica, H. japonica, H. marismortui and H. quadrata; the denitrifying species within the genus Haloferax are Hf. denitrificans and Hf. mediterranei. Halogeometricum borinquense grows anaerobically with nitrate and forms nitrite and gas (Grant et al., 2001). Hf. denitrificans grows as a facultative anaerobic in the presence of nitrate. This archaeon is a complete nitrate denitriﬁer when supplied with 0.1% nitrate (Tindall et al., 1989). However, N2O accumulates in a culture when the nitrate concentration is increased to 0.5%. In addition, some nitrite is found. Py. aerophilum is a complete denitriﬁer and produces N2 from nitrite together with traces of N2O and NO (Vo¨lkl et al., 1993). In both instances the gases were identiﬁed by gas chromatography. 8.3.2. Archaeal N2O Reductase The 3.13-Mb chromosome I of H. marismortui, the larger one of two chromosomes, carries the nos genes, with nosZ being a part of an NGC that also includes the homologous bacterial genes nosDFY (Table 2). Several more copies of putative nosF and nosY genes are located elsewhere on the chromosome, and putative nosD and nosL genes are located on the 0.41-Mb plasmid pNG700, the largest of seven plasmids of H. marismortui (Baliga et al., 2004). nosR and nosX homologues are not detectable. This coincides with the ﬁndings from bacteria that these genes may be absent. The deduced nosZ product exhibits the CuZ domain of N2OR, with the set of conserved histidine residues, and the CuA domain carrying the canonical Cu ligands. It can be modelled structurally on the bacterial N2OR fold. N2OR of

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H. marismortui belongs to the distinct group of Z-type enzymes where the histidine next to the CuA ligand Cys622 is replaced by a serine residue. Like its bacterial counterpart, the archaeal N2OR is inhibited by acetylene (Tomlinson et al., 1986). Analysis of the signal peptide indicates no clear Tat features, although archaea do have the Tat translocation pathway and haloarchaea exhibit a clear preference for it (Dilks et al., 2005). Chromosome I of H. marismortui is at the DNA level 65–70% identical with the large chromosome of Halobacterium NRC1 (Baliga et al., 2004). Nevertheless, only homologues of nosF and nosY were found in the latter by a BLAST search, but no nosZ gene. The physiology of strain NRC1 with respect to dissimilatory nitrate metabolism is not known; most Halobacterium spp. are strict aerobes. Haloferax volcanii, which is the type species of the genus Haloferax, is reported to reduce nitrate only to nitrite (Torreblanca et al., 1986; Grant et al., 2001). Its nitrate reductase has been isolated and studied. The genome indicates that this archaeon is a denitriﬁer, as it reveals genes for the Cu-type nitrite reductase, NirK, the NorZ-type NO reductase, and the nos genes nosD, -F, -Y, and -L. Though the latter are clearly correlated with a CuS centre, no corresponding nosZ gene is evident in Hf. volcanii (Table 2). The genome of Py. aerophilum represents a similar case (Fitz-Gibbon et al., 2002). Although physiological evidence for N2 formation indicates an N2OR, and the genome reveals several accessory genes (Table 2), a homologue of nosZ is not detectable. Py. aerophilum has been described as a complete denitriﬁer that does not grow on N2O as the sole electron acceptor (Vo¨lkl et al., 1993). Since such a limitation is also known in Bacteria, it cannot be taken as an argument for the absence of an N2OR. Adjacent to the nos genes a short ORF translates into a protein exhibiting the CuA domain (annotated as cytochrome oxidase subunit II) that, however, cannot be fused with vicinal ORFs, including alternative reading frames, to represent a nosZ gene. Thus, for this archaeon we have physiological evidence for N2O reduction but cannot extract structural information from the genome. To resolve this discrepancy, we envisage an N2OR with a CuS centre (to justify the presence of the accessory functions), but ligated to a protein scaffold sufﬁciently different from the Z-type enzyme not to be recognized by homology search.

8.3.3. N2O Generation from NO as Intermediate Reports of NO formation by H. marismortui and Py. aerophilum place NO as putative intermediate in the denitriﬁcation pathway. NO reductase is the

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generator of N2O by reducing NO derived from a nitrite reductase. The genomes of Py. aerophilum, Hf. volcanii and Sulfolobus solfataricus reveal gene products with homology to bacterial NO reductases. Speciﬁcally, their NO reductases are of the haem–Cu oxidase type that belong to the NorZ subgroup and obtain their electrons from quinol other than members of the NorB subgroup, which interact with a haem protein (Zumft, 2005a). The NO reductase of Py. aerophilum has been isolated and biochemically characterized (de Vries et al., 2003). An internal peptide of 30 amino acids is identical to the gene-deduced NorZ sequence. A signiﬁcant variation is the presence of haem o versus the otherwise commonly observed haem b.

8.3.4. Two Types of Nitrite Reductases in Archaea Nitrite is both formed and reduced by archaea as it accumulates transiently during nitrate denitriﬁcation. As found with Bacteria, Archaea have their denitrifying capability based on two types of respiratory nitrite reductases. The cytochrome cd1 enzyme is encoded by nirS in the genome of Py. aerophilum (Fitz-Gibbon et al., 2002), and bioinformatic evidence identiﬁes the nirK gene, encoding the Cu-containing nitrite reductase, on chromosome I of H. marismortui (Baliga et al., 2004). The deduced NirK sequence is most similar to the homologue from N. gonorrhoeae (41%). As this is higher than that with other NirK proteins, lateral gene transfer has been suggested (Ichiki et al., 2001). The Cu-containing nitrite reductases have been characterized biochemically from Hf. denitrificans and H. marismortui and exhibit bacterial properties (Inatomi and Hochstein, 1996; Ichiki et al., 2001).

8.3.5. Respiratory Nitrate Reductase The best-studied process of dissimilatory N-oxide metabolism in the Archaea is nitrate respiration. Nitrate reductases have been isolated and characterized from the Haloferax species Hf. denitrificans (Hochstein and Lang, 1991), Hf. volcanii (Bickel-Sandko¨tter and Ufer, 1995), and Hf. mediterranei (Alvarez-Ossorio et al., 1992; Lledo´ et al., 2004), and also from H. marismortui (Yoshimatsu et al., 2002) and Py. aerophilum (Afshar et al., 2001). Although some structural and topological differences exist (the enzyme seems to face the outside of the cell, not the cytoplasm), archaeal respiratory nitrate reductases are homologues to the bacterial enzyme encoded by the narG operon. Nitrate reduction among the archaea was recently reviewed in the context of inorganic N metabolism (Cabello et al., 2004).

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8.4. Gram-Positive Bacteria Denitriﬁcation is found among several genera of gram-positive bacteria with the bacilli being the most prominent (Zumft, 1992). The enzyme, however, has not yet been isolated from any representative. Complete denitriﬁcation was described for the moderately thermophilic Geobacillus thermodenitrificans, but termination by N2O in Virgibacillus halodenitrificans (Denariaz et al., 1989; Nazina et al., 2001). Preliminary data indicate that N2O reduction of B. azotoformans is menaquinol dependent and that it is a membranebound activity (Suharti and de Vries, 2005). The question of the biochemical nature of N2OR in gram-positive bacteria can be answered for the ﬁrst time via genome analysis of D. hafniense. In the original description of D. hafniense by Christiansen and Ahring (1996), the bacterium was said to be gram-negative. However, it had been shown for Desulfitobacterium frappieri (a synonym of D. hafniense) by electron microscopy that, although the Gram stain is negative, its morphotype is gram-positive (Bouchard et al., 1996). D. hafniense forms endospores and belongs with its DNA G+C content of 46–47 mol% to the group of low G+C gram-positive bacteria (Niggemyer et al., 2001). The bacterium was shown to reduce nitrate to nitrite and ammonia, but not to denitrify (Christiansen and Ahring, 1996). Its genome, however, reveals a nosCZ orf nosDLFY cluster (Table 2), and is clearly predicted to encode the Z-type N2OR of gram-negative bacteria. All CuZ-ligating histidine residues are conserved, so is the CuA domain. We had previously translated two partial NosZ sequences from the incomplete genome sequence (Simon et al., 2004), which were later amended as a single ORF. The primary sequence exhibits a putative signal peptide, which is relatively short and carries a twin lysine but not the arginine pair diagnostic for the Tat pathway. If cleaved during putative transport by the Tat translocon, the enzyme is devoid of a membrane anchor and requires other means to be immobilized at the outside. The genome of D. hafniense also encodes the cytochrome bc-type NO reductase, NorB (Zumft, 2005a), which indicates a wider capability to handle N oxides in a dissimilative way.

8.5. Plasmid-Encoded nos Genes Occasionally, nos genes are encoded on a plasmid. The ﬁrst analysis of such a situation was done with C. necator, whose capability for chemolithoautotrophy resides on a 452-kb megaplasmid. The nos genes are adjacent to nor genes for NO metabolism which is based on the quinol-dependent

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NorZ-type reductase (Schwartz et al., 2003). Genes for nitrate reduction, nap and nar, take up two further loci on the plasmid, whereas the genes for cytochrome cd1-dependent nitrite respiration are chromosomally encoded. In the diazotrophic S. meliloti the nos genes form part of the pSymA plasmid required for the bacterium–plant symbiosis (Chan and Wheatcroft, 1993). pSymA is the smaller of two megaplasmids in this bacterium (Barnett et al., 2001). Another example of a plasmid-borne NGC stems from the phototrophic bacterium Rb. sphaeroides which harbours a 115-kb plasmid (Schwintner et al., 1998). The NGC obtained from a cloned DNA fragment is incomplete (Sabaty et al., 1999) and might harbour additional nos genes in both ﬂanking regions. Genomic sequencing of Si. pomeroyi has revealed the plasmid-encoded nature of its nos genes (Moran et al., 2004). As mentioned above, the nosD and nosL genes are found as separate loci on the plasmid pNG700 of H. marismortui, whereas the principal nosZDFY set is chromosomally encoded. The existence of plasmid-borne NGCs emphasizes the independence of the N2O-respiratory system and the possibility of moving it laterally. In spite of this possibility, we notice in the plasmid-borne cases the coincidence of the NosZ clades with the taxonomic grouping (Fig. 10); acquisition of none of the four known plasmidborne NGCs is necessarily attributable to lateral transfer.

9. TOPOLOGY AND TRANSPORT PROCESSES 9.1. The Tat System Exports NosZ and NosX to the Periplasm N2OR is located in the periplasm where it functions as an electron sink for the anaerobic respiratory electron transfer chain. The ﬁrst indications for its location came from considerations of the proton consumption balance. Protons for N2O reduction are delivered from the periplasmic and not the cytoplasmic side of the membrane (Boogerd et al., 1981). The assumption of a periplasmic enzyme was supported from more direct evidence by demonstrating N2O-reducing activity of the periplasmic cell fraction of Pa. denitrificans (Boogerd et al., 1980; Alefounder et al., 1983) and Rb. sphaeroides (Urata et al., 1982), or from using the periplasm of Rhodobacter capsulatus as the source for enzyme isolation (McEwan et al., 1985). A periplasmic location of N2OR of P. stutzeri was demonstrated by electroimmunoassay of cell fractions (Minagawa and Zumft, 1988) and by immunogold staining and electron microscopy (Ko¨rner and Mayer, 1992). In a few cases a membrane association of N2OR has been described (Jones et al., 1992; Hole et al., 1996; Suharti and de Vries, 2005). The genome-deduced

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sequence of N2OR from Tb. denitrificans exhibits a signal peptide with features of a Tat-directed protein, combined with a C-terminal cysteinyl anchor preceded by a lipobox, LysSerAlaCys (see Section 9.3). The same characteristics are found in the unprocessed N2OR proteins of Azoarcus sp. EbN1 and Burkholderia spp. It is feasible that membrane anchoring of N2OR is realized in these cases by a lipid moiety, because otherwise N2OR does not reveal a speciﬁc membrane anchor or signiﬁcantly increased hydrophobicity that could make it a membrane-bound entity. 9.1.1. Export of N2OR It is important to unravel by which mechanisms the proteins necessary for N2O respiration reach their functional site, and how the maturation process for N2OR is interrelated with protein translocation. Two components of N2O respiration, NosZ and NosX, are exported by the twin arginine translocation pathway (Tat). The Tat system is directed at folded proteins, whether or not they are associated with a nonproteinaceous cofactor (Berks et al., 2003). Export of the nitrous oxide reductase protein, NosZ, has been investigated so far with P. stutzeri and C. necator. N2OR was among the very ﬁrst proteins where a conserved sequence motif was observed in the signal peptide. The ﬁnding was serendipitous from the collaboration of two research groups interested in nitrogen and hydrogen metabolism (Zumft et al., 1992). The distinctive elements contributed from the study of N2O metabolism are detailed below. The seminal studies on export of hydrogenase and the discovery of a ‘hydrogenase-type’ signal peptide with two arginine residues have been summarized by Voordouw (2000). The ﬁrst evidence for the mechanism of NosZ export came from the comparison of NosZ signal peptides, which revealed a conserved sequence motif with an arginine pair in a long presequence of about 50 amino acids (Fig. 12). The same motif appeared in the signal peptide of hydrogenase (Zumft et al., 1992), which was unexpected at that time since Sec-type signal peptides lack positional conservation. Adding a fourth sequence for comparison conﬁrmed the initial observation (Hoeren et al., 1993). The importance of the conserved motif was demonstrated with hydrogenase where substitution of the N-terminal position of the arginine pair prevents protein translocation (Nivie`re et al., 1992). The same phenomenon was found with N2OR. The Arg20Asp substitution in the signal peptide causes NosZ to accumulate in the cytoplasm in dimeric form and without the Cu cofactors (Dreusch, 1995). The search for similar peptides revealed a structurally conserved environment in the signal sequences of mostly periplasmic, sometimes also membrane-bound enzymes, including polysulﬁde reductase, iron

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Figure 12 Sequence logo of the Tat-speciﬁc signal sequence of NosZ. The letter size shows the relative frequency of a residue at that position. The cleavage site, AXA, is indicated by positions –1 –3. The signal peptide consensus is LSRRXFLG. The H region exhibits a dominance of glycine and alanine residues. The ﬁgure was constructed with the web-based application WebLogo v. 2.8.2 according to Crooks et al. (2004).

oxidase, periplasmic nitrate reductase, methylamine dehydrogenase and trimethylamine N-oxide reductase. These ﬁndings from a Ph.D. thesis (Dreusch, 1995) were presented during 1995 at international meetings on N2 ﬁxation in St. Petersburg and on Cu proteins in Sta. Severa, Rome (Zumft and Dreusch, 1995; Beinert, 1996) but were only belatedly published (cf. Fig. 6 in Dreusch et al., 1997). Drawing on an extended databank screening for the new type of signal peptide, and the foregone experience with N2OR (Hoeren et al., 1993), Berks (1996) had the insight to propose the existence of a new bacterial transport system. He deﬁned a heptameric consensus motif of (Ser/Thr)ArgArgXaaPheLeuLys for the cognate signal peptide. Experimental discovery of the bacterial Tat secretory system (Sargent et al., 1998; Weiner et al., 1998) followed immediately when it was found that the maize Hcf106 component of a DpH-driven pathway of protein import into chloroplasts has homologues in bacteria (Settles et al., 1997). Tat transport has become a fascinating ﬁeld of investigation since. The chloroplast model for the cyclical assembly of the translocase system starts with the binding of a Tat substrate to the cpTatC-Hcf106 complex, which is homologous to bacterial TatC-TatB. Subsequent association of the substrate-bound receptor complex with Tha4 (homologous to bacterial TatA) generates the translocase and effects substrate translocation, which is followed by dissociation of the receptor and translocator moieties to close the transport cycle (Mori and Cline, 2001, 2002). The evidence that the bacterial system follows the same principal lines has been reviewed recently (Palmer et al., 2005). The bacterial Tat translocon of E. coli is best understood. It comprises

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TatA, TatB, TatC and TatE. TatC is an integral membrane protein, composed of six transmembrane helices. The other proteins have two a-helices: one helix anchors the protein in the membrane, whereas the second, amphipathic helix is exposed to the cytoplasm. TatA and TatE are homologous and functionally substitutable proteins. A Tat-targeted substrate associates ﬁrst with TatC, followed by interaction with TatB, which then proceeds upon membrane energization to interact with the TatA transport pore (Alami et al., 2003). As shown recently for TatA, the cytoplasmic helix can ﬂip across the membrane and also become accessible from the periplasmic side (Goufﬁ et al., 2004). This provides a conceptual basis of how a supramolecular TatA complex may function as the transport channel. The TatA protein assembles in a ring-like structures of 45–55 A˚ height and 25–30 A˚ width and a molecular mass ranging from 130 to 390 kDa. Twelve to 35 TatA molecules are thought to form the channel structure with the hydrophobic helix in the transmembrane ring and the amphipathic helix, lining the pore (Gohlke et al., 2005). The tat locus of P. stutzeri consists of tatABC, and an additional tatE gene as part of the NGC (see Fig. 9). In C. necator a putative tatA locus is implicated in NosZ translocation from the observation of a mutational loss of N2O-reducing activity of whole cells (Bernhard et al., 2000). The tatE and tatA products are interchangeable, since a tatE mutant is not rendered transport incompetent for NosZ (Heikkila¨ et al., 2001). The Tat system of P. stutzeri or other N2O-metabolizing bacteria is assumed to follow the E. coli paradigm and to be operative usually with TatA. The location in the NGC suggests, however, a Nos-related role for tatE of P. stutzeri. The N2OR content of the denitrifying cells comprises 2–3% of the total protein (Coyle et al., 1985). Under denitrifying conditions the increased demand to transport the Tat-dependent substrates NosZ and NosX across the membrane may be met by TatE, which is co-expressed in response to anaerobic, denitrifying conditions. When NosZ is overexpressed in homologous or heterologous systems, co-expression of TatE results in the complete processing of NosZ (Heikkila¨ et al., 2001; Wunsch et al., 2003). A speciﬁc conformation seems to be required for the NosZ substrate to explain the observations with certain recombinant forms. Substitution of the CuA ligand Cys618 for valine leads to a strong rejection of NosZ by the Tat translocon. Most of the NosZ is left in the unprocessed form (Dreusch, 1995; Heikkila¨ et al., 2001). Valine ﬁts into the side of cysteine in the N2OR structure (Brown et al., 2000b). Presumably, it imposes by its hydrophobic side chain a structural change onto the CuA domain and perhaps the overall enzyme conformation, since CuA and CuZ domains are structurally interdependent (see Section 6.1). The His583Gly recombinant form is to a lesser

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degree incompetent for Tat transport and, other than the Cys618Val form, most of the precursor is processed to mature NosZ. The Met629Cys exchange also affects the translocation efﬁciency. These amino acid substitutions indicate that the Tat system is constrained by distinct conformations in the folded form of NosZ. The phenomenon is related to what has been studied with PhoA as quality control of folding (DeLisa et al., 2003). Re-directing PhoA from the Sec to the Tat system, requires both the swap of the signal peptide, as well as forcing disulﬁde-bridge formation onto PhoA for correct folding in the cytoplasm. Since in NosZ the residues in question are positioned close to the C-terminus, the Tat system seems to involve a temporal element to recognize the mutationally induced structural change in the completely or nearly completely synthesized NosZ, prior to initiating export, or aborting the process on recognizing a misfolded enzyme. The rationale for N2OR export by Tat is open. Obviously, it does not reside in cytoplasmic Cu cofactor acquisition or the distinct b-propeller structure. We have predicted that the Wolinella enzyme, which clearly has an overall b-propeller structure fused to a Sec-type signal peptide, is exported by the Sec system (Simon et al., 2004). We also ﬁnd a similar, eight-bladed propeller of b-sheets in cytochrome cd1 nitrite reductase, yet this structure does not tie the enzyme to the Tat system. The tatC mutant of P. stutzeri continues to export cytochrome cd1 unimpaired to the periplasm (Heikkila¨ et al., 2001). Both N2OR and cytochrome cd1 share the global similarity of dimeric entities with each monomer consisting of an electron transfer domain and a catalytic domain. For the P. stutzeri N2OR we conjecture that folding prior to export is related to several disulﬁde bridges in the enzyme (Dreusch et al., 1996). A folded structure, close to the ﬁnal conformation, is anticipated to have the –SH groups positioned correctly for disulﬁde-bond formation in the oxidizing environment of the periplasm. 9.1.2. Export of NosX NosX is the second Nos substrate with relevance for the Tat pathway. This 34-kDa protein was shown by mutational inactivation to be necessary for N2O respiration by certain bacteria (Chan et al., 1997; Saunders et al., 2000). NosX carries a Tat-speciﬁc signal sequence and is presumed to be a periplasmic component of the N2O respiratory system. Its ﬂavin content (D.M. Dooley, pers. commun.) is congruous with Tat transport. A triplearginine motif is nearly always part of its signal peptide (Fig. 13). However, the NosX proteins of the three Burkholderia spp. have a twin lysine motif instead. In certain cases, the deletion of the arginine pair (Summer et al., 2000) or recombinant Lys–Lys substitutions (Ize et al., 2002) do not abolish

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Figure 13 Tat-speciﬁc signal sequence of NosX. The sequence logo highlights conservation of the triple-Arg or a Lys Lys Arg motif, the hydrophobic segment, and the cleavage site at positions –1 –3. The ﬁgure was constructed with the web-based application WebLogo v. 2.8.2 according to Crooks et al. (2004).

Tat transport activity. It may be that NosX of Burkholderia constitutes a naturally occurring Lys–Lys variation among the Tat signal peptides.

9.2. Sec Export-Dependent Components NosR, NosD, NosY and NosL The Sec system is the principal transport system for bacterial protein translocation (for a review, see Dalbey and Chen, 2004). It acts on unfolded proteins whose cofactors, if present, are processed separately. In addition to NosZ of W. succinogenes a few other NosZ proteins have Sec-type signal peptides and, in contrast to the usual Tat export pathway, seem to be exported by the Sec system (Simon et al., 2004). The Nos components NosR and NosY are integral membrane proteins. Both have Sec-speciﬁc signal peptides. NosR had been thought to have an additional N-terminal membrane anchor preceding the periplasmic domain, which is not the case, however. Mature NosR has been puriﬁed and shown to have lost the N-terminal hydrophobic sequence upon insertion into the membrane (Wunsch and Zumft, 2005). NosD and NosY have not been puriﬁed to reveal processing. NosL is predicted to be a component of the outer membrane since it carries the signal peptide of a lipoprotein (see below). The periplasmic location of NosD is deduced from a reporter gene fusion with phoA (A. Dreusch and W.G. Zumft, unpublished data). Speciﬁc transport studies with any of the Nos components, other than NosZ, have not been done. Nonetheless, it is clear that both the Tat and the Sec translocation system have to cooperate to assemble a functional N2O respiratory system.

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9.3. Lipoprotein Targeting NosL has features of a lipoprotein. Lipoproteins cross the membrane by the Sec system. They require processing by attachment of a diglyceride via thioether linkage to the cysteine of the mature N terminus by the diacyl glyceryl transferase, Lpt (Tokuda and Matsuyama, 2004). Subsequent to modiﬁcation, the signal peptide is cleaved by the Lsp signal peptidase II, which targets the lipobox, Leu(Ala/Ser)(Gly/Ala)Cys (E. coli consensus) (Hayashi and Wu, 1990). Aminoacylation of the cysteine residue by a phospholipid–apolipoprotein transacylase yields the mature lipoprotein. An ABC transporter system, consisting of the LolD ATPase, the innermembrane proteins LolC and LolE and the periplasmic chaperone LolA, releases the lipoprotein from the inner membrane to LolA. The LolA-bound lipoprotein interacts with the outer-membrane receptor LolB, which directs the lipoprotein to the outer membrane. Many NosL proteins exhibit the sequence LeuAla(Ala/Gly)Cys as the lipobox consensus (Table 4). We hypothesize that NosL is targeted to the outer membrane by the Lol system. Position +2 after the C-terminal cysteine is important as the so-called Lol avoidance signal. An aspartate residue at this position retains the protein in the inner membrane (Masuda et al., 2002). Indeed, NosL of P. aeruginosa and P. fluorescens have such a conﬁguration, which might indicate in these cases an inner-membrane anchoring of periplasmic NosL (Table 4). Although NosD, -F and -Y form an ABC transporter system, they have no similarity with the Lol system. We consider it unlikely that they would act on NosL. Prediction of lipoboxes is not unequivocal though for every deduced NosL protein, particularly among the paralogues, and requires further studies. Table 4 Signal peptides of NosL proteins NosL source

N region

H region

C region -4-3-2-1C1+2

A. cycloclastes Br. melitensis C. necator Pa. denitrificans P. stutzeri P. aeruginosa P. fluorescens Rp. palustris Rb. sphaeroides Tb. denitrificans Lipobox consensus2

MRTRLR MKR MRVNRR MRH MNALHR MQRHTLPLRP MIECPLKTGR MRISGH MRR MNLSRPAG

FVLVAAALA ALFLAPFFFA QLLLATCALGAQA ALLLELLL IGAGTLLAVLLAF LLGTLLLGL LLAGLLMCL IALLAAAL LALALL VLIVAAG

LLSAC(20)K LLAGC(18)S ALSAC(24)G DLVAC(16)R GLTGC(24)G LLAGC(24)D ALAAC(24)D LLAGC(19)N LLAAC(13)R ALAAC(21)G (L/A) LA (A/G)C

1 2

Position of cysteine residue in parentheses. Derived from 27 sequences.

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10. CU CENTRE ASSEMBLY On analysing the nos locus it was found that the structural gene, nosZ, was accompanied by several other genes whose mutagenesis affected the Cu content and nature of Cu sites in the enzyme. In studying the enzyme from these chromophore mutants it became clear that N2OR is subjected to a speciﬁc process of metal site assembly. The principal ﬁndings were reviewed by Zumft and Kroneck (1996) and more recently by Zumft (2005b). The essential requirement for N2OR biogenesis is that an assembly system complements the structural gene nosZ. Thus, acquiring N2O-reducing capability in the heterologous host P. putida needs the expression of the assembly genes together with nosZ and nosR (Wunsch et al., 2003). To make the system functional in vivo may require only a functional electron donor pathway in addition. Since we have seen that NosZ is targeted to the Tat system, it seems logical that the enzyme would acquire its Cu cofactors in the cytoplasm. This is not the case and makes N2OR an exception to the concept that cofactor acquisition prior to transport were a stringent feature of the Tat translocon. Several lines of evidence support a periplasmic maturation process for N2OR. (1) Export of NosZ proceeds undisturbed when an assembly gene is inactivated. (2) Cu deﬁciency does not affect the location of N2OR, but only lowers enzyme activity. Exogenous Cu remedies the effect. (3) N2OR assembly involves the periplasmic components NosL and NosD. (4) Mutagenesis of either the twin arginine motif of the signal peptide or the tatC gene retains NosZ in the cytoplasm. This topologically aberrant cytoplasmic enzyme is devoid of Cu. (5) A correctly folded cytoplasmic NosZ is deduced from the possibility of incorporating Cu in vitro into the CuA site of the recombinant Arg20Asp enzyme. (6) NosZ of W. succinogenes, transported by the Sec system, requires periplasmic processing of the Cu cofactors. A scheme of NosZ processing and maturation, integrating the known components and functional sites, is shown in Fig. 14.

10.1. Supplying Cu: NosA, NosL and ScoP Relatively little is known about Cu uptake and cellular routing in N2ORharbouring bacteria. With respect to Cu supply for N2O metabolism, the outer membrane Cu-containing proteins NosA and the NosA homologue, OprC, have been characterized from P. stutzeri JM300 and P. aeruginosa, respectively (Lee et al., 1991; Yoneyama and Nakae, 1996). These proteins are not speciﬁc for denitrifying bacteria but are also found in bacterial

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Figure 14 Model of the membrane topology of components for N2O respiration and N2OR biogenesis. The scheme is representative for P. stutzeri or, considering the NosX protein, also for Pa. denitrificans, whose respiratory chain, however, is more complex. Under oxygen-limited conditions and in the presence of nitrate a cytochrome cbb3 terminal oxidase (Cyt cbb3) and the N2O-respiring system are expressed concomitantly. Components of the respiratory chain consist of NADH dehydrogenase (DH), the Q-cycle (Q/QH2), cytochrome bc1 (Cyt bc1), and a terminal oxidase. Cyt c and Cu, are a periplasmic c-type cytochrome and a cupredoxin, respectively, both accepting electrons from the cytochrome-bc1 complex. Single uppercase letters indicate the products of nos genes necessary for the assembly process of the Cu centres of N2OR. Numbers give the approximate protein masses in kDa. Cu-containing proteins are labelled as such. N2OR (NosZ) is shown in its dimeric state; NosZ is exported by the Tat translocon by cleaving the 5-kDa signal peptide. Tat composition is not detailed other than to show the supportive role of TatE for NosZ export. NosF has ATPase activity. [S] is a sulfur species of unknown chemical nature. NosL is shown with a putative lipid anchor in the outer membrane. Cu may enter the periplasm via NosA or another porin, depicted to the left of NosL. The membrane-bound NosR is shown with the periplasmic ﬂavin cofactor domain and the cytoplasmic Fe–S domain. Overall no inferences about stoichiometries of protein complexes are drawn. For further discussion see the text.

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backgrounds lacking N2O-metabolizing capability, for example P. putida or Yersinia pestis (Wunsch et al., 2003). The role of NosA may be that of an outer-membrane pore for a Cu-uptake system (Lee et al., 1991) under speciﬁc growth conditions. Notably, the nosA gene is activated under anaerobiosis and is repressed by Cu. The most conspicuous feature of the nosL gene is its consistent coexistence with nosDFY. The nosL product is predicted as lipoprotein of the outer membrane (Dreusch et al., 1996). The role of NosL may be to provide Cu in the periplasm for NosZ. The protein from A. cycloclastes has been heterologously expressed in E. coli without its signal peptide and has been puriﬁed (McGuirl et al., 2001). Its structure determination is under way (Taubner et al., 2004). The protein is rich in b-sheet content. Cu binding is thought to involve residue Cys24. This residue, however, is poorly conserved in NosL proteins. The afﬁnity of NosL for Cu is markedly higher for Cu(I) than Cu(II), supporting the role as a chaperone (McGuirl et al., 2001). We have discussed in Section 9.3 how NosL might be targeted to the outer membrane. The biogenesis of CuA of COX in yeast depends on two proteins, Sco1 and the Cu chaperone Cox17. Cu(I) is thought to be transferred from Cox17 to Sco1 for insertion of Cu into subunit II (Heaton et al., 2001; Nittis et al., 2001). Existence of a homologue of Cox17 in bacteria (Banci et al., 2005) and the observation that the Sco1 homologue YmpQ of B. subtilis affects COX but not menaquinol oxidase (Mattatall et al., 2000), show at least the feasibility of a similar scenario for CuA synthesis of N2OR. Homologues of Sco1 coexist nearly always in bacteria that have an NGC, though they are not linked with them. Sco1 proteins and homologues carry a Cys(Xaa)3CysPro motif and a histidine residue, which are important for Cu binding. The soluble domain of the Sco1 homologue PrrC from Rb. sphaeroides has thioldisulﬁde oxidoreductase activity, which can be used for Cu mobilization (McEwan et al., 2002). A catalysed insertion of Cu into CuA is in line with the concept of tight routing of Cu in the cell along discrete pathways. On the other hand, the Sco1 homologue of P. stutzeri, ScoP, was found dispensable for NosZ biosynthesis by mutational evidence (Wunsch et al., 2003). The lack of a recognizable phenotype of a scoP mutant may be due to a ScoP paralogue in this bacterium, or to CuA metallation also occurring spontaneously.

10.2. ABC Transporter We will discuss the arguments that help to deﬁne the role of the NosDFY system. CuZ is a Cu–S cluster whose biogenesis requires sources of Cu and

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sulﬁde. We start with the assumption that the necessary Nos components for N2OR maturation have been identiﬁed. This is supported from random mutagenesis, from heterologous nos gene expression in P. putida, and from comparative genomics. From the three lines of evidence the accessory genes are nosX and nosR (both will be discussed in Section 11.2), and nosDFYL. Further, it is taken as certain that nosDFY encodes an ABC transporter because of the clear signatures in the nosF product as an ABC transportertype ATPase. The structure of NosF can be predicted from the X-ray structures of ATP-binding proteins of ABC transporters (Fig. 15). NosF has a C-terminal extension, which is found elsewhere, for instance, in the MalK protein for maltose transport. This domain interacts with its cognate regulator, MalT (Boos and Shuman, 1998). The C-terminal domain of NosF is structurally different from MalK and homologues, and cannot be predicted on the ABC-type ATPase model. Its function is unknown. NosF has been puriﬁed and experimentally shown to have ATPase activity (Honisch and Zumft, 2003). The integral membrane protein, NosY, with ﬁve predicted transmembrane helices, and the periplasmic component NosD complement cytoplasmic NosF. We also accept that NosL functions as a Cu chaperone given the biochemical evidence for differential Cu binding.

Figure 15 Prediction of the secondary structure of NosF ATPase from P. stutzeri. The structure was modelled on ﬁve-best matching ATP-binding proteins of ABC transporters by SWISS Model of ExPASy. Distinct motifs featured by an ABC-type ATPase are labelled. The MalK ATPase of E. coli (PDB 1Q1E) is shown for comparative purpose. Both NosF and MalK have a C-terminal extension which, however, is not present in the NosF model because of the lack of sequence similarity.

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NosD is a member of the carbohydrate-binding proteins and sugar hydrolases (CASH) protein family. It has not been shown that it would function as a binding protein. A NosD function independent of NosFY is feasible but considered unlikely since there is no exception to a nosDFY cluster in the NGC-carrying bacteria. Presumably, these genes are co-transcribed in most instances. The phylogenetic trajectory of the nosDFY genes follows the same pattern indicating a high, selective pressure from a shared function (Fig. 11). There may be a second function for NosFY in H. marismortui in addition to N2O utilization as this archaeon harbours several paralogous nosF and nosY genes without a nosD complement. Let us ﬁrst examine the question whether a NosDFY complex cooperates with NosL. In favour of this possibility we see that nosL and nosDFY practically always coexist in an NGC. Conspicuously, the nondenitrifying bacterium S. oneidensis exhibits a nosLDFY cluster, suggestive of a common function for these genes (see also Section 8.1). Since the likely role for NosL is that of a Cu chaperone, the function of NosDFY may be acquisition of Cu for the Cu–S catalytic site. However, in P. putida no co-expression of nosL is required for holoN2OR synthesis. Other than nosDFY, nosL is not selected by random mutagenesis, indicating a nonessential function. The observation that a nosL mutant of P. stutzeri (the only case studied thus far) has no phenotype is in line with this. As we have seen in Section 8.1, the phylogeny of NosL follows less stringently the matching patterns observed with NosD, -F and -Y. The NosL function allows for more plasticity in the generation of multiple paralogues (Fig. 11) and probably a functionally independent role. If we assume Cu delivery to NosZ by the concerted action of NosDFY and NosL, we see no reason why NosDFY should be essential but not NosL. More likely are independent functions for S and Cu provision, whereby the Cu supply route may draw from alternatives. The biogenesis of CuA supports this view, since nosDFY mutants can metallate CuA. An increase in exogenous Cu increases occupation of the Cu sites. Also recall that the CuA centre is easily reconstituted from apoprotein in vitro in contrast to the CuZ site (Coyle et al., 1985). The best-ﬁtting concept is that NosDFY is a sulfur (sulﬁde?) transporter. If a sulfur donor molecule is transported it leads to the related problem of how S is liberated. It is unlikely that sulfur transport would be speciﬁc for N2OR, but not liberation of sulfur from the donor. If the NosDFY complex does not provide sulfur, an alternative way has to be found. Sulfur provision would then have to follow a common route not speciﬁc for N2OR since there are no candidate genes with a putative function for sulfur delivery in NGCs. It would also mean that other periplasmic proteins have a requirement for inorganic sulfur and share a common route with N2OR. Currently no such

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proteins are known. We are not aware of a periplasmic assembly of Fe–S proteins with respect to the sulfur. A hypothetical scenario could be transport via a persulﬁde as in the Suf system for Fe–S biogenesis (Barras et al., 2005). However, we do not ﬁnd consensus cysteine residues in NosY or NosD.

11. ELECTRON DONATION AND MAINTENANCE OF ACTIVITY IN VIVO N2O reduction is energy-conserving even though N2OR is a soluble periplasmic enzyme (Urata et al., 1983; Kundu and Nicholas, 1985; Richardson et al., 1991). Energy conservation is dependent on charge separation by respiratory complexes in the pathway of electrons towards N2OR. Thus, involvement of the cytochrome bc1 complex in N2O reduction has been demonstrated for Pa. denitrificans (Boogerd et al., 1980; Parsonage et al., 1986) and the phototrophic denitriﬁers Rb. sphaeroides (Itoh et al., 1989) and Rb. capsulatus (Itoh et al., 1989; Richardson et al., 1989). N2OR is not thought to interact directly with a respiratory complex, but rather with mobile carriers to obtain reducing equivalents similar to what has been shown for respiratory nitrite reductase (Matchova´ and Kucˇera, 1991). Absorbance changes on exposure of whole cells to N2O indicate a general involvement of c- and b-type cytochromes (Matsubara, 1975). Parallel routes of electron transfer and alternative electron carriers seem to be the case for N2OR in different bacterial settings. Candidate proteins mediating between the membrane and the periplasmic enzyme are c-type cytochromes and low-molecular mass cupredoxins. However, genes encoding known c-type cytochromes or cupredoxins as potential electron donors are usually not part of NGCs. Only the pazS gene for pseudoazurin of Pa. denitrificans is located adjacent to the NGC upstream of nosC (Pearson et al., 2003), and a putative cupredoxin-encoding ORF is located upstream of nosZ in H. marismortui (Baliga et al., 2004). Even though viologens are used as artiﬁcial electron donors for assaying the enzyme, N2OR does not require a low-potential electron donor. Catalytic turnover of N2OR is possible with methylene blue (Coyle et al., 1985) or mitochondrial cytochrome c (Berks et al., 1993). The enzyme can be reduced to the blue form, N2OR III (Table 2), with Fe(II)EDTA, Eo0 ¼ +96 mV (Riester et al., 1989). In P. aeruginosa the cytochrome bc1 complex transfers electrons to cytochrome c551 (the nirM gene product) and azurin for respiratory nitrite reduction. A succinate-doped membrane

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fraction of a cytochrome c1 mutant is unable to reduce either protein. Unfortunately, it has not been shown whether these electron carriers would also couple to N2OR (Hasegawa et al., 2003).

