Advances in Microbial Physiology, Volume 54

Advances in

MICROBIAL PHYSIOLOGY VOLUME 54

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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 54

Amsterdam  Boston  Heidelberg  London  New York  Oxford Paris  San Diego  San Francisco  Singapore  Sydney  Tokyo Academic Press is an imprint of Elsevier

ACADEMIC PRESS

Academic Press is an imprint of Elsevier 32 Jamestown Road, London NW1 7BY, UK Radarweg 29, PO Box 211, 1000 AE Amsterdam, The Netherlands Linacre House, Jordan Hill, Oxford OX2 8DP, UK Corporate Drive, Suite 400, Burlington, MA 01803, USA 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA First edition 2009 Copyright r 2009 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 (þ44) (0) 1865 843830; fax (þ44) (0) 1865 853333; email: [email protected]. Alternatively you can submit your request online by visiting the Elsevier web site at http://www.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: 978-0-12-374323-7 ISSN: 0065-2911 For information on all Academic Press publications visit our website at elsevierdirect.com

Printed and bound in the United Kingdom 09 10 11 12 13 10 9 8 7 6 5 4 3 2 1

Contents

CONTRIBUTORS

TO

VOLUME 54 . . . . . . . . . . . . . . . . . . . . . . . . . .

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Complex Regulatory Pathways Coordinate Cell-Cycle Progression and Development in Caulobacter crescentus Pamela J.B. Brown, Gail G. Hardy, Michael J. Trimble and Yves V. Brun Abbreviations. . . . . . . . . . . . . . . . . . . . . . . . . . . . . Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Global Regulation. . . . . . . . . . . . . . . . . . . . . . . . . . Polar Structure Biogenesis and Function. . . . . . . . . . . . Chromosome Replication and Segregation, Cell Division, and Cell Shape . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Concluding Remarks. . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements. . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abbreviations. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anoxygenic Phototrophic Bacteria: Physiology and Taxonomy. Anoxygenic Phototrophic Sulfur Bacteria: Ecology . . . . . . . . Transformations of Sulfur Compounds. . . . . . . . . . . . . . . . Oxidative Sulfur Metabolism: Enzymes and Genes. . . . . . . . . Evolution of Sulfur Metabolism . . . . . . . . . . . . . . . . . . . . Sulfate Assimilation in Anoxygenic Phototrophic Bacteria. . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements. . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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105 106 107 116 118 126 165 167 177 179 179

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Sulfur Metabolism in Phototrophic Sulfur Bacteria Niels-Ulrik Frigaard and Christiane Dahl

1. 2. 3. 4. 5. 6. 7. 8.

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Carbon, Iron and Sulfur Metabolism in Acidophilic Micro-Organisms D. Barrie Johnson and Kevin B. Hallberg 1. 2. 3. 4. 5.

Acidophilic Micro-organisms: Overview. . . . . . . . . . Carbon Metabolism in Acidophilic Micro-organisms. Iron Metabolism by Acidophilic Micro-organisms. . . Sulfur Metabolism by Acidophilic Micro-organisms. . Applied and Ecological Aspects. . . . . . . . . . . . . . . Acknowledgments. . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Chemostat-Based Micro-Array Analysis in Baker’s Yeast Pascale Daran-Lapujade, Jean-Marc Daran, Antonius J.A. van Maris, Johannes H. de Winde and Jack T. Pronk

1. 2. 3. 4. 5. 6. 7. 8.

Abbreviations. . . . . . . . . . . . . . . . . Introduction. . . . . . . . . . . . . . . . . . Experimental Tools. . . . . . . . . . . . . . Specific Growth Rate . . . . . . . . . . . . Context Dependency. . . . . . . . . . . . . Functional Analysis . . . . . . . . . . . . . Evolutionary and Reverse Engineering . Outlook. . . . . . . . . . . . . . . . . . . . . Concluding Remarks. . . . . . . . . . . . . Acknowledgements. . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . .

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A Predatory Patchwork: Membrane and Surface Structures of Bdellovibrio bacteriovorus Carey Lambert, Laura Hobley, Chien-Yi Chang, Andrew Fenton, Michael Capeness and Liz Sockett 1. 2. 3. 4.

Introduction. . . . . . . . . . . . . . . . . . . . . The Role of Flagellar Motility in Predation. The Role of Type IV Pili in Predation . . . . Outer Membrane Proteins . . . . . . . . . . . .

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Autotransporters. . . . . . . . . . . . . . . . . The Role of the Sec System . . . . . . . . . . Membrane Chemistry. . . . . . . . . . . . . . Peptidoglycan Chemistry and Metabolism. Summary. . . . . . . . . . . . . . . . . . . . . . Acknowledgements. . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . .

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AUTHOR INDEX. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363 SUBJECT INDEX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 395 Color Plate Section to be found in the back of this book

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Contributors to Volume 54

PAMELA J.B. BROWN, Department of Biology, Indiana University, Bloomington, IN 47405-3700, USA YVES V. BRUN, Department of Biology, Indiana University, Bloomington, IN 47405-3700, USA MICHAEL CAPENESS, Institute of Genetics, School of Biology, University of Nottingham, Queen’s Medical Centre, Nottingham NG7 2UH, UK CHIEN-YI CHANG, Institute of Genetics, School of Biology, University of Nottingham, Queen’s Medical Centre, Nottingham NG7 2UH, UK CHRISTIANE DAHL, Institut fu¨r Mikrobiologie & Biotechnologie, Rheinische Friedrich Wilhelms-Universita¨t Bonn, D-53115 Bonn, Germany JEAN-MARC DARAN, Department of Biotechnology, Delft University of Technology and Kluyver Centre for Genomics of Industrial Fermentation, Julianalaan 67, 2628 BC Delft, The Netherlands PASCALE DARAN-LAPUJADE, Department of Biotechnology, Delft University of Technology and Kluyver Centre for Genomics of Industrial Fermentation, Julianalaan 67, 2628 BC Delft, The Netherlands JOHANNES H. DE WINDE, Department of Biotechnology, Delft University of Technology and Kluyver Centre for Genomics of Industrial Fermentation, Julianalaan 67, 2628 BC Delft, The Netherlands ANDREW FENTON, Institute of Genetics, School of Biology, University of Nottingham, Queen’s Medical Centre, Nottingham NG7 2UH, UK

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CONTRIBUTORS TO VOLUME 54

NIELS-ULRIK FRIGAARD, Copenhagen Biocenter, Department of Biology, University of Copenhagen, Ole Maaløes Vej 5, 2200 Copenhagen N, Denmark KEVIN B. HALLBERG, School of Biological Sciences, Bangor University, Bangor LL57 2UW, UK GAIL G. HARDY, Department of Biology, Indiana University, Bloomington, IN 47405-3700, USA LAURA HOBLEY, Institute of Genetics, School of Biology, University of Nottingham, Queen’s Medical Centre, Nottingham NG7 2UH, UK. D. BARRIE JOHNSON, School of Biological Sciences, Bangor University, Bangor LL57 2UW, UK. CAREY LAMBERT, Institute of Genetics, School of Biology, University of Nottingham, Queen’s Medical Centre, Nottingham NG7 2UH, UK. JACK T. PRONK, Department of Biotechnology, Delft University of Technology and Kluyver Centre for Genomics of Industrial Fermentation, Julianalaan 67, 2628 BC Delft, The Netherlands. LIZ SOCKETT, Institute of Genetics, School of Biology, University of Nottingham, Queen’s Medical Centre, Nottingham NG7 2UH, UK. MICHAEL J. TRIMBLE, Department of Biology, Indiana University, Bloomington, IN 47405-3700, USA. ANTONIUS J.A. VAN MARIS, Department of Biotechnology, Delft University of Technology and Kluyver Centre for Genomics of Industrial Fermentation, Julianalaan 67, 2628 BC Delft, The Netherlands.

Complex Regulatory Pathways Coordinate Cell-Cycle Progression and Development in Caulobacter crescentus Pamela J.B. Brown, Gail G. Hardy, Michael J. Trimble and Yves V. Brun Department of Biology, Indiana University, Bloomington, IN 47405-3700, USA

ABSTRACT Caulobacter crescentus has become the predominant bacterial model system to study the regulation of cell-cycle progression. Stage-specific processes such as chromosome replication and segregation, and cell division are coordinated with the development of four polar structures: the flagellum, pili, stalk, and holdfast. The production, activation, localization, and proteolysis of specific regulatory proteins at precise times during the cell cycle culminate in the ability of the cell to produce two physiologically distinct daughter cells. We examine the recent advances that have enhanced our understanding of the mechanisms of temporal and spatial regulation that occur during cell-cycle progression.

Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Global regulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Coordination between Oscillating Global Regulators . . . . . . . . . . 2.2. Sigma Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Promoter Architecture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Two-Component Signal Transduction Proteins . . . . . . . . . . . . . . 3. Polar structure biogenesis and function . . . . . . . . . . . . . . . . . . . . . . 3.1. The Flagellum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. The Pili . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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3.3. The Stalk . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4. The Process of Adherence and the Holdfast . . . . . . . . . . . . . . . Chromosome replication and segregation, cell division, and cell shape . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Chromosome Replication Occurs Once and Only Once per Cell Division Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Coordination of Chromosome Segregation and Cell Division . . . . 4.3. Regulation of Cell Division Protein Expression . . . . . . . . . . . . . . 4.4. Cell Shape . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

40 44 60 60 65 74 76 82 83 84

ABBREVIATIONS A22 AFM bps c-di-GMP Cori ECT Flp GDP GFP GTP Hpt IHF NAG BP PBP2 WGA YFP

S-(3,4-dichlorobenzyl)isothiourea atomic force microscopy base pairs cyclic-diguanosine monophosphate C. crescentus origin of replication electron cryotomography fimbrial low-weight protein guanosine diphosphate green fluorescent protein guanosine triphosphate histidine phosphotransferase integration host factor N-acetylglucosamine phosphorylated penicillin-binding protein 2 wheat germ agglutinin yellow fluorescent protein

1. INTRODUCTION Decades of work in eukaryotic biology have shown that many types of cells undergo programmed cell cycles, which consist of a series of invariant steps. The transitions from one stage to the next are mediated by complex regulatory networks that lead to the ordered production, localization,

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and activation of proteins for critical cellular events such as chromosome replication, organelle development, and cell division. The aquatic bacterium Caulobacter crescentus undergoes a programmed developmental cycle that requires integrated regulatory networks that may rival those of eukaryotic cells. Thus, C. crescentus is an excellent model system for studying cellular differentiation (Brun and Janakiraman, 2000; Poindexter, 1964). Progression through the cell cycle results in a sequential series of changes in cellular morphology and requires the coordination of processes including chromosome replication, chromosome segregation, polar morphogenesis, cell growth, and cell division. Unlike many prokaryotes, C. crescentus replicates its chromosome only once during the cell division cycle such that the G1, S, and G2 phases are readily distinguishable (Fig. 1). The cell division cycle is tightly coupled to a series of morphological transitions resulting in the formation of two distinct cell types, a motile swarmer cell and a sessile stalked cell. The swarmer cell has a polar flagellum and pili and is incapable of chromosome replication. After a gap period (G1) equivalent to about one-third of the cell cycle, the swarmer cell differentiates into a stalked cell. Cellular differentiation involves ejection of the flagellum, retraction of the pili, and synthesis of a stalk with an adhesive holdfast, which all occur at the same pole. At the onset of S phase, the stalked cell initiates chromosome replication, cell division, and synthesis of a flagellum at the pole opposite the stalk. Flagellum rotation is activated just prior to cell separation. After cell division, pili are synthesized on the new swarmer cell and the stalked cell undergoes a new round of chromosome replication and cell division. Many genes and gene products important for cell-cycle and developmental regulation have been identified in a number of genetic screens. The ability to synchronize the cells and monitor changes in the transcriptome and proteome as cells proceed through the cell cycle has led to the identification of additional genes and proteins that are important for developmental regulation. In this review, we will highlight recent advances in the study of C. crescentus that have enhanced our understanding of the complex regulatory circuits that are required for cell-cycle progression. We will examine the roles of several regulatory elements and discuss how they influence polar development, maintenance of cell shape, and cell division.

2. GLOBAL REGULATION Most bacteria respond to environmental and physiological changes by using complex global regulatory mechanisms to adjust the transcription levels of

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SW

CtrA CtrA~P DnaA GcrA

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PDE

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Flagellum ejection Holdfast synthesis Stalk synthesis

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G2 Flagellum synthesis

DNA replication

ST G1 Pilus synthesis

DNA methylation

Chromosome segregation

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Figure 1 The C. crescentus cell cycle and the localization of the major regulators CtrA, DnaA, and GcrA. The cell cycle, including the swarmer (SW), stalked (ST), early predivisional (PDE), late predivisional (PDL), and progeny cells, is depicted. The stages of the cell division cycle and cell cycle events are shown below the cell cycle schematic. G1 is the pre-synthesis gap period, S is the DNA synthesis period, and G2 is the post-synthesis gap period. Phosphorylated CtrA (CtrABP) is found in the swarmer cell. As the cell undergoes the swarmer to stalked cell differentiation, CtrA is recruited to the flagellar pole where it is proteolyzed. DnaA is then synthesized and leads to the production of GcrA, the dominant regulator in the stalked cell. In the predivisional cell, GcrA is responsible for the activation of ctrA transcription. CtrA is phosphorylated and represses transcription of gcrA. In the stalked compartment, CtrA is subject to proteolysis at the stalked pole and both GcrA and DnaA accumulate in the new stalked cell. In the swarmer compartment, phosphorylated CtrA is present and it blocks chromosome replication in the new swarmer cell.

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specific genes. The numerous combinations of interactions amongst cis- and trans-acting elements lead to an extensive web of transcriptional regulation that controls critical events such as cell division and cell differentiation. A compilation of earlier studies indicated that the transcription of 72 single genes varies as a function of the cell cycle, including many genes involved in the regulation of the cell cycle and cell differentiation (Laub et al., 2000). The availability of the genome sequence of C. crescentus (Nierman et al., 2001) resulted in the development of methods for global identification of genes that control the cell cycle. The initial global transcription analysis found that 19% of the genome (553 genes) was comprised of genes whose RNA levels varied throughout the cell cycle (Laub et al., 2000). The validity of this approach is supported by the fact that the 72 genes that were previously identified as being cell-cycle regulated are found within the set of temporally regulated transcripts. The use of global transcriptional profiling has provided a tantalizing glimpse into the molecular mechanism(s) involved in cell cycle control (Laub et al., 2000). Several insights have been made from this work, which allow us to begin to address the question of how C. crescentus regulates cell-cycle progression at the molecular level. The observation that the peak expression of cell-cycle regulated genes occurs immediately before or coincident with the event requiring the gene product has led to the notion of ‘‘just-in-time’’ transcription that may allow the cell to grow and differentiate more efficiently. The roles of several regulatory elements known to be important in the cell-cycle progression of C. crescentus are discussed below.

2.1. Coordination between Oscillating Global Regulators At least three transcriptional regulators, CtrA, GcrA, and DnaA, modulate the distinctive cell-cycle and morphological changes that occur during the C. crescentus life cycle. Each regulator has a unique pattern of expression that is coincident with its function in the cell (Fig. 1; for review, see Holtzendorff et al., 2006). Phosphorylated CtrA, a response regulator protein, is found in the swarmer cell where it binds to the origin of replication and inhibits chromosome replication (Quon et al., 1998). In the swarmer cell, CtrA also represses ftsZ, which is required for cell division, and podJ, which is required for polar development. CtrA is rapidly degraded by proteolysis as the cell transitions from a swarmer cell to a stalked cell and DnaA is stablized. DnaA is an essential activator of chromosome replication (Gorbatyuk and Marczynski, 2001) and acts as a transcriptional regulator during early S phase, leading to the synthesis of GcrA (Hottes et al., 2005).

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In the stalked cell, DnaA is responsible for the activation of both ftsZ and podJ. During early S phase, GcrA is the dominant regulator and activates genes required for chromosome replication, cell elongation, and polar development by an unknown mechanism (Holtzendorff et al., 2004). As the cell transitions from the S phase to the G2 phase, ctrA transcription is activated by GcrA. Newly synthesized CtrA is phosphorylated and in turn represses gcrA. Recruitment of CtrA to the stalked pole in the stalked compartment of the predivisional cell leads to dephosphorylation and proteolysis of CtrA. The decrease in active CtrA relieves repression of gcrA and allows GcrA to accumulate. Meanwhile, CtrA remains at a high level in the swarmer compartment of the predivisional cell and blocks gcrA transcription and chromosome replication. The sequential expression of the CtrA/DnaA/GcrA regulators drives the cell-cycle progression of C. crescentus and allows the formation of two distinct progeny cells following cell division. The specific roles of each regulator in cell-cycle progression are described below. 2.1.1. CtrA In C. crescentus, CtrA is an essential response regulator that serves as a major regulatory protein. CtrA contains a DNA-binding domain and recognizes specific sequences that are found upstream of many cell-cycle controlled promoters (Laub et al., 2002; Quon et al., 1996). CtrA acts either as an activator or repressor for the transcription of a number of cell-cycle regulated genes. The CtrA regulon includes genes whose products are required for flagellum assembly and activation, pili biogenesis, holdfast synthesis, DNA methylation, chromosome replication, and cell division. The level of ctrA transcription is influenced by two promoters, the methylation state of the DNA, and the presence of activators (Domian et al., 1997; Quon et al., 1996; Reisenauer and Shapiro, 2002). CtrA activity is controlled by cellular localization, proteolysis, and phosphorylation (Biondi et al., 2006a; Domian et al., 1997; Iniesta et al., 2006; McGrath et al., 2006; Ryan et al., 2002; 2004). As a result, active CtrA appears only at precise times in the C. crescentus cell cycle, including the swarmer and predivisional cells. 2.1.1.1. CtrA Transcription. The expression of ctrA is controlled by two promoters which are active at different times in the cell cycle (Domian et al., 1999). P1 is a relatively weak promoter located 122 base pairs (bps) upstream of the ctrA translation start site and is active only in the stalked cell. The P1 promoter contains a GAnTC methylation site and is activated soon after

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chromosome replication, when the chromosome is hemi-methylated (Marczynski, 1999; Reisenauer and Shapiro, 2002; Stephens et al., 1996). GcrA activates ctrA expression leading to the production of phosphorylated CtrA (CtrABP) (Holtzendorff et al., 2004). In predivisional cells, CtrABP activates the transcription of ccrM, which encodes the adenine DNA methyltransferase that methylates the newly replicated DNA (Quon et al., 1996). When the chromosome is fully methylated, the P1 promoter is repressed allowing minimal ctrA expression prior to a new round of chromosome replication (Domian et al., 1999). P2 is a stronger promoter located 65 bps upstream of the ctrA translational start site and is activated by CtrABP in the predivisional cell. Thus, both promoters are subject to feedback control by CtrA; P1 is repressed and P2 is activated. Combined with regulated phosphorylation and proteolysis of CtrA (see Sections 2.1.1.2 and 2.1.1.3), the feedback control of the ctrA promoters results in the appearance of active CtrA during specific times in the cell cycle (Fig. 1). 2.1.1.2. CtrA Localization and Proteolysis. In addition to transcriptional control, proteolysis plays an important role in the presence of active CtrA. Following the production of CtrA, the predivisional cell becomes compartmentalized and CtrA is degraded from the stalked cell compartment, but not from the swarmer cell compartment prior to cell division (Domian et al., 1997). The daughter swarmer cell inherits CtrA, but the daughter stalked cell does not. CtrA is degraded as the swarmer cell undergoes differentiation into a stalked cell and reappears in the early predivisional cell (Fig. 2). Proteolysis of CtrA is mediated by the essential ClpXP protease (Jenal and Fuchs, 1998). Depletion of either ClpX or ClpP leads to the stabilization of CtrA and arrest of the cell cycle prior to chromosome replication. Since CtrA is not degraded, it is presumed to remain bound to the origin of replication, preventing the initiation of chromosome replication. Even though clpX and clpP are constitutively expressed and ClpXP is present throughout the cell cycle, the activity of ClpXP must be modulated since CtrA is degraded only in the swarmer cell and the stalked compartment of the late predivisional cell. CtrA degradation is controlled by two determinants within the CtrA protein (Ryan et al., 2002). One determinant is present at the C-terminus of CtrA. A proteolytically stable form of CtrA is generated by deleting the last three C-terminal amino acid residues or by modification of the last two residues from alanines to aspartates (Domian et al., 1997). The second determinant is located within the first 56 residues of CtrA. The N-terminus of CtrA is comprised of the receiver domain, including the critical aspartate residue that is phosphorylated by the CckA-ChpT-CtrA phosphorelay as

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CckA

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CpdR

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DivK~P

DivK

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PodJS

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ST

Figure 2 Localization of proteins affecting CtrA phosphorylation and proteolysis during the C. crescentus cell cycle. All stages of the cell cycle, including the swarmer (SW), stalked (ST), early predivisional (PDE), late predivisional (PDL), and progeny cells, are shown. In the swarmer cell, PodJS is localized at the flagellar pole and is responsible for the localization of PleC. PleC dephosphorylates DivKBP leading to the accumulation of delocalized DivK. CckA is also localized to the flagellar pole and controls CtrA prosphorylation (CtrABP), leading to a block in chromosome replication. As the cell undergoes the swarmer to stalked cell differentiation, PodJ is degraded, releasing PleC from the pole. CpdR is localized to the flagellated pole and recruits both CtrA and ClpXP/RcdA to the pole, resulting in the proteolysis of CtrA. In the stalked cell, PodJL is synthesized and localizes to the incipient swarmer pole. DivJ is localized to the stalked pole where it recruits and phosphorylates DivK, leading to accumulation of DivKBP at the pole and in the cytoplasm. In the early predivisional cell, PodJL recruits PleC to the incipient swarmer pole where PleC dephosphorylates DivKBP. Meanwhile, DivJ is localized to the stalked pole where DivK is phosphorylated. Due to the opposing actions of PleC and DivJ, DivK and DivKBP are found in the cytoplasm. In the swarmer compartment of the late predivisional cell, the periplasmic protease PerP is responsible for the cleavage of PodJL to PodJS at the incipient flagellar pole. CckA and PleC remain at the flagellar pole and CtrA is phosphorylated. The absence of DivJ and the presence of PleC in the swarmer compartment lead to the accumulation of unphosphorylated DivK. In the stalked compartment of the predivisional cell, CpdR and ClpXP/RcdA join DivJ and DivKBP at the stalked pole, resulting in the degradation of CtrA and accumulation of DivKBP. In the new stalked cell, PodJL is synthesized, CtrA is completely degraded, and CpdR and ClpXP/RcdA are no longer polarly localized.