11.1. Electron Transfer Components 11.1.1. Phototrophic Denitrifiers and Cytochrome c2 For certain phototrophic denitriﬁers the donor is cytochrome c2 (Itoh et al., 1989; Richardson et al., 1991). Its synthesis is increased under denitrifying conditions (Michalski et al., 1986). Cytochrome c2 is similar to mitochondrial cytochrome c; its reduction potential is around +350 mV (Yamanaka, 1992). In Rb. capsulatus cytochrome c2 was shown to be reduced by alternative routes, involving either the bc1 complex or an ubiquinol oxidoreductase, bypassing cytochrome bc1 (Richardson et al., 1991). 11.1.2. Cytochrome c550 and Cupredoxins as Alternative Carriers in Pa. denitrificans Cytochrome c550, as the principal electron carrier from the bc1 complex to sinks in the periplasm of Pa. denitrificans, is synthesized under all growth conditions. Since mutational inactivation of cytochrome c550 does not affect N2O reduction, the mutual substitution by alternative electron carriers has been suggested (van Spanning et al., 1990). Puriﬁed N2OR from Pa. pantotrophus accepts electrons in vitro from pseudoazurin and cytochrome c551 (Berks et al., 1993). The reactivity with different types of electron carriers manifests a low degree of recognition speciﬁcity at the electron entry site, and by inference also at the donor side. To underline this, horse heart cytochrome c can serve as electron donor in vitro (Berks et al., 1993; Rasmussen et al., 2005). The mutant C010 of Pa. denitrificans PD1222, which lacks cytochrome c550, is not affected in N2O reduction. However, the reaction becomes sensitive to a Cu chelator, which suggests a Cu protein as alternative electron donor to N2OR (Moir and Ferguson, 1994). Pseudoazurin of Pa. denitrificans has been isolated and shown to transfer electrons to cytochrome cd1 nitrite reductase (Koutny´ et al., 1999). N2OR seems to exhibit the same ‘pseudo speciﬁcity’ towards alternative electron donors as discussed for cytochrome cd1 nitrite reductase (Williams et al., 1995). Support for this concept comes from a mutant affected both in cytochrome c550 and pseudoazurin. The double mutant grows poorly under anaerobic conditions and secretes nitrite into the medium (Pearson et al., 2003).

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A theoretical docking study based on the crystal structures of both N2OR and the candidate donor molecules shows the feasibility of such a concept. Complexes were selected for force ﬁeld energy minimization and proximity of haem c or type-1 Cu ligands of the cupredoxin to CuA. A tenable model involves the interaction of seven glutamate and eight aspartate residues of N2OR with 13 lysine residues on cytochrome c550 (Mattila and Haltia, 2005). Complementary hydrophobic patches on both proteins realize the contact between N2OR and cytochrome c550. The patch covers surface areas and residues of both N2OR subunits. The charged residues are thought to promote the formation of the complex and orientation of reaction partners, whereas the hydrophobic patch is more important for binding afﬁnity. The modelling situation is similar when applied to the cupredoxin. It should be noted, however, that the role of a cupredoxin does not seem to be universal throughout the denitriﬁers. Canters and coworkers (Vijgenboom et al., 1997) showed that the expression of azurin of P. aeruginosa is not coupled to denitrifying conditions, but maximal when entering stationary phase and oxidative stress conditions. We found a similar scenario for P. stutzeri where azurin (previously thought to be absent from this bacterium (Matsubara et al., 1982)) is expressed under conditions of Cu supply approaching toxic concentrations, but not under conditions of anaerobic denitrifying growth and low Cu (U. Heimann and W.G. Zumft, unpublished data). 11.1.3. A Putative Novel Cytochrome c The NGCs of Azoarcus sp. EbN1, Burkholderia spp., Cupriavidus spp., De. aromatica, Ca. fetus, Pa. denitrificans, R. solanacearum, Rf. ferrireducens, Tm. denitrificans and W. succinogenes all display one or two nosC genes. In Pa. denitrificans nosC is located in an operon together with the nosR gene, which may indicate a Nos-related function. The deduced gene products of the nosC genes of about 150 amino acids have a conserved CysXaaXaaCysHis motif suggestive of the protein as a c-type cytochrome (Fig. 16). An N-terminal hydrophobic stretch in NosC can function as a membrane anchor or as a signal sequence for periplasmic location. For W. succinogenes it has been postulated that NosC is part of an electron transfer pathway to N2OR (Simon et al., 2004). A redox role in binding or release of Cu, considering the differential afﬁnity of Cu(I/II) to NosL, is also feasible. 11.1.4. Electron Transfer to N2O Reductase in W. succinogenes The C-terminus of N2OR from W. succinogenes has an extension that forms a haem c domain fused to the CuA domain. The same situation is realized in

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Figure 16 Consensus sequence of the heme c-binding region of NosC proteins. For abbreviations see Table 2.

Tm. denitrificans. It is likely that this represents an additional electron transfer domain mediating between an exogenous donor and the CuA centre. The presence of covalently bound haem subjects this N2OR additionally to the maturation apparatus for c-type cytochromes (Kranz et al., 1998). C-terminal extensions of N2OR that fall short of the haem c-binding motif exist in De. aromatica and M. magnetotacticum. A cognate monohaem c-type cytochrome (9.2 kDa) is active with the N2OR of Wolinella and reduces N2O with second-order kinetics (Zhang and Hollocher, 1993). This cytochrome encoded by the Wolinella gene Ws0700 is not a part of the nos cluster (Baar et al., 2004). Cytochrome c551 from P. aeruginosa (Eo0 (pH 7) ¼ +286 mV, Horio et al., 1960) is also active with N2OR from Wolinella, although to a lesser degree. Figure 17 depicts the N2O respiratory system and putative electron donor pathway for W. succinogenes. The gene for the DnrD transcription factor, discussed above, is part of the NGC, giving weight to the concept that a Crp-Fnr factor is the regulator for nos gene expression. The NGC of Wolinella carries homologues of napG and napH, encoding FeS proteins (Table 2). They were termed as nosG and nosH because of their location, and also because W. succinogenes carries genuine napG and napH genes. NosG and NosH are proposed to function like the Nap proteins in a quinol loop,

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Figure 17 The N2O respiration system of W. succinogenes. Genes and gene products are correlated by the same shading; genes encoding assembly factors are cross-hatched; their gene products are not depicted (but see Fig. 14). The genes encoding putative electron transfer components of a quinol-oxidizing system are boxed. The topology of NosH and NosG is adopted from the Nap homologues. dnrD encodes the presumed regulator for nos gene expression. Modiﬁed from Simon et al. (2004).

providing electrons either directly to N2OR or via a cytochrome component (Simon et al., 2004). NosH is predicted as a membrane protein. It exhibits the same cysteine and FeS signatures as NosR and may be a substitute for the polyferredoxin domain of NosR. NosG and NosH homologues are also found in the NGCs of Ca. fetus, De. aromatica, M. magnetotacticum and Tm. denitrificans.

11.2. Role of Flavoproteins NosR and NosX 11.2.1. NosR The nosR gene is found in the NGCs of many N2O-respiring bacteria located mostly adjacent to and upstream of nosZ. nosR was ﬁrst identiﬁed by mutational analysis. Its inactivation results in a complete lack of nos gene expression since it affects the transcription of both nosZ and the nosD operon. Recent studies indicate a broader role in N2O respiration. The

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Figure 18 Topology model of the protein domains of NosR. Transmembrane helix 1 is cleaved upon membrane insertion and is not part of the mature protein. Helices four and six are arbitrarily placed adjacent to emphasize a presumptive interaction (X) of the Cys(Xaa)3CysPro motifs by a metal or –SH redox chemistry. The positions D242 and E392 in the periplasmic domain denote the amino acid codons used for PhoA fusions; the boxed region marks the glycine-rich sequence.

nosR gene product is a membrane-bound protein; it has been genetically engineered with a histidine tag and puriﬁed either as the holoprotein or a ﬂavoprotein fragment (Wunsch and Zumft, 2005). The protein has a complex topology with different cofactors at either side of the cytoplasmic membrane. Figure 18 shows its predicted topology with a ﬁve-helix bundle. The transmembrane portion is N-terminally preceded by a large periplasmic domain, comprising about half of the protein, and followed by a cytoplasmic domain with a polyferredoxin sequence motif. Periplasmic exposure of the N-terminal domain is proven from phoA reporter gene fusions into the codons for Asp242 and Glu392 (Cuypers et al., 1995). The EPR spectrum of puriﬁed NosR supports the presence of FeS clusters (S. Andrade, P.M.H. Kroneck and W.G. Zumft, unpublished data). The Fe content is about eight atoms per 78 kDa, which would account for two [4Fe4S] clusters. The polyferredoxin region is delimited N- and C-terminally by a positively charged region, which is hypothesized to form a docking site at the protein surface (Fig. 19). Two Cys(Xaa)3CysPro motifs conserved in every NosR protein follow transmembrane helices four and six; they may undergo

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reversible metal binding or SH redox chemistry for positioning these two helices in a functionally important interaction. The C-terminal part of NosR has a structural homologue in NapH of E. coli (with the same Cys(Xaa)3CysPro and FeS modules). Reporter gene fusions have indicated a cytoplasmic location of the FeS domain for NapH (Brondijk et al., 2004). The combination of Cys(Xaa)3CysPro motifs and FeS modules is found in several other redox proteins with putative regulatory or sensory function and is anticipated to have a broader signiﬁcance (Wunsch and Zumft, 2005, and references therein). The periplasmic domain of NosR has structural similarity with the FMNbinding domain of the NqrC subunit of the Na+-translocating NADH:quinone oxidoreductase (Yeats et al., 2003). NqrC carries FMN covalently bound as phosphodiester to a threonine residue (Zhou et al., 1999; Hayashi et al., 2001). An invariant threonine is located in the periplasmic ﬂavinbinding region of all NosR proteins (Fig. 19). Given the spectral evidence for ﬂavin and the cofactor-binding consensus sequence, it is assumed that NosR of P. stutzeri carries its ﬂavin covalently bound to Thr163 (Wunsch and Zumft, 2005). The threonine residue is followed by a short a-helix as a structural element in the FMN-binding domain. Following the ﬂavin-binding region, the periplasmic domain has a remarkably conserved glycine-rich region, which is preceded by a short helix and a hydrophobic stretch. In unprocessed NosR of P. stutzeri this motif is positioned at amino acids 289–312 (Fig. 19). Glycine-rich motifs have repeatedly been brought into the context of nucleotide binding. Whether this is related to the ﬂavin content is open, as is the problem for the source of the ﬂavin cofactor. Although it is tempting to speculate about a NosR–NosX relationship towards this aim, it is clearly to be noted that the dissemination of NosR and NosX is only partially overlapping among the NGC-harbouring bacteria. A genuine NosR protein can easily be recognized from the assemblage of the various distinctive structural elements (Table 5). The NosR proteins of the genera Cupriavidus, Ralstonia, Rhodoferax and Thiobacillus are structurally modiﬁed. They carry a C-terminal extension of about 150 amino

Figure 19 Sequence alignments of conserved regions of the NosR protein. (A) Flavin-binding region. The alignment depicts the region around the putative cofactor-carrying threonine (*), which is delimited in the amino acid sequence N- and C-terminally by indels. (B) Glycine-rich region within the periplasmic domain; (C) C-terminal polyferredoxin region. Colour coding as of the default ﬁle of ClustalX 1.83 except for cysteine. For abbreviations see Table 2; Ddeh, Desulfitobacterium dehalogenans; PdenI, Pa. denitrificans NirI; Rbal, Rhodopirellula baltica. (See Colour Plate Section in back of this volume.)

188

Species

Gene product

Flavin-binding region

Threonine motif

Glycine-rich motif

Five-helix bundle

2 C(X)3CP

Polyferredoxin motif

N. gonorrhoeae L. majuscula B. japonicum Pa. denitrificans M. magnetotacticum Rb. capsulatus Rl. baltica D. hafniense

NosR NosR NosR NirI NosR NosRX NosR CprC

+ + + + + + Shortened Absent

TXT TXT TXT TXS TXS TXS TXT TXS

GX5GXGX3GG GX5GXGX3GG Modiﬁed1 Modiﬁed1

+ + + + + + + +

+ + + + + + + +

+ + + + + +

1

See Fig. 19.

WALTER G. ZUMFT AND PETER M.H. KRONECK

Table 5 Structural features in NosR proteins and homologues

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acids not found in other NosR proteins. The extension is characterized by hydrophobic regions, which are predicted to provide two or three further transmembrane domains. NosR of Azoarcus sp. EbN1 represents a unique case where the protein is composed of two subunits of 142 and 724 amino acids. The homologue of the C-terminal extension is encoded here by a short separate ORF which is located upstream of the principal NosR-encoding gene. The major subunit of 724 amino acids carries the ﬂavin-binding region and the FeS polyferredoxin signature. Figure 20 shows the phylogenetic relationship among NosR proteins. We ﬁnd NosR in the clades deﬁned in the NosZ tree that correspond to the a-, b- and g-proteobacteria. It is absent from those lineages, where the

Figure 20 Phylogenetic relationship among NosR proteins and homologues. Greek letters represent the corresponding groups of proteobacteria. For abbreviations see Table 2 and Fig. 19. Asterisks indicate deviations from the taxonomic group. Sequence retrieval and methods as in Fig. 10.

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database is currently limited to a single NGC-harbouring representative. It would be unreasonable to assume that all these entries, stemming from genome projects, represent exceptions of otherwise nosR-harbouring lineages. The observed dissemination thus limits the NosR function to a speciﬁc type of N2O respiratory system whose biochemical basis needs to be deﬁned. The phylogenetic relationships within the NosR clades closely reﬂect the patterns with the other Nos proteins. This is interpreted as co-evolution of this group of proteins. The nosR sequence of the cyanobacterium Lyngbya majuscula stems from a DNA fragment that encodes biosynthetic genes for curacin A, a potent cancer cell toxin (Chang et al., 2004). The deduced cyanobacterial protein exhibits all structural elements of NosR, making it a genuine member of the family (Fig. 19; Table 5). Intriguingly, nosR is followed by a short sequence segment that shows similarity to nosZ. This opens the possibility that Lyngbya will be the ﬁrst-known case of an N2O-reducing cyanobacterium. An N2O-generating NO reductase has been found in Synechocystis sp. (Bu¨sch et al., 2002), which shows that certain cyanobacteria have the capability to metabolize gaseous N oxides. Prior to the knowledge about denitriﬁcation enzymes in cyanobacteria, production of N2O from nitrite has been reported (Weathers and Niedzielski, 1986). Six branches form largely individual lineages in the NosR phylogeny, and the structural features of these NosR proteins are shown in Table 5. N. gonorrhoeae follows structurally the canonical NosR pattern, but does not branch within its cognate group of b-proteobacteria. Strains of N. meningitidis have only short N-terminal fragments of NosR (see Section 8.1), which cluster in the N. gonorrhoeae lineage. NosR of M. magnetotacticum is on a deepbranching lineage together with NosRX from Rb. capsulatus, but otherwise independent from N2O-utilizing species. Both proteins do have the regular NosR elements except the consensus glycine motif. The Nos proteins of M. magnetotacticum are phylogenetically related to the e lineage (Wolinella clade) and to De. aromatica, where no NosR proteins are annotated in the genomes. It is possible that nosR of M. magnetotacticum was acquired laterally as suggested for the other nos genes, but from a different source. NosRX is a singular case of a fusion protein encoded outside the NGC in the Rb. capsulatus SB1003 genome (gene no. RRC03556). The 98.7-kDa protein comprises 925 amino acids, versus 78 kDa and about 735 amino acids for NosR encoded within the NGC. NosX is fused C-terminally to NosR which otherwise exhibits, the usual structural features except for the glycine motif (Table 5). The two NosR proteins of Rb. capsulatus follow quite independent trajectories with NosRX independent from the NGC phylogeny (Fig. 20). While NosX is a periplasmic protein, the fused X domain of NosRX

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is predicted to be exposed to the cytoplasm but with a C-terminal membrane anchor. Whether NosX has the same or a different function in either cell compartment remains to be seen. Lack of the glycine-rich sequence motif and further individual modiﬁcations (Table 5) are observed in three NosR homologues, deeply separated from genuine NosR proteins in the phylogenetic tree. NirI of Pa. denitrificans acts on the transcription of the structural gene for cytochrome cd1 nitrite reductase (Saunders et al., 1999). The ﬂavin-binding sequence is ThrXaaSer, a modiﬁcation which is otherwise noted in NosR of M. magnetotacticum, NosRX and CrpC. NirI cannot be considered as a simple paralogue of NosR in Pa. denitrificans given its independent phylogeny from NosR of this bacterium (Fig. 20). In the genome of Rhodopirellula baltica a nosR gene has been annotated (Glo¨ckner et al., 2003). The bacterium belongs to the Planctomycetes, a group that came into scientiﬁc focus because of its organelle-like inclusion as the site for the Anammox process. Rl. baltica scores several homologues of the NosX–ApbE family, but otherwise there are no genuine nos genes in this bacterium. The NosR protein lacks the polyferredoxin cluster and the ﬂavinbinding domain is somewhat shortened, thereby losing the glycine-rich region. The role of the NosR homologue in this marine bacterium is not known. CprC of Desulfitobacterium dehalogenans is believed to exert a transcriptional effect on genes necessary for o-chlorophenol degradation (Smidt et al., 2000). It is also encoded in the genome of D. hafniense. CprC is structurally and substantially modiﬁed versus NosR with respect to the ﬂavin-binding and polyferredoxin regions (Table 5). Genetically engineered forms indicate that NosR is an indispensable factor in sustaining cellular N2OR activity. Recombinant manipulations of the periplasmic domain or metal-binding centres of NosR cause the loss of whole-cell N2O-reducing activity. This indicates a redox role for NosR with mature N2OR as the target, in addition to its function in nosZ transcription. N2OR proteins synthesized in the genetic background of defective NosR proteins show the UV–vis spectral signatures of CuA and CuZ (Wunsch and Zumft, 2005). The N-terminal, periplasmic domain and the transmembrane and C-terminal domains of NosR have been expressed as separate proteins that complement each other functionally. Certain structural modiﬁcations of NosR transform the CuZ centre to the CuZ* state, similar to what has been observed for the NosX phenotype (Wunsch et al., 2005) or the form of N2OR generated by exposure of frozen cell extract from Pa. pantotrophus to air (Rasmussen et al., 2002). Since CuZ* becomes manifest upon nosR and nosX mutations, it must have physiological status.

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11.2.2. NosX NosX has been shown by mutational evidence to provide an essential function for N2O respiration in S. meliloti (Chan et al., 1997) and Pa. denitrificans (Saunders et al., 2000). The nosX gene is a moderately frequent component of NGCs (Table 2). Where present, it follows immediately downstream of nosL or is located upstream of nosZ or nosC, in opposite transcriptional orientation to nosZ. Pa. denitrificans harbours paralogous nosX and nirX genes, which are part of the nosCRZDFYLX (Saunders et al., 2000) and nirSIX (Saunders et al., 1999) gene clusters, respectively. NosX(NirX) had been proposed to be involved in CuA biogenesis, since unfractionated periplasm of Pa. denitrificans failed to show the hyperﬁne structure in the EPR spectrum diagnostic for CuA (Saunders et al., 2000). However, investigation of the NosX phenotype at the level of isolated N2OR reveals that a catalytically active holoN2OR with spectral evidence for both Cu centres is synthesized in a nosXnirX background. The absence of the NosX function leaves the catalytic centre at the CuZ* state, but a direct role in CuA biosynthesis cannot be conﬁrmed (Wunsch et al., 2005). NosX is a 34-kD ﬂavoprotein with a Tat-speciﬁc signal sequence (Fig. 13). Location of NosX in the outer cell compartment is consistent with a role directed at N2OR. A redox role is likely from its ﬂavoprotein nature. When present, NosX is required for the maintenance of N2OR activity, which may be achieved either independently of or in combination with NosR. A feasible scenario sees NosX functioning in concert with NosR, for instance, in an electron transfer route with NosR as a quinol–NosX oxidoreductase (Wunsch et al., 2005). Cooperation between NosR and NosX may be inferred from existence of the NosRX fusion protein of Rb. capsulatus. HoloN2OR can be expressed from nosRZDFY without co-expressing nosX (Wunsch et al., 2003). Since NosX is not ubiquitous, a certain variant of N2O respiration or type of N2OR does not require this component or cells have a substitute for it. NosX and ApbE are homologues of the same protein family which led to the consideration of ApbE proteins as functional substitutes (Wunsch et al., 2003). This was prior to the knowledge of the ﬂavoprotein nature of NosX. A NosX–ApbE substitution would probably depend on the same cofactor content of both proteins. ApbE is a 36-kDa monotopic innermembrane lipoprotein, which affects FeS centre synthesis in enzymes for thiamine biosynthesis. Periplasmic exposure is necessary for its function but its precise role and cofactor content, if any, is unknown (Skovran et al., 2004). Figure 21 shows a phylogenetic tree of NosX and ApbE proteins. Overall the ApbE family harbours many more members. Selected here were only ApbE proteins in the context of bacteria harbouring an NGC. No major

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Figure 21 Phylogenetic relationship among NosX and ApbE proteins. ApbE was retrieved only from NGC-harbouring species. Thus entries do not represent the entire pool of ApbE family members. The transcriptional direction of the underlying nosX genes with respect to nosZ is opposite in the a and b groups of proteobacteria (Table 2). For abbreviations see Table 2 and Fig. 19; Zmob, Zymomonas mobilis; Sgor, Streptococcus gordonii. Asterisk indicates deviation from taxonomic group. Sequence retrieval and methods as in Fig. 10.

distinctive features exist to differentiate NosX and ApbE. ApbE proteins are often encoded from paralogous genes. The overall positional sequence identity within the NosX–ApbE group is considerably lower compared to that among the other Nos proteins. We have considered gene neighbourhood, distance from nos genes, distribution within the cognate clade and existence and nature of a signal peptide, which results in a conservative attribution of NosX proteins as shown in Fig. 21, and restricts the dissemination of NosX to the a- and b-proteobacteria. Within the NosX group of the a-proteobacteria we ﬁnd the homologue of A. cycloclastes. This bacterium has been discussed in Section 8.1 in the context of lateral gene transfer. Zymomonas mobilis and Streptococcus

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gordonii, which neither denitrify nor harbour nos genes, have nosX homologues annotated in their genomes. It seems that NosX-type proteins have also cellular functions not related to N2O utilization. The S. gordonii nosX is part of a gene region encoding quinol–NADH oxidase. The system is involved in bioﬁlm formation (Loo et al., 2004).

12. A GLIMPSE OF HISTORY Joseph Priestley prepared nitrous oxide in 1772, two years before he described ‘dephlogisticated air’, later called oxygen. The anaesthetic property of N2O was found by S.L. Mitchill in 1795 and again by H. Davy in 1800 (for discussion of the role of Mitchill see Bergman, 1985). Present day usefulness as the only inorganic anaesthetic is being discussed with opposing arguments (Hopkins, 2005; Jahn and Berendes, 2005). Curiously, the mechanism of its anaesthetic property has never been fully clariﬁed. In addition to its medical application, N2O has remained to society an amusement tool with serious adverse side effects resulting in myeloneuropathy. The French scientists Gayon and Dupetit (1886) described the denitriﬁcation phenomenon after several years of investigating nitrate fermentation. Nitriﬁcation, i.e. the appearance of nitrate from ammonia was studied at the same time and it seems that ‘denitriﬁcation’ was coined as an antonym. The term appears in a short paper in 1882 by Gayon and Dupetit, which we surmise is its ﬁrst use. Gayon and Dupetit’s work qualiﬁes for its authoritative and comprehensive nature, having, as often in the scientiﬁc process also predecessors, anticipating or contributing to the discovery. We consider among them Schloesing, Reiset, Meusel (the latter ascribed ﬁrst to bacteria the reduction of nitrate to nitrite) and also Goppelsro¨der, who was, in turn, in a very different setting the experimental forerunner to Tswett, the discoverer of chromatography (Jo¨ssang, 1992). Formation of N2O (‘protoxyde d’azote’ in French) was observed already by Gayon and Dupetit but entered its ﬁrst phase of oblivion until Beijerinck and Minkman (1910) studied again the production and consumption of N2O by bacteria. Using Bacillus stutzeri (now P. stutzeri) and Micrococcus denitrificans (now Pa. denitrificans), these authors suggested a reaction sequence to proceed from nitrate or nitrite to N2O, and further on to N2. The status of N2O as an intermediate in denitriﬁcation, however, did not receive broad acclaim, and passed another long phase of oblivion until Mori and coworkers started to search for the molecular entities responsible for the process in cell-free systems, and addressed N2O metabolism and

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denitriﬁcation in a series of papers (Iwasaki et al., 1963; Matsubara and Mori, 1968). Around the time of positioning N2O as an intermediate ﬁrmly in the denitriﬁcation process, the ﬁrst transition metal N2O complex was prepared (Armor and Taube, 1969). Denitriﬁcation is generally investigated under the axiom of a facultative anaerobic bacterial activity. Gayon and Dupetit (1882) stated the anaerobic nature of the underlying organismal activity and its loss upon air exposure. However, Meiklejohn (1940) found two species of Pseudomonas that completely removed nitrate in an aerated culture, putting ‘aerobic denitriﬁcation’ on the scene. Her study still followed a concept where nitrate should supply the cell with O2, a view that had already been refuted years ago in favour of the parallel functions of O2 and the N oxide, both providing electron acceptors for aerobic and anaerobic respiration, respectively (Kluyver and Donker, 1926). More recently, Robertson and Kuenen (1984) revived the subject of aerobic denitriﬁcation. Until today, it has not found its full molecular explanation, but continues to be of interest for biotechnological applications to remove nitrate from wastewater under aerobic conditions. N2OR was discovered in P. stutzeri ZoBell by pursuing as search principle the trace-metal requirement for N2O utilization (Zumft and Matsubara, 1982). The ZoBell strain, initially introduced under the name Pseudomonas perfectomarinus (sic) by Payne and coworkers (Rhodes et al., 1963), became one of the principal model organisms for denitriﬁcation research. It was also the source for the ﬁrst NO reductase to be recognized and studied biochemically (Heiss et al., 1989). The observation of gas formation from nitrate by halobacteria (ElazariVolcani, 1940) attained signiﬁcance when the gas species were identiﬁed as N2O, NO and N2 in ‘Halobacterium marismortui’. It became the ﬁrst archaeon shown to be capable of denitriﬁcation (Werber and Mevarech, 1978). In an unexpected turn, fungi were found to release N2O (Bollag and Tung, 1972) and the underlying enzyme was identiﬁed as a cytochrome P-450 species (Shoun and Tanimoto, 1991). This extended the denitriﬁcation trait to the eukaryotic domain of life. Nitrate fermentation versus nitrate respiration developed as an interesting evolutionary topic with the debate continuing until today on the reasonable positioning of N-oxide respiratory enzymes in the sequence of biochemical evolution. Broda (1975) discussed these aspects in detail prior to the recognition of the structural relationships among cytochrome oxidase, NO reductase and N2OR (Saraste and Castresana, 1994; van der Oost et al., 1994). N2OR was biochemically and genetically characterized ﬁrst from P. stutzeri (Coyle et al., 1985). Determination of its primary structure (Viebrock and Zumft, 1988) preceded that of all other enzymes involved in respiratory and

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dissimilative nitrate utilization. It was also the starting point to track by molecular means the denitriﬁcation process in the environment by the development of gene probes (Linne von Berg and Bothe, 1992; Smith and Tiedje, 1992). As biochemical analysis of N2OR proceeded, its Cu centres were recognized as novel types of metal complexes among the Cu protein family in the form of the mixed-valent binuclear CuA centre (Kroneck et al., 1988) and the ﬁrst biological active Cu–S cluster known (Rasmussen et al., 2000). With the crystal structures of N2OR from Ma. hydrocarbonoclasticus establishing a tetranuclear Cu cluster for the catalytic centre (Brown et al., 2000b), and a high-resolution structure of the Pa. denitrificans enzyme (Haltia et al., 2003) the ﬁeld has advanced to three-dimensional structural information, with half a dozen different Nos proteins more waiting in line.

13. CONCLUSIONS AND PERSPECTIVES Prokaryotic metabolism of N2O inﬂuences the composition of the atmosphere, its warming and the decomposition of ozone. The sink and source mechanisms for N2O on land and in the oceans are beyond the scope of this review; however, they are of prime environmental importance and have to be seen against the mechanistic background of the process detailed above. The possibility to probe now the environment by molecular means has been a signiﬁcant outcome of the genetic analysis of the process. The biochemistry and genetics of N2O respiration have provided novel insights. Two new types of Cu metal centres were discovered studying N2OR. Unraveling the signal peptide of N2OR stands at the roots of the discovery of a new transport system for proteins in the folded state. Insertion of the prosthetic Cu ions into the protein is a highly specialized process which provides insights in the complex ways the bacterial cell synthesizes a metalloprotein. Genomics has yielded new and unexpected knowledge about the overall process and today allows predictions and conclusions based on the robust body of accumulated evidence from model organisms. In spite of much progress since the discovery of the enzyme, numerous questions remain. Have we already identiﬁed all relevant genes for N2O respiration? In which genomic network are they embedded? How do we reconcile with the observation of N2O metabolism and N2 formation in prokaryotes with the apparent lack of genomic information for the Z-type enzyme? How are other types of bacterial and archaean N2ORs structured? How does internal electron transfer in N2OR proceed? What are the structures of CuZ* and anaerobically isolated CuZ species? What is the mechanism of N2O activation and reduction? How

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are Cu and sulfur delivered to the catalytic site? What are the precise roles of the ﬂavoproteins NosR and NosX? In perspective, there is hardly a biochemical or cell biological aspect where fresh approaches will be redundant. Deﬁning the electron-donation pathways; formulating a consistent mechanism for catalysis; understanding the processes of signalling, regulation and expression of the underlying genes; expanding our views on the organismic variety and on ecological aspects; all are areas for inspiring future activities. One objective in understanding anaerobic N-oxide regulation is to interrupt the cellular network and develop an artiﬁcial aerobic denitriﬁcation process for biotechnological applications. The crucial breakthrough to manipulate the cellular regulatory network is still ahead. Denitriﬁcation is applied in wastewater treatment. Removal of ﬁxed N has become a pressing problem in many areas of the globe. Knowledge about respiratory N-oxide metabolism is necessary to understand those ramiﬁcations and develop useful cleanup strategies. In agriculturally managed ecosystems with high N input, N2O emission is an important concern. Understanding the factors that inﬂuence N2O evolution will help to take measures to mitigate N2O emissions.

ACKNOWLEDGEMENTS We thank our students, coworkers and collaborators, named in the cited references, for their valuable contributions. WGZ is indebted to H. Ko¨rner for bioinformatics assistance, phylogenetic analyses and helpful discussions; PMHK thanks O. Einsle for his assistance with ﬁgures. We also thank D.M. Dooley, R.J.M. van Spanning and E. Yabuuchi for personal communications. Unpublished sequence information incorporated in this article was made freely available by TIGR Microbial Database, The Joint Genome Institute Microbial Genomics Database, The Wellcome Trust Sanger Institute, Centre of Marine Biotechnology of the University of Maryland and ERGO Database of Integrated Genomics Inc. Work in the authors’ laboratories was supported by the Deutsche Forschungsgemeinschaft and Fonds der Chemischen Industrie.

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A Circadian Timing Mechanism in the Cyanobacteria Stanly B. Williams Department of Biology, Life Science Building, University of Utah, Salt Lake City, UT 84112, USA

ABSTRACT Cyanobacteria such as Synechococcus elongatus PCC 7942, Thermosynechococcus elongatus BP-1, and Synechocystis species strain PCC 6803 have an endogenous timing mechanism that can generate and maintain a 24 h (circadian) periodicity to global (whole genome) gene expression patterns. This rhythmicity extends to many other physiological functions, including chromosome compaction. These rhythmic patterns seem to reﬂect the periodicity of availability of the primary energy source for these photoautotrophic organisms, the Sun. Presumably, eons of environmentally derived rhythmicity – light/dark cycles – have simply been mechanistically incorporated into the regulatory networks of these cyanobacteria. Genetic and biochemical experimentation over the last 15 years has identiﬁed many key components of the primary timing mechanism that generates rhythmicity, the input pathways that synchronize endogenous rhythms to exogenous rhythms, and the output pathways that transduce temporal information from the timekeeper to the regulators of gene expression and function.

Copyright r 2007 by Elsevier Ltd. ADVANCES IN MICROBIAL PHYSIOLOGY VOL. 52 All rights of reproduction in any form reserved ISBN 0-12-027752-2 DOI: 10.1016/S0065-2911(06)52004-1

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Amazingly, the primary timing mechanism has evidently been extracted from S. elongatus PCC 7942 and can also keep time in vitro. Mixing the circadian clock proteins KaiA, KaiB, and KaiC from S. elongatus PCC 7942 in vitro and adding ATP results in a circadian rhythm in the KaiC protein phosphorylation state. Nonetheless, many questions still loom regarding how this circadian clock mechanism works, how it communicates with the environment and how it regulates temporal patterns of gene expression. Many details regarding structure and function of the individual clock-related proteins are provided here as a basis to discuss these questions. A strong, data-intensive foundation has been developed to support the working model for the cyanobacterial circadian regulatory system. The eventual addition to that model of the metabolic parameters participating in the command and control of this circadian global regulatory system will ultimately allow a fascinating look into whole-cell physiology and metabolism and the consequential organization of global gene expression patterns.

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1. Circadian Clock Definition and Nomenclature . . . . . . . . . . . . . 1.2. The Cyanobacteria. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3. A Circadian Clock in the Cyanobacteria . . . . . . . . . . . . . . . . . 2. The Cyanobacterial Circadian Clock: The S. Elongatus PCC 7942 Kai Locus. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Sequence, Structure and Function of Clock Proteins and the Kai-Clock Complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. The KaiA Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. The KaiB Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. The KaiC Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4. The SasA Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5. The Kai-Clock Protein Complex . . . . . . . . . . . . . . . . . . . . . . 4. Clock-Controlled Gene Expression . . . . . . . . . . . . . . . . . . . . . . . . 4.1. No Solitary Output Pathway . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Chromosome Compaction . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Clock Input. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1. The CikA Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2. The LdpA Protein. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3. The Pex Gene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6. Other Components: The rpo (Sigma Factor) and cpmA Genes . . . . 7. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1. Kai Genes and Circadian Clock Evolution . . . . . . . . . . . . . . . 7.2. A Cyanobacterial Clock . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.3. A Final Comment. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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1. INTRODUCTION Time, of course, is a fundamental variable in biological systems. Biology requires consideration of time scales ranging from fractions of a second to billions of years. Curiously, when the considered time scale is intuitively comprehensible—per day, for example—it seems to evoke a particular appeal among both laymen and practicing biologists. There are undoubtedly many alluring temporal correlations and causations between non-biological processes and the rates of their associated biological phenomena. Yet most of these relationships have timescales that are missing from tangible human experience. What is one millisecond? What is one million years? In contrast, daily biological rhythms – circadian rhythms – are well within our realm of experience and have evoked a documented fascination among scientists and others for nearly three centuries (deMairan, 1729). There are consistent and therefore predictable daily ﬂuctuations in light (quality and quantity), humidity and temperature (magnitudes) caused, of course, by the presence of the Sun and the rotation of the Earth about its axis. It makes good sense, in scientiﬁc hindsight, that genetic selection on this planet for organisms that regulate their behavioral, metabolic and physiological states in response to these circadian ﬂuctuations would have occurred (Bu¨nning, 1973; Pittendrigh, 1988). This ongoing selection is especially pertinent for organisms like the cyanobacteria that are, in general, obligate phototrophs. As such, their primary energy and reductant source is predictably transient. Fortunately, there are also experimental data supporting this notion of evolutionary selection for temporal expectation. Data from direct competition experiments have clearly demonstrated the selective advantage of the circadian timing system in the cyanobacterium Synechococcus elongatus (S. elongatus) PCC 7942 (Ouyang et al., 1998). One of the more exceptional aspects of this contemporary circadian regulatory system is that it depends upon a timing mechanism that keeps relative time independent of environmental dynamics. (The most phenomenal aspect is discussed below.) Evidently, these daily environmental ﬂuctuations have, over evolutionary time, been ‘‘memorized’’ and duly incorporated into the temporal regulatory mechanisms themselves. Contemporary circadian clocks have clearly evolved as a result of predictable, daily environmental change, yet they are now deﬁned, in part, by their ability to keep time without such external input (Roenneberg and Merrow, 2002; Dvornyk et al., 2003). The S. elongatus PCC 7942 circadian clock has become the bacterial model system for timing studies in vivo (Fig. 1). However, circadian clock (or Kai) protein structural and biochemical studies have, in several cases, also

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Figure 1 Rhythmic gene expression and chromosome compaction in S. elongatus PCC 7942. (A) Bioluminescence (counts per second) recorded over time (hours) from a F(kaiB-luc+) reporter in an otherwise wild-type strain. Three independent data sets are graphed. The gray boxes indicate time without illumination. Such data can be recorded automatically for many days. This technology has allowed rapid growth in the study of cyanobacterial circadian rhythms. (B) Deconvolved ﬂuorescence microscopy images (red-cell autoﬂuorescence; green-DAPI stained DNA) of wild-type cells sampled during a light/dark cycle at ZT ¼ 0 and ZT ¼ 12. Note the diffuse state of the chromosome at ZT ¼ 0 and the arrangement of the DNA into distinct ‘‘nucleoids’’ at ZT ¼ 12. Cells are approximately 5 mm long. (See Colour Plate Section in back of this volume.)

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been conducted with proteins puriﬁed from the thermophilic, unicellular cyanobacterium Thermosynechococcus elongatus (T. elongatus) BP-1, the mesophilic, ﬁlamentous (multicellular) cyanobacterium Anabaena sp. strain PCC 7120 and the mesophilic, unicellular Synechocystis sp. strain PCC 6803. Except for the Anabaena sp., all of these cyanobacteria are known to have bona fide circadian clocks and the associated temporal regulatory systems (Kondo et al., 1993; Aoki et al., 1995, 2002; Onai et al., 2004). However, there are very few functional, genetic complementation data to support the idea that these organisms have circadian clocks that employ similar mechanisms. In fact, the T. elongatus BP-1 kaiB gene did not complement an S. elongatus kaiB strain under conditions where a Synechocystis sp. strain PCC 6803 kaiB gene did (Iwase et al., 2005). Nonetheless, the assumption throughout this review (and within the current scientiﬁc literature) is that all cyanobacteria utilize homologous or similar clock works. Support for this idea comes exclusively from comparative DNA sequence analysis and protein sequence and structural analyses (Dvornyk et al., 2003). Sequence comparisons of individual Kai (circadian clock) proteins from many cyanobacteria are shown in Figs. 3–5. Although there are circadian clockrelated data from research efforts using other cyanobacteria, S. elongatus PCC 7942, T. elongatus BP-1, and Synechocystis sp. strain PCC 6803 are the primary sources of mechanistic data as supported by a growing scientiﬁc literature that includes recent publication of individual Kai-protein structures, a Kai-protein-complex structure, an analysis of kai gene transcription regulation and work showing circadian clock-regulated chromatin dynamics. The most phenomenal data regarding the cyanobacterial circadian clock mechanism are that the mixing, in vitro, of three proteins KaiA, KaiB and KaiC from S. elongatus PCC 7942 and ATP can reconstitute a temperaturecompensated circadian timing system (Nakajima et al., 2005). In fact, the period of this rhythm in vitro is the same as that manifest by the identical Kai proteins in vivo. Wild-type Kai proteins – in or out of the cell – confer a free-running period of about 25 h and several mutant KaiC proteins that confer longer or shorter periods in vivo do the same in this in vitro phosphorylation assay (Nakajima et al., 2005). These data are quite remarkable and have been reproduced in several independent laboratories. Many of the data discussed here may be speciﬁc to the circadian timing systems in S. elongatus PCC 7942, T. elongatus BP-1, Synechocystis sp. PCC 6803 and closely related cyanobacteria. However, functional generalizations to other cyanobacteria, to eukaryotes and perhaps even to other bacteria are very likely as we begin to understand the complex relationships among daily environmental dynamics, the circadian clock mechanism, the subsequent global regulation of gene expression, and all other palpable, clock-regulated

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behaviors, metabolic functions, and physiological states. The current situation in the cyanobacterial clocks ﬁeld is very exciting yet somewhat daunting. We now have a basic timing mechanism – the phosphorylation state of KaiC protein – but how does that process work? How is that phosphorylation state regulated in vivo? How does temporal information get from the Kai-clock to the temporally regulated signal transduction networks? How are phase relationships established among various gene products and their associated functions? Is the common presumption that the clock allows an organism to anticipate imminent sunset or imminent sunrise a valid one? How does the clock oversee and regulate basic cellular functions like transcription and translation? How is initiation of DNA replication regulated by the circadian clock? What about cell division? How do behavioral, metabolic and physiological states then feedback onto clock function? As discussed below, the circadian clock is the master regulator in S. elongatus PCC 7942. It regulates global gene expression patterns as a function of circadian time (CT). Thus, the answers to these questions – and the countless others that will ultimately arise – will help us to better understand the intimate relationships that exist among cell metabolism, physiology and behavior and the unyielding and unending environmental variable known as time.

1.1. Circadian Clock Definition and Nomenclature As mentioned, the fount of any circadian rhythm is a timing mechanism dubbed the circadian clock. To be considered a true circadian clock by chronobiologists a timing mechanism must keep relative time and continue to drive oscillatory behaviors even under constant environmental conditions (Pittendrigh, 1981). The current model organism for circadian biology among the bacteria is the photoautotrophic cyanobacterium S. elongatus PCC 7942. Because this organism is an obligate phototroph, experimental conditions require constant illumination and temperature. Time under these constant conditions is designated as a ‘‘free run’’. The period of any oscillation during the free run is called the free-running period. To synchronize and re-set the S. elongatus PCC 7942 circadian clock, cultures are typically placed in the dark for 6–12 h before releasing them into a constant-light free run. The moment of release is considered Zeitgeber time zero (ZT ¼ 0) ZT is then measured as ordinary time. The ﬁrst half of a free-running cycle is subjective day and the latter half subjective night. Regardless of period length every circadian cycle has 24 circadian hours. Thus, for a mutant strain with a free-running period of 14 h each circadian hour is only 34.9 min long. This is an important point when physiological comparisons among

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strains with different free-running periods are required. Comparisons need to be made at similar relative times during the circadian cycle. circadian time 12 (CT ¼ 12 ) is halfway through the 24 circadian-hour cycle regardless of the amount of ‘‘real’’ time that has passed. In addition to their free running, self-sustained oscillations, true circadian clocks can be entrained. Entrainment is the process by which an environmental rhythm – generally light/ dark cycles – determines the period and phase relationship of an otherwise self-sustained oscillation. In other words, even though S. elongatus PCC 7942 gene expression rhythms have a 25 h free-running period, a culture grown under a 15 h light–15 h dark cycle will have a 30-h period to its gene expression, and other clock-controlled, rhythms. An authentic circadian clock must also temperature compensate such that its free-running period remains constant over a physiologically relevant temperature range. The thermophilic organism T. elongatus BP-1 maintains a 25 h free-running period in its gene expression rhythms at temperatures ranging from 30 to 60 1C (Onai et al., 2004). The nomenclature and definitions discussed above are useful for chronobiologists in interpreting and describing rhythmic patterns. However, they are oversimpliﬁcations. Environmental conditions such as light intensity do affect intrinsic, free-running periods as formalized in Aschoff’s Rule (Dunlap et al., 2004). Normally, as light intensity increases, diurnal organisms have shorter free-running periods and the converse holds for nocturnal species. Temperature compensation should not be confused with temperature insensitivity. In fact, circadian clocks have input signaling pathways that sense and respond to temperature variation. For some organisms rhythmic temperature changes are a strong entrainment cue, often as deterministic as a light/dark cycle (Liu et al., 1998). Again, a circadian oscillator compensates for temperature differences so that gross changes in period do not occur. This is, of course, distinct from a simple chemical or biochemical reaction rates that do change as a function of temperature.

1.2. The Cyanobacteria Cyanobacteria are a monophyletic group of photoautotrophic prokaryotes (Tomitani et al., 2006). They are bacteria not ‘‘blue-green algae’’. Their lineage is among the oldest on Earth, as suggested by the fossilized cyanobacterium-like organisms found in 3500 Ma-old conglomeratic Apex chert (Schopf and Packer, 1987; Schopf, 1993; Brasier et al., 2002). The genetic diversity among extant cyanobacteria, exempliﬁed by comparing the mol% G+C content of their genomes, is noteworthy. For example, Cyanobium sp.

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strain PCC 6707 has nearly 70% G+C, S. elongatus PCC 7942 has 55% G+C, and the Nostoc sp. strain PCC 7524 genome has only 39% G+C (Tandeau de Marsac and Houmard, 1987). Some of those fossilized, antiquated cyanobacteria evidently originated the use of water as a reductant during light-driven respiration (and subsequent photosynthesis). Of course, it was the concomitant oxygen production that allowed them to create our present day oxygen-enriched atmosphere (Kasting and Siefert, 2002). Cyanobacteria and plant chloroplasts are homologous. It is generally accepted that modern plastids (including the land plant chloroplasts) evolved from a free-living cyanobacterium after its sequestration by a primitive eukaryotic-like cell (Cavalier-Smith, 2002; Martin et al., 2002). The success of this particular 1–2 billion-year-old endosymbiotic event was evidently unique, as all extant plastids are considered monophyletic (Douglas and Raven, 2003). The primary plastids, those directly descended from that ﬁrst cyanobiont, still exist among the rhodophyte, chlorophyte, and glaucocystophyte algae (Douglas and Raven, 2003). Evolutionary relationships among cyanobacteria and these plastids remain an intriguing and beguiling area of study (Morden and Golden, 1989; Suzuki and Bauer, 1995; Tomitani et al., 1999; Cavalier-Smith, 2000, 2002; Nobles et al., 2001; Martin et al., 2002; Ting et al., 2002; Raven and Allen, 2003). A glimpse into the genetic diversity among the cyanobacteria can perhaps be gained from their diverse morphologies (Fig. 2). The sizes and shapes of these organisms are diverse and fascinating. Many species, including those within the Aphanacapsa, Chroococcus, Merismopedia, Prochlorococcus, Synechocystis, and Synechococcus genera, grow as ovoid- or rod-shaped unicells ranging from 0.4 mm to 40 mm in diameter (Whitton and Potts, 2000). Obviously, many of these unicellular species live as single cells. However, others remain in tightly grouped cell aggregates after cell division (Paerl, 1996; Palinska et al., 1996). Some species appear to regulate this lifestyle choice based upon prevailing environmental conditions (Palinska et al., 1996). The cell aggregates are often highly organized, perhaps reﬂecting an underlying social order (Paerl and Priscu, 1998; Gorelova, 2000; De Philippis et al., 2005; Fuks et al., 2005). Other cyanobacterial species, including those from the Anabaena, Lyngbya, Scytonema, Spirulina, Stigonema, Tolypothrix, and Trichodesmium genera, are long, relatively thin, multicellular ﬁlaments commonly surrounded by a mucilaginous sheath. Generally, they are several micrometers in diameter and can be several hundred micrometers long (Green et al., 1989; Shi et al., 1995). Filamentous species can often form differentiated cells including hormagonia (motile fragment of a cyanobacterial ﬁlament), akinetes (resting cyanobacterial spores), and terminally differentiated cells called heterocysts, which develop

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Figure 2 Morphological diversity among the cyanobacteria. (A) A deconvolved ﬂuorescence microscopy image (red-autoﬂuorescence; green-DAPI stained DNA) of the ovoid-shaped cyanobacterium, P. marinus MIT9313. (B) As above, but image is of the ﬁlamentous cyanobacterium Anabaena sp. PCC 7120. (C) As above, except image is of a halotolerant Spirulina sp. isolated from The Great Salt Lake, Utah. USA. Each size bar represents 5 mm. (See Colour Plate Section in back of this volume.)

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under nitrogen-limited conditions and essentially function as anaerobic chambers for nitrogen ﬁxation (Golden and Yoon, 1998; Garrity, 2001; Yoon and Golden, 2001). As one might expect from their long evolutionary history, genetic diversity, and morphological malleability, cyanobacteria are liable to be found in every habitat that sunlight inﬁltrates. An extreme example is the cavedwelling Gloeocapsa sp. that survives under light intensities as low as 1 lx (0.02 mmol photon m2 s1) (Cox et al., 1981). Cyanobacteria have also been isolated from geothermal hot springs throughout the world. These thermophilic organisms have maximum growth temperatures ranging from 50 to 74 1C (Garcia-Pichel et al., 1998; Nandi and Sengupta, 1998; Ohto et al., 1999; Abed et al., 2002; Nakamura et al., 2002). T. elongatus BP-1 was isolated from the Beppu hot spring in Japan and has an optimal growth temperature of 57 1C (Yamaoka et al., 1978; Rippka et al., 1979; Stanier, 1980). Mesophilic cyanobacteria are ubiquitous and have been isolated from most dry land ecosystems, including karst and travertine regions. They also ﬂourish in benthic, limnetic, lotic, and pelagic fresh- and salt- water habitats (Paerl, 1996; Carmichael et al., 1997; Martinez et al., 1997; Olson et al., 1998; Ostensvik et al., 1998; Richter et al., 1998; Sano et al., 1998; Atkins et al., 2001; Cuvin-Aralar et al., 2002; Frank, 2002). S. elongatus PCC 7942, formerly known as Anacystis nidulans R2 and Synechococcus leopoliensis, was originally isolated from a freshwater habitat (see http://www.pasteur.fr/ recherche/banques/PCC/). Many mesophiles, including Oscillatoria agardhii, Aphanizomenon flos-aquae, Trichodesmium erythraeum, and Microcystis aeruginosa produce proteinacous gas vesicles that allow them to ﬁnd a ‘‘place in the sun’’ and perhaps avoid predation by regulating their buoyancy and, as a result, relative position in the water column (Damerval et al., 1989; Damerval et al., 1991; Walsby, 1994; Beard et al., 2002). Aphanothece halophytica, Dactylococcopsis salina, Microcoleus chthonoplastes, and Spirulina major, among many other cyanobacterial species, are halotolerant if not true halophilic organisms (Fig. 2). Cyanobacteria have also been isolated from extreme hypersaline environments (Brock, 1976; Ehrlich and Dor, 1985; Davis and Giordano, 1996). Psychrophilic species, like Nodularia harveyana, Phormidum frigidum, and Rivularia minutula, are characteristically the predominant life forms in their low-temperature environments. Their seemingly inhospitable habitats include the tundra, ice shelves, glacial moraines, and polar desert soils of both the Arctic and the Antarctic regions (Wharton et al., 1981; Sheath et al., 1996; Priscu et al., 1998). Cyanobacteria isolated from desert climates have an uncommon and astonishing ability to withstand multiple rounds of desiccation and subsequent re-hydration (Billi and Potts, 2002; Ballal and Apte, 2005; Ohad et al., 2005; Rothrock and

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Garcia-Pichel, 2005; Tamaru et al., 2005). As a general survival strategy in harsh environments, cyanobacteria have evolved mechanisms to ingeniously generate their own macroscopic, insular environments; these include such complex ecosystems as benthic, coastal tidal, hot spring, and hypersaline microbial mats, the environmentally essential (and extremely sensitive) cryptobiotic desert crusts, and even fresh- and salt-water blooms (Grotzschel and de Beer, 2002; Neilan et al., 2002; Urmeneta et al., 2003; Ohad et al., 2005; Rothrock and Garcia-Pichel, 2005). The range of growth rates and metabolic activities in cyanobacteria is also worth mentioning. Typical aquatic Synechococcus sp. have doubling times of several hours. It has been estimated that some cyanobacterial populations in the cold, oligotrophic, dry deserts of Antarctica may have doubling times of nearly 10,000 years (Friedmann et al., 1993; Nienow and Friedmann, 1993). Carbon dating in these polar regions supports one implication of this slow growth rate estimate by showing that living cyanobacterial cells may be over 1000 years old (Bonani et al., 1988). Clearly, many interesting survival strategies have evolved in the cyanobacteria and not all of them are particularly accommodating. Cylindrospermopsis raciborskii, Hapalosiphon fontinalis, Hormothamnion enteromorphoides, Umezakia natans, and most of the aforementioned genera make a surfeit of cyanotoxins as secondary metabolites (Beasley et al., 1989; Yoshizawa et al., 1990; Harada et al., 1991; Harada et al., 1994; Kuiper-Goodman et al., 1999; Ito et al., 2002; Mwaura et al., 2004; Welker et al., 2005; Kellmann et al., 2006; Stewart et al., 2006). These toxic metabolites are species-speciﬁc and include alkaloids, macrolides, and short linear or cyclic peptides that can be cytotoxic, hepatotoxic, or even neurotoxic to many organisms including mammals (Stewart et al., 2006). Cyanobacteria are often the primary producers in their ecosystems. Their metabolic activities such as photosynthesis, which generates the energy and reductant for subsequent carbon and nitrogen ﬁxation, and de novo vitamin and enzyme co-factor biosynthesis that have allowed them to inhabit practically every environment have also made them common participants in symbiotic associations. The Calothrix, Cylindrospermum, Fischerella, and Nostoc genera all include species that form endophytic, epiphytic, and true symbiotic relationships with numerous plants, fungi, sponges, and protists (Janson et al., 1999; Costa et al., 2001; Thomas, 2001; Gorelova and Korzhenevskaia, 2002; Guevara et al., 2002; Rikkinen et al., 2002; Wong and Meeks, 2002; Douglas and Raven, 2003). In fact, recent work has shown that many sponge-related compounds with activity against cancerous human cells are actually the products of the bacterial consortia living within those sponges (Wang, 2006).

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Given their exceptional variety of form, faculty, and function – a spectrum that has only been hinted at in this short overture – what biological properties might deﬁne the cyanobacteria? There are several including, perhaps, the ability to keep time. In addition to timekeeping mechanisms, they contain an intracytoplasmic, thylakoid membrane used to house their photosynthesis machinery (Kaftan et al., 2002). (If exception makes the rule, then consider Gloeobacter violaceous PCC 7421, whose photosystems are located within its cytoplasmic membrane (Rippka et al., 1974; Mangels et al., 2002; Inoue et al., 2004; Mimuro et al., 2005).) The cyanobacteria also utilize both photosystem I and II, as mentioned above, use water as the primary reductant during oxygenic photosynthesis (Fromme et al., 2001; Zak et al., 2001; Zouni et al., 2001). Also, cyanobacteria absorb light energy for photosynthesis by synthesizing and employing the chlorophyll a molecule, phycobiliproteins, and accessory phycobilin pigments, such as phycoerythrin, allophycocyanin, and phycocyanin (Brown et al., 1989; Meyer, 1994). High concentrations of these latter two pigments often make the organisms appear greenish-blue, leading to their previous, and incorrect, designation as ‘‘blue-green algae’’ and current moniker of cyanobacteria. Of course, not all are blue-green in color, and the broadly distributed prochlorophyte species use both chlorophylls a and b as antenna pigments and do not make the elaborate phycobilin, light-harvesting antennae (Palenik and Haselkorn, 1992; Kehoe and Grossman, 1994). In addition to all of these traits, cyanobacteria also appear to possess the circadian clocks, which are the focus of this review (Lorne et al., 2000; Dvornyk et al., 2002; Ditty et al., 2003; Dvornyk et al., 2003; Dvornyk and Nevo, 2004). Support for this speculation comes from data showing that 40 different cyanobacterial species have a kaiC gene (Lorne et al., 2000). Additionally, over 20 different cyanobacterial genomes have now been completely sequenced and multiple kai genes are omnipresent.

1.3. A Circadian Clock in the Cyanobacteria Physiological studies involving oxygen-sensitive nitrogen ﬁxation and oxygen-generating photosynthesis were among the ﬁrst to suggest that cyanobacteria have endogenous timing mechanisms (Golden et al., 1997). It was known that Anabaena sp. strain PCC 7120 and many other ﬁlamentous (multicellular), diazotrophic cyanobacteria could develop microaerobic heterocysts to separate spatially nitrogen ﬁxation from oxygen-evolving photosynthetic metabolism (Wolk, 1996; Kaneko et al., 2001). This knowledge begged the question of how unicellular species, lacking the option of differentiation, might balance these seemingly incompatible processes. Many

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groups subsequently demonstrated that some cyanobacteria are able to time the onset of nitrogen ﬁxation. Despite this consensus, the underlying, physiological and metabolic basis for the time schedule was ascribed to a variety of different cellular functions (Stal and Krumbien, 1985; Grobbelaar et al., 1986; Mitsui et al., 1986; Golden et al., 1997; Golden et al., 1998). Physical evidence of temporal metabolic regulation in bacteria confronted a long-standing belief that genuine circadian rhythms were a phenomenon restricted to eukaryotic organisms. This was primarily caused by models for circadian rhythmicity in eukaryotes that were based upon intercellular communication and nuclear sequestration of key clock components (Young and Kay, 2001). (The slightly paranoid among us also suspect the ever-present opinion among some biologists that bacteria are ‘‘too simple’’ for such ‘‘complex’’ phenomena as circadian rhythmicity.) Furthermore, it was assumed that a unicellular organism that could grow and divide at a rate faster than 24 h would not be able to maintain a circadian regulatory regimen (Edmunds, 1983; Kippert, 1987). Of course, this particular supposition has been refuted (Kondo et al., 1994; Mori et al., 1996). The ﬁrst convincing data demonstrating a prokaryotic circadian clock were reported nearly 16 years ago. These data showed a temperature-compensated 24 h rhythm of cell division in Synechococcus sp. strain WH 7803 (Sweeney and Borgese, 1989). At about the same time, Synechococcus sp. strain RF-1 (PCC 8801) was shown to have a rhythm in nitrogen ﬁxation that fulﬁlled the three criteria for circadian-clock control as outlined for eukaryotic circadian rhythms (Huang et al., 1990). Subsequently, it was demonstrated that strain RF-1 also maintained circadian rhythms in amino acid uptake (Chen et al., 1991). Visual data showing the alternation of nitrogen ﬁxation and photosynthesis in a Cyanothece sp., in which storage granules from the products of each process are visible by electron microscopy, has also been reported (Schneegurt et al., 1994) However, determining the underlying mechanism for cyanobacterial circadian rhythms required their demonstration in a cyanobacterial strain to which modern, molecular genetic tools could be applied, and for which an assay suitable for repeated sampling and high throughput could be developed. S. elongatus PCC 7942 met those needs (Fig. 1) (Kondo et al., 1993, 1994; Ishiura et al., 1998). Although somewhat alluring, prevailing notions that the circadian clock and its overt temporal rhythms evolved in cyanobacteria simply to separate oxygenic photosynthesis from oxygen-sensitive nitrogen ﬁxation are, at best, not the complete story. S. elongatus PCC 7942, the unicellular, photoautotrophic model system for prokaryotic circadian biology, does not ﬁx nitrogen (Herrero et al., 2001). In addition, Trichodesmium, a marine, ﬁlamentous (multicellular), and non-heterocyst-forming genus evidently uses both spatial and temporal

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separation of these incompatible metabolic activities (Berman-Frank et al., 2001). It appears that Trichodesmium sp. balance their rates of aerobic respiration and photosynthetic oxygen production so that no net oxygen is produced during part of the photoperiod, which opens a window of time for nitrogen ﬁxation. None of the strategies described above for the separation of oxygen evolution and nitrogen ﬁxation has an obvious requirement for circadian timekeeping. Nonetheless, the cyanobacterial circadian clock is real and is known to provide a selective growth advantage in S. elongatus PCC 7942 (Ouyang et al., 1998). In fact, the circadian clock and its concomitant circadian rhythms will not be truly understood until their complete description includes clear explanation of that selective growth advantage. Obviously, that explanation must involve comprehension of the proper timing of metabolic activities. It must also involve a description of the instantaneous adaptation of the timing system to predictable and, as importantly, unpredictable environmental ﬂuctuations. It must also involve understanding how the cellular timing mechanism measures and reacts to small changes in the cells, overall physiological state. And, it must explain the oversight of all of these processes by a timing mechanism that can allow or disallow fundamental cellular activities based upon the relative time of day. By comparison, describing the particulars of gene regulation and of protein–protein interaction within this remarkable, timed regulatory system is proving to be relatively simple.

2. THE CYANOBACTERIAL CIRCADIAN CLOCK: THE S. ELONGATUS PCC 7942 KAI LOCUS The kai (circadian clock) locus was ﬁrst identiﬁed using a genetic screen designed to test for phenotypic complementation of an altered gene expression rhythm in a mutant S. elongatus PCC 7942 strain (Ishiura et al., 1998). The altered-rhythm mutant had a 44 h free-running period rather than the wild-type 25 h period. The subsequently identiﬁed, complementing locus also restored wild-type rhythms to a large collection of other mutant strains that had altered period or arrhythmic gene expression patterns (Kondo et al., 1994; Ishiura et al., 1998). Ensuing analyses revealed that the relevant locus encodes three contiguous genes, kaiA, kaiB and kaiC and that these three genes are expressed from two different promoters (Ishiura et al., 1998; Ditty et al., 2003; Kutsuna et al., 2005). The kaiA gene has its own promoter (Ishiura et al., 1998). The kaiB and kaiC genes are expressed as a dicistronic message from a promoter immediately upstream of kaiB (Ishiura et al., 1998;

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Kutsuna et al., 2005). In S. elongatus PCC 7942 each individual kai gene is essential for the generation and maintenance of circadian rhythms in gene expression (Ditty et al., 2003). Data regarding some aspects of kai gene regulation were initially over-interpreted as being consistent with contemporary animal, insect and fungal circadian timing models that include transcription-based negative feedback loops (Ishiura et al., 1998). For example, overexpression of the kaiA gene elevates and makes arrhythmic expression from the kaiBC promoter (Ishiura et al., 1998), and a overexpression of the kaiC gene represses and makes arrhythmic expression from the kaiBC promoter (Ishiura et al., 1998; Nakahira et al., 2004). In fact, overexpression of the kaiC gene – and demonstrated concomitant KaiC protein overproduction – globally represses all gene expression in S. elongatus PCC 7942 (Nakahira et al., 2004). However, further experimentation (and somewhat less speculation) now supports a model whereby transcription information is not directly required for circadian timekeeping in this particular cyanobacterium (Tomita et al., 2005). As the overexpression data illustrate, proper transcription regulation is important for circadian clock control of rhythmic gene expression but circadian timing, as illustrated with puriﬁed Kai proteins, can and does continue without transcription or translation (Tomita et al., 2005). Other data too support a circadian clock model in cyanobacteria that excludes the dogmatic transcription–translation feedback loop as the generator of rhythmicity. Several mutations in S. elongatus PCC 7942 have been identiﬁed that alter the phase relationship between the kaiA and kaiBC gene expression rhythms without disrupting overall circadian timing. These speciﬁc data suggest that the relative transcriptional activity of expression from at least the kaiA promoter is not important for generating circadian rhythms (Katayama et al., 1999; Nair et al., 2002). In addition, expressing kaiC from a consensus-sequence Escherichia coli (E. coli) promoter in a kaiC genetic background can restore circadian rhythmicity to gene expression patterns. These data indicate that neither kai-speciﬁc promoter regions, nor their encoded speciﬁc transcription regulation information, are essential for generating circadian rhythms in S. elongatus PCC 7942 (Xu et al., 2003). One caveat to these data is that this synthetic E. coli promoter is also expressed rhythmically in S. elongatus PCC 7942, and is expressed in a pattern similar to that of the kai genes. To directly test whether transcription timing at the kai locus is required for circadian oscillations, the natural timing of kaiA was circumvented by expressing it from the S. elongatus PCC 7942 purF promoter, such that kaiA expression was delayed 12 h with respect to wildtype expression timing patterns (peaking at subjective dawn rather than the normal subjective dusk (Min and Golden, 2000)). In a kaiA null but otherwise wild-type background, an ectopic kaiA gene restores wild-type

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circadian rhythms of gene expression, whether it is expressed from its native promoter or from the oppositely phased purF promoter (Ditty et al., 2005). Curiously, in genetic backgrounds that increase the intracellular concentration of KaiB or KaiC protein, expressing kaiA from the purF promoter lengthens the free-running period of gene expression rhythms to nearly 33 h, whereas expression of kaiA from the wild-type promoter leaves the period unchanged (Ditty et al., 2005). Therefore, speciﬁc timing of kaiA transcription does play some role in circadian regulation in S. elongatus PCC 7942, for only when the oscillator is perturbed (e.g., elevated levels of KaiB or KaiC) is the timing of kaiA expression important for generating circadian rhythms in gene expression. It seems that transcriptional feedback is necessary for sustaining the circadian timing cycle, rather than being part of the underlying timing mechanism (Ditty et al., 2005). Recent analysis of the promoter region just upstream of the kaiB gene further supports the idea that speciﬁc transcription feedback is not a key aspect of circadian rhythmicity in S. elongatus PCC 7942 (Kutsuna et al., 2005). A transcription start site and, subsequently, a likely RNA polymerase holoenzyme binding site were identiﬁed. There is also an inverted repeat sequence that overlaps with the upstream portion of that putative polymerase binding site. The inverted repeat may bind an additional but so far unidentiﬁed, regulatory protein (Kutsuna et al., 2005). Part of the kaiB promoter region lies within the open reading frame of the upstream kaiA gene. Deletion of this cis-acting, negative regulatory element increases kaiBC gene expression by nearly 50-fold (Kutsuna et al., 2005). However, more mechanistically telling data regarding transcription feedback loops and kaiBC gene expression are that a series of incremental deletions from the 50 -end of the promoter sequence still result in promoter regions that retain rhythmicity and behave as wild type with respect to kaiA and kaiC overexpression (Kutsuna et al., 2005). It does not appear that the Kai proteins directly regulate their own expression or that the kai genes are, in any way, autoregulatory (Maloy and Stewart, 1993). In addition, recall that the Kai proteins generate a circadian rhythm of KaiC phosphorylation state in vitro and that this same phosphorylation rhythm persists in vivo without transcription or translation (Nakajima et al., 2005; Tomita et al., 2005). Obviously, the expression of the kai genes is important for circadian rhythmicity because the proteins must be present. However, cyanobacterial circadian rhythms in gene expression are not necessarily generated by regulatory transcription feedback loops but instead by the biochemistry intrinsic to Kai-protein interactions and the subsequent transduction of that timing information to relevant promoter sequence. Models hypothesizing the connection between Kai protein – that

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is, the circadian clock – function and these promoters must necessarily have additional players and mechanistic detail. Recent work demonstrates the existence of a clock- and thus kai- dependant rhythmic chromosome compaction process (Smith and Williams, 2006). This activity, and the related functional proteins, may turn out to be important parts of that next hypothetical, circadian clock model. Speculation from that work includes the notion that this compaction rhythm may physically conﬁscate promoter elements at certain times of the day and that process, coupled with timed regulatory protein availability, leads to the generation and maintenance of circadian rhythms in gene expression (Smith and Williams, 2006).

3. SEQUENCE, STRUCTURE AND FUNCTION OF CLOCK PROTEINS AND THE KAI-CLOCK COMPLEX In S. elongatus PCC 7942 all gene expression is under circadian clock control and consequently, the clock is considered a global regulator (Liu et al., 1995a; Nakahira et al., 2004; Smith and Williams, 2006). As discussed above, the kai genes A, B and C encode the small set of circadian timing proteins essential for circadian clock regulation of rhythmic gene expression patterns in this single-celled, photoautotrophic organism (Ishiura et al., 1998; Iwasaki et al., 2000). The physical interactions among these three proteins and the presumed or demonstrated, resultant changes in their biochemical actions combine to generate and maintain daily rhythms in global gene expression even, as is required by the circadian clock designation, in the absence of any environmental stimuli (Liu et al., 1995b; Ditty et al., 2003; Nakahira et al., 2004). Among the other clock-related proteins, two sensory kinases, SasA and CikA, play particularly important roles in the generation and maintenance of S. elongatus PCC 7942 circadian rhythms (Iwasaki et al., 2000; Schmitz et al., 2000). Comparative protein sequence analyses and resolution of three-dimensional structures of all these proteins has been an important part of the reductionist approach taken to decipher overall circadian clock function in this fascinating cyanobacterium. Many insights of these efforts are discussed below.

3.1. The KaiA Protein The S. elongatus kaiA gene is 855 bp in length, has a GTG start codon, and encodes 284 amino acyl residues that ultimately form a 32.6 kDa protein.

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The overall fold of this KaiA protein constitutes an amino-terminal pseudoreceiver domain and a carboxyl-terminal four-helix bundle domain (Ye et al., 2004). (Information and images regarding the structure of crystalline KaiA protein are available at http://www.rcsb.org/pdb/Welcome.do using code 1R8J.) The intracellular concentration of the KaiA protein remains fairly constant over time and has been determined to be near 500 molecules per cell (Kitayama et al., 2003). This is about 25 times less than the concentration of either the KaiB or the KaiC protein. Unlike KaiA, the absolute amounts of both of those latter proteins are known to oscillate over the circadian cycle (Kitayama et al., 2003). Physical properties of the KaiA protein suggest that it is a homodimer in solution and remains so upon crystallization (Kageyama et al., 2003; Garces et al., 2004; Uzumaki et al., 2004; Vakonakis and LiWang, 2004; Ye et al., 2004). The KaiA protein interacts directly with KaiC and causes an increase in the rate at which KaiC autophosphorylates (Iwasaki et al., 2002; Williams et al., 2002). This increased rate is modulated (somewhat decreased) by the presence of the KaiB protein (Williams et al., 2002). Interestingly, sequence comparisons among the 13 KaiA proteins shown in Fig. 3 suggested the potential for at least two functional regions (Williams et al., 2002). Notice that all of these proteins have similar sequence in their carboxyl-terminal regions but that the aminoterminal region sequence is highly variable. In fact, sequence related to the amino-terminal part of the S. elongatus PCC 7942 KaiA protein is completely absent in the KaiA proteins from the Nostoc sp. and the two Anabaena sp. listed in Fig. 3. Structure resolution efforts have conﬁrmed the presence of two distinct domains in the S. elongatus PCC 7942 KaiA protein and that the Anabaena sp. PCC 7120 and S. elongatus PCC 7942 carboxylterminal domains are essentially the same (Williams et al., 2002; Garces et al., 2004; Ye et al., 2004). Structure determination studies also revealed that the KaiA protein amino-terminal domain (in effect residues 1–135) folds in a manner consistent with it functioning as a bacterial-type receiver domain (Williams et al., 2002). On the basis of those structure data and the protein interaction data mentioned above, there has been speculation that KaiA acts as a conduit for timing adjustments to the clock as environmental conditions change – as stated above, a deﬁning feature of circadian clocks is their capacity for entrainment by environmental cycles (Pittendrigh, 1981; Williams et al., 2002). Here, of course, clock adjustment refers to changes in the phosphorylation state of the KaiC protein. Note that unlike a generic bacterial receiver domain, the amino-terminal part of KaiA does not have the properly positioned, conserved aspartyl residue that accepts a regulating phosphoryl group (Stock et al., 1995) (Fig. 3). The idea is that the KaiA protein receiver domain interacts directly with one or more environmentally

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Figure 3 A ClustalW (http://www.ebi.ac.uk/clustalw/) derived alignment of the amino acyl residue sequence from 13 putative KaiA proteins. Protein sequence was obtained from public sequence databases using a BLAST algorithm. Structure and function of the KaiA protein is discussed in the text. The bracket indicates a somewhat arbitrary beginning for the carboxy terminal region of these protein sequences. Protein sequences are from: (1) Crocosphaera watsonii WH8501, (2) Synechocystis sp. PCC 6803, (3) Synechococcus sp. PCC 7002, (4) Anabaena sp. PCC 7120, (5) A. variablis, (6) Nostoc punctiformes PCC 773102, (7) T. elongatus BP-1, 8) Trichodesmium erythraeum IMS101, (9) S. elongatus PCC 6301, (10) S. elongatus PCC 7942, (11) Synechococcus sp. PCC 9605, (12) Synechococcus sp. PCC 9902 and (13) Synechococcus sp. WH8102. Numbers on the left indicate the protein source. Numbers on the right indicate protein residue.

responsive proteins – the light-sensitive CikA protein is currently the best candidate – and then alters the conformation of its carboxyl-terminal domain (O’Hara et al., 1999; Uzumaki et al., 2004; Ye et al., 2004). This carboxyl-terminal domain is necessary and sufﬁcient to increase the autophosphorylation rate of KaiC (Williams et al., 2002; Garces et al., 2004; Uzumaki et al., 2004; Ye et al., 2004). Thus, environmental information

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would ﬂow through the KaiA protein to regulate the rate of KaiC autophosphorylation. As mentioned, the phosphorylation state of KaiC protein appears to be a measure of passing time (Ditty et al., 2003; Nakajima et al., 2005). An interesting aspect of considering the KaiA protein as an environmental conduit to the clock comes from contemplation of the circadian clockrelated phenotypes of the large number of kaiA alleles (Table 1). Nearly all missense mutations in kaiA cause either arrhythmic or long-period gene expression rhythms. Gene expression rhythms in a wild-type strain are approximately 25 h (Kondo and Ishiura, 1994). Also, note that the KaiC protein cycles in a wild-type strain (and in vitro) from a non-phosphorylated to a phosphorylated state and then back again (Nakajima et al., 2005; Nishiwaki et al., 2000, 2004). The period of this KaiC ‘‘phosphorylationstate’’ oscillation correlates with the period of clock-regulated, in vivo, gene expression rhythms (Nakajima et al., 2005). In addition, recall that wild-type KaiA protein increases the rate at which KaiC autophosphorylates (Williams et al., 2002). Given the large number and the high percentage of long period phenotypes associated with mutation in kaiA, we will assume that this phenotype results from partial ‘‘loss of function’’ in the KaiA protein. So, partial loss of KaiA function (gene expression is still rhythmic in these mutant strains – extends the period of the gene expression rhythms and thus, presumably, the underlying circadian clock. In other words, these mutant KaiA proteins do not interact as well with KaiC and the rate of KaiC autophosphorylation may therefore decrease. Thus, it should take longer for the KaiC protein to move through its ‘‘phosphorylation-state’’ oscillation and the period of any circadian clock driven oscillation would thereby be extended. From this mechanistic viewpoint, it is fairly simple to imagine how environmental information might travel through KaiA and affect the periodicity of rhythms driven by the clock. Altering the rate of KaiC autophosphorylation via KaiA adjusts the rate at which KaiC moves through its cycle. Most of this argument is pure speculation. A thorough analysis of these mutant KaiA proteins and their affect on KaiC autophosphorylation in vitro is needed to further test this idea and improve the argument. Two other points regarding the kaiA gene are particularly intriguing. First, some KaiA proteins, namely those from the genera Anabaena and Nostoc, are smaller than most and do not have an amino-terminal receiver domain (Fig. 3) (Williams et al., 2002). A search for distinct and separate proteins encoded by the genomes of Anabaena sp. PCC 7120, Anabaena variablis (A. variablis), and Nostoc puntiformes PCC 773102 revealed no proteins with sequence similarity to the amino-terminal region of the S. elongatus PCC 7942, or any other, KaiA protein. It is unclear how these