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described in section 2.1.1.3 (Biondi et al., 2006a). Neither the expression of the receiver domain nor the expression of the C-terminus of CtrA alone is sufficient for cell-cycle degradation (Domian et al., 1997; Ryan et al., 2002). The phosphorylation state of the receiver domain modulates activity (see Section 2.1.1.3), but does not affect proteolysis (Ryan et al., 2002). Fusion of the receiver domain to the 15 C-terminal amino acid residues is sufficient for cell-cycle-regulated proteolysis of CtrA (Ryan et al., 2002). Fusion of the receiver domain and the 15 C-terminal residues of CtrA to yellow fluorescent protein (YFP) is sufficient to confer cell-cycle-dependent degradation to YFP. A series of five hybrid proteins generated using the receiver domain of CtrA homologs from other bacterial genera and the 15 C-terminal residues of CtrA were expressed in C. crescentus. Four of the hybrid proteins were degraded with a temporal pattern similar to native CtrA, indicating that they share a proteolytic signal. Ten candidate residues for a proteolytic signal were identified by a sequence alignment and nine of these residues are predicted to be located on one surface of CtrA. It has been hypothesized that some or all of these residues may form a binding pocket for an activating protein that stimulates degradation of CtrA by ClpXP (Ryan et al., 2002). CtrA localizes to the incipient stalked pole in cells undergoing the swarmer to stalked cell differentiation and to the stalked pole in the stalked compartment of predivisional cells, just prior to its degradation (Fig. 2) (Ryan et al., 2002, 2004). CtrA proteolysis in the stalked cell compartment of the predivisional cell occurs only after a diffusion barrier forms (Judd et al., 2003). The delay in proteolysis until after compartmentalization enables the production of two distinct daughter cells with very different levels of CtrA. These results indicate that there may be a regulatory link between compartmentalization and the initiation of CtrA proteolysis. In this model, polar localization is a prerequisite for proteolysis (Fig. 2). This is consistent with the hypothesis that an activating protein promotes the interaction of CtrA with ClpXP at a specific time and place. The notion of an activating protein for degradation of CtrA by ClpXP is supported by the observation that CtrA and ClpX are found in a polar complex with a third protein, RcdA (McGrath et al., 2006). The localization pattern of RcdA matches that of both CtrA and ClpX; all three proteins appear at the same pole in cells undergoing the swarmer to stalked cell differentiation and in the stalked compartment of predivisional cells (Fig. 2). In cells lacking rcdA, ClpX localization still occurs, but polar CtrA foci are not observed and CtrA remains delocalized throughout the cell cycle. The half-life of CtrA in cells lacking rcdA is 2.6 times greater than the half-life of CtrA in wild-type cells. When cells expressing RcdA are depleted of ClpX,

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RcdA no longer forms polar foci. These results suggest that ClpX localizes to the pole prior to RcdA and that RcdA recruits CtrA to the cell pole. Therefore, the dynamic localization of this complex is responsible for the temporal control of CtrA proteolysis. RcdA is not absolutely required for an interaction between ClpX and CtrA; complexes of nonpolar ClpX-CtrA can be isolated from the cell (McGrath et al., 2006). These complexes are presumed to be nonfunctional in vivo as CtrA is not degraded in an rcdA mutant; however, in vitro studies using recombinant C. crescentus ClpXP indicate that an activating protein is not required for degradation of CtrA by ClpXP (Chien et al., 2007). In vitro studies indicate that ClpXP directly recognizes the C-terminal tag of CtrA. The N-terminal domain required for localization and proteolysis in vivo is dispensable for in vitro degradation. The rates of CtrA degradation by ClpXP are unchanged by the addition of RcdA, suggesting that RcdA is unlikely to act as a typical substrate-specific adaptor for degradation. Kinetic analyses indicate that the rate of unassisted CtrA degradation by ClpXP is sufficient to account for the observed rate of intracellular CtrA degradation. These results show that the N-terminal domain of CtrA and RcdA are not required for the activation of CtrA proteolysis in the in vitro environment. The requirement for RcdA to mediate CtrA proteolysis in vivo, but not in vitro, suggests that there are likely to be additional factors involved in controlling CtrA proteolysis. One possibility is that there are unidentified inhibitors of ClpXP in the swarmer and predivisional cells (Chien et al., 2007). Since the level of ClpXP is limiting compared with the level of CtrA, competition between CtrA and other higher priority substrates could provide a mechanism for inhibition. This is an attractive model since a specific inhibitor is not required. Another possibility is that CtrA may interact with a protein that protects it from degradation in swarmer and predivisional cells. The inhibitor may interact with CtrA and protect CtrA from proteolysis by masking the C-terminal recognition motif either directly or indirectly. In this scenario, RcdA recruitment of CtrA to the pole may result in release of the protease inhibitor. In vitro studies conducted in the absence of the relevant inhibitor may suggest that RcdA is not required for efficient proteolysis. The disparity between in vivo and in vitro results with regard to the role of RcdA in CtrA proteolysis clearly indicates that the regulation of CtrA proteolysis is likely to involve additional factors. How is the timing of CtrA degradation controlled despite the presence of ClpXP through all stages of the cell cycle? To answer this question we turn our attention to the single-domain response regulator, CpdR, which mediates the localization and activity of the ClpXP protease (Iniesta et al., 2006). CtrA is stabilized in a cpdR deletion strain, indicating that

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CpdR is required for CtrA proteolysis. CpdR exhibits the same localization pattern as CtrA, ClpXP, and RcdA throughout the cell cycle suggesting that CpdR may be a localization factor (Fig. 2). Indeed, CpdR forms a complex with ClpX and is required for the specific localization of ClpXP, and subsequently RcdA and CtrA, to the pole at specific times in the cell cycle. Consistent with previous results, localization of both ClpXP and CtrA is a prerequisite for proteolysis. Proper localization of ClpXP requires the nonphosphorylated form of CpdR, which is produced as the cell undergoes the transition from swarmer to stalked cell and in the stalked compartment of the predivisional cell. Interestingly, CpdR phosphorylation is controlled by the same phosphorelay responsible for the phosphorylation of CtrA. Following the localization of ClpXP to the appropriate pole, the ClpXP/ RcdA/CtrA complex forms and CtrA is degraded. In the predivisional cell, CtrA is stabilized following phosphorylation of CpdR by the CckA-ChpT phosphorelay, which is also responsible for the phosphorylation of CtrA (see Section 2.1.1.3). Additional research is likely to elucidate the mechanistic roles of CpdR and RcdA in restricting the proteolysis of CtrA by ClpXP to specific locations and times in the cell cycle. Localization of CtrA to the flagellar pole and proteolysis of CtrA at this pole during swarmer to stalked cell differentiation is influenced by an additional single-domain response regulator, DivK (Hung and Shapiro, 2002). DivK is required for CtrA proteolysis (Hung and Shapiro, 2002) and is localized to the same pole during the swarmer to stalked cell differentiation, but does not recruit CtrA to the pole (Jacobs et al., 2001). A signaling pathway is responsible for controlling the phosphorylation state of DivK and contributes to the regulation of CtrA proteolysis (see Section 2.4.1.2). The histidine kinases, DivJ and PleC, which are responsible for the phosphorylation and dephosphorylation of DivK, respectively, affect the degradation of CtrA in the stalked compartment of the predivisional cell (Judd et al., 2003; Matroule et al., 2004; Ryan et al., 2004). Accumulation of phosphorylated DivK prevents the phosphorylation of CpdR (Biondi et al., 2006a) (see Section 2.1.1.3 for more detail), allowing localization of ClpXP and subsequent proteolysis of CtrA. It will be of interest to determine if this signaling cascade also influences the recruitment of CtrA by RcdA. 2.1.1.3. CtrA Phosphorylation. In addition to the temporal expression of ctrA and the temporal degradation of CtrA, the activity of CtrA is modulated by the phosphorylation state of the protein. CtrA is phosphorylated in vivo and a nonphosphorylatable mutant of CtrA does not complement the lethality of a ctrA deletion when present at low copy (Quon et al., 1996), but is partially active when present at high copy

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(Domian et al., 1997). Furthermore, phosphorylated CtrA has a higher binding affinity for CtrA-regulated promoters and CtrA-binding sites in the origin of replication than nonphosphorylated CtrA (Reisenauer et al., 1999; Siam and Marczynski, 2000). Binding of phosphorylated CtrA to its binding sites in the origin of replication has been shown to promote cooperative binding of phosphorylated CtrA at adjacent binding sites (Siam and Marczynski, 2000). These results suggest that phosphorylation is required for CtrA to act as a transcriptional regulator. While several kinases have been identified as playing a role in controlling the phosphorylation state of CtrA, only the hybrid histidine kinase/response regulator CckA is required for phosphorylation of CtrA in vivo (Fig. 2) (Jacobs et al., 1999). The phosphorylation of CckA correlates with that of CtrA (Jacobs et al., 2003). Like ctrA, cckA is an essential gene. Temperature-sensitive alleles of ctrA and cckA cause remarkably similar phenotypes and result in the differential regulation of a similar set of genes, suggesting that CckA modulates the activity of CtrA (Jacobs et al., 1999, 2003). Interestingly, unlike CtrA, CckA is found at constant levels throughout the cell cycle (Jacobs et al., 1999). How does CckA modulate CtrA levels if it is always present? The key to answering this question is to examine the transient polar localization of CckA in predivisional cells. CckA localizes predominantly to the flagellar pole of predivisional cell and swarmer cells; this localization correlates with the timing of CckA and CtrA phosphorylation (Jacobs et al., 1999). A deletion of the transmembrane domain of CckA is lethal and indicates that membrane anchoring is required for the essential activity of CckA, presumably mediating the phosphorylation of CtrA. When CckA lacking the transmembrane domain is fused to green fluorescent protein (GFP) and expressed in cells containing a wild-type copy of CckA, GFP does not preferentially localize to the nascent flagellar pole of predivisional cells. Taken together, these results suggest that the proper translocation of CckA to the membrane and to the pole is required for the proper temporal activation of CtrA. In addition to the role in CtrA phosphorylation, CckA stabilizes CtrA (Jacobs et al., 2003), indicating that CckA contributes to the control of CtrA by multiple mechanisms. Does CckA phosphorylate CtrA directly or indirectly? Hybrid kinases can directly transfer the phosphoryl group to their cognate response regulators. Alternatively, a phosphorelay cascade may lead to indirect phosphorylation of the response regulator. Phosphorelay typically involves passing the phosphoryl group from the kinase domain to the receiver domain of the hybrid kinase, then onto a histidine phosphotransferase (Hpt), and finally to the cognate response regulator. In order to address this question, phosphotransfer profiling was performed using the purified kinase domain

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of CckA as the phosphoryl-donor and 50 purified response regulators as candidate phosphoryl-acceptors (Biondi et al., 2006a). CckA phosphorylates its own receiver domain as well as three other response regulators, but not CtrA, indicating that phosphorelay is likely necessary for phosphorylation of CtrA by CckA. Although no Hpt proteins had been identified in the genome sequence of C. crescentus (Nierman et al., 2001), a series of structural and functional criteria identified a candidate Hpt protein, ChpT, responsible for the phosphorelay from CckA to CtrA (Biondi et al., 2006a). Depletion of chpT generates a phenotype similar to temperature-sensitive alleles of ctrA and cckA and the gene expression patterns of all three strains are highly correlated, suggesting that ChpT is the intermediate protein required for the phosphorylation of CtrA (Biondi et al., 2006a). Biochemical evidence confirmed that ChpT acts as an Hpt and established the presence of a CckA-ChpT-CtrA phosphorelay. CckA is the only kinase capable of phosphorylating ChpT. ChpT in turn phosphorylates both CtrA and CpdR. CpdR controls the activity of the ClpXP protease, which ultimately is responsible for the degradation of CtrA (see Section 2.1.1.2). These findings explain how CckA leads to both the activation and stabilization of CtrA: phosphorelay from CckA-ChpT-CtrA activates CtrA by phosphorylation and the phosphorelay from CckA-ChpT-CpdR leads to the production of phosphorylated CpdR and prevents the localization of ClpXP and the subsequent proteolysis of CtrA. The histidine kinase DivJ and response regulator DivK have also been implicated in controlling the phosphorylation state of CtrA (Wu et al., 1998). The level of active phosphorylated CtrA is increased in a divJ deletion mutant, indicating that rather than mediating phosphorylation of CtrA, DivJ normally reduces the level of phosphorylated CtrA (Pierce et al., 2006). Suppressors of divJ, including cckA mutants, compensate for the loss of divJ by reducing the amount of CtrABP, but not total CtrA. A temperaturesensitive allele of divK grown at the nonpermissive temperature has increased levels of active CtrA, due in part to the reduction in CtrA proteolysis (Hung and Shapiro, 2002). Accumulation of phosphorylated DivK results in delocalization of CckA, preventing the phosphorylation of CtrA and CpdR (Biondi et al., 2006a). These observations are consistent with a role of DivK in reducing the levels of CtrABP by preventing CckA from mediating the phosphorylation of CtrA and by allowing unphosphorylated CpdR to accumulate, leading to the degradation of any existing pools of CtrABP. Although transcription of divK is regulated by CtrA and peaks late in the predivisional cell, DivK is a stable protein and is found throughout the cell cycle with only a slight increase in late predivisional cells (Hecht et al., 1995; Jacobs et al., 2001). Thus, the phosphorylation of DivK

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late in the cell cycle leads to the inhibition of CckA and CtrA, allowing the cell cycle to reset. The tyrosine kinase DivL has also been implicated in a signal transduction pathway leading to the phosphorylation of CtrA (see Section 2.4.1.4) (Wu et al., 1999). In vitro studies suggest that DivL phophorylates CtrA, but not DivK (Wu et al., 1999) and a conditional mutation in divL results in reduced levels of phosphorylated CtrA (Pierce et al., 2006). Taken together, these results suggest that DivL may regulate CtrA independently of the DivJK phosphorelay. This is supported by the observation that DivL does not impact CtrA proteolysis (Reisinger et al., 2007), suggesting that the mechanism of action differs from that of DivK, which utilizes the CckAChpT phosphorelay to impact both CtrA phosphorylation and proteolysis. It is equally possible that DivL functions within the DivJK pathway since a yeast two-hybrid screen identified DivL as an interacting partner for DivK, which suggests that there may be some crosstalk between the DivJK and DivL phosphorelay pathways (Ohta and Newton, 2003). Two additional histidine kinases, CckN and CckO, were identified as interacting partners for DivK in the screen, but it remains unclear if interactions between DivK and DivL, CckO, or CckN impact the phosphorylation of CtrA or even if the interactions are relevant in vivo. 2.1.1.4. The CtrA Regulon. The extensive regulation of CtrA transcription, localization, degradation, and phosphorylation, along with the essentiality of the ctrA gene, demonstrate its importance in modulating cell-cycle-regulated gene expression. The initial description of CtrA indicated that CtrA regulates promoters of the class II flagellar genes, fliQ and fliL, and the ccrM methyltranferase gene (Quon et al., 1996). FliQ and other flagellar proteins are required early in the predivisional cell when flagellum biosynthesis is initiated, whereas CcrM is required late in the predivisional cell to fully methylate newly replicated DNA. Additional studies have identified several genes that are regulated by CtrA. CtrA positively regulates the expression of the major chemotaxis operon in predivisional cells and three additional genes involved in flagellar filament production (Jones et al., 2001; Leclerc et al., 1998). In the predivisional cell, CtrA activates the transcription of pilA, which encodes the major pilin subunit and leads to the synthesis of pili on the daughter swarmer cell (Skerker and Shapiro, 2000). Transcription of the early cell division gene ftsZ is repressed by CtrA in swarmer cells (Kelly et al., 1998). In contrast, CtrA activates the transcription of the late cell division genes ftsQ and ftsA (Wortinger et al., 2000). These results suggest that CtrA influences the transcription of a diverse array of genes throughout the cell cycle.

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Microarray experiments indicate that CtrA regulates the transcription of nearly a quarter of all the cell-cycle-regulated genes (Laub et al., 2000). CtrA influences the transcription of 144 genes; however, only 55 genes (with an additional 40 genes in potential operons) are directly regulated by CtrA (Laub et al., 2000, 2002). Genes that are directly regulated by CtrA were identified by combining microarray analysis and genome-wide location analysis, which maps the in vivo CtrA-binding sites using a modification of chromatin-immunoprecipitation (Laub et al., 2002). Among the 95 genes that are directly regulated by CtrA, 29 are repressed and 66 are activated by CtrA (Laub et al., 2002). Most of the genes that are repressed are maximally expressed during swarmer to stalked cell differentiation, coincident with CtrA proteolysis. Conversely, most of the genes that are activated by CtrA are maximally expressed after the accumulation of CtrA in predivisional cells. CtrA directly regulates at least 14 regulatory genes, including operons encoding 10 two-component signal transduction proteins and two sigma factors. rpoN, which encodes the alternative sigma factor, s54, is activated by CtrA and is required for both flagellum and stalk formation, as well as proper cell division (Brun and Shapiro, 1992). sigT, which encodes another alternate sigma factor, is repressed by CtrA, leading to maximal expression of sigT during the swarmer to stalked cell transition suggesting that SigT may regulate gene expression early in the cell cycle (Laub et al., 2000, 2002). CtrA also activates additional regulators of unknown function. A number of regulatory genes, including the response regulators cheY, cheYII, cheYIII, and the histidine kinase cheA, are activated by CtrA and are known to affect polar morphogenesis (Laub et al., 2002). CtrA-dependent genes required for polar morphogenesis encode proteins required for the biosynthesis and activation of flagella, production of chemotaxis machinery, pili biogenesis, and holdfast synthesis (see Section 3) (Laub et al., 2002). Each of these processes occurs in the swarmer compartment of the predivisional cell in which CtrA binds the origin of replication, demonstrating the importance of CtrA in the formation of the nonreplicative, motile daughter swarmer cell. CtrA directly regulates essential cell processes including DNA methylation and cell division (Laub et al., 2002). The genes for DNA methylation include ccrM and metK, which encodes the S-adenosylmethionine synthetase that produces the substrate used for CcrM-dependent DNA methylation. CtrA influences the transcription of genes required for cell division initiation and progression by repressing ftsZ and activating ftsA, ftsQ, ftsW, and ftsI. The proteins encoded by these genes are part of a core set of proteins that localize to the division plane where they perform specific functions in cell division (see Section 4.3). The remaining 39 genes directly regulated by CtrA have no known function (Laub et al., 2002).

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Most of the genes positively regulated by CtrA are activated in the predivisional cell; however, the genes are not transcribed at the exact same time. For example, fliQ is activated earlier in the predivisional cell than ccrM (Reisenauer et al., 1999). The differential temporal regulation of gene expression occurs because phosphorylated CtrA has different affinities for its binding site (Reisenauer et al., 1999). Disruption of an inverted repeat sequence within the ccrM promoter, which left the CtrA-binding site intact, did not affect the timing of transcription initiation, but significantly reduced the amount of transcript indicating that an accessory factor may act as activator (Reisenauer et al., 1999). Differences in the ability of CtrA to recognize the CtrA-binding site have been attributed to specific features of the CtrA-binding site including the specific CtrA-binding site sequence, the distance between the TTAA elements, and the strength of the downstream promoter (Ouimet and Marczynski, 2000). Promoters strongly bound by CtrA contain the consensus sequence TTAA-N7-TTAA, followed by a poor match to the consensus 10 region (gCTANAWC). In this case, the transcription of the downstream gene is cellcycle regulated. When the upstream TTAA element contains mutations, there is a moderate decrease in transcription levels and transcription remains cellcycle regulated. When the spacing between the TTAA elements is changed by even a single base pair, there is a significant decrease in transcription levels and transcription remains cell-cycle regulated. In contrast, when the downstream TTAA element contains mutations, there is a drastic decrease in transcription levels and transcription is no longer cell-cycle regulated. When both TTAA elements contain mutations, the presence of a strong 10 promoter element restores cell-cycle transcription. The pilA promoter contains four CtrA-binding sites (Skerker and Shapiro, 2000) indicating that the number of CtrA-binding sites may also impact CtrA-binding affinity and affect the timing and levels of transcription. Naturally occurring promoters in C. crescentus contain mutations in the upstream TTAA element ( fliQ) and the downstream TTAA element (ccrM and fliF) indicating that differences in the CtrA-binding site contribute to the sequential activation of genes throughout the cell cycle (Ouimet and Marczynski, 2000). 2.1.2. GcrA The GcrA protein is conserved exclusively among the Alphaproteobacteria, but does not contain any known functional motifs and little is known about the mechanism of regulation by this protein (Holtzendorff et al., 2004). gcrA expression is cell-cycle controlled and GcrA is essential and serves as a second global cell-cycle regulator.