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Table 1 Mutant alleles of the S. elongatus PCC 7942 kai genes Gene

Allelea

Protein change

Period length of gene expression rhythm (h)b

Reference

kaiA

kaiA5

I9T

29

kaiA6

I16F

27

kaiA7

L31P

27.2

kaiA8

S36P

27.1

kaiA9

C53S

NR

kaiA10

V76G

27.8

kaiA11

V76A

28.4

kaiA1

E103K

33

kaiA12

Q113R

33.3

kaiA13

Q117L

26.2

kaiA14

D119E

30.2

kaiA15

D119G

26.2

kaiA16

V131A

27.9

kaiA17

D136V

30.0

kaiA18

D136Y

28.9

kaiA19

Y166C

NR

kaiA4

F178S

NR

kaiA20

F178I

NR

kaiA21

R180H

26.1

kaiA22

M194T

25.7

kaiA23

F224S

NR

kaiA24

F225S

NR

Nishimura et al. (2002) Nishimura et al. (2002) Nishimura et al. (2002) Nishimura et al. (2002) Nishimura et al. (2002) Nishimura et al. (2002) Nishimura et al. (2002) Ishiura et al. (1998) Nishimura et al. (2002) Nishimura et al. (2002) Nishimura et al. (2002) Nishimura et al. (2002) Nishimura et al. (2002) Nishimura et al. (2002) Nishimura et al. (2002) Taniguchi et al. (2001) Nishimura et al. (2002) Nishimura et al. (2002) Nishimura et al. (2002) Nishimura et al. (2002) Nishimura et al. (2002) Nishimura et al. (2002)

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Table 1 (continued ) Gene

kaiB

kaiC

Allelea

Protein change

Period length of gene expression rhythm (h)b

Reference

kaiA25

E239G

27.5

kaiA26

M241T

38.0

kaiA27

D242V

27.1

kaiA28

D242G

27.3

kaiA29

E243A

33.7

kaiA30

F244V

26.4

kaiA31

A245D

25.7

kaiA32

R249H

30

kaiA2

I266V

27.1

kaiA33

H271A

NR

kaiA34

C273Y

30.2

kaiA35

E274K

NR

kaiA3

1–174

NR

kaiA36

175–284

NR

kaiA37

1–139

24

kaiB1

L11F

21

kaiB2

R74W

22

kaiB3 kaiB4 kaiB5 kaiB6 kaiB7 kaiB8 kaiC15

K5A K10A K42A K57A K66A 94–102 K52H

nd NR NR NR NR NR NR

Nishimura et al. (2002) Nishimura et al. (2002) Nishimura et al. (2002) Nishimura et al. (2002) Nishimura et al (2002) Nishimura et al. (2002) Nishimura et al. (2002) Ishiura et al. (1998) Nishimura et al. (2002) Uzumaki et al. (2004) Nishimura et al. (2002) Ishiura et al. (1998) Uzumaki et al. (2004) Uzumaki et al. (2004) Uzumaki et al. (2004) Ishiura et al. (1998) Ishiura et al. (1998) Iwase et al. (2005) Iwase et al. (2005) Iwase et al. (2005) Iwase et al. (2005) Iwase et al. (2005) Iwase et al. (2005) Nishiwaki et al. (2000)

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Table 1 (continued ) Gene

Allelea

Protein change

Period length of gene expression rhythm (h)b

Reference

kaiC16

G71A

LA

kaiC1

A87V

22

kaiC17

G114A

27

kaiC18

G115R

NR

kaiC2

S157C

29

kaiC3

R215C

16

kaiC4

P236S

28

kaiC5

P248S

NR

kaiC6

P248L

NR

kaiC7

R253H

40

kaiC8

M273I

37

kaiC19

K294H

NR

kaiC9

R321Q

21

kaiC10

T409A

27

kaiC11

G421R

44

kaiC20

V422A

PR

kaiC21 kaiC22

T426A S431A

NR NR

kaiC23

T432A

NR

kaiC12

Y442H

60

kaiC24 kaiC13

R468C G460E

55 NR

Nishiwaki et al. (2000) Ishiura et al. (1998) Nishiwaki et al. (2000) Nishiwaki et al. (2000) Ishiura et al. (1998) Ishiura et al. (1998) Ishiura et al. (1998) Ishiura et al. (1998) Ishiura et al. (1998) Ishiura et al. (1998) Ishiura et al. (1998) Nishiwaki et al. (2000) Ishiura et al. (1998) Ishiura et al. (1998) Ishiura et al. (1998) Kiyohara et al. (2005) Xu et al. (2004) Nishiwaki et al. (2004) Nishiwaki et al. (2004) Ishiura et al. (1998) Xu et al. (2003) Ishiura et al. (1998)

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Table 1 (continued ) Gene

Allelea

Protein change

Period length of gene expression rhythm (h)b

Reference

kaiC14

T495A

NR

kaiC25

E77Q E78Q

NR

kaiC26

E317Q E318Q

NR

Ishiura et al. (1998) Hayashi et al. (2004) Hayashi et al. (2004)

a

Allele numbers are consistent with those given in the relevant reference. Alleles with no published number were assigned one. b NR (not rhythmic); PR (altered phase response); LA (low amplitude rhythm).

organisms might entrain their circadian clocks to prevailing environmental cycles. Curiously, the kaiA gene from Anabaena sp. PCC 7120 can genetically complement a kaiA null allele in S. elongatus PCC 7942 (Uzumaki et al., 2004). The assay for complementation was the demonstration of rhythmic, free-running gene expression patterns but, unfortunately, did not include testing for environmental entrainment (Uzumaki et al., 2004). The second intriguing point concerns the presence and location of the kaiA gene. Three cyanobacteria, T. elongatus BP-1, S. elongatus PCC 7942, and Synechocyctis species strain PCC 6803 have demonstrated circadian rhythms and have had their genomes completely sequenced (Aoki et al., 1995; Kaneko et al., 1996; Ishiura et al., 1998; Nakamura et al., 2002; Onai et al., 2004). In each of these organisms, there is a kai locus organized as described above for S. elongatus PCC 7942. The kaiA gene is located immediately upstream of the other two kai genes. However, this is not the end of the story. T. elongatus has an additional kaiB gene and Synechocyctis sp. strain PCC 6803 has two additional kaiB and two additional kaiC genes (Figs. 3–5). The functions of these additional genes are unknown. Several other cyanobacterial sp. also have extra kai genes in addition to the presumed clock-related kaiABC locus. However, no species has yet been identiﬁed that has more than one kaiA gene. In fact, the genus Prochlorococcus evidently does not have any kaiA genes. However, a kaiBC locus is present in the four Prochlorococcus genomes that have been completely sequenced. Unfortunately, the presence of a functional circadian clock in any Prochlorococcus sp. has not been conﬁrmed. Thus it is not yet clear whether kaiA function is the determinant for a cyanobacterium to have a circadian clock and thus

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generate circadian rhythms. These sequence data combine to make KaiA a very intriguing protein. When present, it interacts directly with KaiC and is absolutely required for circadian rhythmicity. Yet, it comes in at least two different functional modes, which differ significantly in size and functional character, and in some organisms it is not present at all. Presumably, a more complete and comparative understanding of the variety of potential circadian clock mechanisms among the cyanobacteria will eventually explain some of these KaiA-associated quandaries.

3.2. The KaiB Protein The S. elongatus PCC 7942 kaiB gene is 309 bp in length and encodes 102 amino acyl residues that ultimately form an 11.4 kDa protein. This KaiB protein likely folds into the alpha–beta meander motif as do similar proteins from other cyanobacteria (Fig. 4). Those similar proteins include the KaiB protein from Anabaena sp. strain PCC 7120 that crystallizes as a dimer, the T. elongatus BP-1 KaiB protein that crystallizes as a dimer of dimers and the Synechocystis sp. strain PCC 6803 KaiB protein that crystallizes as a tetramer (Garces et al., 2004; Iwase et al., 2004; Hitomi et al., 2005; Iwase et al., 2005). (Information and images regarding the structure of crystalline KaiB proteins are available at http://www.rcsb.org/pdb/Welcome.do using codes 1VGL, 1R5P, and 1WWJ.) The intracellular concentration of the KaiB protein oscillates with a circadian rhythm. Its concentration peaks near subjective dusk at about 25,000 molecules per cell and has a nadir just after subjective dawn of about 10,000 molecules per cell (Kitayama et al., 2003). KaiB has no recognizable sequence motifs based upon primary protein sequence comparisons to known proteins (Ishiura et al., 1998; Garces et al., 2004). Curiously, the KaiB protein has no discernible effect on the in vitro KaiC autophosphorylation rate despite the fact that the two proteins directly interact (Iwasaki et al., 1999; Williams et al., 2002; Kitayama et al., 2003). However, it does decrease the autophosphorylation rate of KaiC when the KaiA protein is present (Iwasaki et al., 2002; Williams et al., 2002). In effect, KaiB modulates the interaction between the KaiA and KaiC proteins. Analogous to the compartmentalization of circadian clock proteins in many eukaryotic organisms, KaiB protein is evidently membrane-sequestered during the ﬁrst half (subjective day) of the S. elongatus PCC 7942 circadian cycle (Kageyama et al., 2003). Its controlled release from the membrane during the subjective night is not yet understood but its subsequent interactions with KaiC are an essential part of circadian timing (Ditty et al., 2003; Kageyama et al., 2003; Kitayama et al., 2003; Johnson, 2004a).

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Figure 4 A ClustalW (http://www.ebi.ac.uk/clustalw/) derived alignment of the amino acyl residue sequence from 27 putative KaiB proteins. Protein sequence was obtained from public sequence databases using a BLAST algorithm. Structure and function of the KaiB protein is discussed in the text. Protein sequences are from: (1) A. variablis, (2) Anabaena sp. PCC7120, (3) N. punctiformes PCC773102, (4) T. erythraeum IMS101, (5) Crocosphaera watsonii WH8501, (6) T. elongatus BP-1, (7) A. variablis, (8) Anabaena sp. PCC 7120, (9) Nostoc punctiformes PCC 773102, (10) Trichodesmium erythraeum IMS101, (11) Crocosphaera watsonii WH8501, (12) Synechocystis sp. PCC 6803, (13) Synechococcus sp. PCC 7002, (14) T. elongatus, (15) P. marinus MED4, (16) Prochlorococcus sp. NATL2A, (17) P. marinus SS120, (18) Synechococcus sp. WH8102, (19) Synechococcus sp. PCC 9605, (20) Synechococcus sp. PCC 9902, (21) P. marinus MIT9313, (22) S. elongatus PCC 7942, (23) S. elongatus PCC 6301, (24) Crocosphaera watsonii WH8501, (25) Synechocystis sp. PCC 6803, (26) Synechocystis sp. PCC 6803, and (27) Gloeobacter violaceus PCC 7421. Several species have multiple KaiB-like proteins. Asterisks indicate potentially interesting residues conserved among the 6 putative KaiB-long proteins. See text for discussion. Numbers on the left indicate the protein source. Numbers on the right indicate protein residue.

There is some speculation that KaiB transits from a dimer to a tetramer upon moving from the membrane to the cytoplasm (Hitomi et al., 2005). Primarily, that speculation comes from the fact that the exposed surface of the Synechocystis sp. strain PCC 6803 KaiB dimer is more hydrophobic than that of the tetramer. There are no functional or biochemical data to support

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such an argument (Hitomi et al., 2005). As will be discussed below, the KaiC, KaiA, and SasA proteins form a high order complex during the circadian cycle and this is very likely an important part of the timing process (Kageyama et al., 2003). The transition of KaiB from the membrane to this complex during the subjective night may signal of the deterioration of the complex (Kageyama et al., 2003). This deterioration would then allow the re-formation of new protein complexes as the system moves through a subsequent circadian cycle. Evidence for the membrane sequestration of the KaiB protein during the subjective day suggests the potential for coupling cellular chemiosmotic actions, and thus multiple metabolic parameters, to the circadian clock timing mechanism. For instance, the regulation of protein function through changes in the redox state of thiol groups is well known (Buchanan and Balmer, 2005; Buchanan and Luan, 2005). In fact, this type of regulation was initially discovered in the context of redox proteins that monitor photosynthetic activity (Wolosiuk and Buchanan, 1977). One idea is that KaiB is somehow sensitive to redox conditions and, as they change early in the subjective night (after sundown), KaiB responds by leaving the membrane and interacting with the Kai protein timing-complex. Unfortunately, there are no thiol groups or other obvious redox sensitive, amino-acyl residues or other ligands associated with the typical, S. elongatus PCC 7942-like, KaiB protein (Fig. 4). Thus, we need to invoke a more indirect mechanism for regulating KaiB membrane sequestration. Perhaps, the ﬁrst six KaiB-like sequences shown in Fig. 4 provide a smidgen of credence and even some possibility to this notion of KaiB redox control. Compare sequences 1–6 to the other 21 listed in Fig. 4. These ﬁrst six KaiB-like sequences are from the genera Anabaena, Nostoc, Trichodesmium, Crocosphaera, and Thermosynechococcus. (Other than these six proteins, there are no obvious or documented relationships among these genera to set them apart, as a group, from the other cyanobacteria.) Each organism has a KaiB-like sequence that is over twice the length of the ‘‘typical’’ S. elongatus PCC 7942 KaiB sequence and this additional sequence would form an amino-terminal region not present in the other, shorter KaiB proteins. Also note that the particular organisms that have these KaiB-long proteins also have the shorter, more typical KaiB protein. The interesting feature that these six proteins share, other than their relatedness to one another and to KaiB, is that within their amino-terminal additions they contain conserved redox sensitive residues. All six have three conserved cysteinyl residues and there are a number of other cysteinyl and histidyl residues that are conserved among four or ﬁve of the six proteins (Fig. 4). As notable are the four, evenly spaced, tryptophanyl residues

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conserved within a 100 residue stretch in all six proteins. (By comparison, the 519 amino acyl residue KaiC protein has just three of the relatively rare tryptophanyl residues.) The functional meaning of these residues is not yet clear. Nevertheless, an obvious speculation derived from the presence and conservation of the potentially redox-sensitive or ligand-binding, cysteinyl and histidyl residues is that they function as redox sensors. No data exist regarding function for any of these six proteins so speculation is easy. So, what are these proteins trying to tell us? Perhaps they sequester the typical, shorter KaiB proteins to the membrane by forming hetero-oligomeric protein complexes. They could then release KaiB-short as a function of redox potential or metabolic state. For example, the ratio of NADP(H) to NAD(H) in S. elongatus PCC 7942 controls the reversible dissociation of a protein complex that consists of phosphoribulokinase, the protein CP12, and glyceraldehyde-3-phosphate dehydrogenase (Tamoi et al., 2005). That particular complex is involved in the Calvin cycle but those data provide precedence for protein complex dissociation as a function of metabolic state. It is also noteworthy that the KaiB protein has a three-dimensional fold that is reminiscent of the redox-sensitive thioredoxin protein (Vakonakis et al., 2004a). Clearly, many speculative possibilities exist regarding KaiB protein function. Of particular interest will be the relationship between that function and cellular metabolism. The primary sequence of the amino-terminal region of the longer KaiB proteins (1–6 in Fig. 4) may also help to identify proteins that potentially membrane-sequester KaiB in those strains that do not have the KaiB-long protein. However, searches for proteins in S. elongatus PCC 7942 with sequence similar to that of the KaiB-long extended amino-terminal region have not yielded positive results. Although a number of kaiB alleles have been characterized, no clear phenotypic patterns exist (Table 1). Once interactions among the Kai and Sas proteins have been further characterized the phenotypes caused by mutation in KaiB will be more easily interpreted.

3.3. The KaiC Protein The S. elongatus PCC 7942 kaiC gene is 1560 bp in length and encodes 519 amino acyl residues that ultimately form an approximately 58 kDa protein. (Information and images regarding the structure of crystalline KaiC protein are available at http://www.rcsb.org/pdb/Welcome.do using code 1TF7 or 1U9I.) The intracellular concentration of the KaiC protein oscillates with a circadian rhythmicity. It peaks just after subjective dusk at about 15000 molecules per cell and the nadir is near 5000 molecules per cell just after

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subjective dawn (Kitayama et al., 2003). Overall the monomeric S. elongatus PCC 7942 KaiC protein is dumb-bell shaped with two distinct, but very similar, domains, called CI and CII, separated by a short linker region (Pattanayek et al., 2004). Each domain is a mostly parallel-stranded, twisted b sheet consisting of seven strands surrounded by eight a helices (Pattanayek et al., 2004). There is significant similarity between the ﬁrst and second halves of the KaiC protein sequence suggesting that KaiC protein is comprised of tandemly duplicated domains (Iwasaki et al., 1999). In fact, the CI (amino-terminal) domain and the CII (carboxyl-terminal) domain are structural analogs (Pattanayek et al., 2004). The cyanobacterial kaiC gene most likely arose via a tandem gene duplication event and unduplicated ancestral type genes remain in many Archaea and Proteobacteria (Leipe et al., 2000; Dvornyk et al., 2003). Both its fold and its primary sequence place the KaiC protein in the E. coli RecA/DnaB family of proteins (Story et al., 1992; Story and Steitz, 1992; Leipe et al., 2000; Pattanayek et al., 2004). This family consists of proteins that bind and hydrolyze ATP in order to do work. That work typically includes DNA manipulation. The RecA protein is a recombinase whereas DnaB is a prototypical bacterial helicase (Leipe et al., 2000). In the presence of ATP the S. elongatus PCC 7942 KaiC protein and the T. elongatus BP-1 KaiC protein are each known to form a hexameric complex (Mori et al., 2002). These hexamers look like two donuts stacked on top of one another (Mori et al., 2002; Hayashi et al., 2003). The KaiC monomers are oriented such that the CI domains form one end of the double donut-like structure and the CII domains form the other (Pattanayek et al., 2004). The hexameric complex has a tapered, cone-like central channel. A dozen arginyl and histidyl residues line the channel’s inside, which is also capped at the CII end by arginyl residues. This suggests that, like a helicase, the hexamer could form around or somehow associate with, negatively-charged DNA molecules (Pattanayek et al., 2004). In fact, the ATP-dependant KaiC hexamer does interact with a small forked-DNA substrate (Nishiwaki et al., 2000; Mori et al., 2002; Hayashi et al., 2003; Pattanayek et al., 2004). This type of DNA substrate is standard in many helicase activity assays (Mori et al., 2002). Despite all of these obvious similarities to a helicase, KaiC protein does not appear to unwind the double-stranded, forked-DNA substrate (Weng et al., 1996; Weng et al., 1998; Mori et al., 2002). Primary motif searches revealed that KaiC contains two sets of putative P-loop ATP binding motifs (Walker box A motif), two imperfect Walker box B motifs, and two putative catalytic carboxylate glutamyl residues (E78 and E318 in CI and CII, respectively) that are present in many other ATPbinding proteins. Those two glutamyl residues and the surrounding Walker

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box motifs are conserved in all 20 of the KaiC protein sequences listed in Fig. 5. In addition, the KaiC protein carries two putative DXXG motifs, which are typically conserved in GTPase proteins (Leipe et al., 2000). Biochemical characterization revealed that KaiC binds ATP in vitro and, with much lower afﬁnity, GTP (Nishiwaki et al., 2000; Mori and Johnson, 2001a). Site-speciﬁc mutation in the Walker box A motif within the CI domain of KaiC (K52H) resulted in a recombinant protein with markedly reduced ATP-binding activity in vitro. The reciprocal mutation in the CII domain (K294H) had no effect on nucleotide binding (Nishiwaki et al., 2000). The molar ratio of ATP per KaiC protein was calculated at 22.6 +/ 0.8 pmol of ATP per 12.5 pmol of KaiC protein, suggesting that each KaiC protein binds two ATP molecules (Hayashi et al., 2003). In vivo, the K52H change resulted in arrhythmic patterns of gene expression, whereas the K294H change lengthened the period of those rhythms (70 h) and reduced their amplitude (Nishiwaki et al., 2000) (Table 1). Both of these two lysinyl residues are conserved among the KaiC protein sequences listed in Fig. 5. As discussed above, the S. elongatus PCC 7942 KaiC protein also has an autokinase activity and autophosphorylates from bound ATP molecules. At least three CII domain amino acyl residues (T426, S431, and T432) can accept a phosphoryl group from ATP and all of these residues are also conserved among KaiC proteins (Iwasaki et al., 2002; Hayashi et al., 2004; Xu et al., 2004). This kinase activity and each of the three residues are essential for circadian clock function in S. elongatus PCC 7942. As mentioned, the CI domain, which forms the wider end of the central channel, also binds ATP but does not appear to autophosphorylate. Evidently, the

Figure 5 A ClustalW (http://www.ebi.ac.uk/clustalw/) derived alignment of the amino acyl residue sequence from 20 putative KaiC proteins. Protein sequence was obtained from public sequence databases using a BLAST algorithm. Structure and function of the KaiC protein is discussed in the text. Speciﬁc residues discussed in the text are highlighted in gray. Protein sequences are from: (1) Crocosphaera watsonii WH8501, (2) Synechocystis sp. PCC 6803, (3) A. variablis, (4) Anabaena sp. PCC 7120, (5) Nostoc punctiformes PCC 773102, (6) Trichodesmium erythraeum IMS101, (7) Crocosphaera watsonii WH8501, (8) Synechocystis sp. PCC 6803, (9) Synechococcus sp. PCC 7002, (10) T. elongatus BP-1, (11) Synechococcus elongatus PCC 6301, (12) Synechococcus elongatus PCC 7942, (13) P. marinus MED4, (14) P. marinus SS120, (15) Prochlorococcus sp. NATL2A, (16) Synechococcus sp. PCC 9605, (17) Synechococcus sp. WH8102, 18) Synechococcus sp. PCC 9902, (19) P. marinus MIT9313 and (20) Synechocystis sp. PCC 6803. Several species have multiple KaiClike proteins. See text for discussion. Numbers on the left indicate the protein source. Numbers on the right indicate protein residue.

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resultant energies from ATP binding by the CI domain are primarily used to regulate hexamer formation (Hayashi et al., 2004). Both the KaiA and KaiB proteins interact with KaiC and alter the rate at which it autophosphorylates (Williams et al., 2002; Kitayama et al., 2003) KaiA stimulates that rate and KaiB negatively modulates the stimulation (Williams et al., 2002; Kitayama et al., 2003). Each S. elongatus PCC 7942 KaiC protein has two putative KaiA-binding sites but detailed functional data regarding selective or cooperative interactions remains uncollected (Taniguchi et al., 2001). Both binding regions were identiﬁed using a yeast two-hybrid assay. The CI domain KaiA-binding region is located within residues 206–263 of KaiC and the CII domain KaiA-binding region is located within residues 418–519 (Taniguchi et al., 2001). Again, these are wellconserved regions of KaiC protein sequence (Fig. 5). However, the two different regions do not share obvious sequence similarity with one another (Vakonakis et al., 2004b). Interestingly, only the CII KaiA-binding region (residues 473–518) derived from the T. elongatus BP-1 KaiC protein affects the NMR spectra of the KaiA180C peptide (Vakonakis et al., 2004b). This latter peptide is a truncated version of the T. elongatus KaiA protein comprised of residues 180–283. Thus, it now appears that KaiA may stimulate KaiC autophosphorylation by direct interaction with the KaiC-CII kinase domain. It should be stated that the putative KaiC-CI KaiA-binding region is not probably accessible to KaiA when KaiC is in its hexameric state whereas the CII region is highly exposed at one end of the complex (Vakonakis et al., 2004b). Despite their described interactions, no KaiB binding regions on the KaiC protein have been identiﬁed (Iwasaki et al., 1999).

3.4. The SasA Protein The S. elongatus PCC 7942 sasA gene is 1164 bp in length and encodes 387 amino acyl residues that ultimately form an approximately 43 kDa protein. (Information and images regarding the solution structure of the aminoterminal domain of the SasA protein are available at http://www.rcsb.org/ pdb/Welcome.do using codes 1T4Y and 1T4Z.) The concentration of the SasA (Synechococcus adaptive sensor) protein does not oscillate over the circadian cycle; however, SasA co-immunoprecipitates with the aforementioned higher order Kai-protein complex. Peak interaction with KaiC appears between circadian time 16 (CT ¼ 16) and CT ¼ 22 of the circadian cycle (Kageyama et al., 2003). Hence SasA, although not essential for all aspects of circadian rhythmicity, is in close physical association with the

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Kai-protein oscillator complex. The SasA protein seems to play a key role in transducing circadian-time output. It likely serves as a conduit by which the Kai-protein complex converts temporal information into downstream rhythmic gene expression patterns (Iwasaki et al., 2000; Smith and Williams, 2006). The sasA gene was originally identiﬁed in 1993 not for its circadian functions, but from a genetic screen seeking cyanobacterial sensor kinases (Nagaya et al., 1993). In high copy number the sasA gene complements mutant E. coli strains that lack the histidine protein kinase sensors of prototypical two-component regulatory systems (Nagaya et al., 1993; Stock et al., 1995, 2000). Two independent lines of research later identiﬁed the SasA protein as integral to the generation and maintenance of S. elongatus PCC 7942 circadian rhythms in gene expression (Iwasaki et al., 2000). Protein sequence comparisons revealed that the amino terminal domain of SasA has strong resemblance to the full-length KaiB protein. These protein sequences share 36% identity and 58% similarity across 77 residues (Iwasaki et al., 2000; Vakonakis et al., 2004a). The carboxy terminal sequence of SasA is very similar to that of histidine protein kinases and it undergoes autophosphorylation in vitro (Nagaya et al., 1993). All of the conserved N-, D/F-, and G-boxes associated with histidine protein kinases are found in the SasA protein sequence (Stock et al., 1995, 2000). The rate at which SasA autophosphorylates is increased by the presence of the KaiC protein (Smith and Williams, 2006). Disruption of the sasA gene in S. elongatus PCC 7942 dramatically reduces the amplitude of the expression rhythm from the kaiBC promoter but does not completely abrogate circadian rhythmicity. However, the residual kaiBC expression rhythm does have a slightly shorter free-running period length. Expression from all other – non-kai – tested promoters is arrhythmic in a sasA null background (Iwasaki et al., 2000). These other promoters do vary in terms of their expression levels. Some are constitutively high, some low, and some at midline levels of relative (compared to wild type) expression (Iwasaki et al., 2000). To determine the role of SasA autophosphorylation in circadian timing, the conserved catalytic histidyl residue was substituted with a glutaminyl residue, H162Q. The resultant phenotype was as the sasA null (Iwasaki et al., 2000). SasA protein was independently identiﬁed as interacting physically with the KaiC protein in a yeast two-hybrid screen (Iwasaki et al., 2000). The ﬁrst 97 residues of SasA (the domain with similarity to KaiB) were pulled out as ‘‘prey’’ using the KaiC protein as ‘‘bait’’. Subsequent in vitro assays conﬁrmed that either full-length SasA or just SasA residues 1–97 alone, would interact with KaiC. Curiously, either the CI or CII half of KaiC protein is sufﬁcient for these interactions. The stoichiometry of KaiC-SasA protein interaction has not been determined.

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A cognate response regulator that would receive a phosphoryl signal from SasA has not yet been identiﬁed, so the pathway for SasA communication with downstream genes remains elusive (Stock et al., 2000). The circadian clock in the sasA null background entrains normally to 12-h dark and temperature pulses, but cells are impaired when grown in a light/dark cycle (Stock et al., 2000). One possible interpretation of this data is that SasA acts on an input pathway, affecting interpretation of light and dark signals. However, disruption of sasA globally affects downstream genes, and the light/dark defect could arise indirectly from altered expression of any one of them. Because the sasA light/dark defect is not shared with any of the kai null strains, it may result from a function of SasA that is unrelated to its role in the clock. The strongest evidence for SasA as an output pathway component is that, whereas association with KaiC stimulates SasA autophosphorylation, SasA has no effect on KaiC autophosphorylation (Smith and Williams, 2006). Thus, information ﬂow seems to be from the Kai complex to SasA, rather than the other way around. Continuous overproduction of SasA in S. elongatus PCC 7942 completely represses expression from the kaiBC promoter; however, transient pulses of SasA affect kaiBC promoter expression differently as a function of the point in the circadian cycle at which SasA is elevated. When expressed during subjective day, SasA production causes a phase delay in kaiBC expression, and during subjective night a phase advance, consistent with a role in close association with, but that is not an integral part of, the Kai-protein circadian oscillator (Iwasaki et al., 2000). NMR-based structure analyses of the ﬁrst 105 amino acyl residues of the S. elongatus PCC 7942 SasA protein indicates a secondary structure of bababba, which then folds into a thioredoxin-like tertiary structure (Vakonakis et al., 2004a). Recall that this is the part of the SasA protein necessary and sufﬁcient for KaiC interaction and that its sequence resembles that of KaiB (Fig. 6). Also, note that the rate at which SasA autophosphorylates is increased by KaiC interaction (Smith and Williams, 2006). The effect of KaiC on the SasA autophosphorylation rate is diminished by the presence of the KaiB protein (Smith and Williams, 2006). These latter points have been used as a basis from which to speculate that SasA and KaiB might compete for binding space on the KaiC protein. An alternative idea is that one of these proteins smay interact with the KaiC–CI domain and that the other would then interact with the KaiC–CII domain. In either case, interactions between KaiB and KaiC could affect SasA autophosphorylation and thus its subsequent functions as a two-component, signal-transducing, sensor kinase. Interestingly, when this N-SasA structure was compared to the Anabaena sp. PCC 7120 KaiB protein structure, no overt structural

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Figure 6 (A) Structure and (B) protein architecture of N-SasA from S. elongatus PCC 7942 and KaiB from Anabaena sp. strain PCC 7120. See text for discussion of comparative function. Figure modiﬁed from Vakonakis et al. (2004). (See Colour Plate Section in back of this volume.)

similarities that might suggest competition for the same KaiC binding site were apparent (Vakonakis et al., 2004a). Obviously, a more direct biochemical approach is needed to determine the relationships among these three proteins. Simple kinetics experiments should tell us if KaiB is a competitive or non-competitive inhibitor of the KaiC–SasA interaction.

3.5. The Kai-Clock Protein Complex The three Kai proteins form timing complexes that change in composition, size, phosphorylation state, and quantity as a function of CT (Kageyama et al., 2003; Hayashi et al., 2004). Recall that in vitro these three proteins and ATP will produce a circadian rhythm in the KaiC phosphorylation state (Nakajima et al., 2005). Organization of these timing complexes within S. elongatus PCC 7942 is dependent upon the KaiC protein (Kageyama et al., 2003). During the subjective day (the ﬁrst half of a circadian cycle), KaiC protein forms a 360 kDa homo-hexameric structure and then supports creation of an even larger (550 kDa) heterogeneous protein complex during the subjective night (the latter half of a cycle) (Mori et al., 2002; Kageyama et al., 2003). Throughout the circadian cycle KaiA and KaiB proteins are present as homo-multimeric species but also become part of the larger, heterogeneous, KaiC-dependent subjective-night, protein complex (Kageyama et al., 2003). As mentioned above, KaiA is probably dimeric and KaiB may alternate between dimeric and tetrameric states (Garces et al., 2004; Iwase et al., 2004; Ye et al., 2004; Hitomi et al., 2005; Iwase et al., 2005). The number of KaiC and KaiB molecules oscillates over a circadian cycle whereas the number of KaiA molecules per cell remains relatively constant

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over time (Kitayama et al., 2003). In vivo, the ratio of phosphorylated to non-phosphorylated KaiC proteins also increases throughout the subjective day and early subjective night (Nishiwaki et al., 2000). This KaiC ‘‘phosphorylation status’’ ratio is the one that maintains a rhythmic pattern when puriﬁed Kai proteins are mixed in vitro (Nakajima et al., 2005). Thus a general description of circadian timing can be made based upon the known functions of the Kai proteins. Transcription of the kai genes and subsequent kai-message translation starts a circadian cycle. Early in the subjective day, KaiC binds ATP, forms a hexamer and begins to autophosphorylate. Later in the subjective day, the autophosphorylation rate is modulated by increased interaction with KaiA and possibly the KaiB protein. The amount of both phosphorylated and non-phosphorylated KaiC protein increases over time and eventually, in the late subjective night, nearly all KaiC protein is phosphorylated. KaiB protein also accumulates during the subjective day and is primarily membrane-associated. During the subjective night, KaiB is found as a soluble protein that has transited into the KaiC-based timing complex. Some aspect of this transition may begin Kai-complex degradation. The degradation of these complexes and of the individual Kai proteins signiﬁes the end of a circadian cycle. Because KaiC protein negatively regulates expression of both the kaiB and kaiC genes, its degradation allows de-repression of kai gene expression and the cycle begins anew (Ishiura et al., 1998; Ditty et al., 2003). SasA protein co-immunoprecipitates with the aforementioned, subjective evening 550-kDa Kai-protein complexes. Peak interaction between SasA and KaiC appears during the interval from CT ¼ 16 to CT ¼ 22 (Kageyama et al., 2003). Like most bacterial sensory kinases, SasA autophosphorylates using ATP as a phosphoryl group donor (Smith and Williams, 2006). The presence of KaiC increases the rate at which SasA autophosphorylates by about 20-fold (Smith and Williams, 2006). The other Kai proteins modulate the maximum autophosphorylation rate of SasA but only if KaiC protein is present. It is not yet known whether the KaiA and KaiB proteins act as intercalary proteins and directly rework KaiC and SasA interaction, or act by altering the phosphorylation state of KaiC that in turn reworks the interaction (Williams et al., 2002). Obviously and most importantly, there is a clear KaiC-dependent modulation of the autophosphorylation rate of SasA. So, it appears that SasA may mediate circadian-timed gene expression by having the Kai-protein complex, via KaiC, regulate its rate of autophosphorylation. As stated, the 43 kDa SasA protein partitions more heavily into the Kaiprotein timing-complex during the subjective night. It has moved there from

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a smaller, non-Kai, subjective-day complex with a mass of about 140 kDa (Kageyama et al., 2003). In a kaiC genetic background, the SasA protein remains within that smaller subjective-day complex. Sensory kinase proteins like SasA have been found almost exclusively as dimers and SasA demonstrates homotypic interaction (Iwasaki et al., 2000). The size of the SasAprotein daytime complex seems to then suggest interaction with uncharacterized components – most likely a Sas response regulator (Iwasaki et al., 2000; Williams et al., 2002; Wolanin et al., 2002). Sensory kinases have cognate response regulators that typically function as DNA-binding, transcription activating proteins (Stock et al., 2000). Thus, transcription timing information evidently ﬂows from the Kai-protein timing complex via direct interactions between KaiC and SasA. This information regulates SasA autophosphorylation and, we propose, subsequent transfer of its phosphoryl group to an awaiting response regulator. This phosphorylated response regulator protein would then control gene transcription as a function of time – a function of CT, as monitored by the physical state of the Kai-protein clock complex. Curiously, KaiC stimulates SasA autophosphorylation but SasA is present in the Kai-protein complex primarily during the subjective night when gene expression levels are, in general, decreasing. One might expect gene expression levels to be high during the time that SasA is autophosphorylating at its highest rate and then, presumably, acting to stimulate gene expression via that cognate response regulator. More data regarding the KaiC/SasA interactions and identiﬁcation and characterization of the putative ‘‘SasR’’ cognate response regulator should help clarify this current quandary. Perhaps SasR is a repressor of gene expression or phosphorylated KaiC protein is not the best stimulator of SasA autophosphorylation. Also, SasA protein does not alter the phosphorylation rate of KaiC (Smith and Williams, 2006). Thus, information ﬂows from Kai to Sas but not vice versa. Another interesting aspect of the interaction between SasA protein and the Kai-protein complex may be most evident in a kaiB null genetic background. The distribution of SasA protein is shifted dramatically away from its small subjective-day complex and toward the large KaiCbased, subjective-night complex suggesting increased association with KaiC in the absence of KaiB (Kageyama et al., 2003). Recall that the aminoterminal region of SasA, which interacts with KaiC, has clear sequence and structural similarity to the entire KaiB protein, which also interacts directly with KaiC (Iwasaki et al., 2000; Garces et al., 2004; Vakonakis et al., 2004a). Again, kinetic examination of the KaiC–SasA–KaiB functional relationships should prove enlightening.

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4. CLOCK-CONTROLLED GENE EXPRESSION 4.1. No Solitary Output Pathway Although we can now describe several biochemical processes that are essential for circadian rhythm generation and maintenance in S. elongatus PCC 7942, the question of how those processes are connected to circadian clock regulation of global gene expression still looms. The circadian timing mechanism continues to function in a sasA genetic background because daily rhythms in expression levels continue from a F(kaiB-luc+) reporter. The amplitude and period length are altered but the circadian clock is still running and robust (Iwasaki et al., 2000; Smith and Williams, 2006). However, nearly 20 different and unrelated promoter-luc+ (or promoter-lux) gene fusions, excepting those using a kai promoter, are not rhythmically expressed in a sasA null strain (Iwasaki et al., 2000). Thus, even though the circadian timing mechanism is functioning, temporal information is not getting from the clock – the Kai-protein complex – to the regulatory systems controlling clock-regulated gene expression. So, it is clear that SasA protein functions to mediate circadian-timed gene expression. Yet, careful consideration of all the existent data makes it difﬁcult to conclude that Kai and Sas protein functions and interactions are sufﬁcient for circadian clock regulation of global gene expression in S. elongatus PCC 7942. For example, the KaiC protein also interacts with DNA and its over-production reduces gene expression levels on a global scale (Mori et al., 2002; Nakahira et al., 2004). Also, clock regulated gene expression patterns include at least two major temporal classes (Liu et al., 1995b; Min and Golden, 2000; Min et al., 2004). The major class includes kaiB and is illustrated in Fig. 1. The minor class has a phase angle that is shifted, under constant illumination and relative to the major class, by 1801 (so-called opposite phase expression) (Liu et al., 1996; Min and Golden, 2000; Min et al., 2004). A lack of identiﬁable cis- or transacting elements that would explicitly determine phase angle also provokes speculation that SasA is not the only carrier of temporal output. Something else is going on. In fact, it has been suggested by several research groups that chromosome arrangement or DNA topology may be the phase determinant for gene expression rather than simple cis-acting regulatory sequences (Min and Golden, 2000; Min et al., 2004; Nakahira et al., 2004; Johnson, 2004b). In addition, there are data showing that heterologous promoters from noncircadian clock containing bacteria drive rhythmic gene expression in S. elongatus PCC 7942 (Ditty et al., 2003).