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2.1.2.1. GcrA Transcription. The pattern of gcrA transcription and subsequent protein production is strikingly out of phase with that of ctrA (Fig. 1). The maximal level of gcrA transcription occurs in the stalked cell where CtrA levels are low and gcrA transcript levels decrease in the predivisional cell as ctrA expression is reactivated (Collier et al., 2006; Holtzendorff et al., 2004). The reciprocal oscillation of CtrA and GcrA suggested that these two proteins might regulate the transcription of one another. Indeed, gcrA transcription is negatively regulated by CtrA whereas GcrA is required to activate the P1 promoter of ctrA (Holtzendorff et al., 2004). Phosphorylated CtrA binds directly to the promoter of gcrA and the level of gcrA transcription is increased in temperature-sensitive mutants of CtrA and CckA at the restrictive temperature, confirming a role for CtrA and CckA in the repression of gcrA transcription (Holtzendorff et al., 2004). Mutagenesis of the CtrA-binding site in the gcrA promoter does not eliminate the cell-cycle regulation of gcrA expression (Collier et al., 2006). This observation suggests that the proteolysis of the CtrA repressor is not the only regulatory element controlling gcrA expression. DnaA has been shown to bind the gcrA promoter and induce gcrA transcription (Collier et al., 2006; Hottes et al., 2005). Mutation of the DnaA box in the gcrA promoter or depletion of DnaA leads to a decrease in gcrA transcription (Collier et al., 2006). When the gcrA promoter is mutated such that it is either CtrA-independent or DnaA-independent, temporal regulation of gene expression is maintained (Collier et al., 2006). When the gcrA promoter is mutated to be both CtrA-independent and DnaAindependent, the temporal regulation is attenuated (Collier et al., 2006). The maximum change in the transcriptional activity throughout the cell cycle is 1.5-fold for cells with the CtrA- and DnaA-independent promoter compared with fivefold for cells with the wild-type promoter. These results indicate that the temporal regulation of gcrA transcription is directly mediated by both CtrA-dependent repression and DnaA-dependent activation. Since the cell-cycle regulation of the gcrA promoter is not completely abolished when both the CtrA and the DnaA-binding sites are compromised, additional regulatory elements may also contribute to gcrA regulation. For example, full methylation of the gcrA promoter has been proposed to partially repress gcrA transcription (Collier et al., 2006). 2.1.2.2. GcrA Proteolysis. The regulation of gcrA transcription is not the only mechanism for regulating the level of GcrA in the cells. When gcrA is transcribed constitutively in cells, GcrA abundance varies throughout the cell cycle (Collier et al., 2006). GcrA accumulates only in the stalked and predivisional cells (Fig. 1). The half-life of GcrA in stalked cells is about 40

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min whereas the half-life of GcrA in swarmer cells is about 10 min. These results suggest that GcrA is subject to cell-cycle-regulated proteolysis, but the mechanism of GcrA proteolysis remains unknown. 2.1.2.3. GcrA Regulon. Using microarray analysis, it was determined that GcrA depletion altered the gene expression of 125 genes, including 49 cellcycle-regulated genes (Holtzendorff et al., 2004). Furthermore, chromatin immunoprecipitation assays indicate that GcrA interacts with promoter sequences, including the ctrA promoter (Holtzendorff et al., 2004). GcrA could interact directly with the promoter DNA or, alternatively, could interact with a protein bound to the promoter DNA. Since GcrA lacks any detectable functional motifs, if an interaction with DNA occurs it is likely to be via a novel mechanism (Crosson et al., 2004). If GcrA does not interact directly with DNA, the mechanism of transcriptional regulation by this protein is likely to be novel. The GcrA-regulated genes are involved in a vast array of functions including motility, polar development, cell wall biogenesis, amino acid metabolism and transport, chromosome replication, repair, and recombination. Given the maximal expression of gcrA in the stalked cell, when chromosome replication is initiated, it is not surprising that GcrA regulates a number of genes involved in DNA metabolism. The regulation of gcrA transcription and the initiation of chromosome replication share a number of common features (Collier et al., 2006). Both the C. crescentus origin of replication (Cori) and the gcrA promoter are repressed by CtrA binding, activated by DnaA binding, and contain DNA methylation sites suggesting that similar mechanisms couple the initiation of gcrA transcription and the initiation of chromosome replication (Collier et al., 2006; Holtzendorff et al., 2004; Marczynski and Shapiro, 2002). This is consistent with the observation that GcrA represses genes encoding chromosome replication initiation factors but activates genes encoding proteins involved in the progression of chromosome replication and segregation (Holtzendorff et al., 2004). 2.1.3. DnaA DnaA is an essential bacterial chromosome replication initiation factor. In Escherichia coli, DnaA binds to a specific binding site (the DnaA box) in the origin of replication and unwinds the two DNA strands, allowing the replication machinery to assemble on each DNA strand (for review of this process, see Messer, 2002). In C. crescentus, DnaA binds to a DnaA box in Cori (Marczynski and Shapiro, 2002), and depletion of DnaA leads to a block in chromosome replication and cell division (Gorbatyuk and

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Marczynski, 2001). The transition from swarmer to stalked cell is not affected by the depletion of DnaA and all stalked cells become filamentous. This observation indicates that transcription and protein synthesis processes remain intact; however, transcription of a subset of genes was inhibited, which suggests that DnaA may have an additional regulatory role. Indeed, DnaA has been shown to be a global transcriptional regulator (Hottes et al., 2005) that is subject to proteolysis at specific times in the cell cycle (Fig. 1) (Gorbatyuk and Marczynski, 2005). 2.1.3.1. DnaA Transcription. Unlike ctrA and gcrA, transcription of dnaA does not appear to be dynamically regulated. Transcription of dnaA occurs throughout the cell cycle, but reaches a maximal level (twofold higher than the minimal level) just prior to the swarmer to stalked cell transition (Zweiger and Shapiro, 1994). Transcription of dnaA has been shown to be DnaA-dependent, although the absence of a DnaA box in the promoter region suggests that the autoregulation of dnaA transcription may be indirect (Hottes et al., 2005). 2.1.3.2. DnaA Proteolysis. DnaA synthesis occurs throughout the cell cycle, with a twofold increase in the swarmer cells; however, DnaA is not a stable protein and it is targeted for proteolysis during the cell cycle (Gorbatyuk and Marczynski, 2005). DnaA is subject to proteolysis by ClpP throughout the cell cycle, but proteolysis occurs twofold faster in swarmer cells. DnaA degradation is not reduced when ClpA is absent or ClpX is inactive. The presentation of DnaA to the ClpP protease likely requires an unidentified ATP-dependent chaperone or other specificity factor. The combination of DnaA synthesis and proteolysis ensures that newly synthesized DnaA is present as the cell undergoes the swarmer to stalked cell transition and initiates chromosome replication (Fig. 1). 2.1.3.3. The DnaA Regulon. In addition to its role as an initiator of chromosome replication, DnaA has been shown to be a transcriptional regulator (Hottes et al., 2005). Using microarray analysis, transcription of 40 genes expressed during the swarmer to stalked cell differentiation was shown to be DnaA-dependent. Thirteen of these genes have putative DnaA boxes in the promoter region, indicating that DnaA directly regulates them. These genes encode nucleotide biosynthesis enzymes, chromosome replication machinery components, GcrA, the polar localization factor PodJ, and the cell division protein FtsZ. Gel-shift assays demonstrated that DnaA binds to the gcrA, podJ, and ftsZ promoters confirming the role of DnaA

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as a transcriptional regulator. The dual role of DnaA as a chromosome replication initiator and transcriptional regulator of components for chromosome replication, polar development, and cell division allows the coordination of multiple processes that are necessary for proper cell-cycle progression (Hottes et al., 2005).

2.2. Sigma Factors The three global regulators described above account for regulation of roughly 30% of the cell-cycle-regulated genes, indicating that additional levels of regulation must mediate cell-cycle progression in C. crescentus. One possibility is that sigma factors may alter gene expression throughout the cell cycle. The genome sequencing of C. crescentus revealed the presence of 16 putative extracytoplasmic function sigma factors, which typically lead to changes in gene expression in response to periplasmic or extracellular stimuli (Nierman et al., 2001). Only six of the sigma factors, rpoD (s73), rpoN (s54), rpoH (s32), sigF (sF), sigT (sT), and sigU (sU) have been studied previously (Alvarez-Martinez et al., 2006, 2007; Brun and Shapiro, 1992; Malakooti and Ely, 1995; Reisenauer et al., 1996; Wu and Newton, 1996). Elucidation of the roles of the remaining sigma factors is likely to aid in understanding the complex regulation of gene expression throughout the cell cycle. 2.2.1. s73: The Principal Sigma Factor rpoD is constitutively expressed throughout the cell cycle and encodes the principal sigma factor, s73 (Malakooti and Ely, 1995). s73 recognizes a consensus promoter sequence, TTGaCgS (N10-14) GCtANAWC, which is found in the promoter region of a number of biosynthetic and housekeeping genes (Malakooti and Ely, 1995; Malakooti et al., 1995). The consensus sequence for this sigma factor has been confirmed recently (see Section 2.3; McGrath et al., 2007). s73 also recognizes E. coli s70-dependent promoters (Malakooti and Ely, 1995; Wu et al., 1997). This observation is interesting since the two sigma factors do not share a consensus sequence (Malakooti and Ely, 1995; Wu et al., 1997). Although the 35 region of s73 promoter is similar to the consensus 35 region recognized by E. coli, s70, the 10 region of the s73 promoter is not recognizably similar to 10 region of the E. coli s70 promoter. In addition, the space between the 35 and 10 regions is smaller in the s73 promoters, when compared to the s70 promoters. These observations suggest that s73 has less promoter specificity than s70.

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2.2.2. s32: The Heat Shock Sigma Factor Following heat shock, a transient increase in both rpoH transcription and s32 protein levels is observed in C. crescentus (Reisenauer et al., 1996; Wu and Newton, 1996). The rpoH promoter has two promoter elements; P1 is dependent on s73 and P2 is autoregulated by s32, which recognizes the following consensus sequence, TNNCNCCCTTGAA (Wu and Newton, 1997). Transcription from P2 increases in response to heat shock. Heat shock also influences the expression of about 20 genes, including clpXP and genes that encode molecular chaperones (Gomes et al., 1986; Osteras et al., 1999). Transcription of clpP, which encodes an essential protease, is enhanced by heat shock, whereas transcription of clpX, which encodes an ATP-dependent chaperone, is repressed by heat shock (Osteras et al., 1999). The activation of clpP transcription by s32 may explain why s32 cannot be inactivated, even at low temperatures (Osteras et al., 1999; Reisenauer et al., 1996). One of the chaperones regulated by s32, DnaK, has been shown to negatively regulate the heat shock response (da Silva et al., 2003). After heat shock, cells express a high level of DnaK and the level of s32 is transiently high; however, the heat shock proteins are not induced. This result suggests that DnaK inhibits the activity of s32 and may stimulate its degradation. It has been proposed that competition between s73 and s32 for RNA polymerase may be responsible for the down regulation of the heat shock response during the recovery phase (da Silva et al., 2003). 2.2.3. s54: The Alternative Sigma Factor for Polar Development The s54-RNA polymerase holoenzyme recognizes the consensus TGGCNCCGNCCTTGCA promoter and requires activator proteins in order initiate transcription (Brun and Shapiro, 1992; McGrath et al., 2007). In C. crescentus, the transcription of s54 is cell-cycle regulated, with an increase in expression just prior to both flagellum biosynthesis and stalk biosynthesis (Brun and Shapiro, 1992). rpoN mutants are nonmotile, stalkless, and display aberrant cell division indicating that s54 regulates genes involved in flagellum and stalk biosynthesis, as well as cell division (Brun and Shapiro, 1992). Indeed, s54 is specifically responsible for the transcription of the class III and class IV flagellar genes (Anderson et al., 1995; Fig. 3). 2.2.3.1. s54 Activators. In the absence of an activator protein, the s54-RNA polymerase holoenzyme is unable to form an open complex and transcription is not activated. s54 activator proteins typically belong to the NtrC family of response regulators and function as transcriptional

Class III

22

Class II

Class IV Filament FljP, FljK, FljL

Hook FlgE, FlgK, FlgL

L-ring

FlgH

OM

P-ring

FlgI

PG Basal Body

Distal rod FlgG E-ring FlaD Proximal rod FlgB, FlgC, FlgF FliF

IM

MS-ring C-ring/ Switch FliG, FliM, FliN

Export Apparatus FlhA, FlhB, FlhE, FliQR, FliP, FliL, FliJ

FliX

Filament Assembly

Basal Body/Hook Assembly IHF

FlbD~P σ54 CtrA~P

Class III Genes flgF operon, flgBC, flgI operon, flgK, hook operon, flbT, flaF

Class II Genes flhA, fliF operon, fliLM fliOP, fliQR, fliLJ

IHF

Class IV Genes fljP, fljK, fljL, fljL, fljM,fljN, fljO

FlbT

FlaF

FljK

PAMELA J.B. BROWN ET AL.

MS ring/Switch/Export Apparatus Assembly

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activators in the phosphorylated state. The genome sequence of C. crescentus revealed the presence of four possible activators based on homology to the highly conserved central domain of NtrC (Nierman et al., 2001). In E. coli, s54 has been shown to interact specifically with the threonine residue of the GAFTGA motif present in s54 activator proteins (Bordes et al., 2004). The two activators which have been characterized, FlbD and TacA, have the complete GAFTGA motif that is required for the interaction with s54. The remaining two possible activators do not have the complete motifs required for interacting with s54 and have not been characterized. Identifying the function of the remaining two activators and determining if they interact with s54 may further elucidate the role of the alternative sigma factor, s54, in polar development and cell division. FlbD is required for the transcription of the class III and class IV flagellar genes in the swarmer compartment of the predivisional cell (see Section 3.1.1.1; Ramakrishnan and Newton, 1990). FlbD is present throughout the cell cycle, but is phosphorylated exclusively in the predivisional cells when the class III flagellar transcription is initiated (Wingrove et al., 1993). The kinase responsible for the phosphorylation of FlbD has not been identified. In addition to activating late flagellar genes, FlbD represses early flagellum assembly genes, indicating the ability of this transcriptional regulator to affect multiple steps in flagellum assembly (Fig. 3). Mutants in flbD are not defective in stalk biosynthesis, indicating that another activator interacts with s54 to regulate stalk synthesis. Figure 3 Transcriptional regulation, translational regulation, and assembly of flagellar proteins. The presence of phosphorylated CtrA (CtrABP) leads to the transcription of the class II flagellar genes, which include flbD, which is part of the fliF operon. FlbD, a s54-dependent transcriptional activator, is repressed by FliX until assembly of the class II flagellar proteins, including the export apparatus, C-ring/switch, and MS-ring, is complete. The assembly of class II flagellar proteins is detected by FliX through an unknown mechanism and leads to the activation of FlbD by phosphorylation in a FliX-dependent manner. FlbD phosphorylation results in the transcriptional activation of the class III and class IV flagellar genes. Maximal transcription of the class III and class IV flagellar genes also requires IHF. The class III flagellar proteins are then assembled to form the basal body and hook of the flagellum. Although the class IV genes are transcribed, they are subject to posttranscriptional regulation by FlbT and FlaF, which prevents translation of at least one class IV flagellin message, fljK, until assembly of the basal body and hook is complete. Following assembly of the basal body and hook, fljK transcripts are stabilized by FlaF. The presence of FljK, the major flagellin required for motility, enables filament assembly. With the addition of the filament, flagellum assembly is complete. Solid lines depict pathways involved in transcriptional control and dashed lines indicate pathways for translational control. IM, inner membrane; PG, peptidoglycan; OM, outer membrane.

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TacA was initially identified in an effort to find s54 activators by PCR amplification of conserved domains (Marques et al., 1997). The TacA promoter contains a CtrA-binding site and the transcription of tacA is temporally regulated; maximal transcription of tacA occurs in the predivisional cell (Biondi et al., 2006b; Marques et al., 1997). Following chromosome replication initiation in the stalked cell, CtrA activates the transcription of rpoN and perhaps tacA; however CtrA binding to the tacA promoter has not been demonstrated. TacA is phosphorylated through a phosphorelay cascade (Biondi et al., 2006b). ShkA, a hybrid histidine kinase, autophosphorylates and transfers the phosphoryl group to its own receiver domain. From there, the phosphoryl group is transferred onto ShpA, an Hpt, which transfers the phosphoryl group to TacA. Phosphorylated TacA interacts with the RNA polymerase-s54 holoenzyme to enable the transcription of genes required for the regulation of stalk biogenesis, including StaR, a regulator of stalk length (see Section 3.3.3). It remains to be determined if TacA activity is spatially restricted. 2.2.4. sF: Alternative Sigma Factor for Oxidative Stress during Stationary Phase sF is not required during normal growth. However, sF is specifically required for a response to oxidative stress during stationary phase (Alvarez-Martinez et al., 2006). In response to oxidative stress during stationary phase, sF activates the transcription of eight genes, including those which encode methioine sulfoxide reductase, superoxide dismutase, and carbonic anhydrase, which are involved in protection against oxidative stress. The function of the remaining proteins in the sF regulon remains unknown. Interestingly, transcription of sigF is reduced as the cell goes into stationary phase; however, sF protein levels increase in stationary phase. In exponential cells, sF is degraded either directly or indirectly by the protease FtsH. During stationary phase, sF is less susceptible to degradation by FtsH. It has been suggested that an accessory protein may interact with sF during exponential phase and lead to increased degradation of this sigma factor (AlvarezMartinez et al., 2006). 2.2.5. sT: Alternative Sigma Factor for Osmotic and Oxidative Stress Transcription of sigT occurs specifically during the swarmer to stalked cell differentiation and is regulated by CtrA (Laub et al., 2000, 2002). Transcription of sigT is induced by growth in minimal media, heavy metal stress, and osmotic stress (Alvarez-Martinez et al., 2007; Hottes et al., 2004;

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Hu et al., 2005). sT is not essential under normal growth conditions; however, sT is essential for survival under conditions of osmotic or oxidative stress (Alvarez-Martinez et al., 2007). Interestingly, sigT transcription is not induced under oxidative stress. This observation suggests that the normal levels of sT present in the cell are sufficient for protection against oxidative stress. The sT regulon includes genes predicted to encode proteins involved in the biosynthesis or structure of the cell envelope, stress response, and electron transfer suggesting the sT is involved in mediating a general stress response and cell envelope functions. The sT regulon revealed that sT regulates its own transcription as well as two additional sigma factors, sigR and sigU. This observation indicates that sT likely initiates a regulatory cascade allowing the indirect regulation of additional genes. 2.2.6. sU: Alternative Sigma Factor which may be Involved in Stress Response The transcription of sigU is positively regulated by sT and as a result, sigU is induced by growth in minimal media, heavy metal stress, and osmotic stress (Alvarez-Martinez et al., 2007; Hottes et al., 2004; Hu et al., 2005). sU is not required for growth under normal condition or under any environmental conditions tested thus far (Alvarez-Martinez et al., 2007). It has been proposed that sT activates sigU leading to the regulation of another distinct set of genes perhaps required for resistance to an unidentified growth condition (Alvarez-Martinez et al., 2007). Determining the function of this sigma factor, as well as the remaining sigma factors predicted by the genome sequence, may help identify genes with specific functions cell survival, stress response, or perhaps in cell-cycle progression.

2.3. Promoter Architecture Using an Affymetrix array, promoter motifs were identified by searching upstream of the transcriptional start sites of 14 gene clusters comprised of genes that are coexpressed throughout the cell cycle of C. crescentus (McGrath et al., 2007). A total of 14 promoter motifs were identified within 10 different groups of coexpressed genes. Seven of the 14 promoters had been previously described including a s54-dependent promoter and a CtrAdependent promoter. The remaining seven promoter motifs are found in gene clusters that are expressed at particular times in the cell cycle. This type of analysis demonstrates that there are likely to be more unknown regulatory elements that effect cell-cycle progression. For example,

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one motif, cc_9, with a consensus sequence of GACACNNTGTCGCA, was identified upstream of a CtrA-binding site in a set of genes maximally expressed early in predivisional cells. It is likely that the element that binds to cc_9 impacts the transcription of genes involved in polar development.

2.4. Two-Component Signal Transduction Proteins In addition to global regulators and sigma factors, two component signal transduction proteins are known to dramatically impact cell-cycle progression. The genome sequence of C. crescentus revealed the presence of 105 genes encoding two-component signal transduction proteins including 34 histidine kinases, 44 response regulators, and 27 hybrid histidine kinase/ response regulators (Nierman et al., 2001). About one-third of the histidine kinases are located adjacent to response regulators suggesting that these comprise functional pairs of signal transduction proteins. While twocomponent signal transduction proteins are typically involved in mediating responses to environmental changes, a number of the two-component proteins have been shown to function in cell-cycle regulation in C. crescentus. A systematic deletion analysis has shown that at least 39 of the twocomponent genes are required for cell-cycle progression, growth, or morphology and nine of the genes are essential for cell survival (Skerker et al., 2005). Potential cell-cycle or cell-growth regulatory genes were identified from mutants with decreased motility in swarm agar and an increase in generation time. This strategy identified divJ, flbD, and tacA, which encode proteins previously known to be involved in cell-cycle progression. Two regulators, ShkA and CpdR, were identified and have since been shown to play significant roles in regulation of cell-cycle progression. A single deletion of any of seven regulators, including PhoB and six uncharacterized regulators, results in a phenotype indicative of prolonged swimming suggesting that these regulators may play a role in the swarmer to stalked cell differentiation. This observation suggests that there is much more to be learned about the complex regulation of cell-cycle progression. 2.4.1. Subcellular Localization of Signal Transduction Proteins While global regulators and sigma factors primarily contribute to the temporal regulation of gene expression, the signal transduction proteins are typically involved in the activation of cognate genes or proteins. The spatial organization of signal transduction proteins in C. crescentus plays an

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important role in preparing each compartment of the predivisional cell for the formation of distinct cell types (Fig. 2). 2.4.1.1. CckA: The Mediator of CtrA Phosphorylation. As described in Section 2.1.1.3, the histidine kinase, CckA, is necessary for the phosphorylation and activation of CtrA via the CckA-ChpT phosphorelay. In the predivisional cell, CckA is localized to the nascent swarmer pole leading to increased levels of CtrABP that block chromosome replication in the new swarmer cell (Fig. 2; Jacobs et al., 1999). Conversely, the absence of CckA in the stalked compartment of the predivisional cell prevents the phosphorylation of CtrA and allows a new round of chromosome replication to be initiated in the new stalked cell. Thus, the dynamic localization of CckA contributes to the formation of distinct daughter cells. 2.4.1.2. PleC-DivJ-DivK-PleD Multicomponent System. The histidine kinases, PleC and DivJ, are localized to opposite cell poles in the predivisional cells (Fig. 2; Wheeler and Shapiro, 1999) and have opposing actions on the response regulators, DivK and PleD. DivJ phosphorylates both DivK and PleD, whereas PleC acts as a phosphatase for DivKBP and prevents the formation of PleDBP, perhaps by acting as a phosphatase although the mechanism has not been experimentally determined (Aldridge et al., 2003; Hecht and Newton, 1995). DivJ is responsible for the dynamic localization of DivK throughout the cell cycle (Fig. 2; Lam et al., 2003). In the swarmer cell, the presence of PleC at the flagellated pole results in low levels of DivKBP, which stabilizes CckA and allows CtrABP to remain bound to Cori, thereby preventing chromosome replication in the new swarmer cell following cell division (Biondi et al., 2006a). As the cell undergoes the swarmer to stalked cell differentiation, PleC delocalizes and DivJ is synthesized and binds to the stalked pole. DivJ recruits DivK to the stalked pole; the kinase activity of DivJ causes the level of DivKBP to increase; and DivKBP localizes to the flagellar pole of the predivisonal cell (Lam et al., 2003), which leads to the delocalization of CckA and prevents the phosphorylation of CtrA (Fig. 2). The remaining pools of CtrA are dephosphorylated and degraded, allowing the initiation of chromosome replication to occur (Biondi et al., 2006a). As the predivisional cell is formed, DivKBP is targeted to both the flagellar and stalked poles (Lam et al., 2003). In the predivisional cell, PleC is localized to the swarmer pole and dephosphorylates DivK. The unphosphorylated DivK freely diffuses in the cytoplasm to the stalked pole where it is phosphorylated again by DivJ (Matroule et al., 2004). The localization and activities of both PleC and DivJ at opposite cell poles result in a rapid exchange of DivK and DivKBP at

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the cell poles and result in bipolar localization of DivKBP (Matroule et al., 2004). Following cell division, DivK is completely delocalized in the swarmer cell compartment and the swarmer to stalked cell transition begins (Matroule et al., 2004). The difference in the level of DivKBP in each of the daughter cells contributes to the distinct physiologies of the new stalked and swarmer cells. Localization of DivJ and PleC also impact the phosphorylation state and activity of PleD, a response regulator that contains a diguanylate cyclase domain. PleDBP is required for ejection of the flagellum and elongation of the stalk during the swarmer to stalked cell differentiation and for preventing flagellum activation in the predivisional cell (Aldridge and Jenal, 1999; see Section 3.1.4; Fig. 4). The presence of PleC in the new swarmer cell prevents the phosphorylation of PleD and premature ejection of the flagellum. As the cell undergoes the swarmer to stalked cell transition, DivJ is localized to the incipient stalked pole allowing the phosphorylation of PleD, which is then sequestered at the pole, resulting in subsequent ejection of the flagella and elongation of the stalk (Aldridge et al., 2003; Paul et al., 2004). Proper temporal control of holdfast synthesis during the swarmer to stalked cell differentiation is also dependent on PleDBP (Levi and Jenal, 2006). PleDBP has di-guanylate cyclase activity capable of converting two molecules of guanosine triphosphate (GTP) into cyclic-diguanosine monophosphate (c-di-GMP) (Paul et al., 2004). Since activated PleDBP is found only at the stalked pole, and c-di-GMP can serve as a second messenger, it has been proposed that production of c-di-GMP may serve as a signal for polar development (Jenal and Malone, 2006). The structure of PleD suggests that the efficient production of c-di-GMP requires PleD dimerization and that feedback inhibition by the product is likely to limit the concentration of c-di-GMP in the cell (Chan et al., 2004; Wassmann et al., 2007). Indeed, biochemical analyses have shown that PleD is activated by dimerization and that dimerization occurs when the protein is phosphorylated (Paul et al., 2007). It will be of particular interest to determine the specific role of c-di-GMP signaling in the development of the stalked pole in C. crescentus. Recent work has demonstrated that DivK acts as an allosteric response regulator to determine cell fate (Paul et al., 2008). During the swarmer to stalked cell transition, DivK activates the kinase activity of both DivJ and PleC. The activation of DivJ results in a positive feedback loop leading to stimulation of DivK kinase activity and polar localization of DivK. In addition, DivK switches PleC from a phosphatase into an autokinase leading to the phosphorylation of PleD and the accumulation c-di-GMP. Remarkably, DivK modulates the activity of both PleC and DivJ despite spatial separation of these kinases.

PleD

Swarmer compartment

Stalk to predivisional cell differentiation

PleD~P TipN

c-di-GMP

TipF

pGpG

c-di-GMP

DgrA

DgrA-c-di-GMP

TipF

SW Flagellum activation

Flagellum assembly

Flagellum ejection Holdfast formation ST Stalk formation

SW

GTP

PleD~P

Flagellum assembly PDE

PDL

ST c-di-GMP PleD~P

c-di-GMP

GTP

Swarmer to stalk cell differentiation

Stalked compartment

Figure 4 Cyclic-di-GMP signaling throughout the C. crescentus cell cycle. Each stage of the cell cycle, including the swarmer (SW), stalked (ST), early predivisional (PDE), late predivisional (PDL), and progeny cells, is shown. In the swarmer cell PleD is delocalized. As the cell undergoes the swarmer to stalked cell differentiation, dimers of PleDBP are localized to the flagellar pole leading to the production of cyclic-di-GMP. The increase in c-di-GMP levels promotes flagellum ejection, holdfast formation, and stalk formation. Flagellum assembly in the early predivisional cell is achieved by a reduction in c-di-GMP levels through degradation into linear diguanylate (pGpG) by TipF and occurs at the pole marked by the presence of TipN. After flagellum biosynthesis, TipN is briefly delocalized. In the late predivisional cell, just prior to cell division, flagellum activation occurs as c-di-GMP levels decrease due to binding by proteins such as DgrA. TipF is localized to the pole opposite the flagellum throughout the cell cycle, until the onset of cell division, when both TipN and TipF localize to the mid-cell and mark the new pole following cell separation.

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2.4.1.3. PodJ: The Localization Factor. Localization is a critical element for spatial regulation by some two-component systems. Little is known about how CckA and DivJ are localized to the stalked pole, but a localization factor, PodJ, has been identified for PleC (Hinz et al., 2003; Viollier et al., 2002b). PodJ is subject to complex regulation throughout the cell cycle and impacts swarming motility and the formation of pili and holdfast. podJ transcription is repressed by CtrA, activated directly by DnaA, and activated, either directly or indirectly, by GcrA (Crymes et al., 1999; Holtzendorff et al., 2004; Hottes et al., 2005). As a result of this transcriptional regulation, podJ is transcribed following the swarmer to stalked cell differentiation and full length PodJ (PodJL) is then localized to the incipient flagellar pole of the predivisional cell (Hinz et al., 2003; Viollier et al., 2002b). PodJL is required for the localization of both PleC and CpaE, a protein required for pili assembly, to the incipient flagellar pole (Fig. 2; Hinz et al., 2003; Viollier et al., 2002b). Following cell division, PodJL, is processed into a short form (PodJS) by the periplasmic protease, PerP (Chen et al., 2006). PodJS remains localized at the flagellar pole following cell division and is required for chemotaxis and holdfast formation, presumably due to a role in localizing proteins required for these functions (Fig. 2; Hinz et al., 2003; Lawler et al., 2006; Viollier et al., 2002b). During the swarmer to stalked cell transition, PodJS is cleaved from the membrane by the protease MmpA and degraded completely by an unknown protease prior to the synthesis of new PodJL (Chen et al., 2005). How can PodJ influence pili formation, holdfast formation, swarming motility, and PleC localization? PodJ has been shown to be a modular protein with functional domains that are important for specific functions (Lawler et al., 2006). PodJ has a single transmembrane domain with the N-terminus in the cytoplasm and the C-terminus in the periplasm (Viollier et al., 2002b). The cytoplasmic portion of PodJ contains three coiled-coil domains that are typically involved in protein–protein interactions and has been shown to be required for holdfast production and swarming motility (Crymes et al., 1999; Lawler et al., 2006; Smith et al., 2003). The periplasmic portion of PodJ is required for pili formation and contains three tetratricopeptide repeats, which are also involved in mediating protein– protein interactions, and a peptidoglycan-binding domain (Crymes et al., 1999; Lawler et al., 2006; Viollier et al., 2002b). The region immediately preceding the transmembrane domain is necessary for the localization of PodJ to the correct pole throughout the cell cycle, whereas the region immediately following the transmembrane domain is required for localization of PleC (Lawler et al., 2006). Some PodJ mutants that are not able to localize PleC can still produce holdfast, indicating that PleC does not have

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to localize to be active. In contrast, it has been suggested that PodJ localization is required for proper function (Lawler et al., 2006). The necessity of polar localization for proper function is likely to be a common feature for some of the cell-cycle-regulated proteins in C. crescentus. 2.4.1.4. DivL: An Essential Tyrosine Kinase. DivL, a tyrosine kinase, is required for cell survival and impacts cell-cycle progression of C. crescentus (Wu et al., 1999). DivL autophosphorylates on a tyrosine residue and is capable of passing the phosphoryl group to CtrA in vitro (Wu et al., 1999). Reduced levels of phosphorylated CtrA are observed in conditional mutants of divL (Pierce et al., 2006). Together these results indicate that DivL is involved in the activation of CtrA. DivL is dynamically localized throughout the cell cycle (Sciochetti et al., 2005). Following cell division, DivL is absent from both the swarmer cell and the stalked cell. Shortly after cell division, DivL appears in the stalked cell and is localized to the pole opposite the stalk. The swarmer cell does not acquire DivL until after differentiation into the stalked cell. As the predivisional cell is formed, DivL remains localized at the incipient flagellar pole and is occasionally observed at the stalked pole. The polar foci of DivL are dispersed late in the predivisional cell, prior to cell division. The polar localization of DivL is dependent, either directly or indirectly, on the presence of DivJ. However, the kinase activity of DivL is not dependent on its polar localization (Sciochetti et al., 2005). Interestingly, it is not clear if the kinase activity of DivL is important for its function in cell-cycle progression. Mutation of the tyrosine residue in DivL results in a phenotype that is much less severe than DivL depletion, suggesting that DivL has a kinase-independent function in cell-cycle regulation (Reisinger et al., 2007). DivL affects CtrA phosphorylation but not proteolysis suggesting that DivL acts independently from the CckAChpT phosphorelay. One model is that DivL may protect CtrABP rather than acting as a CtrA kinase (Reisinger et al., 2007). Irrespective of the mechanism of promoting CtrA phophorylation, DivL has another function in cell-cycle control since deletion of divL in a strain expressing a phosphomimetic ctrA allele results in defects in chromosome replication and cell division. DivL has also been shown to modulate the localization and phosphorylation of DivK (Reisinger et al., 2007). DivL may promote DivK localization by regulating DivJ localization and kinase activity or by direct interactions with DivK at the flagellar pole, or both. Thus, kinaseindependent activities of DivL are likely to be responsible for the modulation of both CtrA and DivK. It remains to be determined if the kinase-independent activities of DivL are dependent on its localization.

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3. POLAR STRUCTURE BIOGENESIS AND FUNCTION The regulatory mechanisms described above function primarily to impart polarity to the predivisional cell and allow the formation of two different types of daughter cells – the swarmer cell and the stalked cell. The polar structures found on each cell type are distinct; the swarmer cell contains a flagellum and pili whereas the stalked cell has an extension of the cell membranes and peptidoglycan layer called a stalk or prostheca and a polysaccharide containing adhesin at the tip of the stalk called a holdfast. The holdfast is found in both swarmer and stalked cells, but primarily serves to attach the stalked and predivisional cell to a surface. The role and biosynthesis of each of the polar structures is discussed below.

3.1. The Flagellum C. crescentus produces a single polar flagellum that is responsible for fast and efficient swimming in swarmer cells (Li and Tang, 2006). The regulation of flagellum biosynthesis and assembly has been studied extensively and has been the subject of a number of recent reviews (Aldridge and Hughes, 2002; England and Gober, 2001; Gober and England, 2000; Jenal, 2000). The following sections focus on the most recent advances that have enhanced our understanding of the most characterized aspect of C. crescentus polar development, flagellum biogenesis. 3.1.1. Biosynthesis and Assembly of Flagella The biosynthesis of flagella is a complex process, requiring a vast array of structural, regulatory, and force-generating proteins. In C. crescentus, this process is temporally and spatially constrained by a number of molecular mechanisms including both transcriptional and translational regulation. 3.1.1.1. Transcriptional and Translational Regulation of Flagellar Genes. The temporal transcriptional regulation of about 50 flagellar genes is mediated by a complex hierarchy that is tied to both cell-cycle progression and assembly of the flagellum (Fig. 3; for review see Gober and England, 2000). CtrABP is responsible for the transcription of the class II flagellar genes in stalked cells following the initiation of chromosome replication. The class II flagellar genes encode the MS-ring, flagellar switch, and flagellum export apparatus, which are the first flagellar components to be assembled. Class II flagellar genes also

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encode regulatory proteins including the transcriptional activator FlbD, the trans-acting factor FliX, and s54, which impact the transcription of the class III and class IV flagellar genes. Maximal transcription of a subset of class III and class IV flagellar genes also requires the sequence-specific binding protein integration host factor (IHF) (Muir and Gober, 2005). In E. coli, binding of IHF induces a bend into DNA which brings RNA polymerase bound to the promoter into close proximity with transcriptional activators bound at distant site (Hoover et al., 1990). It has been proposed that IHF-induced bending of DNA may enhance the interaction of FlbD with other molecules of FlbD to promote oligomerization, which is thought to enhance the ability of FlbD to isomerize RNA polymerase from a closed complex to an open complex (Muir and Gober, 2005). The transcription of the class III and class IV flagellar genes does not occur until assembly of the class II flagellar proteins is complete (Fig. 3). Prior to class II protein assembly, the trans-acting factor FliX interacts directly with FlbD, preventing FlbD from binding to enhancer sequences and subsequent s54-dependent transcription of the class III and class IV flagellar genes (Dutton et al., 2005; Muir and Gober, 2001, 2002, 2004; Muir et al., 2001). Once assembly of the class II flagellar proteins is complete, FliX inhibition of FlbD is relieved, FlbD is phosphorylated by an unknown kinase, and transcription of the class III and class IV flagellar genes is initiated (Muir and Gober, 2001, 2002; Muir et al., 2001; Wingrove et al., 1993). It remains unclear how FliX senses the completion of assembly. The class III genes encode structural components of the flagella including the basal body, the L-ring, and the P-ring. The assembly of the class III structural components into the basal body hook complex requires the lytic transglycosylase, PleA (Viollier and Shapiro, 2003). pleA mutants are nonmotile and lack flagella. Thus, it has been proposed that PleA is responsible for the hydrolysis of peptidoglycan at the swarmer pole, allowing the penetration of structural components of the flagellum through the peptidoglycan layer. In addition to structural components of the basal body hook complex, the class III flagellar genes include flbT, which encodes a regulatory protein responsible for translational control of the class IV flagellar genes encoding the flagellins (Anderson and Gober, 2000; Mangan et al., 1999). In mutants unable to assemble the hook complex, flagellin mRNA is present at low levels due to messenger instability. When a flbT mutation is introduced into a strain which cannot assemble the hook complex, flagellin mRNA is stabilized indicating that FlbT is a negative regulator of flagellin expression (Mangan et al., 1999). The mechanism of FlbT repression of flagellin gene

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expression was explored using the fljK transcript, which encodes one of the major flagellins (Anderson and Gober, 2000). Studies using translational fusions revealed that only 63 nucleotides of the 5u untranslated region and the first 14 codons of fljK mRNA are necessary for FlbT-mediated repression. This region of the fljK mRNA is predicted to fold into two different secondary structures, including one structure that is likely to be unfavorable for translation since the ribosome-binding site is blocked by base pairing. Using cell extracts, FlbT was shown to bind to a loop structure of the fljK mRNA, likely stabilizing the secondary structure that prevents translation. Based on these observations, it has been suggested that in the absence of a complete basal body hook complex, FlbT binds to flagellin mRNA and blocks translation leading to destabilization of the flagellin mRNA (Fig. 3; Anderson and Gober, 2000). Following the assembly of the class III flagellar proteins into the basal body hook structure, FlbT repression is relieved by an unknown mechanism and fljK transcripts are stabilized leading to translation of the flagellins and assembly of the filament (Fig. 3). It remains unclear how the completion of basal body hook assembly is detected by FlbT. The stabilization of flagellin mRNA requires FlaF, a protein required for motility. The flaF gene is directly downstream from the flbT gene and is required for fljK transcript stabilization (Llewellyn et al., 2005). In the absence of flaF, the basal body hook complex is assembled, but flagellin mRNAs are not translated. This observation indicates that FlaF is involved in the transition between completion of the basal body hook complex assembly and the initiation of flagellin translation and filament assembly. It has been proposed that throughout the cell cycle, the regulation of flagellin translation and filament assembly is controlled by the opposing actions of FlbT, which destabilizes flagellin mRNA, and FlaF, which stabilizes flagellin mRNA (Llewellyn et al., 2005). Both flbT and flaF are conserved among swimming bacteria belonging to the Alphaproteobacteria indicating that this regulatory mechanism, which delays flagellin translation and filament assembly until after the completion of the basal body hook complex assembly, may constitute a conserved checkpoint in flagellum assembly. 3.1.1.2. Spatial Control of Flagellum Assembly. How is the location of flagellum assembly determined? To answer that question, it is necessary to first consider how polarity is determined in the cell (Fig. 4). TipN, a ‘‘birth scar’’ protein, marks polarity in new daughter cells (Huitema et al., 2006; Lam et al., 2006). TipN is found exclusively at the new pole in both the swarmer and stalked cells and marks the site of future flagellum assembly. TipN remains at the new pole as the flagellum is assembled in the

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predivisional cell. Following the biosynthesis of the flagellum, TipN is briefly delocalized before localizing to the division site (Huitema et al., 2006; Lam et al., 2006). Following cell division, TipN localization serves as a molecular beacon, marking the new pole of the daughter cell. TipN is required for proper localization of TipF, a c-di-GMP phosphodiesterase protein required for flagellum assembly (Huitema et al., 2006). In cells lacking TipN, the flagellum is assembled, but is frequently misplaced; however, in cells lacking TipF, the flagellum fails to assemble. These results suggest that presence of TipF, but not its localization to the pole, is required for flagellum assembly and that the interaction of TipN and TipF is responsible for assembly of the flagellum specifically at the new pole. Mutational analysis indicates that the TipF phosphodiesterase domain, which likely degrades c-di-GMP, is required for its function in flagellum assembly (Huitema et al., 2006). This result suggests that reducing the level of c-di-GMP may facilitate flagellum assembly. 3.1.2. Flagellum Biosynthesis is Coupled to Cell Division Assembly of the class II flagellar structures is required for proper cell division; mutants in class II flagellar genes result in the formation of filamentous cells (Gober et al., 1995; Yu and Shapiro, 1992; Zhuang and Shapiro, 1995). Such a checkpoint is logical, delaying cell division until the assembly of the flagellum has initiated on the swarmer pole of the predivisional cell ensures that the swarmer cell will have a functional flagellum. Two links between flagellum biogenesis and cell division have recently been uncovered. First, the regulation of FlbD activity by FliX influences the transcription of late flagellar genes and the completion of cell division (Muir et al., 2005). Negative regulation of FlbD activity by FliX is responsible for the cell division defect in class II flagellar genes. Identifying the genes activated by s54 and FlbD will reveal if FlbD is required for the transcription of any cell division genes. Second, once the predivisional cell is mature and the flagellum has been assembled, TipN and then TipF are localized to the division site (Huitema et al., 2006; Lam et al., 2006). The localization of TipN and TipF at the division site depends on FtsZ ring formation and constriction suggesting that these proteins may also have a role in cell division (Huitema et al., 2006; Lam et al., 2006). 3.1.3. Flagellum Activation Cell division defects result in nonmotile cells that have an assembled flagellum indicating that assembly is not sufficient for flagellum rotation and

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function (Degnen and Newton, 1972; Muir et al., 2005; Ohta et al., 1997; Quardokus and Brun, 2003). FliL is required for flagellum rotation, but is not part of the flagellar transcriptional hierarchy and is not a component of the assembled flagellum (Jenal et al., 1994). The mechanism of motor control by FliL remains unknown, but recent results suggest that the level of the second messenger, c-di-GMP, may modulate motility and specifically reduce the levels of FliL (Fig. 4; Aldridge et al., 2003; Christen et al., 2007). Motility is blocked in C. crescentus cells with an increased level of c-di-GMP (Christen et al., 2007). The block in motility is caused by a loss of flagellum function and can be bypassed in a dgrA mutant (Christen et al., 2007). DgrA is a member of the diguanylate receptor proteins and has a high affinity for binding c-di-GMP. Cells lacking DgrA are motile in the presence of high levels of c-di-GMP and have reduced levels of FliL. However, the levels of other motility proteins including FliM, which is cotranscribed with FliL, are not affected. This result indicates that changes in FliL protein levels are likely to be due to translational regulation or decreased protein stability. The simplest model to explain these results is that when DgrA is bound to c-di-GMP, FliL is repressed and cannot promote flagellum rotation (Fig. 4). 3.1.4. Ejection of the Flagellum During the swarmer to stalked cell transition, the flagellum is ejected prior to the formation and elongation of the stalk. The ejected flagellum is largely intact, containing the filament, the hook, and part of the rod, indicating that the breakpoint occurs at the junction between the MS-ring and the rod (Kanbe et al., 2005). Indeed, flagellum ejection coincides with the degradation of FliF, which forms the MS-ring that anchors the axial components of the flagellum to the cell (Aldridge and Jenal, 1999). Biochemical evidence suggests that the MS-ring, but not the rod, of C. crescentus flagella is particularly susceptible to protease activity (Kanbe et al., 2005). The ClpAP protease recognizes the hydrophobic tail of FliF and is responsible for the degradation of FliF during the swarmer to stalked cell differentiation (Grunenfelder et al., 2003, 2004). The response regulator, PleD, is also involved in the regulation of flagellum ejection. In a pleD mutant, FilF is stabilized and the flagellum is not ejected (Aldridge and Jenal, 1999). Furthermore, the GGDEF (Gly-Gly-Asp-Glu-Phe) output domain of PleD is responsible for the production of c-di-GMP and is required for flagellum ejection (Fig. 4; Aldridge and Jenal, 1999; Paul et al., 2004). This result suggests that c-di-GMP signaling mediated by PleD is responsible for the degradation of the MS-ring.