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4.2. Chromosome Compaction Recent analysis of chromosome compaction has exposed the process as a potential mechanism for control of global gene expression by the cyanobacterial circadian clock (Smith and Williams, 2006). The chromatin in S. elongatus PCC 7942 was visualized by ﬂuorescence microscopy and was observed compacting, decompacting, and compacting again, over the course of a circadian cycle (Smith and Williams, 2006). This chromosome-compaction rhythm continues under free-running (constant light) conditions and is dependant upon the Kai-protein circadian clock (Fig. 7). Both kaiA and kaiC strains are unable to rhythmically compact their chromosomes and neither of these strains demonstrates rhythmic patterns of gene expression (Smith and Williams, 2006). An S. elongatus PCC 7942 strain that generates gene expression rhythms with a 14-h period was also shown to have a chromosome compaction rhythm with that same periodicity (Smith and Williams, 2006). Interestingly, in a sasA genetic background the chromosome-compaction rhythm continues but, as discussed above, gene expression patterns are not rhythmic in this background. Thus, it seems that both rhythmic chromosome compaction and properly timed regulatory protein function are required for the generation and maintenance of circadian rhythms in gene expression. The idea is that diffuse, transcription-competent chromatin is present during the subjective day and that compacted, transcription-resistant chromatin is present during the subjective night. If the proper regulatory proteins are available during the transcription-competent period then wild-type gene expression rhythms are generated. Of course, the experimental test for this model will come when an otherwise wild-type strain loses the ability to compact its chromosome. Will a functional circadian clock generate gene expression rhythms in the absence of chromosome compaction? Several other features of the chromosome-compaction data are interesting. Note in Fig. 7 that during the light/dark cycle the chromosome was decompacted at times 0 and 24 and recognize that these cells were sampled in the darkness immediately before the lights came on. Also, the cells sampled at time 12 were taken before the lights went off and those cells had compacted chromosomes. S. elongatus PCC 7942 can evidently anticipate transitions from light to dark and vice versa. Other than some obvious conjecture regarding the regularity of daily oscillations in light quality and quantity, in temperature variation, and in humidity levels, the selective pressures that have guided the evolutionary progression of a timing mechanism in cyanobacteria remain enigmatic (Roenneberg and Merrow, 2002).

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For obligate photosynthetic organisms like S. elongatus PCC 7942 speculation about these pressures has included suggestion of a clock-based anticipatory timing strategy that allows the organism to ready itself for renewed inter-species competitions over energy acquisition and utilization just ahead of imminent sunrise. These compaction data demonstrate and support the idea of an anticipatory character underlying circadian clock function (Smith and Williams, 2006). In addition, some degree of chromatin compaction was consistently noted about midway through the subjective day. There may also be an expeditious compaction cycle that takes place during the cell division cycle. The clock runs independently of cell division but does gate (permit) cell division by allowing it only at speciﬁc circadian times – around the time we generally see this compaction (Mori and Johnson, 2001b). Although rhythmic chromosome compaction requires the KaiC protein, it is not directly responsible for all of the observed compaction. Some level of chromosome compaction is observed even in a kaiC genetic background (Smith and Williams, 2006). Presumably, other proteins are more (or equally) involved in the physical process of compacting the DNA. The fundamental nature of the Kai-protein circadian clock may be this rhythmic ‘‘breathing’’ of the entire S. elongatus PCC 7942 chromosome – compact, de-compact, compact, de-compact. Although the chromosome is absolutely accessible to transcription during part of the circadian cycle, if the proper regulatory elements, as controlled by clock-output proteins such as SasA and the relevant sigma factors, are not available then important temporal information is lost and gene expression patterns are arrhythmic. While chromosome compaction is expected to somehow sequester most promoter elements – because compaction is greatest during the subjective

Figure 7 Chromosome compaction in S. elongatus PCC 7942 as demonstrated with deconvolved ﬂuorescence microscopy images (red-autoﬂuorescence from S. elongatus PCC 7942; green -DAPI stained DNA). Cultures of wild-type and kaiC strains were sampled at the indicated times during a light/dark cycle (A and C) or a free-running cycle (B and D). The chromosome arrangement image shown for each time point is representative of 499% (n ¼ 300) of the cells from a sample. For the wild-type strain note that in each of the cycles (A and B) there is a slow arrangement of the DNA into distinct ‘‘nucleoids’’ and then a return to the time 0 diffuse state. For the kaiC strain note that at any given time point within each experiment some variation in the extent of chromosome compaction was evident. This variation is also reﬂected in the gene expression rhythms from that strain (Smith and Williams, 2006). However, no rhythmic compaction patterns are observed in this genetic background (see text for discussion). Cells are approximately 5 mm long. (See Colour Plate Section in back of this volume.)

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night while gene expression levels are low – it could also newly expose other promoter elements as a result of the presumed dramatic changes in chromosome topology. This speculation suggests a mechanism for the minor class of genes with increased expression during the subjective night when the chromosome is compacted (Min and Golden, 2000; Min et al., 2004). The challenge of relating these large-scale chromosome dynamics to individual promoter function should be an interesting one.

5. CLOCK INPUT 5.1. The CikA Protein The S. elongatus PCC 7942 cikA gene is 2265 bp in length and encodes 754 amino acyl residues that form an approximately 84 kDa protein. (an NMRderived solution structure of the carboxyl terminal, 133-residue pseudo-receiver domain of the S. elongatus PCC 7942 CikA protein will soon be published. Chemical shift assignments for the various nuclei have been made (Gao et al., 2005).) The CikA protein is a key player in the molecular mechanism for coupling diurnal environmental signals to the circadian clock, i.e., it is part of the input pathway (Schmitz et al., 2000; Mutsuda et al., 2003; Zhang et al., 2006). The cikA (circadian input kinase) gene was ﬁrst discovered in a genetic screen for mutants of S. elongatus PCC 7942 that had defects in light-responsive regulation of a photosystem II gene, psbAII (Schmitz et al., 2000). Analysis of temporal gene expression patterns in a cikA background subsequently revealed a putative circadian clock-related phenotype. Gene expression patterns, assayed like those shown in Fig. 1, in a cikA mutant from a number of reporter fusions (kaiA, kaiB, and psbAII) have a reduced amplitude and a period length that is shortened by approximately 2 h; some of these reporter genes also show unusual phasing. However, the most prominent phenotype of a cikA strain is that its circadian clock is practically unresponsive to environmental stimuli such as light and temperature (Mutsuda et al., 2003). A lack of phase resetting in the cikA mutant background in response to a 5-h dark pulse demonstrates that the CikA protein is involved in an input pathway to the Kai-protein circadian clock (Schmitz et al., 2000). The deduced amino acid sequence of the CikA protein reveals three motifs consistent with its involvement in clock input: a potential chromophore binding motif, a histidine protein kinase motif, and a cryptic two-component receiver domain (Mutsuda et al., 2003). The CikA chromophore-binding

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motif is similar in sequence to a so-called GAF motif. (GAF motifs are common among plant and cyanobacterial phytochromes and among other ‘‘phototransducing’’ proteins.) The acronym is derived from the fact that cGMP-speciﬁc phosphodiesterases, Anabaena adenylate cyclases and E. coli FhlA all contain the motif. No particular function can be ascribed to a GAF domain based solely on sequence (Aravind and Ponting, 1997; Anantharaman et al., 2001; Anantharaman and Aravind, 2005). True bacterial and plant phytochromes possess a lyase activity for covalent bilin attachment to their GAF domains. However, the GAF motif of CikA is atypical because it lacks the conserved cysteinyl or histidyl residues expected for covalent bilin binding (Hughes and Lamparter, 1999). A histidyl-tagged CikA (CikA6His) will ligate phytochromobilin and phycocyanobilin chromomophores in vitro, indicating lyase activity even in the absence of conserved bilin-binding residues. However, even though the adduct is attached, it is not photoactive. Deleting the GAF motif sequence from CikA reduces, but does not eliminate, the ability of the protein to form a covalent adduct with phycocyanobilin. Oddly, isolation of CikA6His from the cyanobacterium did not reveal a covalently bound bilin, and expression in a recombinant E. coli strain that express phycocyanobilin or biliverdin, which could have detected a photoactive noncovalent holoprotein, yielded negative results. Overall, these results suggest that the enigmatic bilin binding by CikA in vitro is unlikely to reﬂect a biliprotein complex in S. elongatus PCC 7942 (Mutsuda et al., 2003). Downstream of the GAF motif, CikA possesses a conserved histidyl residue that is the site for autophosphorylation (Mutsuda et al., 2003). Examination of CikA protein sequence reveals all of the conserved N-, D/F-, and G-boxes associated with histidine protein kinases (Stock et al., 1995; Stock et al., 2000). In addition, the CikA protein has demonstrated in vitro autophosphorylation activity. Chemical stability of the phosphorylated protein is consistent with phosphoryl linkage at a histidyl residue. Changing the H393 codon to encode an alanyl residue destroys the autophosphorylation activity. Removal of the amino terminal portion or the GAF domain from CikA protein drastically reduces autokinase activity, suggesting that these domains modulate CikA autophosphorylation (Mutsuda et al., 2003). Within the carboxyl terminal region of CikA is a motif that is similar to the receiver domains found in response regulators involved in two-component signal transduction systems (Stock et al., 1995; Stock et al., 2000). However, this particular receiver domain is cryptic in that it does not contain the conserved aspartyl residue that, in a response regulator, would be phosphorylated by a cognate histidine protein kinase. The potential for an unusual phosphoryl group transfer reaction from phosphohistidyl-CikA to this putative receiver domain was tested by using a CikA

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donor that lacks the carboxyl terminal receiver domain but retains autokinase activity and a recipient H393A-CikA protein that contains the receiver domain but is defective for autokinase activity. No phosphoryl group transfer occurred between these two proteins. Thus, the CikA receiver domain is now considered a pseudo-receiver domain. Truncation of this pseudoreceiver domain from CikA greatly increases the in vitro autophosphorylation activity of the truncated protein relative to wild type CikA (Mutsuda et al., 2003) Also, over-production of just the 133 amino acyl residues CikA pseudo-receiver domain shortens, by 2 h, the period of in vivo gene expression rhythms from a kaiBC reporter system (Zhang et al., 2006). Interestingly, this pseudo-receiver domain is also necessary for the proper intracellular localization of CikA (Zhang et al., 2006). Full-length CikA protein localizes at a cell pole (Zhang et al., 2006). The CtrA and other signal transducing proteins in Caulobacter crescentus also use pseudo-receiver domains as implements to direct polar localization (Ausmees and Jacobs-Wagner, 2003; McAdams and Shapiro, 2003; Stephens, 2004; Sciochetti et al., 2005; Holtzendorff et al., 2006). Recent work also identiﬁed the cikA gene as having a role in S. elongatus PCC 7942 cell division (Miyagishima et al., 2005). Exactly how CikA protein functions in cell division or as a Kai-protein clockinput device remains to be elucidated. The cumulative data regarding CikA protein function suggest that it is part of an input pathway to the Kai-protein circadian clock. It functions as an auto-regulated kinase in which the pseudo-receiver domain is playing two roles. First, the domain negatively regulates CikA autokinase activity and second, it interacts with other proteins to localize CikA at a cell pole. Perhaps interaction of the pseudo-receiver domain with localization proteins keeps it from inhibiting CikA kinase activity and CikA becomes an active kinase only when properly localized (Zhang et al., 2006). Interestingly, the LdpA protein co-puriﬁes with both CikA and KaiA suggesting that all of these proteins may form a large, localized timing-input complex (Ivleva et al., 2005). This complex may be truly multifarious, as ﬁve additional CikA protein interactive partners have been recently identiﬁed from a yeast 2-hybrid screen (Zhang et al., 2006).

5.2. The LdpA Protein The S. elongatus PCC 7942 ldpA gene is 1059 bp in length and encodes 352 amino acyl residues that ultimately form an approximately 38 kDa protein. The functions encoded by the ldpA gene (light-dependent period) affect input to the clock somewhat differently than those encoded by the cikA

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gene. For example, the ldpA gene was identiﬁed from a transposon insertion mutation harbored within a strain showing skewed response to a phasealtering dark pulse (Katayama et al., 2003) Although insertions at the ldpA locus were originally classiﬁed as having an altered phase response phenotype, the actual circadian-related phenotype of the ldpA null allele is a bit more complicated: it has a conditional 1-h shortening of the free-running period in gene expression rhythms (Katayama et al., 2003). As mentioned near the beginning of this review, the bioluminescence rhythm from many cyanobacterial reporters adheres to Aschoff’s Rule, meaning that the period length varies with an inverse relationship to light intensity (Nair et al., 2002; Katayama et al., 2003). In the ldpA null background, these typical period variations are abrogated (Katayama et al., 2003). The deduced amino acid sequence encoded by the ldpA gene allowed prediction that LdpA is a soluble protein that contains two Fe-S centers (Katayama et al., 2003). Both EPR (electron paramagnetic resonance) and UV-visible absorption spectrometry have been used to conﬁrm the presence and redox sensitivity of two [Fe4S4]+1 clusters per molecule of LdpA protein (Ivleva et al., 2005). These data are consistent with LdpA being part of a redox-sensing, signal transduction pathway. Perhaps the pathway transduces a measure of photosynthetic activity to the circadian timing mechanism (Katayama et al., 2003) Over-production of His-tagged LdpA protein does not significantly alter the period of gene expression rhythms (Ivleva et al., 2005). More interestingly, the LdpA6His protein was over-produced in S. elongatus PCC 7942 to determine if any clock-related proteins would co-purify. Amazingly, the CikA, KaiA and SasA proteins all co-purify with the LdpA6His protein (Ivleva et al., 2005). Even in the absence of the KaiB or KaiC protein, LdpA6His still co-puriﬁes with KaiA and CikA (Ivleva et al., 2005). Moreover, KaiA is not required for the CikA–LdpA protein interaction. There are no shared sequence motifs within these three clock-related proteins that would make an obvious LdpA binding region and these proteins do not interact with LdpA in a yeast 2-hybrid assay. Presumably, there are intermediate proteins that enable this group, CikA, KaiA and SasA, to interact with LdpA6His (Ivleva et al., 2005). Only a fraction of the population of CikA and KaiA molecules is interacting with LdpA at any given time. Curiously, the amount of CikA associated with LdpA remains constant over CT whereas the amount of KaiA associated with LdpA is dynamic. KaiA appears to have a higher afﬁnity for LdpA near the middle of the subjective day (Ivleva et al., 2005). The LdpA protein is clearly playing a role in getting redox information into the Kai-protein circadian clock. Determining the source of that information will provide further explanation of the relationship between cellular metabolism and the circadian clock.

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5.3. The Pex Gene The S. elongatus PCC 7942 pex gene is 447 bp in length and encodes 148 amino acyl residues that ultimately form an approximately 17 kDa protein. The pex (period extender) gene was originally identiﬁed as ‘‘apparently complementing’’ a mutation that caused a 22-h, short period circadian rhythm (Kutsuna et al., 1998). In fact, the mutant phenotype was caused by the kaiC1 allele (Table 1). Further analysis showed that the ectopic copy of pex was acting as an extragenic suppressor of the 22-h period by extending the period of circadian rhythms in gene expression by about 2 h. Overproduction of Pex protein in either wild type or various period-mutant backgrounds extends the period and decreases the amplitude of gene expression rhythms (Kutsuna et al., 1998). The pex gene encodes no obvious protein sequence motifs. However, Pex protein sequence does have some similarity to the functionally diverse PadR-family of transcription regulators (Gury et al., 2004; Huillet et al., 2006). Inactivation of pex results in a slightly short period phenotype with respect to gene expression rhythms but does not appear to result in any growth defects. Interestingly, inactivation of pex greatly increases kaiA gene expression. Pex protein seems to act as either a direct or indirect repressor of kaiA expression. Recall the Kai-protein, circadian clock model whereby KaiA adjusts circadian period length by regulating the rate at which KaiC autophosphorylates. The pex-dependent phenotypes ﬁt this model. Over-production of Pex extends the circadian period because KaiA, whose production is thereby repressed, is not present to stimulate KaiC phosphorylation and allow a normal rate of progression through the circadian cycle. Inactivation of pex increases KaiA protein production, due to de-repression at kaiA, and causes shortened circadian periods by over-stimulating KaiC autophosphorylation. Speculative or not, no other speciﬁc role in an input pathway can be deﬁnitively assigned to the Pex protein.

6. OTHER COMPONENTS: THE RPO (SIGMA FACTOR) AND CPMA GENES Because the circadian oscillator pervades expression of the entire S. elongatus PCC 7942 genome, the regulatory mechanisms that couple timing information to downstream genes are likely to be global in nature and, of course, tied to fundamental metabolic processes (Liu et al., 1995a). The chromosome compaction rhythm demonstrates the global dynamics of temporal

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regulation by the circadian clock. However, it is clear in the sasA genetic background that chromosome dynamics are not the whole story behind rhythmic gene expression (Smith and Williams, 2006). Regulatory proteins also have to be in the right place at the right time to get the precise, temporal control of gene expression that is characteristic of the circadian clock. Some evidence points to an underlying rhythmicity in the basic transcription machinery (Tsinoremas et al., 1996; Nair et al., 2002). Insertion mutations in the S. elongatus PCC 7942 rpoD2 gene, which encodes a group 2 sigma factor, results in low-amplitude, gene expression rhythms from a small subset of cyanobacterial promoters (Tsinoremas et al., 1996). Recall that sigma factors are subunits of the prokaryotic RNA polymerase holoenzyme and that they confer promoter speciﬁcity (Gross et al., 1998). Cyanobacteria are somewhat atypical in that they contain multiple, closely related, sigma factors that are not essential for growth. These are known as the group 2 sigma factors. S. elongatus PCC 7942 also has an essential housekeeping sigma factor encoded by its rpoD1 gene (Tanaka et al., 1992). Further analysis of sigma factor encoding genes showed that inactivation of any of the four known group 2 sigma factor genes (rpoD2, rpoD3, rpoD4, and sigC), either singly or in pairs, alters circadian expression patterns from the psbAI promoter (Nair et al., 2002). Inactivation of the sigC gene, which consistently lengthens the period of psbAI expression by 2 h, has little or no effect on the period of expression from either the purF (opposite phase) or kaiBC promoter fusions. Interestingly, expression from the kaiBC promoter is affected only by mutations in rpoD2, or by pair-wise mutations in rpoD3/rpoD4 and rpoD2/rpoD3, suggesting that the kaiBC promoter is relatively insulated from most regulatory pathways. Because each sigma factor dissimilarly affects transcription from speciﬁc cyanobacterial promoters, the working model contends that the cyanobacterial transcription apparatus oscillates in a circadian manner. In fact, oscillations in the RpoD4 protein level have been demonstrated. Also, the composition of the transcription-active, RNApolymerase holoenzymes changes over the circadian cycle due to temporal exchange by the holoenzyme of individual group 2 sigma factors (Nair et al., 2002). Nonetheless, no direct link between the SasA protein – an important output device from the Kai-protein clock – and the expression or activity of any group 2 sigma factor has been demonstrated. It is not clear how any regulatory pathway, sas or rpo, might act in conjunction with the clockdriven compaction rhythm to generate rhythmic patterns of global gene expression. The cpmA gene (circadian phase modiﬁer) may also have a role in output from the S. elongatus PCC 7942 circadian clock. Inactivation of cpmA drastically alters the relative phasing of a subset of cyanobacterial transcription

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reporters. Most interestingly, a cpmA null allele affects the phase angle of the expression rhythm from the kaiA promoter, but has no effect on the kaiBC promoter. This causes a relative phase angle difference of nearly 10 h between the expression rhythms of the two kai cistrons. Despite lack of kai transcription coordination in this cpmA genetic background, circadian timing and rhythmic gene expression remain intact (Katayama et al., 1999).

7. CONCLUSIONS 7.1. Kai Genes and Circadian Clock Evolution Subsequent to conﬁrmation that the three kai genes are essential components for the circadian timing mechanism in S. elongatus PCC 7942 (Ishiura et al., 1998), the genomes of nearly 20 phylogenetically diverse cyanobacteria have been completely sequenced and each contains the genetic basis for a circadian clock: at least one kai locus. Functional clock-driven circadian rhythms have been tested, and subsequently demonstrated with genetic techniques, only in species of the unicellular genera Synechococcus, Synechocystis and Thermosynechococcus (Sweeney and Borgese, 1989; Huang et al., 1990; Chen et al., 1991; Kondo et al., 1993, 1994; Aoki et al., 1995; Aoki et al., 2002). However, over 40 diverse cyanobacterial strains from widely divergent genera have at least one kai gene (Lorne et al., 2000). In addition to the typical phycobilisome-containing cyanobacteria, kai genes are found in four prochlorophyte, chlorophyll b-containing strains, Prochlorococcus marinus sp. MED4, MIT 9313 and SS120, and Prochlorococcus sp. NATL2A; all have the kaiB and kaiC genes, but lack an obvious kaiA ortholog. Although circadian rhythms, and their requisite functional circadian oscillator, have not been demonstrated directly in any of these strains, Prochlorococcus sp. strain PCC 9511 displays distinct diel patterns of gene expression (Holtzendorff et al., 2001). Diel patterns do not necessarily indicate the existence of a circadian clock, but the absence of the diel rhythms would have suggested absence of a circadian clock. As was discussed brieﬂy above, the S. elongatus PCC 7942 KaiA protein plays an essential role in circadian rhythm generation. However, the amino-terminal receiver-like domain is missing in some species, and the entire protein is absent from prochlorophyte species that are very likely to have circadian rhythms. Lack of ubiquity of KaiA is not really surprising across the vast evolutionary distance of the cyanobacteria, especially given the diverse mechanisms these organisms use for harvesting and interpreting light (Bryant, 1994; Bibby

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et al., 2001; Boekema et al., 2001). If the role of KaiA is really as a conduit for information from the input pathways, this function may be ﬁlled by completely unrelated proteins in other species. The chief requirement for any KaiA functional analog is an ability to receive environmental information and dock with the clock complex, inducing a conformational change. It will be interesting to ﬁnd out if any Prochlorococcus sp., lacking a ‘‘true’’ KaiA, can generate circadian rhythms. If so, their clock-input pathways could shed some light on the origins of the cyanobacterial circadian clock entrainment and environmental input mechanisms. Molecular phylogenetic analysis of the kai locus suggests that the kaiBC operon is nearly 2 billion years old (Dvornyk et al., 2003). It seems likely that the common ancestor shared by cyanobacteria and plastids also had prokaryotic-like circadian kai genes. So, where are the kai gene homologs in the plastid lineage? It is not clear. Plastid genomes from algae such as the glaucocystophyte Cyanophora paradoxa, the rhodophyte Porphyra purpurea, and the chlorophytes Chlamydomonas reinhardtii and Chlorella vulgaris are among over two dozen that have been completely sequenced (Reith and Mulholland, 1995; Wakasugi et al., 1997; Glockner et al., 2000; Maul et al., 2002); none of these encodes obvious kai orthologs. Perhaps kai genes will be found as more plastid genomes are sequenced or else they may be genuinely absent in plants, reﬂecting something fundamental about the prokaryotic circadian clock and endosymbiosis. Plastid genome evolution has evidently been a process of size reduction, as most plant genes considered cyanobacterial in origin have been incorporated into the nuclear genome (Martin and Herrmann, 1998; Race et al., 1999; Douglas and Raven, 2003). Cyanobacterial genomes range from about 1.8 to 9 million base pairs, whereas plastid genomes are much smaller, ranging from around 35 to 200 kb (Delwiche and Palmer, 1997). As a result, in A. thaliana about 18% of nuclear genes appear to be of cyanobacterial origin (Martin et al., 2002). Although there is no clear correlation between the evolutionary origin of an A. thaliana gene and the cellular localization of its functional product, many cyanobacterial-like nuclear genes encode products that act exclusively in chloroplast photosynthetic biochemistry (Martin and Herrmann, 1998; Martin et al., 2002). Perhaps in A. thaliana, circadian regulation of incorporated cyanobacterial genes by an existing nuclear-encoded clock (Hennessey and Field, 1992; Roenneberg et al., 1995; Bognar et al., 1999; Sai and Johnson, 1999; Thain et al., 2000, 2002) was sufﬁcient for circadian control of chloroplast development and photosynthetic activity Giuliano et al., 1988; Beator and Kloppstech, 1993; Anderson et al., 1994). As plastid genes were relocated to the nuclear genome, probably for regulatory reasons, the plastid clock became redundant, in light of the fact that the plastid clock was

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not likely amenable to temporal signals from the plant’s tissue-speciﬁc circadian oscillator(s) and input mechanisms. A particular, fascinating plant circadian system may shed more light on the prokaryotic (kai) circadian clock mechanism. Organisms from several genera of single celled eukaryotic green algae, such as Acetabularia, Chlorella, and Gonyaulax, generate circadian rhythms in a variety of physiological functions including photosynthetic metabolism and individual protein abundances (Sweeney et al., 1967; Driessche and Bonotto, 1968; Vanden Driessche and Bonotto, 1969; McMurry and Hastings, 1972; Schweiger and Schweiger, 1977; Woolum, 1991; Hastings, 2001). Incredibly, the nucleus can be removed from the rest of the Acetabularia mediterranea (A. mediterranea)cell and circadian oscillations in photosynthesis and protein abundance continue, under constant conditions, for many weeks in the enucleated cell (Sweeney, 1974). When present, the nucleus may impart phase information upon rhythms because, when the nucleus is entrained differentially relative to the rest of the cell and then added back to the enucleated cell, the nucleus affects phasing of cellular rhythms (Sweeney, 1974; Schweiger and Schweiger, 1977). The underlying timing mechanism in this single-celled eukaryotic alga may resemble that of the prokaryotic Kai clock. Current eukaryotic circadian mechanism models hold that delayed re-entry of transcription regulators into the nucleus is a key temporal component of circadian oscillators (Young and Kay, 2001). In contrast, models that have been proposed to account for most of the circadian phenomena in A. mediterranea include the partitioning of an essential, rhythm-driving protein across an organellar membrane (instead of the nuclear membrane) to monitor time. Theoretically, properties of the membrane, integral membrane transport proteins, or a proposed delayed-release protein, would be altered as a function of time and thereby participate in the A. mediterranea circadian timing mechanism (Sweeney, 1974; Schweiger and Schweiger, 1977). Recent work (see above) that suggests KaiB sequestration to the membrane and delayed re-entry into the cytoplasm as a function of cell metabolism is reminiscent of this 30-year-old hypothesis (Sweeney, 1974; Schweiger and Schweiger, 1977; Kitayama et al., 2003).

7.2. A Cyanobacterial Clock A comprehensive model for the S. elongatus PCC 7942 Kai-protein, circadian clock data reviewed above is illustrated in Fig. 8. At subjective dawn or sunrise, CikA protein, either through its own chromophore or the chromophores of other real or putative interactive proteins such as LdpA, recognizes light input information, and relays that information to the Kai clock

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Figure 8 A schematic depiction of one circadian cycle in the cyanobacterium S. elongatus PCC 7942. Genes or their products that were discussed prominently in the text are included in this highly speculative scheme. High levels of transcription begin the cycle. KaiC protein forms a hexamer as ATP is bound to the CI domain. This subjective day Kai-protein complex is then shown autophosphorylating due to a CII domain-associated KaiA protein. CikA and LdpA are set to regulate this autokinase activity via KaiA. CikA is presumably monitoring light quality and LdpA is somehow tuned into the cell’s redox state. As the dusk approaches the chromosome compacts, transcription levels from most promoters slows, and the Kai-protein complex continues to autophosphorylate at a carefully regulated rate. KaiB protein is associated with KaiB-long or some other membrane associated protein. A redox signal is accepted by KaiB-long and KaiB is released to interact with the soluble Kai-protein complex. The KaiA, KaiB and SasA proteins are all associated with this pre-dawn KaiC-dependent complex. The complex disappears and the chromosome begins to de-compact. A de-compacted chromosome and arrival of the proper regulatory proteins (Rpo, CpmA, Pex) leads to another cycle of temporally regulated gene expression. Many more details are provided in the text.

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for synchronization to local time. The daily light (and temperature) information is integrated into the Kai mechanism by the amino-terminal, pseudo-receiver domain of KaiA. The information relay is probably via a protein–protein interaction, with an as-yet unidentiﬁed protein(s), given the structural similarity of amino-terminal KaiA to bacterial receivers that propagate conformational changes based on information from protein interactions (O’Hara et al., 1999; Roche et al., 2002). At dawn and during the subjective day, the chromosome is diffuse throughout the cell and transcription of the kaiA and kaiBC operons and subsequent translation of KaiA, KaiB, and KaiC monomers occurs. KaiB protein is sequestered to the cell membrane by an as-yet unidentiﬁed, membrane-associated or membrane-bound KaiB-interactive protein. Perhaps sequestration is mediated by a protein that is related to the longer KaiB proteins found in some cyanobacteria (Fig. 4). Meanwhile, KaiC protein monomers bind ATP, inducing the hexamerization of KaiC, and the carboxy-terminal domain of KaiA aids in KaiC autophosphorylation. By subjective dusk, the chromosome has started to compact, KaiC is fully phosphorylated and associated with KaiA, and kaiBC transcription has reached its peak level. KaiB protein is still membrane localized. Interactions between the amino terminus of SasA (which has high sequence similarity to full-length KaiB) with KaiC may be occurring at this point, allowing for the autophosphorylation of SasA to occur while KaiB is still localized to the cell membrane. Phosphorylated SasA could then transfer its phosphoryl group to activate its cognate, as-yet unidentiﬁed, response regulator (SasR), which transmits timing information to downstream genes. Additionally, the putative SasR protein could react, either directly or indirectly, with the kaiBC promoter, to negatively inhibit transcription. At about CT ¼ 20, KaiB is released from the membrane, and association of KaiB with the KaiC/KaiA/SasA complex is initiated, reducing the autophosphorylation of KaiC, and subsequently the autophosphorylation of SasA. Introduction of KaiB to the clock complex itself, or the action of inhibiting KaiC phosphorylation, may induce dissociation of the Kai clock complex by the subsequent subjective dawn.

7.3. A Final Comment Exploiting the genetically ﬂexible cyanobacterium S. elongatus PCC 7942 as a model for the elucidation of a circadian mechanism has been fruitful. In the decade since luciferase reporter rhythms were ﬁrst utilized, core components from input pathways, output pathways, and the oscillator itself have been identiﬁed, and many key features of the underlying biochemistry

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of timekeeping have been established. It is clear that the Kai oscillator is a physical complex: an entity that is built, modiﬁed, and re-generated over a 24-h time span. The cyanobacterial timing mechanism is fundamentally different from models for eukaryotic timing in this respect. In eukaryotic systems, circadian timing seems based upon two separately phased, but intertwining, feedback loops that involve partner switching between positive and negative elements (Dunlap, 1999; Harmer et al., 2001; Van Gelder et al., 2003). In S. elongatus PCC 7942, it is the intricate relationship between the Kai proteins and SasA, and timed modiﬁcations to these proteins by phosphorylation, that constitute the oscillator. Transcriptional autoregulation occurs, but is not an essential part of the timing mechanism and thus plays a minor role in the timepiece. On the other hand, the dynamics of chromosome compaction are similar to those observed in the chloroplast of the green alga Chlamydomonas reinhardtii where genome-wide ﬂuctuations in DNA topology and superhelicity are used to regulate large-scale gene transcription patterns (Salvador et al., 1998). Even the circadian clocks found in insects and mammals evidently use rhythmic histone acetylation and deacetylation for chromatin re-modeling and gene transcription regulation (Curtis et al., 2004; Naruse et al., 2004; Smolen et al., 2004; Brown et al., 2005; Ripperger and Schibler, 2006). As is evident from the projected model of Kai timing, there are still many missing pieces of the circadian-clock mechanism waiting to be found. A comprehensive functional genomics project is under way (see http:// www.bio.tamu.edu/synecho/). The goal of this project is to assay the effect of inactivating each of the 2200 loci in the S. elongatus PCC 7942 genome on circadian rhythms. This project should ultimately identify all of the circadian clock components. Perhaps knowing all of the important players will allow us to piece together the intricate functional relationships that exist among cell metabolism, physiology, behavior and the global regulation of gene expression. Of course, the really big picture will then include tying all of these intracellular functional relationships to the unyielding and unending environmental variable known as time. There is an old saying about time: Time is that thing which keeps everything from happening at once. Lately, it does not seem to be working.

ACKNOWLEDGEMENTS Takao Kondo of Nagoya University, Carl H. Johnson of Vanderbilt University and Susan S. Golden of Texas A&M University have overseen the

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development of the cyanobacterium S. elongatus PCC 7942 as a model system for studies of circadian biology. Their efforts and insights are duly noted. Rachelle M. Smith provided digital photographs of DAPI-stained cyanobacteria. Lory Mattucci provided excellent administrative assistance. During the writing of this review, the author’s time, effort, and laboratory research were supported by the University of Utah.

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Author Index Page numbers in italics indicate where a reference is given in full. Names beginning de, van and von have been listed under their respective alphabets.