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Notably, the only flagellar protein required for efficient removal of FliF prior to flagellum ejection is FliL. FliL is known to be modulated by DgrA when it is bound to c-di-GMP (Aldridge and Jenal, 1999; Christen et al., 2007). The mechanism by which PleD and c-di-GMP trigger the degradation of FliF remains unknown. Determining the interactions among proteins that generate, bind, and degrade c-di-GMP and a subset of flagellar proteins will lead to a greater understanding of the temporal regulation of flagellar assembly, activation, and ejection. 3.1.5. Function of the Flagellum C. crescentus lives in oligotrophic aquatic environments and the ability to swim is certainly important for tracking nutrients and cell survival. Despite structural similarities between the flagella of E. coli and C. crescentus, C. crescentus swims nearly 10 times more efficiently than E. coli (Li and Tang, 2006). The increase in swimming efficiency may reflect an adaptation to low nutrient environments (Li and Tang, 2006). The flagellum of C. crescentus also plays a role in the establishment and escape from biofilms under hydrodynamic conditions (Entcheva-Dimitrov and Spormann, 2004). C. crescentus forms biphasic biofilms consisting of a monolayer biofilm interspersed with large mushroom-like structures. Swimming motility mediated by the flagellum enhances the initial attachment event leading to eventual biofilm formation (Fig. 5; Bodenmiller et al., 2004; Entcheva-Dimitrov and Spormann, 2004). In a monolayer biofilm formed under hydrodynamic conditions, cell division produces new swarmer cells that swim away from the established biofilm (Entcheva-Dimitrov and Spormann, 2004). The release of swarmer cells from the biofilm can be viewed as a dispersal mechanism, allowing the establishment of new biofilms in other locations. In a mushroom-like biofilm structure, which is comprised primarily of clonal growth arising from established microcolonies, the flagellum is involved in retaining new swarmer cells within the threedimensional structure and may function as an adhesin (Entcheva-Dimitrov and Spormann, 2004). The dual function of the flagellum in the two types of biofilm formations is intriguing and suggests that the role of the flagellum in C. crescentus may not be confined to cell motility.

3.2. The Pili Immediately after cell division, Flp (fimbrial low-weight protein) pili are synthesized at the flagellar pole of the swarmer cell. Like the flagellum, the

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Flagellum A F B Pili

C

D

E

Stalk Holdfast

Figure 5 Attachment of a C. crescentus cell to a surface. The stages of initial attachment are shown, A–F. The surface for attachment is shown as a gray box and the polar structures are labeled. Contact with the surface is mediated by flagellar motility, which overcomes repulsive forces (A–B). The flagellum enhances the initial attachment of C. crescentus to the surface (C). Following flagellum ejection, the interaction of the pili with the surface may properly position the cell prior to a more permanent attachment (D). As the swarmer cell differentiates into a stalk cell, the holdfast is exported and the pili are hypothesized to retract (E). The biosynthesis of the stalk brings the holdfast in contact with the surface (F). The holdfast is responsible for the remarkably strong adhesion of C. crescentus to surfaces.

biosynthesis of the pili is temporally and spatially restricted and is subject to transcriptional regulation. Despite the similarity in the timing and location of flagellum and pilus biogenesis, these two processes share few regulatory elements. Whereas the flagellar assembly machinery is similar to type III secretion machinery, the pilus assembly machinery more closely resembles the type II Tad (tight adherence) macromolecular transport system, indicating that these two structures are assembled independently from one another (Skerker and Shapiro, 2000; Tomich et al., 2007). In this section, an overview of the most recent advances regarding the regulation, assembly, and function of the pili is provided. 3.2.1. Transcription of the Pili Genes DNA microarray experiments have shown that the genes required for pilus biosynthesis are maximally transcribed just prior to the assembly of the pili at the flagellar pole of the swarmer cell (Laub et al., 2000). First the genes encoding the pilus secretion machinery, cpaBCDEF, are transcribed and then the cpaA gene that encodes the prepilin peptidase is transcribed. Transcription of the pilA gene, which encodes the major protein subunit in the pilus filament, occurs last. The transcription of pilA is under the control of CtrA, linking pilus biogenesis to cell-cycle progression

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(Skerker and Shapiro, 2000). When the transcription of pilA is under the control of a constitutive promoter, pili are assembled prematurely in predivisional cells. This observation indicates that the temporal regulation of pilA transcription by CtrA prevents premature pilus biogenesis. 3.2.2. Assembly of the Pili How are the pili assembled at the proper time and place? PodJL binds to the future site of pilus biogenesis and serves as a localization factor for the pilus assembly protein, CpaE (Viollier et al., 2002b). The presence of CpaE at the incipient swarmer cell pole triggers the localization of the pilus secretion channel, CpaC, to the same pole (Viollier et al., 2002a). The localization of CpaC is required for pore formation and subsequent pilus assembly. The formation of the CpaC secretion channel depends on the presence of PleA, a transglycosylase, indicating that localized peptidoglycan hydrolysis is required for the formation of the channel (Viollier and Shapiro, 2003). This is consistent with the observation that pleA mutants lack pili. Following cell division, PilA accumulates and is polymerized to form the filament, which is secreted through the CpaC pore, thus completing pilus biogenesis (Skerker and Shapiro, 2000; Viollier et al., 2002a). 3.2.3. Function of the Pili The Flp pili enhance the initial binding of C. crescentus cells to a surface (Fig. 5; Bodenmiller et al., 2004; Entcheva-Dimitrov and Spormann, 2004). Although, retraction of Flp pili has never been demonstrated in any bacterium, it is presumed that the C. crescentus pili retract, since pili or pilin does not accumulate in the media following loss of pili from the cell surface (Lagenaur and Agabian, 1977). Further support for pili retraction is provided by the observation that shortly after the addition of phage, the phage can be visualized along the pili by TEM; however when more time is allowed, the phage are observed only at the cell pole, suggesting that the pili have retracted (Skerker and Shapiro, 2000). As the swarmer cell differentiates into a stalked cell, retraction of the pili is followed by the synthesis of the stalk with the adhesive holdfast at the tip. It has been proposed that the initial attachment mediated by the pili may properly position the cell and allow a more permanent surface attachment following contact between the holdfast and the surface (Bodenmiller et al., 2004; Janakiraman and Brun, 1999). The pili are also involved in the maintenance of complex biofilms (Bodenmiller et al., 2004; Entcheva-Dimitrov and Spormann, 2004).

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On coverslips, cpaA mutants fail to form dense monolayer biofilms compared to wild-type cells after overnight incubation (Bodenmiller et al., 2004). Under hydrodynamic conditions, a deletion mutant lacking genes for pilus assembly and for the pilin subunits forms loose microcolonies, but does not develop the dense mushroom-like structures formed by wild-type cells within 96 hours (Entcheva-Dimitrov and Spormann, 2004). These results suggest that pili may function in mediating cell-to-cell contact and function in biofilm maturation.

3.3. The Stalk The synthesis of a polar stalk as the swarmer cell differentiates to form a stalked cell results in a striking change in cellular morphology. Following flagellum ejection, stalk biosynthesis occurs at the same pole, culminating in the extension of a stalk one-fifth of the diameter of the cell body. The synthesis of the stalk is temporally and spatially constrained, indicating that this process must be regulated throughout the cell cycle. In this section, the structure, biosynthesis, regulation, and function of the stalks of C. crescentus will be discussed. 3.3.1. Structure of the Stalk The stalk is a true extension of the cell envelope, containing the inner and outer membranes, peptidoglycan, and periplasmic space, but is seemingly devoid of DNA, ribosomes, and cytoplasmic material (Ireland et al., 2002; Poindexter and Bazire, 1964; Wagner et al., 2006). The stalk contains crossbands, intermittent electron dense rings perpendicular to the long axis, which appear to be synthesized once during the cell cycle and can be used to determine the age of a stalked cell (Poindexter and Staley, 1996). It has been proposed that crossbands compartmentalize or stabilize the stalk (Jones and Schmidt, 1973). 3.3.2. Stalk Biosynthesis Stalk elongation occurs as new cell envelope material is incorporated at the junction between the cell body and the stalk (Aaron et al., 2007; Schmidt and Stanier, 1966; Seitz and Brun, 1998; Smit and Agabian, 1982). The biosynthesis and elongation of the stalk is dependent on at least three proteins, which are also required for elongation of the cell body. Depletion or inactivation of penicillin-binding protein 2 (PBP2), RodA, or MreB leads to

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defects in both cell body elongation and stalk elongation (Divakaruni et al., 2007; Seitz and Brun, 1998; Wagner et al., 2005). RodA or MreB depletion generates bulbous or short wide stalks at the proper pole (see Section 4.1.1; Fig. 8; Wagner et al., 2005). Cells depleted, and then repleted for RodA and MreB have multiple mislocalized stalks indicating the presence of ectopic poles (Wagner et al., 2005). Similar results were obtained when cells were briefly exposed to the PBP2 inhibitor mecillinam. The phenotype of multiple mislocalized stalks demonstrates that PBP2, RodA, and MreB contribute to cell polarity and stalk localization (Wagner et al., 2005). The FtsH protease is also required for stalk elongation, but is not required for cell body elongation (Fischer et al., 2002). Depletion of ftsH leads to filamentous cells with either no stalk or with short stalks at the proper pole. Some of the short stalks contain up to four adjacent crossbands indicating a defect in stalk elongation. The role of FtsH in stalk elongation has not been elucidated; however, FtsH does not appear to modulate the activity of s54. One possibility is that FtsH may control the activity of one or more of the s54 activators. The observation that depletion of FtsH results in filamentous cells indicates that FtsH directly or indirectly contributes to cell division. Proper stalk biogenesis also requires the cell division protein FtsZ, which is responsible for the formation of crossbands in the stalk (Divakaruni et al., 2007; see Section 4.2.2 for a description of FtsZ). FtsZ controls peptidoglycan synthesis at the division site; however, the requirement of FtsZ for crossband synthesis indicates that FtsZ may be required for multiple forms of peptidoglycan synthesis. FtsH and FtsZ impact both stalk biosynthesis and cell division indicating that the two processes may be coupled. Since the stalk is found only at a specific location and is considerably narrower than the cell body, there must be stalk-specific proteins or mechanisms that determine the location and size of the stalk. Thus far, all mutations that impact stalk biosynthesis are pleiotropic in nature and often also impact cell division. Identification and characterization of proteins that are required for stalk biosynthesis specifically will illuminate the processes which allow a local extension of the cell envelope. Since stalks are not required for cell survival, the study of stalk biosynthesis is an extremely tractable experimental system. The study of stalk biosynthesis is expected to reveal basic principles that apply to other morphological changes (Wagner and Brun, 2007). 3.3.3. Regulation of Biogenesis and Stalk Length Mutations in a number of regulators involved in cell-cycle progression, including CtrA, CckA, PleC, PleD, s54, and TacA, result in defects in stalk

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biogenesis (Aldridge and Jenal, 1999; Biondi et al., 2006b; Brun and Shapiro, 1992; Jacobs et al., 2003, 1999; Quon et al., 1996; Wang et al., 1993). The exact role of CckA and PleC in mediating stalk biosynthesis is unclear. One possibility is that the stalk defect of CckA and PleC is indirect and related to the role of these proteins in modulating CtrA activity, which is responsible for coupling stalk biogenesis to cell-cycle progression. CtrA is responsible for the precisely timed transcription of rpoN and tacA (Laub et al., 2000, 2002). TacA is activated by phosphorylation and then binds to enhancer sequences, enabling s54-dependent gene expression for stalk biogenesis (Biondi et al., 2006b). A phosphorelay cascade involving the hybrid histidine kinase, ShkA, and the Hpt protein, ShpA, mediates the activation of TacA. The signal responsible for initiating the ShkA-ShpATacA phosphorelay remains unknown. Microarray analysis identified 30 candidate genes downstream of TacA (Biondi et al., 2006b). One of the TacA targets, staR, is predicted to encode a putative transcriptional factor of the Cro/CI family, suggesting that a transcriptional cascade may control stalk biosynthesis (Biondi et al., 2006b). Although deletion of staR does not eliminate stalks, the stalks are considerably shorter than the stalks of wild-type cells. Determination of which genes are regulated by StaR may lead to increased understanding of how stalk length is regulated. Since a tacA deletion generates stalkless cells and a staR deletion does not, it is likely that additional downstream targets of TacA contribute to the control of stalk biosynthesis. While stalk biogenesis seems to be controlled largely by cues mediating cell-cycle progression, stalk length is known to be influenced by the extracellular environment of the cell. In the natural, nutrient-depleted environment of C. crescentus, the stalks are remarkably long, up to 20 times the length of the cell body (Brun and Janakiraman, 2000; Poindexter, 1964). Limitation of organic phosphate or inactivation of the genes required for phosphate uptake leads to a dramatic increase in the rate of stalk elongation (Gonin et al., 2000; Schmidt and Stanier, 1966). By analogy with work in E. coli (Wanner, 1996), a model for the role of the Pho regulon in phosphate uptake and stalk elongation of C. crescentus has been proposed (Gonin et al., 2000). The PstSCAB proteins comprise a high-affinity phosphate transporter that controls the autophosphorylation of the histidine kinase, PhoR. When phosphate is in excess, PhoR remains unphosphorylated. Under phosphate-limiting conditions, PhoR is released from the Pst complex, autophosphorylates, and then transfers the phosphoryl group to PhoB, its cognate response regulator. Phosphorylated PhoB then directly or indirectly activates the expression of genes required for stalk elongation.

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3.3.4. Function of the Stalk A number of possible functions have been attributed to the stalk of C. crescentus, including adhesion and nutrient acquisition. The localization of the adhesive holdfast to the distal tip of the stalk is often taken as an indication that the stalk may play a role in adhesion. Several lines of evidence indicate that there is an adhesion-independent function for the stalk. First, stalk and holdfast synthesis are not coupled. Stalk synthesis occurs after holdfast synthesis and does not depend on the presence of the holdfast (Bodenmiller et al., 2004; Levi and Jenal, 2006). Similarly, holdfast synthesis does not depend on stalk synthesis (Janakiraman and Brun, 1999). Second, in closely related organisms such as Asticaccaulis excentricus and A. biprosthecum, the holdfast is associated with the cell body and not the stalk (Poindexter, 1964). Lastly, the swarmer cell, which lacks a stalk, initiates the adhesion process (see Fig. 5; Bodenmiller et al., 2004; Levi and Jenal, 2006). These observations suggest that there is a selective advantage for stalk synthesis, independent of adhesion. The observation that stalks are considerably longer in nutrient-depleted environments suggests that stalk elongation may provide a means to increase cellular surface area and nutrient uptake ability (Gonin et al., 2000; Poindexter, 1964). A number of recent results have provided evidence in support of a role for the stalk in nutrient uptake. First, in diffusionlimited environments, like the natural oligotrophic environment where C. crescentus lives, the rate of nutrient uptake should be proportional to the length of the cell, rather than to the surface area of the cell (Wagner et al., 2006). Thus, the elongation of the stalk rather than the entire cell body is likely to be bioenergetically favorable because synthesizing a long thin cell envelope extension requires less energy than elongating the cell by the same length. Second, stalks purified from C. crescentus are capable of transporting and hydrolyzing an organic phosphate-containing molecule (Wagner et al., 2006). Isolated stalks of A. biprosthecum are also able to transport glucose and all 20 amino acids (Larson and Pate, 1976; Tam and Pate, 1985). Thirdly, the predicted functions of the proteins associated with the stalk suggest a role for the stalk in nutrient uptake (Ireland et al., 2002; Wagner et al., 2006). A large number of outer membrane transporters and periplasmic-binding proteins for a variety of nutrients, including phosphate, are found in the stalk (Ireland et al., 2002; Wagner et al., 2006). Each of these observations provides support for a model that the stalk of C. crescentus is a morphological adaptation that enables survival in an oligotrophic environment (for review see Wagner and Brun, 2007).

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3.4. The Process of Adherence and the Holdfast Many bacteria possess the ability to adhere to surfaces. What are the potential advantages of adherence? It may be bioenergetically favorable for bacteria to attach to a surface and absorb passing nutrients rather than to actively swim in pursuit of nutrients. This may be because nutrients, as well as the enzymes the bacteria need to assimilate the nutrients, tend to be concentrated near solid surfaces in a dilute aqueous environment (ZoBell, 1937). In addition, the association of the bacteria with larger extracellular structures, such as other organisms or surfaces of ships and pipes, may provide a physical barrier and prevent predation (Costerton et al., 1987). Adhesion mediates the formation of biofilms, in which bacteria are more resistant to antibiotics and are better protected from the environment, including changes in pH, desiccation, and shear forces in an aqueous environment (Fux et al., 2005; Jefferson, 2004; Stoodley et al., 2002). C. crescentus, like many bacterial organisms, can vigorously adhere to both biotic and abiotic surfaces (Poindexter, 1964). While the pili and the flagellum facilitate the initial process of adherence and biofilm maturation, it is the holdfast, a polarly secreted adhesin comprised of both proteins and polysaccharides, which mediates a more permanent attachment (Fig. 5; see Sections 3.1.5 and 3.2.3). C. crescentus cells are often associated with each other in large groups exclusively via their holdfasts, a structure that is referred to as a rosette. During early studies, C. crescentus was isolated based on its ability to adhere to glass slides and an extracellular structure at the tip of the stalk, termed holdfast, was hypothesized to be responsible for this attachment (Henrici and Johnson, 1935; Poindexter and Bazire, 1964). Identification and characterization of several groups of holdfast mutants determined that the holdfast contains acidic polysaccharides (Umbreit and Pate, 1978). Synthesis and export of the C. crescentus holdfast is cell cycle regulated (Janakiraman and Brun, 1999). Here, we will discuss the process of adherence in C. crescentus and the role of the holdfast. We will also examine the properties and regulation of the holdfast, as well as the holdfasts of other prosthecate bacteria. 3.4.1. Role of Polar Structures in Adherence and Biofilm Formation For C. crescentus, several components are necessary for the initial stages of attachment, including flagella, pili, and motility (Fig. 5; Bodenmiller et al., 2004; Levi and Jenal, 2006). To initiate an association with a surface, the cell must overcome repulsive forces, possibly including hydrophobic and

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electrostatic forces (Marshall, 1972; ZoBell, 1943). Freshwater bacteria, such as C. crescentus, may have a relatively hydrophobic cell surface and, therefore, the aqueous phase at the water–solid interface may serve as a barrier to attachment (Marshall, 1972). Both the bacterial cell surface and most attachment surfaces carry a net negative charge leading to repulsion due to the electrostatic forces. Brownian motion is not sufficient to overcome these forces (Marshall, 1972); however, motility via the flagellum likely aids in overcoming the repulsive forces. The initial attachment may be facilitated directly by both the flagella and the pili, which help to orient the cell properly for more permanent adherence that is only achieved with the presence of the holdfast (Cole et al., 2003; Merker and Smit, 1988). After the early stages of attachment, C. crescentus can create a more complex biofilm structure (Entcheva-Dimitrov and Spormann, 2004; Smit et al., 2000). The biofilm begins with microcolonies that develop into mushroom-shaped structures and a monolayer biofilm forms between these structures. The holdfast is the most important structure for proper biofilm formation, although the pili and the flagella were also found to be critical for the proper development and maintenance of the mushroom structures within the biofilm (Entcheva-Dimitrov and Spormann, 2004). 3.4.2. Holdfast Composition and Properties The C. crescentus holdfast is comprised of both protein and polysaccharide. N-acetylglucosamine (NAG) was identified as a component of the holdfast polysaccharide based on the ability of the holdfast to bind wheat germ agglutinin (WGA) lectin and on the sensitivity of the holdfast to chitinase and lysozyme, which are specific for NAG polymers (Merker and Smit, 1988). Lysozyme and chitinase cleave the b1-4 linkages between N-acetyl-Dglucosamine residues in polysaccharides, indicating that some of the linkages in the C. crescentus holdfast are b1-4 linkages. NAG polymers that function as adhesins have been identified in other bacteria including E. coli (PGA) (Wang et al., 2004) and Staphylococcus sp. (PIA) (Cramton et al., 1999; Mack et al., 1996). The PGA and PIA adhesins of E. coli and Staphylococcus sp. are linear polymers of b1-6 NAG (Wang et al., 2004). PIA and PGA are important for biofilm formation and ultrastructure and the PIA adhesin is also important for the pathogenicity of Staphylococcus sp. through protection from the innate immune response (Cramton et al., 1999; Go¨tz, 2002; Vuong et al., 2004; Wang et al., 2004). In addition to NAG, holdfast-associated proteins may play an integral role in adherence, since one group of proteins associated with the holdfast, the holdfast attachment proteins (HfaA, HfaB, and HfaD, see Section 3.4.3.2),

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have similarity to the curli proteins of E. coli, which are involved in bacterial attachment. Recent biophysical analysis of the holdfast has shown that it is an elastic, gel-like substance, with impressive adhesive properties. Initial indications that the holdfast had strong adhesive properties came from the failure of laser tweezers with a maximum working force on the order of 10 pN to detach single C. crescentus cells from a glass surface (Tsang et al., 2006). In order to determine the force of adhesion of single C. crescentus cells, a micromanipulation method was developed. Cells were allowed to attach to thin flexible pipettes whose force constants had been determined by atomic force microscopy (AFM). A suction pipette was used to grab the cell body and pull it in a direction perpendicular from the flexible pipette. The force required to break the cell-to-pipette contact was determined by measuring the displacement of the flexible pipette at the time the contact was broken. The force of adhesion of individual cells ranged from 0.11 to 2.26 mN with an average of 0.59 mN (Tsang et al., 2006). The large variation in force of adhesion is thought to be due to the variation in the size of holdfasts, the different breaking points, and the angle of pulling. Cells attached to a glass surface were subjected to a strong jet of water and examination of a glass surface by AFM following cell detachment indicated that the contact between the cells and the surface broke most often within the stalk or at the stalk-to-holdfast junction. Since the holdfast-to-surface junction remained intact, the force calculated for single cell adhesion is an underestimate of the force of holdfast adhesion. The holdfast spreads over a larger area (B411 nm) than the diameter of the stalk tip (B119 nm); therefore, the stress endured by the holdfast was calculated using finite element analysis, an engineering method used to determine stress in mechanical systems. This analysis revealed that the maximum stress at the holdfast–surface interface was 68 N/mm2, indicating that holdfast covering a 1 cm2 surface could theoretically hold a weight of B700 kg, making the holdfast the strongest biological adhesin described thus far (Tsang et al., 2006). Interestingly, such a force is in the right range to resist the force caused by the passage of an air–liquid interface, which can be up to 0.2 mN for a microscopic object. C. crescentus cells may be subjected to such forces when they are on surfaces that are hit by waves in an aquatic environment. Individual C. crescentus cells adhered to a surface are constrained by both the stalk and the holdfast. Microscopic observation of attached cells in liquid medium shows that the attached cells are subject to Brownian motion that slightly displaces the cells, resulting in a deformation of the stalk and/or holdfast. This deformation introduces a restoring force, which returns the cells to their original equilibrium position. Mathematical analysis of this

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behavior has been used to study the biophysical properties of the holdfast (Alipour-Assiabi et al., 2006; Li et al., 2005). After displacement of a cell by Brownian motion, the restoring force would be expected to result from the properties of both the holdfast and the stalk. The restoring force was determined by measuring the displacement of individual cells using optical microscopy. The force constant of the holdfast and stalk assembly was determined by fitting the displacement distribution to a modification of the Boltzmann equation, which typically describes the distribution of particles in a fluid (Li et al., 2005). If deformation of the stalk contributes significantly to the elastic nature of the stalk–holdfast assembly, the force constant would depend on stalk length since the ease of bending an object increases with its length. The analysis showed no correlation between the force constant and stalk length, indicating that the stalk is stiff as compared to the holdfast and that the force constant is a measure of the elasticity of the holdfast. A similar fluctuation analysis of the displacement of attached cells using a higher frequency of image recording was used to study pairs of cells attached to a surface by a shared holdfast (Alipour-Assiabi et al., 2006). This analysis showed that in some pairs, the elastic coupling between the two cells was stronger than their respective coupling to the surface, suggesting that the two cells attached to each other at the swarmer cell stage and subsequent holdfast synthesis allowed a strong crosslinking of the two cells, perhaps mediated by their respective Hfa proteins (see Section 4.4.3.2). In other pairs, one cell was tightly attached to the surface, whereas the other cell was coupled more tightly to the first cell than to the surface, suggesting that the cells attached to each other after holdfast synthesis, resulting in a less efficient melding of the two holdfasts. Digestion of the holdfast with lysozyme reduces the force constant of attached cells, indicating that the NAG polymer is important for the elastic properties of the holdfast (Li et al., 2005). Furthermore, lysozyme digestion drastically reduced the force of adhesion of the holdfast, such that cells were aspired by the suction pipette without causing a displacement of the flexible pipette in cell-pulling experiments (Tsang et al., 2006). Notably, the adhesion force was still sufficient to resist detachment using laser tweezers with a force of 10 pN, suggesting that components other than NAG polymers also contribute to the force of adhesion of the holdfast. 3.4.3. Genetics of Holdfast Synthesis and Attachment in C. crescentus Several loci involved in the biosynthesis, regulation, and anchoring of the holdfast in C. crescentus have been identified. Four classes of holdfast mutants have been characterized by transposon and UV mutagenesis: I) cells

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that make no holdfast, II) cells that make reduced levels of holdfast, III) cells that make holdfast that does not remain attached to the cell, and IV) cells with pleiotropic mutations that result in holdfast and polar development defects (Mitchell and Smit, 1990; Ong et al., 1990; Smith et al., 2003). 3.4.3.1. Holdfast Secretion and Biosynthesis Loci. The mutations in the class I and class II holdfast mutants that produce no holdfast or reduced amounts of holdfast, respectively, reside within the holdfast export and biosynthesis locus. Holdfast export is predicted to be accomplished by the products of three adjacent genes, hfsD (CC2432), hfsA (CC2431), and hfsB (CC2430) (Smith et al., 2003). The genes involved in holdfast polysaccharide biosynthesis are adjacent to those involved in holdfast export and are comprised of hfsC (CC2429), hfsE, hfsF, hfsG, and hfsH (CC2425-28) (Fig. 6B; Toh et al., 2008). hfsA, hfsB, and hfsC are transcribed convergently with the predicted biosynthesis genes, while hfsD is transcribed divergently from hfsA, hfsB, and hfsC (Fig. 6B; Smith et al., 2003). Deletion of the holdfast export genes, hfsD, hfsA, or hfsB, result in complete loss of holdfast polysaccharide synthesis. Based on sequence similarity, holdfast polysaccharide export is likely to occur by a wzy-dependent mechanism similar to group I capsule biosynthesis of E. coli (Fig. 7) (Whitfield, 2006). HfsD is a 25-kDa lipoprotein (C.S. Smith and Y.V. Brun, unpublished) with similarity to the Wza secretin of E. coli, an octomeric outer membrane lipoprotein required for Type I capsule polysaccharide secretion (Drummelsmith and Whitfield, 2000). In the type I capsule export system of E. coli, Wza and Wzc form a complex that spans the inner membrane, the periplasm, and the outer membranes (Collins et al., 2007). HfsA is a predicted 55-kDa protein with similarity to Wzc, a member of the membrane periplasmic auxiliary (MPA-1) family of polysaccharide transport proteins (Paulsen et al., 1997; Smith et al., 2003). In E. coli, Wzc functions as a tetramer (Collins et al., 2007). The Wzc proteins of Gramnegative bacteria typically have a cytoplasmic region that contains two ATP-binding motifs (Walker A and B; Whitfield and Paiment, 2003). HfsA does not have either motif; however, HfsB is a predicted 25-kDa protein that contains putative Walker A and Walker B motifs and has some amino acid similarity to Wzc protein homologs in both Gram-positive and Gramnegative bacteria (J.W. Javens and Y.V. Brun, unpublished; Smith et al., 2003). Interestingly, in Gram-positive bacteria, Wzc homologs involved in capsule translocation are comprised of two proteins: the periplasmic membrane translocator, similar to HfsA, and a separate cytoplasmic protein with an ATP-binding motif, similar to HfsB (Cozzone et al., 2004; Paulsen et al., 1997). Therefore, it is possible that the C. crescentus Wzc

Figure 6 Organization of the hfa and hfs loci in C. crescentus, Caulobacter sp. K31, M. maris and O. alexandrii. Each arrow represents a gene and the direction of transcription for that gene. Cc, C. crescentus CB15; K31, Caulobacter sp. K31; Mm, M. maris MCS10; Oa, Oceanicaulis alexandrii HTCC2633. (A) Comparison of the holdfast attachment loci for C. crescentus, M. maris, and O. alexandrii. The holdfast attachment locus is composed of hfaA, hfaB, hfaD, and hfaC. The corresponding homologs for hfaA (black), hfaB (dark gray), and hfaD (light gray) in Caulobacter sp. K31, M. maris, and O. alexandrii are shown below the hfa locus for C. crescentus. The gene numbers are indicated in the arrow representing the gene for Caulobacter sp. K31, M. maris, and O. alexandrii. Predicted open reading frames surrounding the hfa loci are represented with white arrows. There is a conserved rhomboid protease (rhom) upstream of many of the hfa loci. M. maris has two hfa loci. HfaC is not present in any of the other hfa loci. (B) Comparison of the holdfast biosynthesis loci from C. crescentus, Caulobacter sp. K31, M. maris, and O. alexandrii. The holdfast biosynthesis and transport loci comprise of hfsE, hfsF, hfsG, hfsH and then hfsD, hfsA, hfsB and hfsC. The gene numbers for each gene are indicated in the arrow representing the gene for Caulobacter sp. K31, M. maris and O. alexandrii. Both M. maris and O. alexandrii have additional genes that may function in holdfast biosynthesis, which are shown as white arrows. Caulobacter sp. K31 has additional genes within the holdfast biosynthesis and secretion loci (shown in white), but their suggested functions do not appear to be related to polysaccharide biosynthesis. gt indicates a predicted glycosyl transferase; mt, is a predicted mannosyl transferase; gh, a predicted glycosyl hydrolase; celD indicates a gene involved in cellulose biosynthesis. The gene function assignments are based on the annotations of TIGR or JGI and blastp analysis. O. alexandrii has a homolog to hfsE that is located elsewhere in the genome as indicated by the gene number.

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homologues, HfsA and HfsB, function similarly to those of Gram-positive bacteria. The holdfast biosynthesis genes are comprised of hfsC hfsE, hfsF, hfsG, and hfsH (CC2425-29) (Fig. 6B; Toh et al., 2008). HfsC is a predicted 46-kDa protein with similarity to ExoQ from Sinorhizobium meliloti, a member of the Wzy protein family of polysaccharide polymerases (Collins et al., 2007; Smith et al., 2003). However, mutational analysis indicates that an hfsC-deletion mutant is not deficient in holdfast synthesis or surface adhesion (Smith et al., 2003). Recently, a paralog of hfsC, hfsI, was identified in C. crescentus indicating that HfsC and HfsI may work in concert to polymerize the holdfast polysaccharide (Toh et al., 2008). This situation would be analogous to lipopolysaccharide biosynthesis in Pseudomonas aeruginosa, which involves two O-antigen polymerases, Wzya and Wzyb (Kaluzny et al., 2007). HfsE, a predicted integral membrane protein, exhibits significant similarity to initiating glycosyltransferases that catalyze the first step in polysaccharide biosynthesis. HfsF has similarity to polysaccharide flippases. HfsG has similarity to the group 2 family of glycosyltransferases and HfsH has similarity to polysaccharide deacetylases. Interestingly, the NAG of the PGA and PIA adhesins of E. coli and Staphylococcus spp. are deacetylated to varying degrees (Mack et al., 1996; Wang et al., 2004). The exact polysaccharide content and structure of the C. crescentus holdfast and the role of deacetylation in adherence have yet to be determined. Figure 7 Model of holdfast biosynthesis and attachment in C. crescentus. HfsE transfers N-acetylglucosamine (NAG) from UDP-NAG to undecaprenol (black oval). HfsG transfers NAG to the first sugar on the undecaprenol. HfsH deacetylates some of the NAG on the growing polysaccharide chain to glucosamine. HfsF is a flippase and transfers the small saccharide repeat unit linked to undecaprenol across the cytoplasmic membrane. HfsC and HfsI are polymerases that link the saccharide repeat units together to create the holdfast polysaccharide. HfsA, HfsB, and HfsD translocate the holdfast polysaccharide across the outer membrane so it can be anchored to the cell surface via the holdfast attachment proteins. An HfsA tetramer and HfsD octomer create the translocator and HfsB is an ATPase important for phosphorylation and signaling. For the holdfast attachment proteins, HfaA and HfaD are translocated via the Sec system across the cytoplasmic membrane to the periplasm where the signal sequence is cleaved by signal peptidase (SP). After processing, HfaA and HfaD interact with HfaB, the outer membrane secretin. HfaB translocates HfaA and HfaB across the outer membrane where they create the holdfast anchor and associate with the holdfast polysaccharide (only drawn for HfaA) by an unknown mechanism. Homologs to each of the Hfs and Hfa proteins are shown in parentheses adjacent to each protein. UDP, uridine diphosphate; P, phosphate; dark gray hexagon, NAG; light gray hexagon, glucosamine; IM, inner membrane; OM, outer membrane; CW, cell wall; SS, signal sequence.

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One possible model for polysaccharide biosynthesis based on the above gene function similarities is shown in Fig. 7 (Toh et al., 2008). HfsE transfers NAG or possibly other sugars onto the undecaprenol phosphate lipid carrier in the inner membrane. HfsG transfers additional sugars to the lipid intermediate undecaprenol phosphate and serves to lengthen the repeat unit. HfsH removes acetyl groups from some of the sugars in the polysaccharide repeat unit attached to undecaprenol phosphate. HfsF, the putative flippase, translocates the lipid-linked polysaccharide units across the inner membrane of the cell. HfsI and HfsC assemble the polysaccharide chain from the repeat units transferred by HfsF. The polysaccharide oligomers are then translocated outside the cell via the holdfast export proteins: HfsD, HfsA, and HfsB as described above. 3.4.3.2. Holdfast Attachment Locus. The class III holdfast mutants, which produce holdfasts that do not remain cell associated, are known as holdfast shedding mutants. The mutations responsible for the holdfast-shedding phenotype have been mapped to three genes in the holdfast attachment (hfa) locus: hfaA (CC2628), hfaB (CC2629), and hfaD (CC2630) (Fig. 6A) (Cole et al., 2003; Kurtz and Smit, 1992; Kurtz and Smit, 1994; Smith et al., 2003). HfaA is a 12-kDa protein with limited amino acid similarity (28%) to the E. coli curlin monomer CsgA and other fimbrial family adhesins (Cole et al., 2003; Hardy et al., submitted for publication). CsgA is a bacterial amyloidlike protein that is the structural component of the curli fibrils involved in E. coli attachment to surfaces (Olse´n et al., 1989; Prigent-Combaret et al., 2000). HfaB is a 36-kDa lipoprotein with amino acid similarity (41%) to the E. coli curli secretin CsgG, which is a lipoprotein required for the secretion of the curli monomer CsgA and the nucleator protein CsgB (Robinson et al., 2006). HfaD is a 41-kD membrane protein with low-level amino acid similarity (20–40%) to other adhesins and collagen triple helix proteins. The hfaC gene (CC2631) is found downstream of hfaD and encodes a predicted 66 kDa protein with similarity to ABC transporters. hfaC was initially thought to be involved in holdfast attachment based on the erroneous mapping of an hfaD transposon insertion to hfaC (Kurtz and Smit, 1994); however, mutations in hfaC do not affect holdfast anchoring or cell adhesion (Cole et al., 2003). In addition, none of the other prosthecate genomes examined in this review have homologs of HfaC associated with putative holdfast attachment loci (Fig. 6A). Taken together, this suggests that HfaC is not involved in anchoring the holdfast. Insertions within hfaA, hfaB, or hfaD result in different holdfast-shedding phenotypes, reduced adherence, and decreased WGA lectin binding. Insertions within hfaB are the most severe resulting in very low levels of

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lectin binding (5%, as compared to 75% for the parental strain CB15), low levels of adherence, no rosette formation, and high levels of holdfast shedding. hfaA insertions result in moderate levels of lectin binding (30%) and adherence, reduced levels of rosette formation, and some holdfast shedding. hfaD insertions result in moderate levels of lectin binding (43%) and adherence, rosette formation is comparable to wild-type cells, and there is a high level of holdfast shedding (Cole et al., 2003). HfaA, HfaB, and HfaD have a predicted signal sequence suggesting that they are translocated across the cytoplasmic membrane. HfaB and HfaD are associated with the membrane fraction of C. crescentus and examination of isolated stalks and cell bodies indicates that HfaB and HfaD reside in the stalk of the cell (Cole et al., 2003). One possible model for an Hfa-mediated holdfast anchor is that HfaB is an outer membrane secretin that translocates HfaA and HfaD across the outer membrane where they form a complex that interacts with the holdfast polysaccharide to anchor it to the cell (Fig. 7). If HfaB translocates HfaA and HfaD then deletion of hfaB would prevent their translocation to the cell surface, possibly explaining why an hfaB mutant has the most severe phenotype. Deletion of either hfaA or hfaD result in moderate phenotypes, so HfaA and HfaD may contribute equally to anchoring the holdfast. HfaA and HfaD could associate with each other in several combinations: 1) HfaD could be a nucleator protein and HfaA could be a fibrillar adhesin that mediates most of the association with the polysaccharide; 2) HfaA and HfaD could form independent adhesin structures in close proximity on the cell surface; or 3) HfaD could act as the main adhesin and HfaA could be a tip adhesin. 3.4.4. Regulation of Holdfast Synthesis The holdfast is synthesized during specific times in the developmental cycle. The holdfast is synthesized in the mid to late swarmer stage at the flagellar pole and throughout swarmer cell differentiation, and then remains at the tip of the stalk for the rest of the cell cycle (Levi and Jenal, 2006). Of the three holdfast loci that have been identified, only the cell cycle regulation of the hfa locus has been examined directly (Janakiraman and Brun, 1999). Transcription of the hfa operon begins in the swarmer compartment of the predivisional cell and then decreases during swarmer to stalk cell differentiation (Janakiraman and Brun, 1999). The promoter region of the hfa operon contains both a putative s54 promoter and CtrA-binding sites. While s54 is not required for transcription of the hfa operon, deletion of rpoN, the gene encoding s54, results in increased transcription of the hfa operon suggesting that either s54 or a s54 regulated gene product negatively

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regulates the hfa operon (Janakiraman and Brun, 1999). Transcription of the hfa operon is decreased in a ctrA mutant suggesting that CtrA acts as a transcriptional activator for this operon (Laub et al., 2002). The fourth class of holdfast mutants is pleiotropic resulting in a variety of phenotypes in addition to the loss of holdfast (Merker and Smit, 1988; Smith et al., 2003; Yun et al., 1994). The class IV mutations were mapped to podJ and pleC (Smith et al., 2003). Mutation of podJ and pleC (see Sections 2.4.1.3 and 2.4.1.2) both result in loss of holdfast production and severely reduced binding to surfaces (Hinz et al., 2003; Smith et al., 2003). Functional domain analysis of PodJ indicates that the cytoplasmic region between amino acids 589 and 639 is important for holdfast biosynthesis (Lawler et al., 2006). PleD (see Section 2.4.1.2) has been implicated in the regulation of holdfast synthesis. Deletion of pleD results in a 70% reduction in C. crescentus adherence to polystyrene (Levi and Jenal, 2006); however, adherence levels and holdfast production are only affected during the swarmer to stalk cell differentiation. The reduced adherence of a pleD mutant during the swarmer cell differentiation results from a delay in holdfast biosynthesis. This result suggests that PleD is necessary for the appropriate timing of holdfast biosynthesis during cell cycle development and possibly in response to environmental signals (Levi and Jenal, 2006). pleD mutants also remain motile after differentiation of swarmer cells (Sommer and Newton, 1989) indicating that synthesis and degradation of c-di-GMP by PleD probably regulates the transition between motility and cell adhesion. Motility and attachment are divergently regulated using c-di-GMP in Salmonella sp. and E. coli. In Salmonella enterica serovar Typhimurium and E. coli, GGDEF domain proteins, such as AdrA, increase cellulose biosynthesis, an extracellular polysaccharide (1-4-b-glucose polymer) important in biofilm formation and curli biosynthesis (Kader et al., 2006; Simm et al., 2004; Zogaj et al., 2001). Increased levels of c-di-GMP also result in shedding of the flagella, and expression of adhesins (Jenal and Malone, 2006; Romling, 2005; Simm et al., 2004; Tamayo et al., 2007). In contrast, YhjH, an EAL phosphodiesterase domain protein, results in decreased levels of c-di-GMP and enhanced motility (Simm et al., 2004). Similarly, in C. crescentus, the phosphodiesterase activity associated with the TipF EAL domain decreases the level of c-di-GMP and results in flagellar biogenesis (Huitema et al., 2006), while increased diguanylate cyclase activity mediated by PleD results in flagellar shedding and activation of holdfast biosynthesis (Jenal and Malone, 2006; Levi and Jenal, 2006; Paul et al., 2004). Together, PleD and TipF regulate timing of expression of polar structures and holdfast elaboration to mediate permanent attachment (Fig. 4). The signals for the switch between motile and sessile lifestyles have not been identified, but may

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be mediated by the initial association with surfaces (Jenal and Malone, 2006). C. crescentus has numerous GGDEF and EAL containing proteins that could be involved in the regulation of the transition between a planktonic lifestyle and a sessile state associated with biofilms, which could be regulated by a variety of cellular and extracellular signals. In addition, a c-di-GMP receptor protein, DgrA (CC1599), as well as its paralog, DgrB (CC3165), are involved in the control of motility (Christen et al., 2007). Both DgrA and DgrB or other c-di-GMP receptor proteins in C. crescentus may be important for the control of holdfast production through signal transduction based on the ability to sense the concentrations of c-di-GMP and affect holdfast production by repressing or activating holdfast-related genes. C. crescentus attachment is also regulated by a photosensory twocomponent system, LovK and LovR (Purcell et al., 2007). These proteins contain an LOV (light, oxygen, or voltage) domain that regulates blue-lightdependent processes and many bacterial proteins containing these domains have been identified in bacteria (Crosson et al., 2003). LovK is a histidineprotein kinase and LovR is a single-domain response regulator (Purcell et al., 2007). Deletion of lovK (CC0285) or lovR (CC0284) results in decreased adherence in C. crescentus, and overexpression of LovK and LovR results in increased rosette formation indicating that LovK and LovR are involved in regulation of adherence associated genes. Maximal transcription of lovK and lovR occurs during swarmer cell differentiation (Purcell et al., 2007), which corresponds to holdfast synthesis and loss of pili and flagella in C. crescentus. 3.4.5. Holdfast in Other Caulobacter Species and Prosthecate Bacteria Other prosthecate bacteria including Hyphomonas, Hyphomicrobium, Asticaccaulis, and a variety of marine bacteria and other freshwater Caulobacter, have adhesive holdfasts (MacRae and Smit, 1991; Moore and Marshall, 1981; Quintero and Weiner, 1995; Umbreit and Pate, 1978; Yun et al., 1994). Although these holdfasts are not as well characterized as the C. crescentus holdfast, some insights have been gained from the basic characterizations that have been performed. The freshwater bacterium Asticcacaulis biprosthecum undergoes a dimorphic lifecycle resulting in a motile swarmer cell and a sessile stalked cell. The stalked cell of A. biprosthecum has two stalks that extend laterally from each side of the cell. The holdfast of A. biprosthecum is localized to the pole of the cell, is not associated with either side stalk, and is composed of acidic polysaccharide based on Ruthenium red binding (Umbreit and Pate, 1978). A. biprosthecum holdfast mutants were isolated by chemical