Aasa, R., 86, 135, 138, 141, 144 Abdelal, A., 40 Abdelal, A.H.T., 175, 177 Abdelal, A.T., 40 Abed, R.M., 238 Abergel, C., 23 Aboelella, N.W., 149 Abola, A.P., 168 Abrahamson, J.L., 13 Abramson, J., 139 Achenbach, L.A., 123 Acker, G., 116, 118, 123 Ackerley, D.F., 22 Adachi, O., 12, 18, 20–22, 25, 28–29 Adam, E., 277 Adams, E.E., 238 Adman, E.T., 133, 137 Afshar, S., 166 Agger, S.A., 83 Ahring, B.K., 167 Ai, J.Y., 132, 148, 151 Aiba, H., 261 Akimitsu, N., 4 Alami, M., 171 Albracht, S.P., 9 Alefounder, P.R., 168 Allen, H.L., 42, 126 Allen, J.F., 236 Allen, J.W.A., 81 Al-Obaidi, A.H.R., 137 Altendorf, K., 16, 116, 118 Altman, E., 47 Alton, J.D., 4 Altschul, S.F., 158 Alvarez, M.L., 132, 143, 148, 151

Alvarez, S., 149 Alvarez-Ossorio, M.C., 166 Alves, T., 79, 84–85, 91 Alzari, P.M., 76–77 Amann, R., 124, 191 Ambler, R.P., 22–23 Ameyama, M., 18, 20–22, 25, 28–29 Anantharaman, V., 271 Anders, H.-J., 117 Anderson, B.E., 244 Anderson, S.L., 277 Andersson, B., 240 Andersson, C.R., 231, 241–243, 245, 249–253, 264, 275–276 Andreasson, L.E., 86 Andrew, C.R., 139, 146 Anemu¨ller, S., 145 Ang, M.C., 137, 144, 146 Angelskar, S.M., 96 Anjum, M.F., 118 Anraku, Y., 8, 28 Anthamatten, D., 25–26 Antholine, W.A., 136, 138, 196 Antholine, W.E., 137–139, 141–143, 147–149 Anthony, C., 16 Anto´n, J., 124 Aoki, S., 233, 241–243, 245, 249–253, 264, 274, 276 Aoyama, H., 139, 141, 143 Apicella, M.A., 78, 93 Appia-Ayme, C., 23 Appleby, C.A., 25–26 Apte, S.K., 238 Arai, H., 21, 36–39, 117, 155–157, 181

298

Arai, T., 241 Araiso, T., 31, 90 Aralar, E.V., 238 Aravind, L., 257, 271 Arciero, D., 79, 84, 88 Arciero, D.M., 79, 84, 86–89, 96, 119 Arents, J.C., 18 Arko, R.J., 93 Armesto, J.J., 239 Armor, J.N., 114, 195 Arp, D.J., 119–120 Arvai, A.S., 253–254, 263 Asamizu, E., 252 Ascenzi, P., 45 Askeland, R.A., 44 Atack, J.M., 73 Atkins, R., 238 Atlas, R.M., 33 Atlung, T., 8 Augier, V., 34 Auling, G., 115–117 Aullo´n, G., 149 Ausmees, N., 272 Ausubel, F.M., 4 Avarbock, D., 30 Averill, B.A., 139 Avgusˇ tin, G., 162 Axtell, K.M., 19 Baar, C., 117, 183 Baba, I., 116, 118 Bader, R., 18 Baillon, M.-L.A., 77–78 Baird, C., 36 Bais, H.P., 3 Baker, L.G., 13 Baker, L.M.S., 76 Balderston, W.L., 115, 131 Baldwin, M.J., 145 Baliga, N.S., 164–166, 180 Ballal, A., 238 Ballou, D.P., 12 Balmer, Y., 255 Banci, L., 177

AUTHOR INDEX

Banks, R.G.S., 112–113 Baratta, D., 180–181 Barber, D., 136 Barber, J., 277 Barloy-Hubler, F., 168 Barnett, M.J., 168 Baron, A., 170 Barquera, B., 9, 24, 27–29, 187 Barr, M.E., 137, 140, 143 Barras, F., 180 Barrell, B.G., 81, 95 Barrett, C.P., 136 Barrett, J., 22, 98 Barrett, T.E., 247, 280 Bartha, R., 33 Barthelmebs, L., 274 Bartlett, D.H., 123 Bartnikas, T.B., 156 Basham, D., 81, 95 Bass, J.A., 48 Bateman, A., 187 Bater, A.J., 14 Batut, J., 26 Bauer, C.E., 26, 236 Bauer, M., 191 Baumann, B., 115 Baur, H., 40 Baxter, D.A., 281 Bayer, A.S., 6, 48 Bazylinski, D.A., 38, 117 Beard, S.E., 19 Beard, S.J., 238 Beasley, V.R., 239 Beator, J., 277 Beattie, K.A., 238 Beaudet, R., 167 Beaumont, H.J.E., 119 Becher, J., 137 Beck, A., 115, 191 Beck, C., 19, 26 Becker, G.E., 117 Becker, K., 238 Bedmar, E.J., 155, 157 Bedzyk, L., 35

AUTHOR INDEX

Bedzyk, L.A., 19 Beijerinck, M.W., 194 Beinert, H., 39, 132, 136–137, 141, 170 Belas, R., 124, 168 Bell, L.C., 35, 156, 180–181 Bell, P., 238 Bellamy, H., 137 Bellon, G., 5–6, 42 Belyaev, S.S., 126, 167 Bendall, D.S., 91 Bender, K.S., 123 Benghezal, M., 4 Bengrine, A., 23 Benlloch, S., 124 Benoit, S., 77 Benson, D., 238 Bentley, S., 187 Berendes, E., 194 Berg, A., 143 Berg, C.A., 4 Berger, J., 5–6, 42 Berger, U., 116 Bergho¨fer, J., 39, 156, 185 Bergman, B., 239, 242 Bergman, N.A., 194 Berks, B.C., 33, 35, 111, 129–132, 147–148, 151–152, 169–171, 180–181, 191, 196 Berman-Frank, I., 242 Berna, B., 168 Bernet, N., 116, 119 Bernhard, M., 171 Berry, E.A., 20–21 Berry, S., 240, 277 Berry, S.M., 147 Bertini, I., 143, 177 Bertsova, Y.V., 9, 23, 187 Besson, S., 38, 79, 84, 91, 130–131, 133, 139, 147–150, 171, 196 Best, A., 195 Beyeler, M., 44 Bibby, T.S., 277 Bickel-Sandko¨tter, S., 166 Bigotti, M.G., 45

299

Bill, E., 145 Billi, D., 238 Bilous, P.T., 170 Binder, B., 241 Bindoff, L.A., 19 Bird, T.H., 26 Birrer, P., 5–6, 42 Bisaillon, J.-G., 167 Bizouarn, T., 19 Blackburn, N.J., 137, 140 Blasco, F., 34 Blazina, M., 236 Blevins, W.T., 12 Blondeau, R., 95 Blumberg, W.E., 142 Blumer, C., 29, 43–45 Bobo, T., 156 Boddy, L., 7, 35, 42 Boekema, E.J., 240, 277 Boetzel, R., 177 Bogachev, A.V., 9, 23, 187 Bognar, L.K., 277 Bognar, S., 28 Bogorad, L., 281 Bogsch, E.G., 170 Bollag, J.-M., 195 Bollinger, J.A., 150–151, 177 Bonani, G., 239 Bonete, M.J., 166 Bonifacio, C., 84–85 Bonnard, G., 22, 183 Bonneau, R., 164–166, 180 Bonnefoy, V., 23 Bonotto, S., 278 Boogerd, F.C., 168, 180 Boos, W., 178 Borgese, M.B., 241, 276 Borovik, A.S., 137 Borriello, G., 6 Borzym, K., 191 Bothe, H., 162, 196 Bottomley, P.J., 159, 162 Botzenhart, K., 5–6, 42

300

Bouchard, B., 167 Boucher, R.C., 5–6, 42, 126 Bouchez, T., 116, 119 Bourgeois, D., 84–85 Bowser, L., 168 Bradley, A.L., 88–89 Bramlett, M.R., 272–273 Brandt, U., 20–21 Bras, A.M., 98 Brasier, M.D., 235 Braun, C., 37 Bren, K.L., 143 Brenner, S.E., 170, 173 Brenot, A., 76 Brettar, I., 116, 162 Breznak, J.A., 120 Brinkac, L., 124, 168 Brinkac, L.M., 125 Brinkman, F.S., 3, 33 Brittain, T., 31, 152, 181 Brock, T.D., 238 Broda, E., 195 Brody, L.L., 3, 33 Brondijk, T.H.C., 187 Brondsted, L., 8 Brosch, R., 76–77 Brown, E.C., 149 Brown, E.D., 13 Brown, K., 38, 130–133, 139, 146–150, 171, 196 Brown, K.R., 22–24 Brown, R.M., 236 Brown, R.S., 238 Brown, S.A., 281 Brown, S.B., 240 Brumfeld, V., 238–240 Brune, A., 119 Brunner, F., 40 Brunner, J., 171 Brunori, M., 36, 135 Bruschi, M., 23 Bruseth, L.J., 96 Bruyant, F., 276 Bryan, B.A., 39, 117

AUTHOR INDEX

Bryant, D.A., 240 Bryk, R., 76–77 Brysk, M.M., 44 Brzezinski, P., 139 Bubacco, L., 143 Buchan, A., 124, 168 Buchanan, B.B., 255 Buchwald, S.L., 113 Buer, J., 7, 11, 41 Bunch, A.W., 43, 45 Bundy, J.G., 45 Bu¨nning, E., 231 Burgess, B.K., 22 Burggraf, S., 125, 164–165 Burghardt, J., 126 Bu¨rgisser, D.M., 170 Burke, J.F., 4 Burns, B.P., 239 Burris, R.H., 113, 116 Bu¨sch, A., 190 Busch, J.E., 36–37, 182 Buse, G., 138, 140–141 Bush, D.R., 170 Bushnell, D., 35, 42 Butt, J.N., 130, 148, 151, 191 Butzler, J.P., 81 Buxton, G.V., 113 Bylund, J., 47 Byrne, J.H., 281 Cabello, P., 166 Cabrito, I., 38, 130–133, 149–150 Cahors, S., 156, 168 Calderwood, S.B., 4 Calhoun, M.W., 24 Camara, M., 44 Cambillau, C., 38, 130–133, 139, 146–150, 171, 196 Campanaro, S., 123 Campbell, J.I., 6, 48 Campbell, J.J., 18 Campbell, M.E., 49 Campos, E.G., 98 Canales, S.R., 244 Cannata, N., 123

AUTHOR INDEX

Cannon, C.L., 4 Canters, G.W., 36–37, 137, 142–144, 146, 182 Cao, J., 27 Caparon, M.G., 76 Capela, D., 168 Caradoc-Davies, T.T., 22 Carlson, C.A., 33–34, 39, 117 Carlton, J., 124, 168 Carmichael, W.W., 236, 238–239 Carpenter, E.J., 239 Carr, G.J., 37 Carter, D.C., 22 Carterson, A.J., 49–50 Caru, M., 239 Casalot, L., 116 Cascio, D., 143 Casey, M., 4 Cashmore, A.R., 277 Castets, A.M., 238 Castillo, F., 16, 18, 35 Castresana, J., 26, 37–38, 163, 195 Castric, P.A., 29, 43, 46 Catalan-Sakairi, M.A.B., 119 Catlow, C.R.A., 113 Caughey, W.S., 113 Cavalier-Smith, T., 236 Cavanaugh, C.M., 235 Cavazza, C., 23 Cavin, J.F., 274 Cayol, J.-L., 116 Cecchini, G., 11 Cekici, A., 5–6, 42 Cestaro, A., 123 Cha, M.-K., 77–78 Chakrabarty, A.M., 48–49 Chakraborty, R., 123 Chakravarti, D., 281 Chan, C., 181, 275 Chan, J.M., 150–151 Chan, R., 42 Chan, S.I., 141 Chan, Y.-K., 155, 168, 172, 192 Chanal, A., 172

301

Chandonia, J.-M., 170, 173 Chang, Z., 190 Chapman, S.K., 143 Charnock, J.M., 131–132, 135, 139–140, 145–146 Charoenlap, N., 78 Chatelain, R., 116 Chatterjee, S., 98 Chauvatcharin, N., 78 Cheesman, M.R., 37, 147–148 Chelikani, P., 76 Chen, D., 19 Chen, F., 35, 42 Chen, K.-Y., 116 Chen, L., 143 Chen, M., 173 Chen, M.-Y., 116 Chen, P., 130, 147, 149–151 Chen, T.-H., 241, 276 Chen, T.-L., 241, 276 Chen, X., 134 Chen, Y.B., 242 Chenivesse, S., 23 Chervin, C., 113 Chillingworth, T., 81, 95 Chin, S.M., 75 Chippaux, M., 23, 34 Chisholm, S.W., 236 Chistoserdov, A., 83 Chistoserdov, A.Y., 83 Chistoserdova, L., 18 Chistoserdova, L.V., 83 Chiu, Y., 164–166, 180 Cho, B.H., 18 Cho, Y., 270 Chobot, S.E., 88–89 Choi, J., 77 Choi, S., 77 Chong, S., 31, 78 Chorus, I., 239 Choukair, M.K., 143 Chow, T.J., 241, 276 Christen, R., 116, 162 Christensen, H.E., 23

302

Christiansen, N., 167 Chu, K.K., 47 Churcher, C., 81, 95 Ciabatti, I., 36 Cioﬁ-Baffoni, S., 177 Ciofu, O., 6, 48 Cipollone, R., 45 Clark, M.A., 175, 177 Clawson, B.J., 43 Clemente, A., 143 Clements, M.O., 78 Cline, K., 170, 172 Cnockaert, M.C., 126 Coates, J.D., 123 Coats, J.R., 79, 84, 96 Codd, G.A., 238 Coddington, A., 34 Coenye, T., 126 Cohen, H., 113 Cohen-Bazire, G., 240 Cole, J., 33, 35, 79, 93–94, 96 Cole, J.A., 35, 170, 187 Cole, J.R., 120 Cole, K.A., 123 Cole, K.M., 238 Cole, S.T., 76–77 Collins, F.S., 4 Collyer, C.A., 143 Colombo, I., 19 Comolli, J.C., 8, 25–28, 30, 45 Comtois, S.L., 78 Conover, R.C., 77 Conrad, L.S., 23 Constantinidou, C., 77–78 Contopoulou, R., 159 Conway, B.A., 47 Cook, G.M., 8, 24, 30 Cook, W.O., 239 Cooper, A., 85, 90–92 Cooper, M., 8, 25, 30–31, 45 Copie´, V., 177 Cornelsen, S., 236, 277 Corpe, W.A., 43 Cosper, N., 177

AUTHOR INDEX

Cosseau, C., 26 Cosson, P., 4 Costa, C., 85, 92 Costa, J.L., 239 Costello, C.M., 42 Costerton, J.W., 42 Cotton, N.P., 19 Coulter, S.N., 3, 33 Couves, J.W., 113 Covert, J.S., 124 Cox, G., 238 Cox, G.M., 31, 97 Cox, M.M., 257, 263, 266 Cox, R., 117 Coyle, C.L., 38, 127, 129–131, 133, 139, 141, 144–145, 147, 150, 171, 179–180, 195 Cramer, S.P., 142, 144 Cramm, R., 37–38, 168, 190 Crawford, J.A., 155 Crofts, A.R., 20–21 Cromack, K., 159, 162 Crooks, G.E., 170, 173 Crossley, H., 78 Crowley, P.B., 91 Cruce, D.D., 93 Cruz-Ramos, H., 30 Csekes, J., 28 Cui, L., 277 Cunningham, L., 28–29, 44–45 Cuppoletti, J., 6, 42 Curtis, A.M., 281 Curty, L.K., 4 Cusanovich, M.A., 91 Cutruzzola, F., 36 Cuvin-Aralar, M.L., 238 Cuypers, H., 37, 39, 137, 148, 154, 156, 169, 185 Czaplinski, K., 257 Dabert, P., 116, 119 Dahlem, A.M., 239 Dalbey, R.E., 173 Daldal, F., 26

AUTHOR INDEX

Damas, A.M., 22–23 Damerval, T., 238 D’Angelo, M., 123 Daniel, M., 23 D’Aoust, J.Y., 125 D’Argenio, D.A., 4 Date, S.V., 164–166, 180 Daugherty, S.C., 124–125, 168 Davidsen, T.M., 125 Davidson, V.L., 83, 143 Davies, K.J., 7, 35, 42 Davies, R.M., 81, 95 Davis, J.S., 238 Davis, R.W., 168 Davis, S.H., 49–50 Davy, S.L., 177 Davydov, R., 137 Dawes, S.S., 30 de Beer, D., 239 de Boer, A.P., 24, 26, 29, 96 de Boer, A.P.N., 155, 157, 163, 191–192, 195 de Gier, J.W., 24, 26, 29 de Gier, J.-W.L., 163, 195 De Hou, Y., 77 De La Mora-Rey, T., 83 de la Rosa, F.F., 166 de Leeuw, E., 171 De Philippis, R., 236 De Smet, L., 31, 84–87, 90–91 de Vos, W.M., 191 de Vries, S., 137–142, 144, 146, 166–168, 172, 192 de Weert, S., 155, 191–192 Dean, R.T., 6 DeBeer George, S., 142, 144 Deboy, R.T., 124–125, 168 Deckwer, W.D., 48 Defago, G., 46 Degobbis, D., 236 Deitermann, S., 171 DeKievit, T.R., 42, 126 del Prado, A., 120 Delgado, M.J., 155, 157

303

Delgenes, J.P., 116, 119 DeLisa, M.P., 172 Dell, R.M., 113 Delon, C., 91 Delwiche, C.F., 277 deMairan, J.J., 231 Deming, J.W., 125 DeMoss, J.A., 34 Denariaz, G., 117, 167 Deng, T.S., 277 Denner, E.B.M., 124 Dennison, C., 137, 143–144 dePamphilis, C.W., 277 Deretic, V., 47 Dermastia, M., 37 Deruelles, J., 238 Deutsch, E.W., 164–166, 180 Devlin, F., 139, 148 Devreese, B., 79, 83–85, 90–92, 130–131, 133, 150 Deziel, E., 3 Di Bernardo, S., 9 Diamantis, A.A., 114 Dias, J.M., 84–85 Dibrov, P.A., 9 Dickerson, R.E., 16, 18 Didonato, S., 19 Diekmann, H., 115–116 Dilks, K., 165 Dimroth, P., 9 DiRita, V.J., 96, 155 DiSpirito, A.A., 79, 84, 96, 119 Ditty, J.L., 240, 242–245, 248, 253, 264, 266, 273, 275 Divies, C., 274 Djinovic-Carugo, K., 38, 130–133, 139, 143, 146, 149–150, 196 D’Mello, R., 8, 30 Dodson, R.J., 124–125, 168 Do¨hler, K., 117–118, 126–127 Doi, M., 117 Dombroski, A., 275 Dong, A., 113 Dong, G., 270, 272

304

Donker, H.J.L., 195 Donohue, T.J., 8, 25–28, 30, 45 Dooley, D.M., 38, 129, 131–132, 139, 143, 147–148, 150–151, 177, 196 Dor, I., 238 Doran, P.T., 238 Doring, G., 4–6, 42 Dorsch, M., 117 Dos Santos, J.P., 34 Doudoroff, M., 7, 13, 159 Douglas, A.E., 236, 239, 277 Downs, D.M., 192 Drake, H.L., 116, 118, 123 Drenkard, E., 4 Dreusch, A., 119, 129, 131–132, 135, 137, 139–140, 145–146, 148, 169–172, 177 Drew, R.E., 247, 280 Driessche, T.V., 278 Drikos, G., 113 Drobner, E., 125, 164–165 Drummond, J.T., 113 Duarte, L.C., 79, 84–85, 91 Dubourdieu, M., 34 Duffy, B., 46 Duine, J.A., 14, 16, 83 Dunford, H., 90 Dunford, H.B., 31 Dunlap, J.C., 235, 281 Dupetit, G., 194–195 Durand, M., 117 Durham, B., 31, 92 Durkin, A.S., 124–125, 168 Durley, R., 143 Dutton, P.L., 134 Dvornyk, V., 231, 233, 240, 257, 277 Dwarte, D.M., 238 Eady, R.R., 130, 135 Eberl, L., 3 Echalier, A., 84, 87 Edmunds, L.N., 241 Eftekhar, F., 6, 48 Egli, M., 251, 256–258

AUTHOR INDEX

Eglinton, D.G., 136 Ehrlich, A., 238 Ehrlich, G.D., 6 Eiamphungporn, W., 78 Einarsdo´ttir, O., 113 Einsle, O., 113, 128, 152, 159, 167, 172–173, 182, 184 Eisem, J.A., 124, 168 Eisenmann, E., 168 Eisensamer, B., 277 Eisner, G., 171 Eitinger, T., 168 Elazari-Volcani, B., 124, 195 Elias, M., 18 Elias, M.D., 12, 18 ElKurdi, A.B., 94 Ellfolk, N., 31, 79, 86, 90 Ellington, M.J., 35 Elliott, S.J., 88–89 Ellis, H.R., 76–77 Elsen, S., 26 Enoch, H.G., 34 Entsch, B., 12 Epel, B., 139, 141 Eppinger, M., 117, 183 Eraso, J.M., 27 Erman, J.E., 78 Errington, N., 90–92 Erwin, A.L., 3, 33 Eschbach, M., 7, 11, 41 Escudero, K.W., 19 Espersen, F., 6 Evans, W.R., 238 Ezaki, T., 126, 161 Fabian, M., 144 Facciotti, M.T., 164–166, 180 Fairhurst, S., 130, 149–151 Falcinelli, S., 36 Falconer, I., 239 Falkowski, P., 242 Fall, R., 3 Falsen, E., 126 Faraloni, C., 236

AUTHOR INDEX

Fardeau, M.-L., 116 Farmer, S.W., 49 Farrar, J.A., 137–139, 141–143, 147–148 Farver, O., 139, 141, 146 Fastner, J., 238–239 Fauque, G., 79, 84, 91, 130–131, 133, 150 Faurisson, F., 4 Feary, T.W., 12 Federspiel, N.A., 168 Fedorova, N.D., 9 Fedorova, R.I., 131 Fee, J.A., 136–138, 140, 143–144 Feldblyum, T.V., 125 Felix, C.C., 142 Felsenstein, J., 79–80, 158 Feltwell, T., 81, 95 Feng, B., 9, 187 Ferguson, L.P., 33–34 Ferguson, S.J., 22, 33, 35, 37, 79, 81, 95, 111, 117, 156, 168–170, 180–181 Ferguson-Miller, S., 27 Fernandez, C.O., 137 Fernandez, R.O., 46 Ferretti, S., 130, 135 Field, C.B., 277 Figueras, J., 238 Filenko, N., 187 Filiatrault, M.J., 35, 42 Filloux, A., 172 Finel, M., 26 Finkmann, W., 116, 118 Finneran, K.T., 125 Finocchiaro, G., 19 Fischer, H.M., 19, 157 Fisher, K. Dance, 144 Fisher, R.F., 168 Fita, I., 76 FitzGerald, D.J., 12 FitzGerald, G.A., 281 Fitzgerald, J., 239 Fitzgerald, M.X., 42 FitzGerald, P.C., 12

305

Fitz-Gibbon, S.T., 165–166 Fjellbirkeland, A., 96 Flatt, P.M., 190 Fleiszig, S.M., 4 Fletcher, H.M., 77 Fleury-Olela, F., 281 Flores, E., 241 Fluit, A.C., 4 Focken, U., 238 Foglino, M., 44 Folger, K.R., 3, 33 Fonstein, M., 117 Foote, N., 31, 88 Foster, S.J., 78 Fox, S., 143 Frangipani, E., 45 Frank, C.A., 238 Frank, D.W., 4 Frank, J., 16 Franklin, F.C., 13 Fraser, C.M., 125 Freeman, H.C., 143 Frerman, F.E., 19 Friedmann, E.I., 239 Friedrich, B., 37–38, 137, 148, 168–169, 171, 190 Frisk, A., 49–50 Fritsen, C.H., 238 Frixon, C., 34 Fromme, P., 229, 240 Froncisz, W., 136 Frost, L., 170 Frunzke, K., 37, 117, 127, 182, 195 Fuchs, G., 117 Fuhrman, J.A., 124 Fujiki, H., 239 Fujita, M., 246–247, 250, 252 Fujiwara, T., 27, 37, 166 Fukamizo, T., 256 Fuks, D., 236 Fukumori, Y., 27, 37–38 Fu¨lop, V., 31, 79, 83–88, 92, 133, 181 Fuqua, C., 124, 168 Furey, W.R., 22

306

Furuishi, K., 187 Furukawa, Y., 233, 250, 253, 257–258, 263 Fyfe, J.A., 49 Gacesa, P., 47 Gade, D., 191 Gadsby, P.M., 31, 34, 88 Gak, E., 83 Galibert, F., 168 Galimand, M., 39, 73, 97 Gallagher, L.A., 4, 29, 46 Galperin, M.Y., 9 Gamelin, D.R., 142, 144, 146 Gamper, M., 40 Gan, R.R., 164–166, 180 Gandhi, H., 120 Ganeshkumar, N., 194 Gao, T., 270 Gao, Y.-G., 137, 144 Garau, J., 4, 37 Garavaglia, B., 19 Garber, R.L., 3, 33 Garbuglio, N., 19 Garces, R.G., 246–247, 253, 263, 265 Garcia, J.-L., 116–117 Garcia-Gil, J., 238 Garcia-Horsman, J.A., 24, 27–29 Garcia-Pichel, F., 238–239 Garczarek, L., 276 Gardner, P.R., 42, 126 Garner, C.D., 131–132, 135, 139–140, 145–146 Garrison, G., 177 Garrity, G.M., 125 Gatzy, J.T., 5, 42 Gavira, M., 16, 18, 35 Gayon, U., 194–195 Gebauer, G., 120 Gebbie, L., 4 Gellera, C., 19 Gel’man, N.S., 22 Gemeinhardt, S., 37–38 Gennis, R.B., 9, 24, 27–29, 144, 187

AUTHOR INDEX

George, G.N., 136, 177 George, S.J., 142 Georgiou, G., 172 Ge´rard, F., 171–172 Geren, L., 31, 92 Germon, J.C., 155 Gerwick, W.H., 190 Getzoff, E.D., 253–254, 263 Ghosh, S., 130, 147, 150–151 Gibson, Q.H., 31, 136 Gibson, T., 79–80 Gibson, T.J., 158 Gidley, M.D., 78 Giles, S.S., 31, 97 Gill, R.E., 19 Gillon, W., 246–247, 253, 263, 265 Gilmour, R., 79, 84–85 Gime´nez, M.I., 165 Ginger, M.L., 81 Ginzburg, B.Z., 126 Ginzburg, G., 113 Ginzburg, M., 126 Giordano, G., 34 Giordano, M., 238 Giovannoni, S.J., 238 Girio, F.M., 85 Girio, F.M.F., 79, 84, 91 Giudici-Orticoni, M.T., 23 Giuliano, G., 277 Givskov, M., 6, 48 Glo¨ckner, F.O., 191 Glockner, A., 119 Glockner, G., 277 Glusman, G., 164–166, 180 Go, M., 248–249, 251, 260 Godfrey, C., 34 Godon, J.-J., 116, 119 Gohlke, U., 171 Gokce, N., 117 Golden, J.W., 238 Golden, S.S., 231, 233, 236, 240–253, 256, 258, 260–262, 264–266, 270–276 Goldfarb, D., 139, 141 Goldfarb, W.B., 29, 46

AUTHOR INDEX

Goldman, B., 22, 183 Goltry, L., 3, 33 Gomez, L., 4, 37 Gonza´lez, J.M., 124, 168 Goodhew, C.F., 79, 83–85, 87, 90–92, 94 Goodin, D.B., 86, 88 Goodman, S.I., 19 Goodwin, P.M., 16 Gordon, D.A., 238 Gorelova, O.A., 236, 239 Gorelsky, S.I., 130, 147, 150–151 Gorisch, H., 14–16 Gorny, N., 119 Gotoh, T., 240 Gottesman, S., 12 Gottschalk, G., 168 Goudreau, P., 261–262, 265, 271 Goufﬁ, K., 171 Gould, R.O., 22–23 Gouzy, J., 168 Govan, J.R., 47, 49 Graham, S.B., 14 Graichen, M.E., 83 Grant, M.A., 153 Grant, W.D., 124, 164–165 Grassineau, N.V., 235 Gray, H.B., 138, 140, 143–144, 146 Greaves, G.N., 113 Green, J., 30 Green, J.W., 236 Green, O.R., 235 Greenberg, E.P., 117 Greenﬁeld, A.J., 117, 168, 180 Greenway, B., 117 Greenwood, C., 31, 34, 79, 84–88, 92, 135–136 Grewell, C.L., 150–151 Grifﬁn, K.J., 19 Grifﬁn, P., 76–77 Grifﬁth, O., 76 Grifﬁths, D.E., 136, 141 Grifﬁths, L., 35 Grigoryan, A.A., 126, 167

307

Grimsrud, P.A., 83 Grisham, M.B., 42 Grishin, N.V., 257 Grobbelaar, N., 241 Grobben, N.G., 115 Gross, C.A., 275 Gross, R., 117, 183 Grossman, A.R., 240 Grossmann, J.G., 130, 135 Grotzschel, S., 239 Gruber, T., 275 Guasco, S., 23 Guckert, J.A., 145 Guerlesquin, F., 23, 90–92 Guerrero, R., 239 Guest, J.R., 97 Guevara, R., 239 Guglielmi, G., 238 Guidici-Orticoni, M.T., 23 Guigliarelli, B., 34 Guimaraes, B.G., 76–77 Gunsalus, R.P., 11 Gupta, R., 137 Gurjal, M., 168 Gury, J., 274 Guss, J.M., 143 Gyurko, C., 194 Ha, D.T.C., 124 Haas, D., 7, 29, 36, 39–40, 43–46, 73, 97 Haft, D.H., 124, 168 Haidaris, C.G., 35, 42 Hajdu, J., 31, 79, 84–88, 92, 181 Hall, A., 277 Hallberg, K.B., 115 Haltia, T., 38, 130–133, 139, 145–146, 149–150, 182, 196 Hamada, A., 277 Hambrook, J.A., 238 Han, J., 139 Han, S., 31, 78, 253–254, 263 Han, S.H., 18 Hancock, R.E., 3–4, 33, 42 Hancock, R.E.W., 126

308

Ha¨ner, A., 117 Hanlon, S.P., 79 Hansen, A., 236, 277 Haque, M.M., 239 Harada, K., 239 Harada, K.I., 239 Harder, J., 115 Hardin, P.E., 281 Harding, C., 137 Harding, M.M., 22–23 Harding, S., 85, 90–92 Harding, S.E., 85, 92 Harkins, S.B., 137 Harmer, S.L., 281 Harms, N., 83, 181 Harris, E.H., 277 Hart, L.T., 117 Hart, S.E., 91 Hartig, E., 96 Hartmann, A., 155 Hartshorne, S., 81 Hase, C.C., 9 Hasegawa, M., 236, 277 Hasegawa, N., 21, 37, 181 Haselkorn, R., 117, 240 Hasnain, S.S., 130, 135 Hassett, D.J., 6, 42, 48–50, 126 Hastings, J.W., 278 Hathaway, B.J., 143 Hay, M., 137 Hay, M.T., 137, 142, 144, 146 Hayashi, F., 233, 246–247, 250, 252–253, 257–258, 263 Hayashi, M., 9, 187 Hayashi, S., 174 Hayes, P.K., 238 Hayon, E., 113 He, C., 137 Heaton, D.N., 177 Hederstedt, L., 11 Hedman, B., 142, 144, 146 Heerfordt, L., 8 Hegab, A.H., 42 Heichen, R.S., 159, 162

AUTHOR INDEX

Heidelberg, J.F., 124, 168 Heider, J., 115 Heikkila¨, M.P., 135, 171–172 Heiss, B., 37, 195 Heitmann, K., 115, 191 Heizmann, C.W., 170 Helmann, J.D., 78, 155 Helton, M.E., 149 Hemmerich, P., 137 Henderson, R.J., 112–113 Hendrich, M.P., 137 Hendriks, J., 37–38 Hendrixson, D.R., 96 Henkel, G., 140 Henne, A., 168 Hennecke, H., 19, 25–26, 157 Hennessey, T.L., 277 Hennig, J., 13 Hennigan, R.F., 42, 126 Henriksen, J.R., 124, 168 Herb, M., 171, 175, 177, 192 Herbig, A.F., 78 Herdman, M., 238 Hermes-Lima, M., 98 Hernandez-Marine, M., 238 Herren, C.D., 95, 97 Herrero, A., 241 Herrmann, R.G., 277 Herzog, E.D., 281 Hess, W.R., 238, 276 Heurlier, K., 44 Hickey, M.J., 3, 33 Hicks, D.B., 8 Hifney, A., 277 Higashi, T., 16, 18, 36–37, 183 Higgins, D., 79–80 Higgins, D.G., 26, 158 Hijikata, A., 248–249, 251, 260 Hill, B.C., 136, 177 Hill, H.A.O., 146 Hill, S., 8, 30 Hillhouse, G.L., 113 Hilliard, G.M., 6, 42, 126 Hillman, B.C., 42

AUTHOR INDEX

Hindle, Z., 44 Hino, T., 38 Hiraishi, A., 126 Hirosawa, M., 252 Hitomi, K., 253–254, 263 Hobeck, P., 126 Hochstein, L.I., 126, 164–166 Hodgson, K.O., 142, 144, 146 Hoeren, F.U., 169–170 Hoffman, B.M., 86, 88 Hoffman, M.Z., 113 Hoffman, N.E., 277 Hoffman, P., 153 Hoffman, P.S., 76 Hoffmann, R., 114, 144 Ho¨ﬂe, M.G., 116, 162 Ho¨hener, P., 117 Hoglen, J., 37 Hoiby, N., 5–6, 48 Holden, M.T., 44 Holder, I.A., 4, 12 Hole, U.H., 168 Hollocher, T.C., 37–39, 117, 127–128, 131, 150, 168, 183 Holloway, P., 155 Holmes, K., 98 Holroyd, J.A., 240 Holroyd, S., 81, 95 Holt, J.G., 125, 161 Holt, R.A., 79 Holtzendorff, J., 272, 276 Holz, R.C., 132, 143 Holzenburg, A., 260 Holzer, G., 126 Hommes, N.G., 119 Hon, G., 170, 173 Hong, A., 168 Honisch, U., 39, 135, 154–156, 171–172, 178 Honore, N., 76–77 Hood, L., 164–166, 180 Hooper, A., 79, 84, 88 Hooper, A.B., 79, 84, 86–89, 96, 119 Hooser, S.B., 239

309

Hopkins, P.M., 194 Hori, H., 38 Horikoshi, K., 123 Horio, T., 16, 18, 36–37, 183 Hormazabal, V., 238 Horn, M.A., 116, 118, 123 Hornberg, J.J., 172, 192 Horsburgh, M.J., 78 Hosler, J., 27 Hosouchi, T., 252 Houben, E., 156 Houben, E.N.G., 155, 191–192 Houmard, J., 236, 238 Houser, R.P., 137, 140, 142–143, 146 Howe, C.J., 91 Howlett, G., 181 Hu, W., 83, 85 Huang, F., 240 Huang, T.-C., 241, 276 Huber, R., 113, 125, 143, 164–165 Huete, J., 239 Hufnagle, W.O., 3, 33 Hugenholtz, P., 126 Hughes, C.V., 194 Hughes, J., 271 Huillet, E., 274 Huizar, L., 168 Hung, L.-M., 241, 276 Hung, P., 164–166, 180 Hunt, J.C., 16–18 Hunter, D.J., 23, 79 Huss, V.A.R., 126 Huston, A.L., 125 Hutchings, M.I., 156 Hutchins, S.R., 117 Hu¨ttermann, J., 139, 141 Hutton, M.N., 19 Huu, N.B., 124 Hwang, H.J., 146–147 Hwang, S.H., 6, 42 Hyde, J.S., 136, 142–143 Hyman, R.W., 168

310

Ichihara, A., 239 Ichiki, H., 166 Igarashi, Y., 21, 36–39, 117, 155–157, 181 Iglewski, B.H., 3–4, 35, 42, 126 Ihssen, J., 116, 118, 123 Iijima, N., 281 Ikeda, S., 113 Ikeda, Y., 19 Ikemoto, H., 241 Ikeuchi, M., 238, 252 Ilyukhina, N.I., 131 Imada, K., 233, 250, 253, 257–258, 263 Imai, K., 233, 244, 248, 263 Imanishi, S., 239 Imlay, J.A., 75–76, 96 Inagaki, F., 116 Inamura, A., 277 Inatomi, K.-I., 166 Ingham, E., 78 Ingledew, W.J., 11–12, 34 Ingraham, J.L., 7, 33–34, 39, 175, 177 Inkson, T.I., 4 Inman, R.B., 257, 263, 266 Inoue, H., 240 Ioerger, T.R., 246–247, 263 Iriguchi, M., 238, 240, 252 Ishida, C., 238 Ishikawa, A., 240 Ishiura, M., 233, 235, 240–253, 257–258, 260–266, 274–276 Islam, S., 239 Itezono, Y., 239 Ito, E., 239 Ito, H., 233, 244, 248, 263 Ito, M., 19, 277 Ito, S., 277 Itoh, M., 180–181 Itoh, N., 246–247, 250, 252, 258, 263 Itoh, S., 233, 235, 252, 258, 263 Itoh, Y., 14, 240 Iuchi, S., 97 Ivanov, M.V., 126, 167 Ivanova, A.E., 126, 167

AUTHOR INDEX

Ivanova, N., 117 Ivleva, N.B., 272–273 Iwamoto, K., 239 Iwasaki, H., 127, 195, 233, 241–246, 248–258, 260–266, 276, 278 Iwasaki, T., 166 Iwase, R., 233, 250, 252–253, 257–258, 263 Iwata, S., 139, 141, 143 Ize, B., 172 Jaafar, S., 194 Ja¨ckel, U., 116 Jackson, D.W., 49–50 Jackson, J.B., 19, 117, 156, 168, 180–181 Jacobs-Wagner, C., 272 Jacquet, S., 276 Jagels, K., 81, 95 Jagtap, P., 117, 183 Jahn, D., 7, 11, 41 Jahn, U.R., 194 Jahnke, L.L., 165 Jakob, W., 129–131, 133, 141, 144–145, 147, 150, 171, 179–180, 195 Jakubovics, N.S., 78 Jalli, T., 28 James, B.R., 114 James, C.A., 144 Jannasch, H.W., 117 Janowiak, B., 76 Janson, S., 239 Jarman, T.R., 49 Jazairi, J., 177 Jeanmougin, F., 79–80 Jenkinson, H.F., 78 Jennings, M.P., 78, 93 Jensen, B.B., 113 Jensen, H.B., 96 Jensen, P., 6, 48 Jeong, W., 78 Jeoung, J.H., 15–16 Jephcoat, A.P., 235 Jeter, R.M., 39, 117

AUTHOR INDEX

Jetten, M.S.M., 115, 119 Jia, J., 190 Jin, D-Y., 77 Jin, H., 139 Jinkins, P.A., 42 Job, C., 117 Jo¨bsis-VanderVliet, F.F., 136 Joffe, A.M., 4 Johnsen, A.H., 6, 48 Johnsen, C.V., 125 Johnson, C.H., 231, 233, 240–243, 245, 249–253, 256–258, 263–264, 266, 269–270, 274–277 Johnson, E., 166 Johnson, MK., 77 Johnson, N.A., 77 Johnson, S.R., 93 Johri, S., 124, 168 Join-Lambert, O., 4 Jones, A.M., 168 Jones, C.L., 22–23 Jones, D.T., 133 Jones, K., 113 Jones, K.L., 42 Jones, L.H., 83 Jones, R.S., 47 Jones, T., 168 Jongejan, L., 83 Jordan, P., 240 Jo¨ssang, P., 194 Ju, L.K., 35, 42 Jubran, N., 113 Juez, G., 126–127, 165 Jug, K., 112 Juhas, M., 3 Jumel, K., 85, 92 Junemann, S., 8, 24, 29–30 Jupiter, R., 49–50 Ka, J.O., 36 Kadener, S., 281 Kadziola, A., 23 Kaetzke, A., 117

311

Kaftan, D., 240 Kageyama, H., 246, 248, 251, 253–255, 260, 263–265 Kageyama, M., 16 Kahn, D., 168, 280 Kahn, M.L., 168 Kaito, C., 4 Kalman, S., 168 Kamagata, K., 249, 260 Kamani, M.C., 6, 42, 126 Kamekura, M., 124, 126–127, 164–165 Ka¨mpfer, P., 116–117 Kana, B.D., 30 Kaneko, T., 238, 240, 252 Kannt, A., 131–132, 135, 139–140, 145–146 Kano, M., 116, 118 Kao, M.C., 9 Kapatral, V., 117 Kaplan, A., 238–239 Kaplan, H.B., 3 Kaplan, S., 26–27, 30 Kaplan, W.A., 110 Kapoor, M., 277 Kappen, L., 239 Kappl, R., 139, 141 Karlin, K.D., 149 Karlsen, O.A., 96 Karlsson, J.J., 23 Karlyshev, A.V., 81, 95 Karplus, P.A., 76–77 Kas, A., 3, 33 Kaspar, H.F., 117 Kasting, J.F., 236 Kastrau, D.H., 37 Kastrau, D.H.W., 138, 141 Katayama, M., 242–245, 266, 270, 272–273, 276 Katayama, Y., 126 Kates, M., 126–127, 165 Kather, B., 16 Kato, C., 123 Kato, H., 246–247, 250, 252 Kato, I., 119

312

Katoh, H., 238, 252 Katoh, S., 238 Katsari, E., 177 Katsaros, N., 177 Kavanaugh-Black, A., 48 Kawamura, Y., 126, 161 Kawasaki, S., 37 Kawashima, K., 238, 240, 252 Kay, S.A., 241, 277–278, 281 Kaya, K., 238 Keating, D.H., 168 Keatings, V.M., 42 Keegstra, W., 277 Keel, C., 46 Kehoe, D.M., 240 Keizer, H.H., 136 Keller, H., 117, 183 Kellmann, R., 239 Kelly, D.J., 73, 78, 81, 95 Kelly, D.P., 126 Kelly, M., 138, 141, 143, 145 Kemp, M.B., 11–12 Kerkhof, L.J., 159, 162 Kern, J., 229, 240 Ketley, J.M., 77–78, 81, 95, 98 Khan, N.H., 239 Khan, S., 239 Kharazmi, A., 6, 48 Khouri, H., 125 Kibsey, P.C., 4 Kiene, R.P., 124, 168 Kiewitz, C., 3 Kiley, P.J., 39 Kim, A.M., 6 Kim, C.H., 18 Kim, D.-H., 118 Kim, E.J., 48 Kim, H.-K., 77 Kim, H.Y., 49 Kim, I.-H., 77–78 Kim, K., 31, 78 Kim, K.Y., 18 Kim, U.-J., 165–166 Kim, Y.C., 18

AUTHOR INDEX

Kim, Y.H., 18 Kimura, T., 238, 240, 252 Kindingstad, L., 96 King, G.M., 124, 168 King, J., 236 King, K.Y., 76 King, T.E., 136 Kippert, F., 241 Kishida, Y., 238, 240, 252 Kiss, J., 19 Kita, K., 8 Kitayama, Y., 245–246, 248, 251, 253, 256, 258, 260–266, 278 Kiyohara, R., 248, 251 Kiyokawa, C., 238, 252 Klein, U., 281 Kleppe, A.K., 235 Klewer, D.A., 256, 261–262, 265 Kley-Raymann, M., 7, 40 Klimmek, O., 117, 183 Klinman, J.P., 18 Klipp, W., 35 Ka¨llebring, B., 138, 140 Klopotowski, T., 13 Kloppstech, K., 277 Kluyver, A.J., 195 Knapp, S., 143 Knoll, A.H., 235–236 Knowles, C.J., 43, 45 Knowles, M.R., 5, 42 Knowles, R., 115, 168 Knudsen, K., 8 Ko, I.J., 27 Ko, K., 277 Kobayashi, M., 240 Koch, C., 5–6 Koch, H.G., 26, 171 Kodama, T., 36–39, 157 Koefoed, S., 155, 191–192 Koenig, F., 240 Kogan, Y., 117 Koh, M., 91 Kohara, M., 238, 240, 252 Kohler, T., 4

AUTHOR INDEX

Kolber, Z., 242 Kolbesen, B.O., 138 Kolodziej, A.F., 128 Kolonay, J.F., 125 Kolter, R., 3 Komp, C., 168 Kondo, F., 239 Kondo, T., 231, 233, 240–246, 248–258, 260–266, 270, 272–276, 278 Konishi, K., 8 Kontchou, C.Y., 95 Koo, B.S., 18 Koonin, E.V., 257, 271 Koresh, Y., 113 Kornberg, A., 49 Ko¨rner, H., 96, 115, 117–118, 127, 129–133, 135, 139–141, 144–147, 150–151, 156, 168, 171, 179–180, 191–192, 195 Korolik, V., 26 Korzhenevskaia, T.G., 239 Kosako, Y., 126, 161 Kotani, H., 252 Koutny´, M., 181 Kovalinka, J.A., 113 Kowalik, D.J., 3, 33 Koyo, A.O., 239 Kranz, R., 22, 183 Krause, M., 238 Krauss, N., 229, 240 Kretzschmar, U., 15–16 Krieg, D.P., 48 Krieger-Brockett, B.B., 125 Krippahl, L., 85, 92 Krishnapillai, V., 36 Kristjansson, 96, 131, 150 Kroeze, C., 110 Kroneck, P.M., 37 Kroneck, P.M.H., 107, 113, 119, 128–133, 135–148, 150–152, 159, 167–168, 171–173, 175, 177, 179–180, 182, 184, 191–192, 195–196 Kroppenstedt, R.M., 115–116 Kruip, J., 240, 277

313

Krukonis, E.S., 155 Krulwich, T.A., 8 Krumbein, W.E., 236 Krumbien, W.E., 241 Kube, M., 115, 191 Kubicek, K., 177 Kucˇera, I., 180–181 Kuenen, J.G., 115, 119, 195 Kuiper-Goodman, T., 239 Kulkarni, R.D., 233, 241, 276 Kumazawa, S., 241 Kumita, H., 38 Kundu, B., 180 Kunisawa, R., 159 Kunz, D.A., 49–50 Kuo, S., 49 Kupper, H., 242 Kuritz, T., 240 Kurun, E., 113 Kusai, K., 16, 18, 37, 183 Kuschel, T., 114, 151 Kushner, D.J., 125 Kutsuna, S., 241–245, 249–253, 264, 266, 274, 276 Kwon, M., 31, 78 Kyrpides, N., 117 Lack, J.G., 123 LaCroix, L.B., 142, 144, 146 Ladner, H., 165–166 Lagrou, M., 3, 33 Laheri, A.N., 23 Lakkis, C., 4 Lam, J., 42 Lam, K., 42 Lamont, I.L., 22 Lamparter, T., 271 Lancaster, C.R., 11 Landin, J.A., 139, 148 Lang, E., 126 Lang, F., 166 Langworthy, T.A., 126 Lanoil, B.D., 238 Lanyi, B., 28

314

Lanz, C., 117, 183 Lanzilotta, W.N., 84, 86–88 Lapidus, A., 117 Lappalainen, P., 135, 138–139, 141–145 Larbig, K., 3, 33 Larsen, H., 124 Larsen, S., 23 Larson, A.D., 117 Larsson, G., 139 Larsson, J.T., 30 Larsson, S., 138, 140 Latiﬁ, A., 44 Lauf, J., 120 Lauhon, C.T., 192 Lauinger, C., 44 Lauro, F.M., 123 Lazdunski, A., 44 Le, L., 4 Lebedeva, N.V., 245, 266 LeCloux, D.D., 137 Lederer, H., 117, 183 Lee, A., 240, 276 Lee, C., 248, 251 Lee, H.S., 175, 177 Lee, Y.P., 19 Leech, A., 177 LeGall, J., 79, 117, 167 Lehnert, N., 114, 151 Leipe, D.D., 257 Leisinger, T., 13–14 Leister, D., 236, 277 Leitao, J.H., 6, 48 Lemanceau, P., 155 Lemberg, R., 22 Lemieux, L., 144 Le´pine, F., 167 Leroy, G., 23 Lessie, T.G., 12 Lester, R.L., 34 Leung, Y.-C., 181 Levit, M., 246, 261, 271 Lewin, A., 177 Lewis, M., 124–125, 168 Li Calzi, M., 76–77

AUTHOR INDEX

Li, F., 120 Li, P.M., 141 Li, X., 83 Liang, J., 113, 116 Liddington, R.C., 18 Lidstrom, M., 83 Lidstrom, M.E., 18, 83 Liebl, U., 26 Liesack, W., 236 Lill, R., 22, 183 Lillehaug, J.R., 96 Lilly, J.W., 277 Lim, L.W., 143 Lim, R., 3, 33 Lin, E.C.C., 97 Lin, G.-H., 116 Lin, H.Y., 241 Lin, Y.-T., 116 Lindahl, P.A., 272–273 Lindblad, P., 239 Lindley, P.F., 133 Lindqvist, A., 8 Lindsay, J.F., 235 Lindum, P.W., 6, 48 Linke, B., 117, 183 Linker, A., 47 Linn, S., 75 Linnane, S.J., 42 Linne von Berg, K.-H., 196 Lins, T., 236, 277 Lipman, D.J., 158 Lippard, S.J., 137 Lipscomb, J.D., 119 Lipski, A., 116, 118 Litaker, R.W., 238 Little, R., 31 Liu, A., 83 Liu, M.C., 79 Liu, Y., 77, 235, 245, 266, 270, 274 LiWang, A.C., 246–248, 253, 256, 258, 260–265, 270 Lizewski, S.E., 49–50 Lledo´, B., 166 Lloyd, D., 7, 35, 42

AUTHOR INDEX

Lo, B.S., 281 Lobocka, M., 13 Lo¨chelt, S., 137, 148, 169 Loehr, T.M., 139, 146 Loewen, P.C., 76 Lohtander, K., 239 Loiseau, L., 180 Lombardot, T., 191 Long, S.R., 168 Loo, C.Y., 194 Lorne, J., 240, 276 Loros, J.J., 235 Lory, S., 3, 33 Los, T., 117 Losonczy, G., 28 Lovell, R.A., 239 Lovley, D.R., 125 Lowe, D.R., 239 Lowery, M.D., 142, 145 Lu, C.D., 40 Lu, S., 166 Lu, W.-P., 113 Lu, Y., 137, 139, 141–142, 144, 146–147 Luan, S., 255 Lubben, M., 26–28 Lubitz, S.P., 170 Lucas, S.K., 136 Luchinat, C., 143 Ludwig, B., 135, 138–139, 141, 143, 145 Ludwig, W., 116–117, 119, 191 Luethi, E., 40 Lu¨ke, I., 171 Lukayanova, M.A. Dstrovskii, 22 Lukow, T., 115–116 Lukoyanov, D., 147 Lum, R.T., 113 Lundgren, P., 242 Lunsdorf, H., 48 Luthi, E., 40 Lyczak, J.B., 4 Lykidis, A., 117 Lymar, S.V., 6, 42

315

Lynn, J.R., 277 Lysenko, A.M., 126, 167 Ma, E.S.F., 114 Ma, J., 24, 27–29 Machonkin, T.E., 132 Macy, J.M., 117 Madden, T.L., 158 Madupu, R., 124–125, 168 Maeda, M., 187 Magalon, A., 34 Magasanik, B., 12 Magnusson, O.T., 18 Mahnane, M.R., 118 Mahne, I., 162 Maier, R.J., 22–23, 27–28, 30, 77–78 Maitra, R., 240 Maklashina, E., 11 Malacrida, G., 123 Malatesta, F., 135 Malmstro¨m, B.G., 135, 138, 140–141, 143–144, 146 Maloy, S., 14, 244 Mandel, M., 117 Mangani, S., 177 Mangels, D., 240 Manoil, C., 4, 29, 46 Manson, I.A., 22 Marcotte, E., 164–166, 180 Margraf, H., 29, 46 Marhuenda-Egea, F.C., 166 Marie, D., 276 Marimanikkupam, S., 83 Ma´rquez, M.C., 126 Marshall, V.P., 13 Martienssen, R., 170 Martin, W., 236, 277 Martinez, A., 238 Martino-Catt, S.J., 277 Martı´ nez-Espinosa, R.M., 166 Mary, I., 276 Mason, R.P., 42, 126 Masuda, K., 174 Masuda, S., 275

316

Masui, H., 239 Masui, N., 123 Matchova´, I., 180 Mathee, K., 6, 48 Mathews, F.S., 83, 143 Matsubara, H., 36–37 Matsubara, T., 127, 130, 147, 180, 182, 195 Matsumoto, M., 238, 240, 252 Matsuno, A., 238, 240, 252 Matsuno-Yagi, A., 9 Matsuo, M., 116, 118 Matsushima, R., 239 Matsushita, K., 12, 18, 20–22, 25, 28–29 Matsuura, K., 38, 180–181, 239 Matsuura, Y., 16, 18 Matsuyama, S., 174 Matsuyama, S.-I., 174 Mattatall, N.R., 177 Matthews, R.G., 113 Matthies, C., 116, 118, 123 Matthijs, H.C., 236 Mattila, K., 130–131, 133, 139, 146, 149–150, 182, 196 Mattingly, S.J., 48–49 Maul, J.E., 277 Mayer, F., 124, 168 Mazdumar, S., 146 Mazur, M., 117 Mazurier, S., 155 McAdams, H.H., 272 McAlister-Henn, L., 78 McCarthy, J.E.G., 168–170 McCleskey, C.S., 117 McCormick, W., 155 McCormick, W.A., 172, 192 McCracken, D.R., 113 McDermott, T.R., 6, 42, 126 McDevitt, C.A., 171 McEwan, A.G., 78–79, 93, 117, 156, 168, 177, 180–181 McGarvey, J.J., 137 McGenity, T.J., 124, 164–165 McGinnity, D.F., 83, 85, 92

AUTHOR INDEX

McGrath, R.B., 277 McGuirl, M.A., 139, 148, 177 Mchaourab, H.S., 142 McIntire, W.S., 83 McKay, C.P., 238–239 McKee, V., 137 McLoughlin, P., 42 McMurry, L., 278 McNamara, P., 281 McNicholl-Kennedy, J., 98 McRee, D.E., 137 McSween, G., 167 Meeks, J.C., 239 Meganathan, R., 29 Mehrotra, P.K., 144 Mei, H., 92 Meiklejohn, J., 195 Meile, L., 13–14 Melamud, E., 125 Melis, K., 23 Melis, K.A., 22 Melville, H.W., 113 Membrillo-Hernandez, J., 8 Mendz, G.L., 26 Menzel, R., 13 Mercenier, A., 40 Merchant, S., 22, 143, 183 Mergel, A., 162 Merriman, M.E., 22 Merriman, T.R., 22 Merrow, M., 231, 235, 267 Merz, K.M., 144 Mesa, S., 155, 157 Messer, W., 238 Messerschmidt, A., 113, 143 Methe´, B.A., 125 Metz, M., 142, 144 Mevarech, M., 124, 195 Meyer, F., 117, 183 Meyer, K.C., 5–6, 42 Meyer, M.A., 239 Meyer, O., 117 Meyer, T.E., 91, 240 Meyerstein, D., 113

AUTHOR INDEX

Mian, F.A., 49 Miao, V.P., 240, 276 Michaels, R., 43 Michalski, W.P., 181 Michel, H., 131–132, 135, 139–141, 143, 145–146 Michel, K.P., 270–272, 277 Migita, C.T., 12, 18 Mikhailova, N., 117 Milekhina, E.I., 131 Milgram, A., 117 Millar, A.J., 277 Miller, D.J., 119, 181 Miller, I., 236 Miller, J.H., 165–166 Miller, M.A., 31, 92 Miller, T.R., 124, 168 Miller, W., 158, 277 Millett, F., 31, 92 Millington, P., 35 Mills, T., 239 Mimuro, M., 240 Min, H., 243, 266, 270, 273, 275 Minagawa, N., 168 Minard, K.I., 78 Minkman, D.C.J., 194 Mirleau, P., 155 Mitrakul, K., 194 Mitsui, A., 241 Mittag, M., 277 Miyagishima, S.Y., 272 Miyajima, N., 252 Miyake, A., 257–258 Miyashita, H., 236, 240, 243, 245, 266 Miyata, S., 4 Miyazaki, T., 256 Miyoshi, H., 12, 18 Mizoguchi, S.D., 3, 33 Mizrahi, V., 30 Mizuno, T., 16, 261 Mizutani, M., 39, 117, 155–156 Mochizuki, K., 166 Moczydlowski, E., 238 Moe¨nne-Loccoz, P., 166

317

Mogi, T., 28 Moir, J.W., 33, 35 Moir, J.W.B., 111, 118, 181 Molenaar, D., 16 Moletta, R., 116, 119 Molin, S., 6, 48 Molina, L., 14 Momen, B., 125 Mongkolsuk, S., 78 Montgomery, B.L., 270–272 Moog, R.S., 139, 147 Moore, G.R., 84, 177 Moore, G.W., 22–23 Moore, T., 76 Moran, M.A., 124, 168 Morden, C.W., 236 Moreno-Vivia´n, C., 16, 18, 33, 35, 166 Morgan, J., 145 Mori, H., 170, 172 Mori, T., 195, 241, 243, 251, 256–258, 263, 266, 269 Morici, L.A., 49–50 Morishima, I., 38 Morishita, M., 233, 235, 250, 252–253, 258, 263 Morrison, S.M., 44 Moser, C.C., 134 Moshiri, F., 30 Mosier, A., 110 Mossialos, D., 30, 45 Mouawad, L., 280 Moule, S., 81, 95 Moult, J., 125 Mouncey, N.J., 26 Moura, I., 38, 79, 84–85, 90–92, 130–133, 139, 147–150, 171, 196 Moura, J., 85 Moura, J.J., 38, 79, 84–85, 90–92, 130–131, 133, 139, 147–149, 171, 196 Moura, J.J.G., 85, 92, 130–133, 149–150 Moynihan, J.B., 42 Mozen, M.M., 113 Mulder, T.C., 142, 146 Mulholland, F., 98

318

Mulholland, J., 277 Mullakhanbhai, M.F., 124 Mu¨ller, A., 140 Mu¨ller, M., 171 Mulliken, R.S., 144 Mungall, K., 81, 95 Muraki, A., 240, 252 Muramatsu, M., 238 Murata, H., 239 Murayama, Y., 233, 244, 248, 263 Muriana, F.J.G., 166 Muro-Pastor, A.M., 14, 241 Murphy, M.E.P., 133 Murtas, G., 277 Musacchio, A., 144 Musser, J.M., 49 Mutsuda, M., 270–272 Muyzer, G., 238 Mwaura, F., 239 Myers, J.D., 81, 95 Myllykallio, H., 26 Myrold, D.D., 159, 162 Nagai, H., 239 Nagai, T., 277 Nagaya, M., 261 Nagy, F., 277 Nair, U., 243, 273, 275 Najdek, M., 236 Nakada, Y., 14 Nakae, T., 175 Nakagawa, S., 116 Nakahira, Y., 242–245, 266 Nakai, M., 16, 18, 37, 183 Nakajima, M., 233, 243–244, 246, 248, 251, 253, 258, 263 Nakamaru-Ogiso, E., 9 Nakamura, H., 28 Nakamura, S., 12, 18, 116 Nakamura, Y., 238, 240, 252 Nakane, H., 238 Nakano, M., 239 Nakashima, K., 277 Nakashima, R., 139, 141, 143

AUTHOR INDEX

Nakatsu, T., 246–247, 250, 252 Nakayama, N., 239 Nakayama, Y., 9, 187 Nakazaki, N., 238, 240, 252 Namba, K., 233, 250, 253, 257–258, 263 Nandakumar, R., 117, 183 Nandi, R., 238 Nar, H., 143 Naruo, K., 252 Naruse, M., 281 Naruse, Y., 281 Nast, C.C., 6, 48 Nathan, C., 76–77 Na¨ther, C., 114, 151 Navarrete, A., 239 Nayar, S., 241 Nazina, T.N., 126, 167 Nedwell, D.B., 159 Neese, F., 113, 130–132, 135, 138–151, 191–192 Neher, I., 277 Neidhardt, F.C., 7 Neilan, B.A., 239 Nelson, J., 131–132, 135, 137, 139–140, 145–146 Nelson, K.E., 125 Nelson, W.C., 124–125, 168 Nevison, C., 110 Nevo, E., 231, 233, 240, 257, 277 Nevo, R., 238–240 Newell, D.G., 98 Newton, A., 272 Ng, T.C., 23 Ng, W.V., 164–166, 180 Nicholas, D.J.D., 119, 180–181 Nicoletti, F., 135 Niedzielski, J.J., 190 Nield, J., 277 Nienow, J.A., 239 Niggemyer, A., 167 Nilavongse, A., 187 Nilges, M.J., 147 Nishijyo, T., 14 Nishino, T., 238

AUTHOR INDEX

Nishiwaki, T., 233, 244, 246, 248, 250–251, 253, 256–258, 260, 263, 278 Nitschke, W., 23 Nittis, T., 177 Nivie`re, V., 169 Noah, T.L., 42, 126 Nobles, D.R., 236 Nogales, B., 159 Nogi, Y., 123 Nolting, H.-F., 140 Norling, B., 240 Norman, D.C., 48 Norman, R.A., 247, 280 Norman, T.R., 114 Novikova, E.V., 126, 167 Nubel, U., 238 Nunn, D.N., 16 Nutley, M., 85, 90–92 Obata, S., 238 O’Brian, M.R., 22 O’Brien, T., 48 Ocampo-Friedmann, R., 239 Ochsner, U.A., 6, 12, 42, 126 O’Connor, C.M., 42 O’Connor, S.M., 123 O’Donnell, S.E., 22 Oenema, O., 110, 120 Oganesyan, V.S., 130, 149–151 O’Gara, J.P., 27 Ogawa, K., 239 Oh, J.I., 26–27, 30 Oh, T.-K., 126 Ohad, I., 238–239 O’Hara, B.P., 247, 280 Oh-hashi, K., 281 Ohman, D.E., 4, 6, 48 Ohno, T., 235–236 Ohta, N., 272 Ohta, T., 277 Ohtani, I., 239 Ohto, C., 238 Okada, K., 236 Okamoto, K., 233, 235, 252

319

Oksanen, I., 239 Okumura, S., 252 Okunuki, K., 16, 18, 36–37, 183 Olczak, A.A., 77–78 Olesen, P.H., 137 Ollivier, B., 116 Olson, J.B., 238 Olson, J.W., 77–78 Olson, M.V., 3, 33 Olsson, M.H.M., 142, 144 Oltmann, L.F., 181 Onai, K., 233, 235, 250, 252–253, 258, 263 Oppenheimer, C.H., 123 Oren, A., 124, 126 Orme-Johnson, W.H., 128 Orth, P., 229, 240 Osborn, A.M., 159 Osipov, G.A., 126, 167 Ostensvik, O., 238 Osterlund, K., 31 Ostermeier, C., 139, 141, 143 Osteryoung, K.W., 272 Ostrom, N.E., 120 Ostrom, P.H., 120 O’Toole, G., 3 O’Toole, R., 45 Otte, S., 115, 119 Otten, M.F., 26 Oubrie, A., 37–38 Ouyang, Y., 231, 242 Owens, M.W., 42 Oyama, T., 233, 243–245, 248, 253–254, 263, 266 Paces, J., 117 Paces, V., 117 Packer, B.M., 235 Paerl, H.W., 236, 238 Page, C.C., 134 Page, M.D., 22, 180–181 Page, R.D.M., 80, 158 Pai, E.F., 246–247, 253, 263, 265 Pai, G., 124, 168

320

Painter, M., 240, 276 Pakrasi, H.B., 240 Palenik, B., 240 Palinska, K.A., 236 Pallen, M.J., 81, 95 Palleroni, N.J., 7, 13, 159 Palm, C., 168 Palma, N., 92 Palmer, G., 144 Palmer, J.D., 277 Palmer, T., 169–172 Pamplin, C.B., 114 Pan, J.G., 19 Pan, M., 164–166, 180 Panda, S., 281 Paranchych, W., 4 Pardon, P., 274 Park, P., 246, 261, 271 Park, S.F., 78 Park, S.M., 40 Park, Y.-H., 126 Parker, B.C., 238 Parkhill, J., 81, 95 Parsek, M.R., 42, 126 Parsonage, D., 77, 180 Partensky, F., 276 Parvatiyar, K., 6, 42, 126 Pascher, T., 144 Passador, L., 35, 42 Patel, B.K.C., 116 Pattanayek, R., 251, 256–258 Pattanayek, S., 251, 258 Patureau, D., 116, 119 Paul, P.P., 149 Paulat, F., 114, 151 Pauleta, S.R., 85, 90–92 Paulsen, I.T., 3, 33, 124, 168 Paulsrud, P., 239 Payne, W.J., 79, 115, 117, 131, 153, 167, 195 Pearl, L.H., 247, 280 Pearson, A.R., 83 Pearson, B.M., 98 Pearson, I.V., 180–181

AUTHOR INDEX

Pecenkova, T., 117 Pecht, I., 139, 141, 146 Peck, M.C., 168 Pedersen, J.Z., 137 Pedersen, S.S., 6 Peisach, J., 142 Peltz, S.W., 257 Penn, C.W., 77–78, 81, 95 Penner-Hahn, J.E., 142 Penny, D., 236, 277 Perahia, D., 280 Pereira, A.S., 38, 84–85, 90–91, 130–131, 133, 139, 147–150, 171, 196 Pereira, M.M., 24 Perfect, J.R., 31, 97 Perkins, G.H., 93 Pessi, G., 43–44 Peters, J.C., 137 Peterson, J., 31, 136 Petras, G., 28 Petratos, K., 143 Petrunyaka, V.V., 126, 167 Pettigrew, G., 31 Pettigrew, G.W., 22–24, 79, 83–87, 90–92, 94 Pfeifer, F., 238 Pfenninger, S., 142–143 Pfenninger-Li, X.D., 9 Pham, X.Q., 3, 33 Phibbs, P.V., 12, 16–18 Philipp, B., 116, 119 Philippot, L., 155, 161 Piantadosi, C.A., 136 Pibernat, I., 238 Pichinoty, F., 116–117 Pier, G.B., 4 Pierard, A., 7, 40 Pietrzak, S., 120 Pin, C., 98 Pina, S.E., 48–49 Pinckney, J.L., 238 Pinto, M., 120 Pioli, D., 13 Piotrowski, M., 277

AUTHOR INDEX

Pistorius, E.K., 277 Pitcher, R.S., 25 Pitt, A.J., 120 Pitt, M., 28–29, 45 Pittendrigh, C.S., 231, 234, 246 Pizarro, R.A., 46 Plewniak, F., 79–80 Po, C., 12 Pohlschro¨der, M., 165 Poltaraus, A.B., 126, 167 Pommier, J., 34 Pond, J.L., 126 Ponting, C.P., 271 Poole, K., 3 Poole, L.B., 76–77 Poole, R.K., 8, 11–12, 20, 24, 30, 34, 115 Porcelli, I., 171 Postma, P.W., 18 Potenza, J.A., 143 Potter, L., 33, 35 Potter, L.C., 35 Potts, M., 236, 238 Poulos, T.L., 84, 86–88 Powell, D., 137 Praneeth, V.K.K., 114, 151 Pratt, J.M., 112–113 Prazeres, S., 79, 84–85, 92 Preisig, O., 25–26 Prichard, R.K., 98 Priscu, J.C., 236, 238 Prudeˆncio, M., 38, 130–131, 133, 139, 147–150, 171, 196 Pu, L.S., 113 Pullan, L., 171 Puntervoll, P., 96 Py, B., 180 Quail, M.A., 81, 95 Rabus, R., 115, 191 Race, H.L., 277 Rachel, R., 125, 164–165 Raddatz, G., 117, 183

321

Radford, S.E., 181 Radic, J., 236 Radic, T., 236 Radosevich, M., 115 Radzioch, D., 4 Ragsdale, S.W., 113 Rahe, E., 124, 168 Rahme, L.G., 3–4 Rainey, F.A., 115–116, 126 Raitio, M., 28 Rajandream, M.-A., 81, 95 Ramirez, B.E., 140, 146 Ramos, C., 14 Ramos, J.L., 14 Randall, D.W., 142, 146 Randall, G.W., 142, 144 Randell, S., 5–6, 42 Rasko, D.A., 124, 168 Rasmussen, T., 129–132, 147–152, 181, 191, 196 Ratjen, F., 4–5 Raudonikiene, A., 76 Raven, J.A., 236, 239, 277 Ray, A., 21, 31 Read, R.C., 118 Rebrekov, D., 117 Rech, S., 117 Redinbo, M.R., 143 Reed, J.C., 18 Regan, J.J., 146 Rehman, J., 277 Reich, Z., 238–240 Reichardt, J., 136 Reichmann, P., 14 Reid, E., 79, 93–94, 96 Reijnders, W.N., 26, 83, 96 Reijnders, W.N.M., 155–157, 172, 191–192 Reimmann, C., 39, 44, 97 Reinhardt, J., 272 Reinhardt, R., 115, 191 Reith, M.E., 277 Reitzer, L.J., 12 Reizer, J., 3, 33

322

Relimpio, A.M., 166 Relman, D.A., 239 Ren, Q., 124, 168 Renner, E.D., 117 Ressler, C., 44 Rettig, S.J., 114 Revzin, A., 27 Rey, L., 22–23 Reyes, F., 35 Reynolds, A.M., 149 Reynolds, C.M., 76–77 Reznik, G., 117 Rheingold, A.L., 113, 149 Rhiel, E., 236 Rhodes, M., 195 Rice, D., 143 Rich, J.J., 159, 162 Richards, J.H., 137–138, 143–144 Richardson, D., 95 Richardson, D.J., 32–33, 35, 81, 111, 156, 180–181, 187 Richaud, P., 156, 168 Richly, E., 236, 277 Richter, S., 238 Ridout, C.J., 79, 84–88, 92 Riester, J., 129–131, 133, 136, 138, 141, 144, 146, 148, 150–151, 172, 177, 180, 196 Righelato, R.C., 49 Rikkinen, J., 239 Ripperger, J., 281 Ripperger, J.A., 281 Rippka, R., 238, 240 Robb, M., 238 Robbins, R.A., 42 Roberts, M.A., 190 Robertson, L.A., 115, 195 Robin Harris, J., 77 Robinson, C., 170 Robinson, H., 137, 144 Robinson, V., 261–262, 265, 271 Rocap, G., 236

AUTHOR INDEX

Rocha, E.R., 95, 97 Roche, P., 280 Rock, J.D., 118 Rodriguez-Valera, F., 124, 126–127, 165 Roe, F., 6 Roe, S.M., 247, 280 Roenneberg, T., 231, 267, 277 Rogner, M., 240 Rogstam, A., 30 Rojas, A., 18 Rolda´n, M.D., 16, 18, 35, 166 Romanovicz, D.K., 236 Romao, M.J., 84–85 Romualdi, C., 123 Ronnberg, M., 31, 86, 90 Roos, E.E., 113 Rosbash, M., 281 Ro¨sch, C., 162 Rose, T., 238 Rosello´-Mora, R., 124 Rosenthal, A., 277 Rosenzweig, A.C., 132, 139, 148 Rosenzweig, R.F., 167 Rosinus, A., 117, 183 Rosovitz, M.J., 124, 168 Rossi, J.V., 190 Rossi, P., 19 Roth, J., 13 Rothrock, M.J., 239 Rowe, J.J., 6, 42, 126 Rubin, H., 30 Rubin, R.H., 278 Ruckert, A., 15–16 Rudic, R.D., 281 Ruiz, E., 149 Rujan, T., 236, 277 Rumbley, J., 24, 27–29, 144 Rupert, P.B., 113 Rupp, M., 14 Rusnak, F., 130–131, 133, 150 Rutherford, K.M., 81, 95 Ryall, B., 1 Ryde, U., 142, 144

AUTHOR INDEX

Sabaty, M., 156, 168 Sabra, W., 48 Sacchettini, J.C., 246–247, 263, 270 Sa-Correia, I., 6, 48 Saeki, M., 18 Saenger, W., 229, 240 Safari, N., 114 Sai, J., 277 Saibil, H.R., 171 Saier, M.H., 3, 33 Saigo, T., 127 Saiki, K., 28 Saint-Joanis, B., 76–77 Sakaguchi, Y., 119 Sakai, M., 18 Sakamoto, T., 239 Sako, Y., 116 Sakuragi, Y., 240 Sakurai, N., 37 Sakurai, T., 37, 166 Salgado, J., 143 Salinas, M.B., 116 Salvador, M.L., 281 Samama, J.P., 280 Sambongi, Y., 22 Samyn, B., 31, 130–131, 133, 150 Sander, O., 114, 151 Sanders, D., 137–138, 140, 143–144 Sanders-Loehr, J., 129, 131–132, 139, 146–148, 151, 196 Sands, R.H., 136, 141 Sanford, R.A., 120 Sankar, G., 113 Sano, M., 130 Sano, T., 238 Santana, M., 24 Santini, C.L., 34, 171 Saraste, M., 26, 28, 37–38, 130–133, 135, 138–139, 141–146, 149–150, 163, 195–196 Sargent, F., 169–170 Sarma, G.N., 77 Sasagawa, M., 183 Sasamoto, S., 238, 240, 252

323

Sato, E., 42 Sato, S., 238, 252 Satoh, K., 238 Satoh, S., 240 Satoh, T., 168, 180–181 Satomi, Y., 248, 251 Sauer-Eriksson, E., 138, 141, 143 Saunders, N., 83, 85, 156 Saunders, N.F.W., 155, 172, 191–192 Saveliev, S.V., 257, 263, 266 Savvides, S.N., 31, 85 Sawers, G., 30, 35 Sawers, R.G., 39 Sawyer, L., 22–23 Sayavedra-Soto, L.A., 119 Sazanov, L.A., 19 Scala, D.J., 159, 162 Schaechter, M., 7 Scha¨ffer, A.A., 158 Schalk, J., 119 Scharf, B., 124 Scheffer, J., 240, 276 Scherz, A., 240 Schibler, U., 281 Schiek, U.M., 171, 175, 177, 192 Schink, B., 116, 119 Schlarb-Ridley, B.G., 91 Schlesner, H., 191 Schlictman, D., 48–49 Schluter, P.J., 239 Schmidt, I., 119 Schmitt, R., 124 Schmitz, F.J., 4 Schmitz, O., 245, 270 Schneegurt, M.A., 241 Schneider, B., 137, 148, 169 Schobert, M., 7, 11, 16, 41 Scholes, C.P., 136, 147, 156 Scholten, E., 115–116 Scholten, J.J.F., 113 Schopf, J.W., 235 Schramm, A., 116, 118, 123 Schreiber, K., 7, 11, 41

324

Schroder, E., 77 Schro¨der, I., 11, 137, 166 Schrover, J.M., 16 Schugar, H.J., 142–143 Schuller, D.J., 84, 86–88 Schumacher, W., 113 Schurr, J.R., 49–50 Schurr, M.J., 49–50 Schuster, S.C., 117, 183 Schwab, G.M., 113 Schwab, U., 5–6, 42, 126 Schwartz, E., 168 Schwartz, W.J., 281 Schwarzenbacher, R., 18 Schweiger, H.G., 278 Schweiger, M., 278 Schweizer, H.P., 3, 12 Schwintner, C., 156, 168 Sciochetti, S.A., 272 Scolnik, P.A., 277 Scott, R.A., 38, 139, 148, 177 Seaburg, K.G., 238 Sears, H.J., 35 Seaver, L.C., 75–76, 96 Seib, K.L., 78, 93 Seitzinger, S., 110 Sekimizu, K., 4 Selander, R.K., 49 Selengut, J., 124, 168 Sellers, R.M., 113 Sengupta, S., 238 Seo, B.B., 9 Seo, S.B., 281 Settles, A.M., 170, 172 Seyler, R.W., 78 Shadle, S.E., 142 Shankar, S., 48–49 Shankowsky, H.A., 4 Shannon, P., 164–166, 180 Shapiro, L., 272 Shapleigh, J., 27, 153 Shapleigh, J.P., 156 Sharp, M., 275 Shaw, G.M., 170

AUTHOR INDEX

Shaw, G.R., 239 Shaw, J.G., 118 Shaw, R.W., 136 Shearer, N., 156 Sheath, R.G., 238 Sheldon, W.M., 124, 168 Shen, J.R., 257–258 Shepard, W., 76–77 Sherman, D.H., 190 Sherman, D.M., 241 Sherman, L.A., 241 Sherr, B., 115, 131 Sherris, J.C., 40 Shi, L., 236 Shi, Y.-C., 116 Shiba, T., 116–117 Shibata, H., 246–247, 250, 252 Shidara, S., 195 Shigeoka, S., 256 Shim, Y.K., 240 Shimada, K., 168, 180 Shimizu, H., 84, 86–88 Shimpo, S., 238, 240, 252 Shin, W., 77 Shinagawa, E., 18, 20–22, 25, 28–29 Shinzawa-Itoh, K., 139, 141, 143 Shioi, Y., 117 Shirai, M., 239 Shirai, N., 116 Shiro, Y., 38 Shoesmith, J.H., 40 Shoun, H., 115–116, 118–119, 195 Shuman, H., 178 Siblot, S., 155 Siddiqui, R.A., 37–38, 168, 171 Siefert, J.L., 236 Sili, C., 236 Silo-Suh, L., 4 Silvestrini, M.C., 36 Simionati, B., 123 Simmons, G.M., 238 Simon, J., 11, 81, 117, 128, 152, 159, 167, 172–173, 182–184 Simon, J.P., 40

AUTHOR INDEX

Simon, M.I., 165–166 Simonato, F., 123 Simpson, J.A., 6 Simpson, K., 49–50 Simpson, K.L., 42 Sitachitta, N., 190 Sivaraja, M., 86, 88 Skirrow, M.B., 81 Skovran, E., 192 Skulberg, O.M., 238 Slotboom, D.J., 83 Slutter, C.E., 138, 143–144 Slutter, C.S., 139, 141 Sly, L.I., 117 Smidt, H., 191 Smith, B.E., 130, 135 Smith, C.J., 95, 97 Smith, E.G., 30 Smith, G.B., 196 Smith, H., 79, 93–94, 96 Smith, J.M., 98 Smith, K., 3, 33 Smith, K.M., 240 Smith, M., 86, 88 Smith, M.A., 26 Smith, P.H., 137 Smith, R.M., 245, 260–262, 264–267, 269, 275 Smith, S.E., 6 Smodlaka, N., 236 Smolen, P., 281 Smyth, E.M., 281 Snow, M., 114 Snozzi, M., 115 Snyder, S.W., 38, 117, 127 Soballe, B., 20 Soﬁa, H.J., 156 Soininen, R., 31, 79 Sojonen, H., 31 Sokatch, J.R., 13 Sokol, P.A., 4 Soldati, L., 13–14 Solomon, E., 150

325

Solomon, E.I., 130, 132, 142, 144–147, 149–151 Sommer, R.D., 149 SooHoo, C.K., 38–39, 117 Soriano, A., 143 Souchon, H., 76–77 Soulimane, T., 140 Sowa, N.A., 12 Sowa, S., 113 Sparling, P., 76 Sparrow, G.J., 114 Speert, D.P., 4, 6, 47–49 Spencer, B., 137 Spencer, D., 3, 33 Spiro, S., 35, 97, 156 Spring, S., 116, 167 Springer, N., 116, 119 Spro¨er, C., 126 Stach, P., 113 Stackebrandt, E., 116–118, 126, 167 Stafford, W.F., 257, 263, 266 Stal, L.J., 241 Stalon, V., 40 Stamper, D.M., 115 Stanier, R.Y., 7, 13, 238 Stanley, N.R., 170 Stan-Lotter, H., 124 Steele, A., 235 Steffens, G.C.M., 138, 140–141, 145 Stein, L.Y., 110, 120 Steiner, B.M., 93 Steitz, T.A., 257 Stephens, C., 272 Stephens, P.J., 139, 148 Stephenson, J.T., 13–14 Steppe, T.F., 238 Stern, D.B., 277 Sternberg, C., 6, 48 Sternberg, J.A., 4 Stetter, K.O., 125, 164–166 Steuber, J., 9 Stevenson, M.M., 4 Stewart, I., 239 Stewart, P.S., 6