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mutagenesis and many of them had multiple phenotypes suggesting that either there were multiple mutations or that some of the mutations isolated are in genes necessary for polar development as has been seen for C. crescentus, where mutations in pleC and podJ affect holdfast synthesis and are pleiotropic (Hinz et al., 2003; Smith et al., 2003; Umbreit and Pate, 1978). The genus Hyphomonas includes species of marine bacteria that have a biphasic lifestyle, which results in a motile swarmer cell and a sessile prosthecate cell; however, the daughter cell is produced by budding from the tip of the stalk. Hyphomonas adherens has an adhesive polysaccharide capsule that is localized around the entire surface of the mother cell, but is not associated with the stalk or daughter cell. Based on lectin and Calcofluor-binding studies, H. adherens polysaccharide capsule is composed of galactose and N-acetylgalactoscosamine possibly in a b1-4 linkage (Quintero and Weiner, 1995). Similar to holdfasts, the H. adherens capsule binds gold and polycationic ferritin, which may be due to the positive charge of the gold ions and the negative charge of the acidic polysaccharides of the capsule (Quintero and Weiner, 1995). In contrast, Hyphomonas sp. strain VP-6, which is now classified as Hyphomonas rosenbergii (Weiner et al., 2000), has holdfast at the pole of the mother cell that binds coral tree lectin suggesting that the holdfast contains galactose-b-1-4-N-acetylglucosamine linkages. The H. rosenbergii holdfast is expressed in the newly released swarmer cell similar to C. crescentus (Langille and Weiner, 1998; Levi and Jenal, 2006). In addition to the holdfast, H. rosenbergii also has an extracellular polysaccharide capsule comprised of N-acetylated sugar that surrounds the entire cell and contributes to adherence (Langille and Weiner, 1998). While all Hyphomonas sp. studied thus far have some type of polysaccharide adhesin, Hyphomonas neptunium has not been examined for holdfast or capsular polysaccharide adhesin; however, the genome sequence for H. neptunium has been completed (Badger et al., 2006). H. neptunium does not have any orthologs to the holdfast attachment proteins indicating that H. neptunium may use an alternative mechanism to anchor holdfast polysaccharide to the cell or may not have a polar holdfast. There is some limited amino acid similarity between a few of the holdfast biosynthesis and secretion proteins of C. crescentus and H. neptunium, HfsA (HNE_2241), HfsB (HNE_2240), HfsE (HNE_2651), and HfsC (HNE_1196), but the genes are spread out around the genome and there are no homologs for the other known C. crescentus genes involved in holdfast biosynthesis, suggesting that H. neptunium synthesizes and secretes adhesive polysaccharide in a different manner than C. crescentus, perhaps as a polysaccharide capsule adhesin.

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Hyphomicrobium sp. can be found in soil, fresh and salt water and are noted for their ability to utilize one carbon compounds for energy (Sperl and Hoare, 1971). Hyphomicrobium sp. are similar to Hyphomonas sp. in that they produce a daughter cell by budding from the end of the mother cell stalk. In Hyphomicrobium vulgare strain ZV580, which has been renamed Hyphomicrobium zavarzinii (Hirsch, 1989), the holdfast is cell-associated but is not present at the tip of the stalk. As a result, H. zavarzinii forms rosettes with the cell body (Moore and Marshall, 1981). Inhibition of rosette formation by D-mannose and D-galactose, as well as Concanavalin-A, which binds a-linked mannose residues, suggests that the holdfast is comprised of mannose and galactose bound by a-linkages. Protein, in addition to the polysaccharide, was found to be important for H. zavarzinii attachment. Like C. crescentus, most marine Caulobacter sp. and Maricaulis sp. examined have holdfasts comprised of NAG (Abraham et al., 1999; Merker and Smit, 1988) and bind strongly to gold particles. Conversely, lysozyme and chitinase do not affect holdfast in a variety of Maricaulis sp. suggesting that the structure of the holdfast is different from the holdfast of fresh water Caulobacter sp. (Merker and Smit, 1988). Holdfast mutants in Maricaulis washingtonensis MCS 6 generated similar classes of holdfast mutants that were seen in C. crescentus (Yun et al., 1994): I) those that make no holdfast, II) those with less holdfast, and III) those with normal holdfast but with altered adhesion. Five different clusters of genes were identified by Southern blot and complementation, but none of these loci have been further characterized (Yun et al., 1994). Recently, the sequence of the Maricaulis maris MCS10 and Oceanicaulis alexandrii HTCC2633 genomes was completed and are available at the Joint Genome Institute website (http://genome.jgi-psf.org/mic_home.html). O. alexandrii is a member of the Hyphomonadacea family and is closely related to M. maris (Lee et al., 2005). Based on examination of their DNA sequence, both M. maris and O. alexandrii have holdfast biosynthesis, secretion, and attachment loci (Fig. 6). O. alexandrii and M. maris have homologs of hfsD, hfsA, hfsB, hfsC, hfsH, hfsG, hfsF, hfaA, hfaB, and hfaD, but the genes are organized in a slightly different manner than those of C. crescentus (J.W. Javens, E. Toh, G.G. Hardy and Y.V. Brun, unpublished). Interestingly, the hfsE homolog for M. maris is in the same region of the genome as the holdfast biosynthesis genes, but the hfsE homolog of O. alexandrii is not contained within the holdfast biosynthesis gene cluster. Both M. maris and O. alexandrii have additional genes within the holdfast biosynthesis gene cluster that appear to be involved in polysaccharide biosynthesis. There is an additional predicted glycosyltransferase gene between hfsH and hfsC. Two additional genes adjacent to hfsF are

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transcribed in the opposite direction. (Fig. 6B and Table 1). In both M. maris and O. alexandrii, the orf adjacent to hfsF encodes proteins with amino acid similarity to CelD in Agrobacterium tumefaciens, which is involved in cellulose biosynthesis in A. tumefaciens (Matthysse et al., 1995a, 1995b). The second orf of M. maris adjacent to hfsF encodes a protein with amino acid similarity to glycosyl hydrolases and the second orf of O. alexandrii is predicted to encode a protein with amino acid similarity to mannosyl transferases (Fig. 6B and Table 1). Both O. alexandrii and M. maris have homologs of the holdfast attachment genes that are organized in a similar manner to those found in C. crescentus (Fig. 6A). Directly upstream of the hfa promoter and divergently transcribed, there is an orf (CC2627) that encodes a protein with amino acid similarity to rhomboid proteases, which are intramembrane serine proteases (Ben-Shem et al., 2007; Koonin et al., 2003). CC2627 is conserved among C. crescentus CB15, M. maris, and O. alexandrii perhaps indicating that this protease may play a role in regulation of adhesion. Interestingly, M. maris has two different hfa loci (Fig. 6A and Table 1; Hardy and Brun, unpublished). The function of the additional hfa locus is unknown. The differences in organization among the genes predicted to be involved in holdfast biosynthesis, secretion, and attachment suggest that the holdfast structure of O. alexandrii and M. maris is likely somewhat different than that of C. crescentus. Finally, the holdfasts of other fresh water Caulobacter sp. have been characterized and in contrast to Maricaulis sp., they contain a variety of polysaccharides (MacRae and Smit, 1991; Merker and Smit, 1988). The holdfasts of C. henricii and C. vibroides have a-linked N-acetylgalactosamine and NAG. C. subvibroides holdfast contains a-linked mannose, a-linked fucose, and N-acetylgalactosamine. C. leidyia, a member of the Sphingomonadaceae that is closely related to A. biprosthecum (Abraham et al., 1999), contains a- or b-linked N-acetylgalactosamine or galactose and a-linked mannose. As with C. crescentus and the other prosthecates described here, all the fresh water Caulobacter species examined bind gold, a cationic metal, suggesting that the polysaccharides of the holdfast are negatively charged (Merker and Smit, 1988). The variation in sugar composition and holdfast structure may allow fresh water Caulobacter sp. to attach to surfaces with different chemistries depending on the specific environment they colonize. No examination of the genes involved in holdfast synthesis has been performed in the other fresh water Caulobacter sp. The draft genome sequence of Caulobacter sp. strain K31, available at the Joint Genome Institute website (http://genome.jgi-psf.org/mic_home.html), indicates that Caulobacter sp. K31 has the same organization of genes for holdfast biosynthesis, secretion, and attachment to that of C. crescentus CB15 (Fig. 6 and Table 1).

C. crescentus CB15

Caulobacter sp. K31

O. alexandrii HTCC2633

M. maris MCS10

Holdfast gene

Putative genea function

Homolog

% Ident./ Sim.b

Homolog

% Identi./ Sim.b

HfsD HfsA HfsB HfsC

OMP accessory (Wza) MPA (Wzc) ATPase (Wzc) Polysaccharide polymerase (Wzy) Glycosyl transferase

2177 2176 2175 2174

75/86 76/87 73/83 68/81

01484 01489 01494 01499

38/57 27/47 23/40 37/56

1691 1690 1689 1688

43/61 26/43 29/45 39/58

2347c

01504

2545 2170 2169 3380c No homolog 0476c

43/58 (100)d 37/51 38/55 39/55 29/44 NAf

1687

Deacetylase Glycosyl transferase Flippase Cellulose biosynthesis (CelD)e Glycosyl hydrolase

43/55 (45/60)d 83/90 76/85 71/80 28/47 NAf

42/56 (50/63)d 41/54 38/55 40/58 27/46 NAf (100)g 43/60 (49/66)d 49/63 37/54 35/51 37/55 54/68 23/38 29/48

CC2277c HfsH HfsG HfsF CC3689c No homolog CC0469c HfsE HfaA

Dolichol-phosphate Mannosyl transferase Glycosyl transferase Holdfast attachment (CsgA)

2168 0606

86/92 (41/58)d 66/76 75/90

HfaB

Hfa protein secretin (CsgG)

0605

HfaD

Holdfast attachment

0604

01509 01514 01519 01524 No homolog 01529 07474 15215

40/59 (100)d 37/49 34/55

81/89

15210

53/66

58/73

15205

27/39

Homolog

1686 1685 1684 1683 1682 0011c 1681 2747 1083 2748 1082 2749 1081

% Ident./ Sim.b

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Table 1 Comparison of the holdfast biosynthesis, secretion, and attachment proteins.

a

E. coli homolog is shown in parentheses. Percent similarity and identity was determined using BLAST and are a comparison to the gene in CB15. c These genes are not associated with the hfa and hfs holdfast gene clusters and are located elsewhere in the genome. d Percent identity and similarity is in comparison to O. alexandrii HTCC2633. e A. tumifaciens homolog is shown in parentheses. f NA, there was no homolog so percent identity and similarity is ‘‘not applicable.’’ g Percent identity is in comparison to M. maris MCS6. b

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Future studies will be needed to identify the specific functions of the holdfast biosynthesis and attachment proteins as well as the biochemistry of holdfast polysaccharide transport, biosynthesis, and physical attachment. As additional genomes of Caulobacter and other prosthecates are sequenced, comparisons of their holdfast loci may provide insights into the functions of the holdfast genes, their regulation, and the reason for the variation in holdfast composition.

4. CHROMOSOME REPLICATION AND SEGREGATION, CELL DIVISION, AND CELL SHAPE In order for C. crescentus to produce two different cell types after cell division, changes in cell shape must be coupled with chromosome replication and segregation and cell division. During the swarmer to stalked cell differentiation, chromosome replication and cell division are initiated. Each compartment of the predivisional cell must contain a complete chromosome prior to the completion of cell division. The complex developmental steps that are required for changes in chromosome replication and segregation, cell division, and cell shape are examined in this section.

4.1. Chromosome Replication Occurs Once and Only Once per Cell Division Cycle Chromosome replication is subject to extensive regulation during the C. crescentus cell cycle. A combination of mechanisms including the control of CtrA and DnaA abundance and activity and the expression of DNA replication proteins ensures that chromosome replication occurs once and only once per cell division cycle. In this manner, C. crescentus differs from E. coli and many other bacteria that are able to initiate chromosome replication more than once per cell cycle, depending on growth rate. The pattern of DNA methylation was used to precisely assess the frequency of chromosome replication initiation per cell division cycle (Marczynski, 1999). The CcrM DNA methyltransferase is subject to extremely tight control during the cell cycle (Marczynski and Shapiro, 2002). The ccrM gene is transcribed in the predivisional cell (Zweiger et al., 1994) under the positive transcriptional control of CtrA (Reisenauer et al., 1999; Stephens et al., 1995). CtrA binds to the ccrM promoter relatively weakly, ensuring that ccrM transcription is only activated late in the

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predivisional cell (Reisenauer et al., 1999). This late expression of ccrM results in a sudden increase in CcrM concentration and subsequent methylation of the chromosomes (Stephens et al., 1996; Zweiger et al., 1994). CcrM is then degraded by the Lon protease just prior to cell division (Wright et al., 1996) and the two daughter cells are released with fully methylated chromosomes and no CcrM. Chromosome replication results in hemi-methylated DNA and the DNA remains hemimethylated until late in the predivisional cell when CcrM is synthesized. Fully methylated DNA can be distinguished from hemi-methylated DNA by restriction digestion using enzymes that are sensitive to methylation state and whose recognition sequence overlaps with the site of CcrM methylation. Therefore, the appearance of hemi-methylated DNA during the cell cycle was used as a measure of the frequency of chromosome replication initiation per cell division cycle (Marczynski, 1999). Using this strategy, hemi-methylated DNA could not be detected above background levels, indicating that less than one cell in 1000 reinitiates chromosome replication. Since plasmids can replicate in all cell types (Marczynski et al., 1990), mechanisms which restrict chromosome replication to the predivisional cell must exist. It is interesting to note that plasmids have a much higher rate of replication in stalked cells as compared to swarmer cells (Marczynski et al., 1990). This is likely to be due in part to cell-cycle regulation of the expression of DNA replication genes (Laub et al., 2000). In contrast, the control of chromosome replication is regulated by the state of the origin of chromosome replication as described in the next section. 4.1.1. The Origin of Replication The precise location of the C. crescentus origin of replication (Cori) was identified using three different methods. The initial discovery of the Cori region came from experiments in which chromosomal DNA from a synchronized swarmer cell population was radioactively labeled throughout the cell cycle and analyzed by pulse-field gel electrophoresis and autoradiography (Dingwall and Shapiro, 1989). Refinement of this method using a temperature-sensitive DNA replication mutant narrowed the Cori region to a smaller DNA fragment (Marczynski and Shapiro, 1992). DNA fragments within this region were tested for their ability to support autonomous plasmid replication and the minimal Cori region was narrowed down to B500 bp (Marczynski et al., 1995; Marczynski and Shapiro, 1992). The localization of Cori was confirmed using the two-dimensional DNA neutral/neutral agarose gel method that resolves nonlinear DNA fragments caused by the passage of a replication fork (Brassinga and Marczynski,

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Pw hemE

Ps

P3 AT a

AG b

IHF c

DnaA d

e

RP001

Figure 8 The C. crescentus origin of replication. The region required for chromosome replication is shown as a thick black line and is flanked by the hemE and RP001 genes which are depicted as gray arrows that indicate polarity. The black arrows indicate RNA start sites for the transcription promoters, PW, PS, and P3. The AT and purine (AG) rich regions required for chromosome replication are shown as labeled thin solid bars. The binding sites of IHF and DnaA are shown in solid boxes. The CtrA-binding sites are shown in dashed boxes and are labeled with the site designation, a–e.

2001). All these methods determined that Cori is located between the hemE gene and the RP001 gene, both of which are transcribed divergently from Cori (Fig. 8). This gene organization is conserved in at least some Alphaproteobacteria and may be useful in the identification of the origin of chromosome replication in these bacteria. DNA-binding studies with CtrA, its homolog in Rickettsia prowazekii CzcR, and IHF indicate that the binding sites for these proteins are conserved in Cori and in the R. prowazekii origin of replication, suggesting conserved regulatory mechanisms for the initiation of chromosome replication (Brassinga et al., 2002). A number of regulatory sequences have been identified in the minimal Cori sequence including five CtrA-binding sites (sites a–e starting with the hemE-proximal site), one DnaA box, an IHF-binding site, and some conserved motifs of unidentified function (Fig. 8; Marczynski and Shapiro, 2002). CtrA represses the initiation of chromosome replication, DnaA stimulates it, and IHF is thought to also act positively on initiation. A weaker promoter, PW, and a stronger promoter, PS, are located upstream of the hemE gene (Fig. 8; Marczynski et al., 1995). PW is responsible for most of the hemE expression while PS transcription is uncoupled from HemE synthesis. The RNA transcribed from the PS promoter lacks a good ribosome-binding site and the 5u untranslated region has the potential to fold because of the presence of four conserved 8-mer motifs found between PS and PW. The 8-mer motifs are essential elements of Cori function since small deletions in this region are not tolerated in the chromosome or in Cori plasmids (Marczynski et al., 1995). It may be that folding of the RNA in the 8-mer region transcript plays a role analogous to RNAs that regulate plasmid replication and copy number, but this has yet to be determined.

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Another indication that transcription from PS is likely to play a role in the control of replication initiation lies in the fact that PS is regulated by CtrA and is only transcribed in stalked cells (Marczynski et al., 1995). Two of the five CtrA-binding sites in Cori, sites a and b, are adjacent and lie within a 40 bp AT-rich region that also contains PS (Fig. 8). Upstream of the AT-rich region is a 40 bp region rich in purines that is essential for Cori function, although the function of this purine rich region is not known (Marczynski and Shapiro, 2002). Next is CtrA-binding site c, which overlaps with the IHF-binding site. IHF binding reduces CtrA binding to site c and also to CtrA binding site d (Siam et al., 2003). Deletion of CtrA-binding site c and the IHF-binding site is tolerated on the chromosome but cells grow slowly and are intolerant to a CtrA phosphomimetic mutant (Siam et al., 2003). Since IHF expression increases during the swarmer to stalked cell differentiation (Gober and Shapiro, 1990, 1992), at the same time as CtrA concentration decreases, it is possible that IHF displaces CtrA and bends Cori DNA to promote the initiation of chromosome replication (Siam et al., 2003). Finally, CtrA-binding site e is adjacent to a DnaA box. DnaA is essential in C. crescentus and depletion of DnaA using a xylose inducible promoter prevents the initiation of chromosome replication, but not ongoing replication (Gorbatyuk and Marczynski, 2001). The function and regulation of DnaA is described in detail in Section 2.1.3. 4.1.2. Regulation of Chromosome Replication by tmRNA Another regulator of chromosome replication is the hybrid tRNA-mRNA molecule called tmRNA, which is encoded by the ssrA gene (reviewed in Keiler, 2007). One domain of tmRNA folds into an alanyl-tRNA-like structure and is charged by alanyl-tRNA synthetase. tmRNA lacks an anticodon stem-loop and its 3u end contains an open reading frame that encodes the degradation tag that is added to the C-terminal end of incomplete proteins. tmRNA is one of the most abundant RNAs in bacterial cells and it plays important roles in general physiology (Keiler, 2007). The main function of tmRNA is to release ribosomes that are stalled due to the presence of mRNAs without a stop codon, which result from partial degradation at the 3u end. tmRNA adds the SsrA degradation tag at the C-terminal end of the incomplete proteins. The degradation of incomplete proteins eliminates the potential for dominant-negative effects associated with some incomplete proteins. A small protein called SmpB assists tmRNA function by increasing its stability and interaction with the ribosome. Deletion of the tmRNA gene (ssrA) in C. crescentus results in slow growth rate that is caused by a stage-specific reduction of growth rate (Keiler and

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Shapiro, 2003b). Synchronized ssrA swarmer cells take 195 min to complete the cell cycle as compared to 150 min for wild-type cells. This difference in the length of the cell cycle is caused by a marked increase in the amount of time ssrA cells remain in the swarmer stage, from 30 min in wild-type cells to 75 min in ssrA mutant cells. Flow cytometry experiments showed that the prolonged swarmer stage in ssrA cells correlates with a delay in the initiation of chromosome replication (Keiler and Shapiro, 2003b). CtrA degradation normally occurs just prior to chromosome initiation. In an ssrA mutant, CtrA degradation occurs at the proper time; however, there is a 40 min delay before the initiation of chromosome replication. Therefore, tmRNA is required for the proper control of replication initiation through a CtrAindependent process. These results also imply that the degradation of CtrA is not sufficient for the initiation of chromosome replication. Is the requirement for tmRNA for the timing of replication initiation due to its role in protein degradation? Expression of tmRNA which encodes a proteolysis-resistant SsrA tag fails to complement an ssrA deletion mutant and produces a phenotype that is more severe than that of the ssrA deletion mutant (Keiler and Shapiro, 2003b). Furthermore, introduction of the mutation encoding the stable SsrA tag into wild-type cells causes a delay in replication initiation. These results indicate that the tagging of some protein(s) with a wild-type SsrA peptide is required for the timing of replication initiation. As suggested by its involvement in the regulation of replication initiation timing, tmRNA abundance is subject to cell-cycle regulation (Keiler and Shapiro, 2003a). tmRNA is transcribed and produces a stable molecule in swarmer and predivisional cells. A sharp increase in transcription from the ssrA promoter leads to high levels of tmRNA during the swarmer to stalked cell transition; however, tmRNA is rapidly degraded in stalked cells. The cell-cycle regulation of tmRNA stability is controlled by the opposing actions of the 3u to 5u exoribonuclease RNase R and the SmpB protein (Hong et al., 2005). RNase R can degrade tmRNA in vitro and is required for the cell-cycle-dependent degradation of tmRNA in stalked cells. The level of RNase R is relatively constant during the cell cycle, indicating that the cell-cycle variation in tmRNA is not due to the regulation of RNase R expression (Keiler and Shapiro, 2003a). Deletion of the gene encoding the tmRNA binding protein SmpB causes the same delay in replication initiation as the deletion of ssrA (Keiler and Shapiro, 2003b). SmpB has high affinity for tmRNA in vitro and selectively protects it from degradation by RNase R (Keiler and Shapiro, 2003a). SmpB is also required for tmRNA stability in vivo (Keiler and Shapiro, 2003b). SmpB abundance dramatically increases during the swarmer to stalked cell differentiation when tmRNA

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accumulates, and SmpB is rapidly degraded in stalked cells when tmRNA is degraded (Keiler and Shapiro, 2003a). Therefore, the cell-cycle dependent fluctuation in tmRNA stability is mediated by the protective action of SmpB against RNase R degradation. A proteomic identification of tmRNA substrates identified 73 proteins that are tagged by tmRNA (Hong et al., 2007). Proteins involved in DNA replication, recombination, and repair are overrepresented in the subset of proteins identified as tmRNA substrates. This observation is consistent with the model that tmRNA regulates the timing of chromosome replication by controlling the amount or activity of protein complexes involved in the initiation of chromosome replication, chromosome replication, and DNA repair.