326

Stewart, V., 244 Stieritz, D.D., 4, 12 Stingl, K., 16 Stock, A., 261–262, 265, 271 Stock, J.B., 246, 261, 265, 271 Stoebe, B., 236, 277 Stone, F.S., 113 Stork, D.M., 26 Story, R.M., 257 Storz, G., 75–76 Stotland, P.K., 4 Stout, C.D., 22–23 Stouthamer, A.H., 24, 26, 29, 96, 157, 163, 168, 180–181, 195 Stover, C.K., 3, 33 Strampraad, M.J.F., 137, 166 Straume, D., 96 Strayer, C.A., 233, 241, 276 Stres, B., 162 Stura, E.A., 137 Sturr, M.G., 8 Stutts, M.J., 5, 42 Sugimoto, M., 238, 240, 252 Sugimura, Y., 143 Sugita, M., 277 Sugiura, M., 252, 277 Sugiyama, J., 118 Suh, S.J., 4 Sullivan, S.A., 124–125, 168 Summer, E.J., 172 Sun, J., 260 Sundaram, U.M., 132 Sundaramoorthy, M., 84, 86–88 Sundin, G.W., 49 Surette, M.G., 246, 261, 271 Surpin, M.A., 27–28 Surzycki, R., 168 Sutka, R.L., 120 Suzuki, H., 195, 257–258 Suzuki, J.Y., 236 Suzuki, M., 239 Suzuki, Y., 277 Svensson-Ek, M., 139 Sweeney, B.M., 241, 276, 278

AUTHOR INDEX

Swem, D.L., 26 Swem, L.R., 26 Swenson, R.P., 19 Swett, K., 236 Swift, S., 3 Swings, J., 126 Sylvia, A.L., 136 Szilagyi, R.K., 142, 144 Taaffe, L.R., 119 Tabata, S., 238, 240, 252 Taghert, P.H., 281 Tait, A.M., 113 Takahashi, A., 241 Takahashi, H., 275 Takahashi, S., 38 Takai, A., 239 Takai, K., 116 Takani, Y., 239 Takano, T., 16, 18 Takao, T., 248, 251 Takaya, N., 115–116, 118–119 Takazawa, M., 240 Takeuchi, C., 238, 252 Talbo, G., 145 Tamaru, Y., 239 Tamegai, H., 123 Tamoi, M., 256 Tan, M.W., 4 Tanabe, A., 241–243, 245, 249–253, 264, 276 Tanaka, A., 236, 240, 252 Tanaka, K., 19, 275 Tanaka, M., 18, 281 Tanaka, Y., 166 Tandeau de Marsac, N., 236, 238 Taniguchi, Y., 249, 253, 257, 260 Tanimoto, T., 195 Taormino, J.P., 30 Tarran, R., 5–6, 42 Taube, H., 114, 195 Taubner, L.M., 177 Tavankar, G.R., 8, 25, 30–31, 45 Tavares, P., 85, 130–131, 133, 150

AUTHOR INDEX

Taylor, W., 233, 241, 276 Teakle, G.R., 277 Teeling, H., 191 Tegoni, M., 38, 130–133, 139, 146–150, 171, 196 Teixeira, M., 24 Temamoto, M., 248, 251 Teraguchi, S., 128 Terai, H., 127 Terao, K., 239 Terry, J.M., 48–49 Tesarı´ k, R., 181 Thain, S.C., 277 Thibaud, M.C., 113 Thomann, H., 139 Thomas, C.M., 19 Thomas, G.H., 35, 170 Thomas, J.M., 113 Thomas, J.W., 24 Thomas, M.A., 239 Thomas, P., 116 Thomas, S., 14 Thomason, P.A., 265 Thompson, A., 31 Thompson, J., 79–80 Thompson, J.D., 158 Thomson, A.J., 31, 34, 37, 88, 129–132, 136–139, 141–143, 147–152, 181, 191, 196 Thony-Meyer, L., 22, 25–26, 28 Tiedje, J.M., 36, 120, 162, 196 Tiley, P.F., 113 Timmis, K.N., 159 Timoteo, C.G., 85 Tindall, B.J., 126, 164 Ting, C.S., 236 Tiwari, N.P., 18 Toci, R., 34 Toftlund, H., 137 Tokuda, H., 174 Tolentino, E., 3, 33 Tolman, W.B., 137, 140, 142–144, 146, 149 Tomita, J., 243–244

327

Tomitani, A., 235–236 Tomizaki, T., 139, 141, 143 Tomlinson, G.A., 126, 164–165 Tompkins, R.G., 4 Torreblanca, M., 126–127, 165 Toth, L., 28 Tourova, T.P., 126, 167 Toyama, H., 12, 18 Traina, S.J., 115 Tran, N.P., 274 Trapnell, B., 6, 42 Tredget, E.E., 4 Trincao, J., 84–85 Trincone, A., 125, 164–165 To¨rnroth, S., 139 Trogler, W.C., 112 Trumpower, B., 20–21 Trumpower, B.L., 20–21 Trunk, K., 7, 11, 41 Tsatsos, P., 9, 187 Tsay, S.-S., 116 Tseng, H.J., 78, 93 Tsinoremas, N.F., 241–243, 245, 266, 275–276 Tsuchiya, T., 240 Tsuchiya, Y., 252, 258, 263 Tsudzuki, J., 277 Tsudzuki, T., 277 Tsukihara, T., 139, 141, 143 Tsur, T., 238–239 Tu, J., 6, 48, 241, 276 Tuan, D.F.-T., 114 Tufﬂi, C.F., 278 Tullman, D., 172 Tummler, B., 3 Tung, G., 195 Tuovinen, O.H., 115 Tupy, J., 275 Tura´nek, J., 181 Turk, T., 37 Turner, R., 31 Turner, R.J., 170 Turner, S., 79, 93–94, 96 Tyekla´r, Z., 149

328

Ubbink, M., 91 Uchiyama, H., 118 Ucurum, Z., 157 Ufer, M., 166 Ulbrich, P., 117 Ulrich, M., 5–6, 42 Ulstrup, J., 23 Underdal, B., 238 Unemoto, T., 9, 187 Urata, K., 168, 180 Urbani, A., 37–38 Urmeneta, J., 239 Utamapongchai, S., 78 Utterback, T.R., 125 Uzumaki, T., 233, 246–247, 250, 252–253, 257–258, 263 Vakonakis, I., 246–248, 253, 256, 258, 260–265 Vakoufari, E., 143 Valdameri, G., 19 Valentin, K., 277 Valentine, W.M., 239 Vallaeys, T., 274 Valle, G., 123 Van Beeumen, J., 31, 79, 84, 90–92, 130–131, 133, 150 Van Beeumen, J.J., 83–87, 90–91 van Belzen, R., 9 van Cleemput, O., 110 Van Craenenbroeck, K., 31 van de Kamp, M., 143 Van Delden, C., 3–4 van der Meer, J.R., 115 van der Oost, J., 24, 26, 29, 96, 137–141, 144–145, 157, 163, 191, 195 van der Palen, C.J., 83 van der Rest, M.E., 16 Van der Wauven, C., 7, 40 Van Driessche, G., 83, 85 Van Gelder, R.N., 281 Van Horen, E., 31, 85 Van Kranendonk, M.J., 235 van Leest, M., 191

AUTHOR INDEX

Va¨nngard, T., 137, 142 van Pouderoyen, G., 146 van Spanning, R.J., 24, 26, 29, 83, 96 van Spanning, R.J.M., 119, 130, 147, 150–151, 155–157, 163, 172, 180–181, 191–192, 195 van Verseveld, H.W., 168, 180 van Vliet, A.H.M., 77–78, 81, 95 van Wielink, J.E., 16 Vancanneyt, M., 126 Vandamme, P., 126 Vanden Driessche, T., 278 Vandenberghe, I., 31, 79, 84, 91 Vander Wauven, C., 40 Vanngard, T., 86 vanSpanning, R.J.M., 119 Vasieva, O., 117 Vasil, M.L., 12 Vattanaviboon, P., 78 Vaughan, G.A., 113 Vaulot, D., 276 Vega, J.M., 128 Velasco, L., 155, 157 Velge, P., 274 Venables, W.A., 13–14 Ventosa, A., 124, 126–127, 164–165 Vergnes, A., 34 Verhoef, J., 4 Verkhovskaya, M.L., 9, 187 Verkhovsky, M., 145 Vermeglio, A., 156, 168 Vernon, D.I., 240 Vernon, M., 117 Veselov, A., 156 Vezzi, A., 123 Viebrock, A., 163, 195 Viebrock-Sambale, A., 39, 154 Vijgenboom, E., 36–37, 137, 144, 182 Vila, A.J., 137 Vilbois, F., 281 Vilchez, S., 14 Villalain, J., 79 Villemur, R., 167 Villeval, D., 40

AUTHOR INDEX

Vincenzini, M., 236 Vinogradova, O., 231, 233, 240, 257, 277 Vinyard, W.C., 238 Viollier, P.H., 272 Vis, M.L., 238 Visca, P., 45 Vitello, L.B., 78 Vitulo, N., 123 Vivanco, J.M., 3 Vlcek, C., 117 Voisard, C., 46 Volcani, B.E., 126 Vo¨lkl, P., 125, 164–165 Vollack, K.U., 39, 96, 154, 156, 168 Volpel, K., 4 von Wachenfeldt, C., 30, 138, 141 Voordouw, G., 169 Voulhoux, R., 172 Wada, T., 252 Wagner, M., 116 Wagner, V.E., 35, 42 Wahl, G., 119 Wainwright, L.M., 115 Wakasugi, T., 277 Walker, D., 177 Walker, T.S., 3 Wallace-Williams, S.E., 144 Waller, D., 113 Walsby, A.E., 238 Walton, J.P., 78 Wan, P.T., 247, 280 Wan, X-Y., 77 Wang, A.H.-J., 137, 144 Wang, B.C., 22 Wang, G., 77–78, 239 Wang, H., 142, 144 Wang, H-L., 77 Wang, J., 256–257 Wang, T., 35 Wang, X., 137 Wang, Y., 83, 156 Wanner, G., 124

329

Wansell, C., 181 Ward, N., 124, 168 Warmerdam, G., 143 Warne, A., 144 Warrener, P., 3, 33 Wastell, S., 156 Watanabe, A., 238, 240, 252 Watanabe, H., 4 Watanabe, M., 239 Watanabe, M.F., 239 Waterbury, J.B., 238, 240 Watmough, J., 79 Watmough, N.J., 19, 25, 135, 152, 181 Watson, R.J., 155, 172, 192 Weathers, P.J., 190 Weaver, B., 124–125, 168 Webb, P.M., 239 Weber, I.T., 257 Wei, Y.-H., 136 Weidenhaupt, M., 19 Weidman, J.F., 125 Weiner, J.H., 170 Weinstein, E.A., 30 Weiss, T., 5–6, 42 Weissgra¨ber, S., 140 Welker, M., 239 Wells, D.H., 168 Wells, J.M., 98 Weng, R.S., 164–166, 180 Weng, Y., 257 Werber, M.M., 124, 195 Werner, E., 6 Westbrock-Wadman, S., 3, 33 Westerhoff, H.V., 26, 96, 119, 155–157, 172, 191–192 Westgate, E.J., 281 Wetzstein, H.G., 117, 168 Wexler, M., 170 Wharton, D.C., 136, 141 Wharton, R.A., 238 Wheatcroft, R., 168 Whitehead, S., 81, 95 Whitman, W.B., 124, 168 Whitton, B.A., 236

330

Wiberg, K.B., 144 Widdel, F., 115 Wieland, H., 171, 175, 177, 192 Wikstro¨m, M., 145 Wild, J., 13 Wilderman, P.J., 12 Willett, G., 144 Williams, F., 44 Williams, H.D., 1, 8, 21, 25, 28–31, 43–45 Williams, K.R., 146 Williams, P., 3, 44 Williams, P.A., 137, 181 Williams, R., 19 Williams, S.B., 229, 240, 242–248, 253, 256, 258, 260–262, 264–267, 269–270, 275 Wilmanns, M., 138, 141, 143 Wilmot, C.M., 83 Wilson, I.B., 79 Wilson, K.S., 143 Winge, D.R., 177 Winkler, J.R., 140, 146 Winteler, H., 40, 44 Wissing, F., 43 Withers, H., 3 Witt, H., 135 Witt, H.T., 229, 240 Wittershagen, A., 138 Wittung, P., 138, 140, 144 Woelﬂi, W., 239 Wofsy, S.C., 110 Wolanin, P.M., 265 Wolk, C.P., 240, 272 Wolosiuk, R.A., 255 Wong, F.C., 239 Wong, G.K., 3, 33 Wong, S.-L., 169 Wong, S.M., 4 Wood, A.P., 126 Wood, J.M., 13–14 Wood, P.M., 37 Wood, T., 177 Wood, Z.A., 76–77

AUTHOR INDEX

Woodruff, W.H., 140, 144 Woolum, J.C., 278 Worlitzsch, D., 5–6, 42 Wouters, J., 239 Wozniak, D.J., 6, 42, 126 Wrage, N., 120 Wren, B.W., 81, 95, 98 Wu, G., 30, 115 Wu, H.C., 174 Wu, L.-F., 171–172 Wu, M., 125 Wu, N., 246–247, 253, 263, 265 Wu, T., 260 Wu, Z., 3, 33 Wunsch, P., 130, 135, 147, 150–151, 155, 171–173, 175, 177, 185, 187, 191–192 Xavier, A.V., 79 Xia, Q., 35, 42 Xia, Y., 270 Xie, Z., 49 Xiong, J., 272–273 Xu, C.-A., 155, 157 Xu, Y., 243, 251, 256–258, 263, 266 Yabuuchi, E., 126, 161 Yagi, T., 9 Yair, M., 238–239 Yakushevska, A.E., 277 Yamada, M., 12, 18, 21–22, 28–29, 238, 252 Yamada, M., Yasuda, M., 240 Yamaguchi, A., 248–249, 251, 260 Yamaguchi, H., 139, 141, 143 Yamakawa, H., 252, 258, 263 Yamamoto, A., 113 Yamanaka, T., 27, 36–37, 181 Yamaoka, T., 238 Yamashita, E., 139, 141, 143 Yamulki, S., 120 Yan, Z-Y., 77

AUTHOR INDEX

Yankaskas, J.R., 5–6, 42 Yano, T., 9 Yaono, R., 139, 141, 143 Yasuda, M., 252 Yasui, M., 187 Ye, R.W., 35–36 Ye, S., 246–247, 263 Ye, W., 124, 168 Yeates, T.O., 143 Yeats, C., 187 Yeh, K.-C., 168 Yi, M.N., 144 Yin, Q.Q., 238 Yonekura, K., 257–258 Yonetani, A., 170 Yoneyama, H., 175 Yoon, H.S., 238 Yoon, J.-H., 126 Yoon, S.S., 6, 42, 126 York, J.T., 149 Yoshida, T., 239 Yoshikawa, S., 139, 141, 143 Yoshimatsu, K., 166 Yoshinaga, K., 277 Yoshinari, T., 115–117 Yoshioka, H., 281 Yoshizaki, Y., 143 Yoshizawa, S., 239 Young, B., 275 Young, C.C., 43 Young, M.W., 241, 278 Young, V.G., 137, 143 Youngblood, D.S., 77 Yuan, Y., 3, 33 Yum, D.Y., 19 Yung, Y.L., 110

331

Zahn, J.A., 79, 84, 96 Zak, E., 240 Zakharov, L.N., 149 Zannoni, D., 7, 22, 29 Zech, B., 239 Zehnder, A.J.B., 115 Zeng, A.P., 48 Zeyer, J., 117 Zhang, C.-S., 128, 183 Zhang, J., 158 Zhang, M., 172 Zhang, X., 125, 270–272 Zhang, Z., 158 Zhang, Z.H., 137 Zheng, M., 76 Zhou, L., 125 Zhou, W., 9, 187 Zhou, Y., 77 Zhou, Z., 119 Zickermann, V., 138, 145 Zimmermann, A., 36, 39–40, 97 Zlosnik, J.E., 30 Zlosnik, J.E.A., 1, 43, 45 ZoBell, C.E., 123 Zouni, A., 240 Zufferey, R., 25–26 Zulianello, L., 4 Zumft, W., 132, 148, 151 Zumft, W.G., 24, 26, 29, 32, 36–39, 96, 107, 110–111, 115, 117–119, 126–133, 135–148, 150–156, 159, 163, 166–173, 175, 177–180, 182, 184–185, 187, 191–192, 195–196

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Subject Index Note: The page numbers taken from ﬁgures and tables are given in italics.

A. anaerobicus 119 A. cycloclastes 161 A. evansii 117 A. faecalis 115, 119 A. vinelandii genome sequence 23, 29–30 A. xylosoxidans 135 Ac. cycloclastes 151 aerobic respiration in P. aeruginosa 7–31 Alcaligenes spp. 117 alginate 47 Alkyl-hydroperoxide reductase (Ahp) 76–78 allo-hydroxy-D-proline dehydrogenase 14 amino acid dehydrogenases 12–14 D-amino acid dehydrogenase 12–13 ammonium-oxidizing bacteria 119 An. dehalogenans 159 anaerobic metabolism in the cystic ﬁbrosis lung 42 anaerobic respiration 31–40 Archaea 79, 257 nitrous oxide (N2O) to dinitrogen respiratory transformation by 108–197, see also under nitrous oxide arginine deiminase (ADI) pathway 40 Azoarcus sp. 115 B. B. B. B. B.

azotoformans 117 cenocepacia 47 fragilis 95, 97 halodenitrificans 117 subtilus 11

bacteria, formation and consumption of N2O by 114–120 denitriﬁcation pathway with N2O as intermediate 114–117 truncated denitriﬁcation 117–118 bacterial cells, CCPS roles in 93–98 detoxiﬁcation of periplasmic hydrogen peroxide 93–94 substrates and multiple enzymes 95–96 bacterial cytochrome-c peroxidases (bCCPs), see also under cytochrome-c peroxidases calcium-binding sites 85 catalysis by, mechanistic aspects 88–90 dimer formation in 84–85 electron donors and electron transport in 90–92, see also under electron donors expression, regulation 96–98 haem sites in 85–87, see also under haem sites oxidative and peroxidative stress, enzymic mechanisms to combat 74–78 phylogenetic analysis 79–82 reaction mechanism for 89 structure 83–88 structure, mechanism and physiological roles 73–98 bacterioferritin-comigratory protein (Bcp) 76–78 C. basilensis 115 C. jejuni genome 77, 94–96

334

C. mucosalis 94 C. nephridii 117 C. violaceum 43–44 catalase 76 CF transmembrane regulator (CFTR) 4–7 CikA protein 270–272 deduced amino acid sequence of 270–271 circadian timing, see also kai (circadian clock) locus circadian clock deﬁnition and nomenclature 234–235 clock input 270–274 kai genes and circadian clock evolution 276–278 clock-controlled gene expression 266–270 chromosome compaction 267–270 no solitary output pathway 266 Co. psychrerythraea 125 Cu centre assembly 175–180 supplying Cu, NosA, NosL and ScoP 175–177 CuA centre, in N2O reductase 136–147 Cu–Cu interaction and metallic bond 143–144 cupredoxin fold and loop-directed mutagenesis 137–138 electron transfer 146–147 EPR spectra 140 recombinant CuA centres of N2O reductase 144–146 spectroscopic properties and electronic structure 138–143 cupredoxin fold and loop-directed mutagenesis 137–138 ribbon representation 138 CuZ centre, in N2O reductase 147–152 development of the ﬁeld 147–149 electron transfer and activation of N2O 150–152 spectroscopic properties and structure 149–150

SUBJECT INDEX

cyanide oxygenase (CNO) 46 cyanide production by P. aeruginosa 46–47 genetics 44 physiology 43–44 tolerance 44–46 cyanide-insensitive terminal oxidase (CIO) 29–31 cyanobacteria, see also under circadian timing; kai (circadian clock) locus; S.elongatus PCC 7942 as primary producers 239 circadian timing mechanism in 229–282 clock-controlled gene expression 266–270, see also separate entry morphological diversity among 237 Rpo (sigma factor) and cpmA genes 274–276 cystic ﬁbrosis (CF) lung 1–50, see also under P. aeruginosa cytochrome-c peroxidases (CCPs) 73–98, see also bacterial cytochrome-c peroxidases and MauG proteins, relationship between 80 genome sequencing 79 reaction mechanism 90 roles of CCPS in bacterial cells 93–98, see also under bacterial cells sequence alignment of the N-terminus of 82 cytochromes 21–24 c-type cytochromes 21–22 cytochrome aa3 oxidase 27–28 cytochrome bc1 complex 20–21 cytochrome bd quinol oxidases 30 cytochrome bd 8 cytochrome bo3 quinol oxidase 28 cytochrome bo3 quinol oxidase 28 cytochrome bo3 8 cytochrome c peroxidase (CCP) 31, see also separate entry

SUBJECT INDEX

cytochrome c4 and c5 22–24 cytochrome c551 36 cytochrome cbb3-type oxidases 25–27 cytochrome cd1 36 D. aromatica 108 D. hafniense 159, 162 dadA gene 3 De. aromatica 123, 161 De. denitrificans 123 denitriﬁcation 32, 167 denitriﬁcation genes, regulation 39–40 termination 118 truncated denitriﬁcation 117–118 E. coli 8, 11–12, 271 aerobic respiratory chain 8 bacterial Tat translocon of 170 CCP in 97 cytochrome bo3 of 28 E. coli, Ahp 77 FNR in 96 glucose dehydrogenase in 18 kaiC protein in 243, 257 membrane-bound Nar in 33–34 peroxide generation in 75 Tpx in 77 ubiquinone biosynthesis pathway in 20 electron donors and electron transport in bCCPs 90–92, see also under hydrogen peroxide electron transfer components, in nitrous oxide (N2O) 181–184 cytochrome c 182 cytochrome c550 and cupredoxins as alternative carriers in 181–182 phototrophic denitriﬁers and cytochrome c2 181 electron transfer ﬂavoproteins (ETF) 19 Embden–Meyerhof pathway 16 Entner–Doudoroff pathway (EDP) 12

335

F. hepatica 97 ﬂavohaemoglobin 115 formate dehydrogenase (FDH) 94 GAF motif 271 genomics and N2O respiration 120–127 halorespirers 120–123 mutualistic and pathogenic relationships 126 taxonomic transfers 126–127 glucose dehydrogenase 18 glutathione (GSH) 76–78 glutathione peroxidases (Gpx) 76–78 glycerol kinase (glpK) 12 gram-positive bacteria 167 H. caldilitoris 118 H. inﬂuenzae 6 H. marismortui 124, 159, 164–166 H. pylori 77 Ha. volcanii 124 haem sites in bCCPs electron transport between 92 high potential haem site in 86 low potential haem site 86–87 halophiles 124–125 halorespirers 120–123 hcnB mutant cioA-lacZ expression 30 heterocysts 236 Hf. volcanii 165 hydrogen peroxide (H2O2) 75 as an electron acceptor 94–95 periplasmic hydrogen peroxide 93–94 hypoxic mucus hypothesis 5 kai (circadian clock) locus 242–245 ClustalW derived alignment of kaiA protein 247 ClustalW derived alignment of kaiB protein 254 ClustalW derived alignment of kaiC protein 259 kaiA protein 245–263 kaiB protein 253–256

336

kaiB protein, membrane sequestration of 255 kaiC protein 256–260 kai-clock protein complex 263–265 molecular phylogenetic analysis of 277 mutant alleles of kaiA protein 249–252 sequence, structure and function of clock proteins and 245–265 L. mephitis 118 lactate dehydrogenases 9–12 ldpA protein 272–273 lipoprotein targeting 174 Lyngbya majuscula 190 M. aerodenitrificans 119 M. capsulatus 96 M. magnetotacticum 108, 161–162 M. succiniciproducens 83 Ma. aquaeolei 124 Ma. hydrocarbonoclasticus 132 malate dehydrogenase (MDH) 16 malate quinone oxidoreductase (MQO) 16 MauG proteins 83 membrane-bound Nar 33–34 methylamine dehydrogenase (MADH) 83 mucoid conversion 47–50 mucoidy and cyanide production, link between 49–50 N0 , N00 -dicyclohexylcarbodiimide (DCCD) 49 N. europaea 79, 81, 84, 86–89, 119 N. gonorrhoea 93, 96, 161–162 N. meningitidis 93, 118, 162 N2O reductase, novel Cu centres in 136–152 CuA centre 136–147, see also separate entry

SUBJECT INDEX

CuZ centre 147–152, see also separate entry nos genes, gene expression, regulation, organization 152–157, see also under nos genes to cytochrome oxidase, relationship 163 N2O reductase, properties 127–131 crystal structure and subunit interactions 133–135 enzyme identiﬁcation by classical microbiological strategy 127–128 N2OR homodimer structure 134 novel Cu centres in 136–152 structure 131–136 surface charge 135–136 Z-type N2O reductase 128–131 NADH dehydrogenases 9 NADH-linked peroxidases 76–78 Neisseria meningitidis 76, 93 Neisseria spp. 161 nictotinamide nucleotide transhydrogenase 19–20 nitrate reductase (Nar) 32–36 membrane-bound Nar 33–34 periplasmic nitrate reductases (Nap) 34–36 nitric oxide reductase (NOR) 32, 37–38 nitrite reductase (NIR) 32 in archaea, types 166 nitrogen cycle, biogeochemical 111 nitrogen ﬁxation 240 nitrous oxide (N2O), see also piezophiles archaeal N2O reductase 164–165 as a green house gas 110 bacteria, formation and consumption of N2O by 114–120, see also separate entry bacterial N-oxide transformations involving 116 chemical reduction of 112–113 chemistry of 112–114

SUBJECT INDEX

Cu centre assembly 175–180, see also separate entry electron donation and maintenance of activity in vivo 180–184, see also individual entry evolutionary aspects and phylogenetic relationships 157–168, see also under NosZ ﬂavoprotein formation and reduction 164 formation, by ammonium-oxidizing bacteria 119 genomic and organismal resources 114–127 genomics and N2O respiration 120–127, see also under genomics gram-positive bacteria 167 in enrichment cultures 117 inorganic metabolism of N2O by Archaea 163–166 metabolism 110 N2O generation from NO as intermediate 165–166 N2O reductase, properties 127–131, see also separate entry N2O–transition metal complexes 114 nitrite reductases in archaea, types 166 nitrous oxide reductase (N2OR or Nos) 32, 38–39 NosR and NosX ﬂavoproteins 184–194 oxygen and N2O utilization 119 respiratory nitrate reductase 166 to dinitrogen respiratory transformation, by bacteria and archaea 107–197 topology and transport processes 168–174, see also individual entry nos gene cluster (NGC), gene expression, regulation, organization 152–157 gene expression and regulation 154–157

337

NO signalling via crp-fnr transcription factors 156–157 organizational patterns 121–122 plasmid-encoded nos genes 167–168 promoter studies and anaerobic gene expression 154–156 transcriptional organization 154 NosF ATPase, structure 178 NosL proteins, signal peptides of 174 NosR ﬂavoprotein 184–194 protein domains of 185 structural features in 188 structural features in 189 Nostoc punctiforme 96 NosX ﬂavoprotein 184–194 phylogenetic relationship among 193 NosZ ﬂavoprotein and NosX export to periplasm 168–173 NosD, NosF, NosY and NosL proteins, phylogenetic relationships 160 NosX export 172–173 NosX, Tat-speciﬁc signal sequence 173 of Br. melitensis 159 of W. succinogenes 159 phylogenetic relationships among 157–163 unrooted phylogenetic tree of 158 oxygen, in P. aeruginosa bioﬁlm physiology 6, see also under P. aeruginosa P. aeruginosa, oxygen, cyanide and energy generation in 1–50 aerobic respiration in 7–31 alginate protecting 6 anaerobic metabolism in the cystic ﬁbrosis lung 42 anaerobic respiration 31–40 anaerobic respiratory capability of 6 arginine deiminase pathway of 40 cyanide-insensitive respiration in, 45

338

cytochrome bc1 complex 20–21 cytochrome c peroxidase (CCP) 31, see also under cytochromes denitriﬁcation pathways of 32 description and importance 3–5 energy generation means in 7 energy inhibitors 49 ethanol oxidation and intermediary metabolism in 15 fermentation 40–41 glucose oxidation in, multiple peripheral pathways 17 in cystic ﬁbrosis (CF) 4 in immunosuppression 4 mucoid conversion of 47–50 oxygen 47–48 P. aeruginosa nitrate reductases 33–39, see also under nitrate reductase (Nar) phosphate 48–49 pyruvate fermentation by 41 quinones 20 respiratory chains of 10 respiratory dehydrogenases of 9–19 respiratory inhibitor hydrogen cyanide in, synthesis 43–47 terminal oxidases in 24, 25–31, see also separate entry transhydrogenase 19–20 P. chlororaphis 117 P. denitrificans 79, 84, 91–92, 96, 115 P. fluorescens 18, 96, 117 P. gingivalis 77 P. nautica 91 P. pantotrophus 79, 84, 87, 91 P. profundum 96 P. stutzeri 33, 37, 84, 96, 111, 115, 117, 123, 129 P. succinatimandens 118 P. taiwanensis 118 Pa. denitrificans 9, 11, 27, 33, 37, 135, 142, 151 Pa. macerans 11 Pa. pantotrophus 159

SUBJECT INDEX

pex gene 274 Ph. profundum 123 piezophiles 123–124 PQQ-containing dehydrogenases 14–19 electron transfer ﬂavoproteins (ETF) 19 ethanol oxidation system 14–16 glucose and gluconate dehydrogenases 16–17 malate dehydrogenase 16 proline dehydrogenase (ProDH) 13 Pseudomonas sp. 123 psychrophiles 125–126 PutA protein 13 Py. aerophilum 125, 165–166 pyrroloquinoline quinone (2,7,9, -tricarboxypyrroloquinoline quinone, PQQ) 14–19, see also under PQQ-containing dehydrogenases quinol fumarate oxidoreductase (QFR) 11 quinones 20 R. capsulatus 79, 84, 86–87, 91 R. denitrificans 117 R. eutrophus 37 R. sphaeroides 26–27 Rh. ferrireducens 125 Rpo (sigma factor) and cpmA genes 274–276 Rusticyanin 23 S. aureus 6 S. elongatus PCC 7942 230–282, see also circadian timing; cyanobacteria chromosome compaction in 268 CikA protein 270–272 cyanobacterial clock 278–280, see also under cyanobacteria kai locus 242–245, see also under kai (circadian clock) locus ldpA protein 272–273

SUBJECT INDEX

NMR-based structure analyses of 262–263 pex gene 274 rhythmic gene expression and chromosome compaction in 232 SasA protein 260–263 S. pyogenes 76 S.longatus PCC 7942 229 Sa. ruber 124 SasA protein 260–265 Sh. denitrificans 162 Sh. oneidensis 162 Si. pomeroyi 123 sn-glycerol-3-phosphate (G3P) dehydrogenase 12 succinate dehydrogenase (succinate quinone oxidoreductase) 9–11 superoxide dismutase (SOD) 75 T. aromatica117 T. elongatus BP-1 229, 233 T. mechernichensis 115 T. pantotropha 95 T. selenatis 117 Te. roseum 125–126, 159 terminal oxidases in P. aeruginosa 24, 25–31 B. japonicum 25, 27 cyanide-insensitive terminal oxidiase (CIO) 29–31 cytochrome aa3 oxidase 27–28

339

cytochrome bo3 quinol oxidase 28 cytochrome cbb3-type oxidases 25–27 thermophiles 125–126 thiolperoxidase (Tpx) 76–78 thioredoxin 76–78 Tm. denitrificans 159, 161, 183 topology and transport processes 168–174, see also twin arginine translocation ABC transporter 177–180 N2O respiration and N2OR biogenesis, membrane topology of 176 N2OR export 169–172 NosZ and NosX export to periplasm 168–173 Sec system 173 transhydrogenase 19–20 tri-haem proteins 79–82 tryptophan tryptophylquinone (TTQ) 83 twin arginine translocation (Tat) system 168–173 sequence logo 170 ubiquinone 20 W. succinogenes 108, 117, 128, 159 electron transfer to N2O reductase in 182–184 N2O respiration system of 184

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Colour Plate Section

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Plate 1 (A) Stereoview of the structure of the N2OR homodimer from Pa. denitrificans. (B) Views of the CuA electron transfer centre and the catalytic centre CuZ. (C) The N2OR monomer seen from the contact side with the other subunit. In this orientation the outermost b-sheets run counterclockwise (Protein Data Bank ﬁle 1FWX; drawn with MolScript and Raster3D Softwares). (See page 134, this volume)

Plate 2 Schematic diagram of the engineered CuA site in azurin from P. aeruginosa. (A) View of the Cu2S2 rhomb; (B) side view of the CuA site (PDB 1CC3). (See page 139, this volume)

Plate 3 Unrooted phylogenetic tree of NosZ sequences. The sequences were retrieved from the NCBI database by identifying respective entries in the annotations and by BLAST searches (Altschul et al., 1997) of the core nucleic acid, protein and genome databases. Ongoing genome projects were screened by FASTA or BLAST algorithms depending on the hosting database server of TIGR Microbial Database, The Joint Genome Institute Microbial Genomics Database, The Wellcome Trust Sanger Institute, and ERGO Database of Integrated Genomics Inc. The data set excludes environmental N2OR fragments obtained from nosZ genes ampliﬁed by PCR. Sequences were processed by the multiple-alignment program ClustalX 1.83 (Thompson et al., 1994), using the Gonnet series as protein weight matrix and parameters set to 10 for gap opening, 0.2 for gap extension and divergent sequences delay at 30%. Subsequently, a protein distance matrix was calculated by the use of ProtDist component of the PHYLIP v. 3.65 package (Felsenstein, 1989). Phylogenetic afﬁliation was calculated by the neighbour-joining method (Neighbour program of PHYLIP), and displayed by the TreeView 1.6.6 software (Page, 1996). Numbers on branchings indicate percentages of partitions in an extended majority rule consensus tree based on a bootstrap with 1000 replicates, carried out with PHYLIP’s SeqBoot and Consense applications. The scheme of colours for the clusters of branches indicates the taxonomic group on the class or phylum level based on 16S rRNA. For strain speciﬁcations and abbreviations see Table 2; Mhyd, Ma. hydrocarbonoclasticus. (See page 158, this volume)

Plate 4 Phylogenetic relationships of NosD, NosF, NosY and NosL proteins. The trees were constructed from species that harbour a nos gene cluster. For discussion of the positions of Neisseria spp. and A. cycloclastes see the text. For abbreviations see Table 2; Halosp, Halobacterium sp. NRC1; for methods and colour code see Fig. 10. (See page 160, this volume)

Plate 5 Sequence alignments of conserved regions of the NosR protein. (A) Flavinbinding region. The alignment depicts the region around the putative cofactor-carrying threonine (*), which is delimited in the amino acid sequence N- and C-terminally by indels. (B) Glycine-rich region within the periplasmic domain; (C) C-terminal polyferredoxin region. Colour coding as of the default ﬁle of ClustalX 1.83 except for cysteine. For abbreviations see Table 2; Ddeh, Desulfitobacterium dehalogenans; PdenI, Pa. denitrificans NirI; Rbal, Rhodopirellula baltica. (See page 186, this volume)

Plate 6 Rhythmic gene expression and chromosome compaction in S. elongatus PCC 7942. (A) Bioluminescence (counts per second) recorded over time (hours) from a F(kaiB-luc+) reporter in an otherwise wild-type strain. Three independent data sets are graphed. The gray boxes indicate time without illumination. Such data can be recorded automatically for many days. This technology has allowed rapid growth in the study of cyanobacterial circadian rhythms. (B) Deconvolved ﬂuorescence microscopy images (red-cell autoﬂuorescence; green-DAPI stained DNA) of wild-type cells sampled during a light/dark cycle at ZT ¼ 0 and ZT ¼ 12. Note the diffuse state of the chromosome at ZT ¼ 0 and the arrangement of the DNA into distinct ‘‘nucleoids’’ at ZT ¼ 12. Cells are approximately 5 mm long. (See page 232, this volume)

Plate 7 Morphological diversity among the cyanobacteria. (A) A deconvolved ﬂuorescence microscopy image (red-autoﬂuorescence; green-DAPI stained DNA) of the ovoid-shaped cyanobacterium, P. marinus MIT9313. (B) As above, but image is of the ﬁlamentous cyanobacterium Anabaena sp. PCC 7120. (C) As above, except image is of a halotolerant Spirulina sp. isolated from The Great Salt Lake, Utah. USA. Each size bar represents 5 mm. (See page 237, this volume)

Plate 8 (A) Structure and (B) protein architecture of N-SasA from S. elongatus PCC 7942 and KaiB from Anabaena sp. strain PCC 7120. See text for discussion of comparative function. Figure modiﬁed from Vakonakis et al. (2004). (See page 263, this volume)

Plate 9 Chromosome compaction in S. elongatus PCC 7942 as demonstrated with deconvolved ﬂuorescence microscopy images (red-autoﬂuorescence from S. elongatus PCC 7942; green -DAPI stained DNA). Cultures of wild-type and kaiC strains were sampled at the indicated times during a light/dark cycle (A and C) or a free-running cycle (B and D). The chromosome arrangement image shown for each time point is representative of 499% (n ¼ 300) of the cells from a sample. For the wild-type strain note that in each of the cycles (A and B) there is a slow arrangement of the DNA into distinct ‘‘nucleoids’’ and then a return to the time 0 diffuse state. For the kaiC strain note that at any given time point within each experiment some variation in the extent of chromosome compaction was evident. This variation is also reﬂected in the gene expression rhythms from that strain (Smith and Williams, 2006). However, no rhythmic compaction patterns are observed in this genetic background (see text for discussion). Cells are approximately 5 mm long. (See page 268, this volume)

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