4.2. Coordination of Chromosome Segregation and Cell Division Prior to cell division, newly replicated chromosomes must be partitioned into the newly formed daughter cells. This important process ensures that each daughter cell contains one copy of the chromosome that it is not truncated by the premature closing of the division site. This section describes the mechanisms that contribute to chromosome segregation in C. crescentus and its coordination with cell division. 4.2.1. Replisome Movement and Chromosome Organization In swarmer cells, the origin of replication is located at the flagellar pole and the terminus is located at the opposite pole (Fig. 9). Following the swarmer to stalked cell differentiation, chromosome replication is initiated and one copy of the origin rapidly migrates to the opposite pole, resulting in bipolar localization of the origins, and the terminus migrates to the mid-cell (Jensen and Shapiro, 1999). The position of the replisome during the cell cycle was monitored using GFP fusions to the DnaB, HolB, and HolC proteins, which are known components of the replisome, in strains where the gene fusions are the only copies of these genes (Jensen et al., 2001). The replication proteins are distributed throughout the swarmer cell prior to the initiation of chromosome replication. At the time of replication initiation, the replication proteins localize to the flagellar pole in a DNA replication-dependent fashion. As replication proceeds towards the terminus, the replisome gradually moves to the center of the cell and is disassembled once DNA replication is complete (Jensen et al., 2001). Movement of the replisome depends on DNA replication and it has been hypothesized that

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PAMELA J.B. BROWN ET AL.

FtsK

ParB

Origin ( ) Terminus ( )

SMC

MreB

Figure 9 Localization of replication origin and terminus and chromosome segregation proteins in C. crescentus. In each cell cycle, the light gray oval represents a non-replicating chromosome and the theta structures represent a replicating chromosome. The origin of replication and the terminus (square and triangle in the middle cell cycle, respectively) are at opposite poles in swarmer cells, with the origin at the flagellated pole. A copy of the origin is rapidly moved to the opposite pole early in chromosome replication. The terminus progressively moves to the midcell and the two copies of the terminus are decatenated prior to cell division. The various proteins involved in chromosome segregation are represented by shaded shapes. FtsK localizes to the pole opposite the flagellum and the stalk in swarmer and stalked cells. During constriction, FtsK localizes to the mid cell where it remains until the cell divides to stabilize the Z-ring and help complete chromosome segregation. ParB colocalizes with and tracks the origin of replication. The SMC protein has a random or spiral-like localization throughout the cytoplasm; however, during cell division, the SMC proteins additionally form discreet foci at the poles. MreB forms a spiral in swarmer cells that coalesces into a band at the mid cell during division. MreB then returns to a spiral near the end of cell division.

accumulation of newly replicated DNA near the stalked pole passively displaces the replisome towards the mid-cell (Jensen et al., 2001). Analysis of the position of over 100 loci on the C. crescentus chromosome revealed that the chromosome is highly organized (Viollier et al., 2004).

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Each locus has a precise location in the cell, with loci organized in a linear order along the long axis of the cell that recapitulates their physical location on the chromosome. Loci closest to the origin of replication are closest to the pole where the origin is localized and those closest to the terminus are closest to the opposite pole. During DNA replication, as each locus is replicated, it rapidly moves to its proper cellular address. Therefore, the chromosome is not randomly compacted within the cell but is organized with impressive precision. Whether this organization has functional or regulatory consequences is not yet known. Part of the mechanism responsible for the precise organization of the chromosome lies in the fact that the origin and terminus are localized to opposite poles of the cell (Fig. 9). This polar localization is also relevant to chromosome segregation and to its coordination with cell division, as is discussed in the next sections. Much of this coordination occurs by regulating the localization of the cell division initiation protein FtsZ. 4.2.2. Formation and Structure of the Z-Ring FtsZ, a tubulin-like GTPase, is the most conserved protein involved in bacterial cell division and is required for the initiation of cell division (reviewed in Lutkenhaus, 2007). Prior to the initiation of cell division, FtsZ localizes around the circumference of the mid-cell, forming a structure called the Z-ring, and recruits other cell division proteins to this site (Fig. 10). In C. crescentus, the timing of Z-ring formation is dependent on cell-cycle progression. FtsZ concentration is low in swarmer cells and increases sharply after the swarmer to stalked cell differentiation and in predivisional cells. FtsZ is then degraded in late predivisional cells (Kelly et al., 1998; Quardokus et al., 1996, 2001). Mid-cell Z-rings begin to form shortly after the swarmer to stalked cell differentiation (Kelly et al., 1998; Quardokus et al., 2001), coincident with the increase in FtsZ concentration. Fluorescence microscopy analysis of FtsZ in many bacteria has led to the popular model that FtsZ forms a continuous ring at the site of cell division; however, the resolution of light microscopy is not sufficient to determine if FtsZ molecules are polymerized into a continuous ring around the cell circumference. Experiments in which the fluorescence recovery of FtsZ-GFP after photobleaching is observed indicate that FtsZ molecules rapidly exchange between the Z-ring and the soluble cytoplasmic pool, challenging the validity of the continuous Z-ring model (Anderson et al., 2004). Electron cryotomography (ECT) of C. crescentus cells was used to probe the structure of the Z-ring (Li et al., 2007). ECT represents a major technological advance in microscopy that is revolutionizing our understanding of cellular

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PAMELA J.B. BROWN ET AL.

FtsK

A

B

FtsQ

CtrA

DnaA

ftsZ

MipZ FtsZ FtsA

ftsQA

C

Cell cycle progression

Figure 10 Localization and regulation of cell division proteins in C. crescentus: (A) In each cell, the light gray oval represents a non-replicating chromosome and the theta structures represent a replicating chromosome. FtsZ (circle) initially localizes to the pole opposite the flagellum along with FtsK (star) in the swarmer cell. MipZ (diffuse shaded circle) resides at the flagellar pole. As the cell cycle is initiated, MipZ tracks with the newly replicated origins and rapidly localizes to both poles. Arrival of MipZ to the pole opposite the stalk displaces FtsZ, which migrates to the midcell (Z-ring, shaded band at the mid-cell). As the cell elongates, FtsA and FtsQ initially localize in a spiral-like pattern (not shown) and FtsK localizes to the mid-cell. In late predivisional cells, a preponderance of FtsQ (square) and FtsA (hexagon) join the Z-ring. (B) Timing of transcription of ftsZ, ftsQ, and ftsA and variation in abundance of the transcriptional regulators CtrA and DnaA during the cell cycle. The solid lines indicate the protein levels of the global regulators CtrA and DnaA. The dashed lines represent the transcription pattern of ftsZ and ftsQA as the cell proceeds through the cell cycle. (C) The diagram depicts how multiple short filaments of FtsZ form the Z-ring at the mid-cell. The cell is represented by the cylinder and the FtsZ filaments are represented by the gray curved lines.

ultrastructure (Jensen and Briegel, 2007). ECT is particularly good at revealing the three-dimensional ultrastructure of small cells in a life-like frozen-hydrated state generated by rapid freezing (Jensen and Briegel, 2007). By tilting the specimen in an electron cryomicroscope, a 3D reconstruction

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of the sample at ‘‘molecular’’ resolution can be obtained (Jensen and Briegel, 2007). C. crescentus cells are ideal specimens for ECT because their small size relative to many bacteria improves the resolution. ECT analysis demonstrated that during cell division in C. crescentus, FtsZ forms multiple short arc-like filaments at the mid-cell. The filaments average B100 nm in length and are found nearly perpendicular to the long axis of the cell, approximately 16 nm inside the inner membrane (Fig. 10; Li et al., 2007). Notably, no complete Z-rings were observed in this study. Only a few filaments were present in a given cell and analysis of cells at different stages of cell division revealed that there is no obvious pattern in the number, position, or configuration of FtsZ filaments during cell division (Li et al., 2007). Filaments are B5 nm in diameter, suggesting that they consist of single or double protofilaments. The FtsZ filaments appear in straight and curved conformations and often appear to extend into and through the inner membrane, possibly interacting with the peptidoglycan (Li et al., 2007). The portion of the filaments extending through the membrane is probably composed of other cell division proteins. In vitro, FtsZ filaments bound to GTP are straight and become curved when GTP is hydrolyzed to guanosine diphosphate GDP (Lu and Erickson, 1999; Lu et al., 2000). An iterative pinching model has been proposed to explain how the short FtsZ filaments could drive constriction (Li et al., 2007). GTP-bound FtsZ polymerizes into straight filaments that attach to the inner membrane through anchor proteins. As GTP hydrolysis occurs, the filaments bend, drawing the membrane inward in the region of the filament. While having only a small local effect for each individual filament, the repetition of this process thousands of times would be sufficient to drive cell constriction, as long as some process, presumably local peptidoglycan synthesis, stabilizes the small constriction prior to FtsZ depolymerization. For simplicity and despite the fact that a continuous Z-ring does not appear to exist, we will continue to refer to the Z-ring in the remainder of the text to describe the ring-like localization of FtsZ as seen by fluorescence microscopy. 4.2.3. Coordination of Z-ring Formation with Development and Chromosome Replication In addition to the importance of coordinating cell division with chromosome replication and segregation, C. crescentus imposes an additional regulatory layer that coordinates cell division with the developmental program, ensuring that cell division occurs once the asymmetry of the predivisional cell has been established. In this section, we describe how the

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timing of cell division is coordinated with development and with chromosome replication and segregation. Z-ring formation is not solely driven by an increase in FtsZ concentration. Ectopic expression of FtsZ in swarmer cells does not cause premature Z-ring formation, nor does it cause earlier initiation of cell division (Quardokus et al., 2001). Overexpression of FtsZ in predivisional cells causes the formation of additional constrictions near the mid-cell and a delay in cell separation (Din et al., 1998; Quardokus et al., 2001). In these cells, Z-ring localization is constrained to the sites of constriction, suggesting the existence of a mechanism to constrain Z-ring formation at the mid-cell. These results suggest that a cell-cycle or developmental cue is required for mid-cell Z-ring formation. Mid-cell Z-ring formation requires the initiation of chromosome replication; however, DNA replication per se is not required for the formation of Z-rings (Quardokus and Brun, 2002). When replication initiation is blocked, Z-rings still form but at subpolar regions of the cell, and cells constrict in an extended area, mostly away from, but sometimes over, the nucleoid (Quardokus and Brun, 2002). These observations suggest that early stages of chromosome replication are a major determinant for positioning of Z-rings in the cell, but not for Z-ring assembly (Quardokus and Brun, 2002). Identification of the terminus of replication (Jensen, 2006) allowed an examination of the timing of separation of the replicated termini during cell constriction. Careful fluorescence microscopy with a synchronized population of C. crescentus, determined that the invagination of the inner membrane clearly occurs before separation of the termini (Jensen, 2006). In fact, the replicated termini remain associated with the deeply constricted division site for an extended period after the completion of DNA replication and termini separation occurs just before the final cell separation step (Jensen, 2006). This indicates that nucleoid occlusion does not occur in C. crescentus, since cell constriction occurs in the presence of nonsegregated DNA (Jensen, 2006). However the Z-ring appears to form preferentially at sites where there is less DNA present (Quardokus and Brun, 2002). The results described above indicate that Z-ring formation is influenced by chromosome replication. Whether this influence is due to a mechanism directly monitoring chromosome replication is not known. However, it is becoming clear that the state of chromosome segregation affects Z-ring formation through the concerted action of ParA, ParB, and MipZ, as described in the next section. It is, therefore, possible that the effect of perturbations in chromosome replication on Z-ring formation occur indirectly through their effect on segregation.

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4.2.4. Coordination of Z-Ring Formation with Chromosome Segregation Homologs of the plasmid partition genes parA and parB are found in most prokaryotic genomes. Type I ParA and ParB are involved in plasmid partitioning, while type II ParA and ParB are involved in chromosome segregation (Gerdes et al., 2000; Mohl and Gober, 1997; Thanbichler and Shapiro, 2006b). The type II chromosome segregation genes parA, parB, and the ParB binding site, parS, of C. crescentus all reside in a single operon near Cori (Mohl and Gober, 1997). ParA and ParB localize at the poles where ParB binds to the parS cis-acting region on the chromosome (Fig. 9; Mohl and Gober, 1997; Thanbichler and Shapiro, 2006b). ParB consists of three domains: an N-terminal domain that interacts with ParA, a central DNA binding helix-turn-helix domain, and a C-terminal dimerization domain (Figge et al., 2003). The N-terminal domain of ParB regulates the ATPase activity of ParA by facilitating nucleotide exchange (Easter and Gober, 2002; Figge et al., 2003). ParA-ATP displaces ParB from parS (Easter and Gober, 2002). ParA and ParB are essential for viability; they are both required for cell division and for chromosome segregation (Mohl et al., 2001; Mohl and Gober, 1997). Perturbation of ParA and ParB protein levels prevents Z-ring formation and arrests cell division without affecting the expression of ftsZ, ftsQ, or ftsA (Mohl et al., 2001; see Section 4.3.2). Bipolar ParB foci appear prior to Z-ring formation, suggesting that polar localization of a chromosome segregation complex provides a checkpoint that coordinates cell division and chromosome segregation (Figge et al., 2003; Mohl et al., 2001). Identification of a member of the ParA superfamily of P loop ATPases, MipZ, has provided the link between ParB function and Z-ring formation (Thanbichler and Shapiro, 2006b). MipZ is highly conserved in Alphaproteobacteria; it is essential for viability and regulates Z-ring formation in C. crescentus (Thanbichler and Shapiro, 2006b). Depletion of MipZ causes cell filamentation and random localization of FtsZ in structures that are unable to form a functional divisome. Overexpression of MipZ causes dispersion of FtsZ in the cytoplasm and is localized to both poles. In vitro, MipZ interacts directly with FtsZ and interferes with its polymerization (Thanbichler and Shapiro, 2006b). By itself, FtsZ polymerizes into long straight polymers, whereas it forms short and highly curved polymers in the presence of MipZ. Careful observations from fluorescence microscopy suggest that FtsZ and MipZ cannot co-exist in the same area of the cell, indicating that MipZ controls FtsZ positioning (Thanbichler and Shapiro, 2006b). FtsZ is localized to the pole opposite the flagellum in swarmer cells and then migrates to the mid-cell during the swarmer to stalked cell transition.

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MipZ has the same pattern of localization as the origin of replication and ParB during the cell cycle (Thanbichler and Shapiro, 2006b). Indeed, MipZ binds to ParB in vitro and the two proteins form a ternary complex with a parS DNA fragment, with ParB providing the parS binding activity (Thanbichler and Shapiro, 2006b). By tracking the chromosome origins with ParB, MipZ coordinates Z-ring formation with chromosome movement (Thanbichler and Shapiro, 2006b). MipZ colocalizes with the origin of replication by binding to ParB, itself bound to parS sites located close to the origin, at the flagellar pole of swarmer cells. After the initiation of chromosome replication, one origin-ParB-MipZ complex remains at the same pole, while another MipZ-ParB complex associates with the second copy of the origin. This second origin-ParB-MipZ complex migrates to the opposite pole, where MipZ displaces FtsZ. Therefore, MipZ localization to both poles prevents FtsZ localization to those sites. In addition, the interaction of MipZ with ParB is highly dynamic and is regulated by the ATPase activity of MipZ, resulting in the formation of a gradient of MipZ with its highest concentration at the poles (Thanbichler and Shapiro, 2006b). This MipZ gradient results in a low concentration of MipZ at the mid-cell, where FtsZ can polymerize productively. Interestingly, an earlier study using immunogold transmission electron microscopy showed that FtsZ can localize to the stalked pole (Quardokus et al., 2001). This localization is consistent with the requirement of FtsZ for crossband synthesis at the stalked pole (Divakaruni et al., 2007). Since MipZ is always present at the stalked pole where it should inhibit FtsZ polymerization, a mechanism has to exist to allow polymerization of a small amount of FtsZ for crossband synthesis. Alternatively, polymerization of FtsZ is not required at the pole for crossband synthesis. 4.2.5. Role of MreB in Chromosome Segregation The actin-like cytoskeleton protein MreB that is required for cell shape determination (see Section 4.4 for a discussion of the role of MreB in cell shape determination) has been implicated in chromosome segregation in a number of bacteria (Thanbichler and Shapiro, 2006a). When synchronized C. crescentus cells are treated with the MreB inhibitor A22 (S-(3,4dichlorobenzyl)isothiourea) prior to chromosome replication initiation, replication still occurs, but origin-proximal regions fail to segregate (Gitai et al., 2005). This inhibition of segregation is rapidly reversed when A22 is removed, and does not occur when A22 is administered to an mreB mutant that is resistant to A22. When chromosome segregation is instead allowed to begin before wild-type cells are treated with A22, the origin-proximal

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regions segregate normally. The inhibition of chromosome segregation is specific to origin-proximal regions, since origin-distal regions still segregate normally when cells are treated with A22. Chromatin immunoprecipitation indicates that MreB associates, most likely indirectly, with origin-proximal regions but not with origin-distal regions (Gitai et al., 2005). These results indicate that MreB only controls the separation of the genes near Cori and probably only affects the initial steps of chromosome separation. Whether MreB is directly involved in chromosome segregation or whether the effect of MreB disruption on segregation is due to effects on cell shape remains controversial. For example, deletion of mreB in a cyanobacterium, Anabaena sp. PCC 7120, or of mreBCD in E. coli causes no defect in chromosome segregation (Hu et al., 2007; Karczmarek et al., 2007). To determine if cell shape perturbation causes defects in chromosome segregation, the effects of A22 treatment and mecillinam (amdinocillin) treatment, which targets PBP2, were compared in E. coli (Karczmarek et al., 2007). Mecillinam treatment results in the formation of round cells without disrupting MreB. Altered chromosome segregation was observed following either mecillinam or A22 treatment. This observation suggests that altered cell shape, rather than a specific function of MreB, is responsible for the chromosome segregation defect (Karczmarek et al., 2007). 4.2.6. Other Proteins Involved in Chromosome Segregation Chromosome segregation is also controlled by the structural maintenance of chromosome (SMC) proteins (Graumann, 2001; Jensen and Shapiro, 1999). The SMC proteins consist of N- and C-terminal domains with characteristic Walker A and B boxes, respectively, that are separated by a coiled-coil domain with a non-coiled ‘‘hinge’’ patch in the middle (Lo¨we et al., 2001; Melby et al., 1998). SMC dimerizes in an anti-parallel fashion with the opposite adjoining terminal domains interacting with one another (Melby et al., 1998). A null smc mutant in C. crescentus causes aberrant nucleoid localization, similar to what is seen in E. coli and B. subtilis (Jensen and Shapiro, 1999). Interestingly, SMC displays punctate localization throughout much of the cell cycle and roughly 35% of predivisional cells have brighter foci of SMC at the poles, with fainter foci throughout the cell (Fig. 9) (Jensen and Shapiro, 2003). The brighter foci may stem from the increase in smc transcription from the swarmer to stalk cell transition (Jensen and Shapiro, 2003). There are roughly between 1,500 and 2,000 SMC molecules per cell, enough for one protein to 6,000–8,000 bp of chromosomal DNA (Jensen and Shapiro, 2003). While it is unclear if all the SMC molecules are present in the foci, multiple SMC proteins must bind the

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chromosome since they are likely to play a role in condensing replicated DNA (Jensen and Shapiro, 2003; Thanbichler and Shapiro, 2006a). The coordination of chromosome segregation and cell division also involves FtsK (Bigot et al., 2007; Wang et al., 2006). FtsK is a member of the FtsK/SpoIIIE/Tra family of proteins. FtsK consists of an N-terminal domain required to localize FtsK to the mid-cell and for its function in cell division and a cytoplasmic C-terminal ATPase domain required for chromosome translocation (Fig. 9; Bigot et al., 2007). FtsK interacts with topoisomerase IV, a protein complex involved in chromosome decantenation, and the XerCD site-specific recombinase that resolves chromosome dimers (Bigot et al., 2007). Finally, FtsK translocates the separated chromosomes to their respective side of the septum (Bigot et al., 2007). In C. crescentus, FtsK localizes to the site of cell division early in the division process and is required for the assembly or maintenance of the Z-ring (Wang et al., 2006). FtsK is essential for viability and its C-terminus is required for proper segregation of the chromosome terminus. One of the roles of FtsK in chromosome segregation involves the ParC subunit of topoisomerase IV (Wang et al., 2006). The parC and parE genes encoding the topoisomerase IV subunits are essential for viability and required for proper chromosome segregation and the polar localization of the origin of replication in C. crescentus (Wang and Shapiro, 2004; Ward and Newton, 1997). ParC colocalizes with the replisome and, even though ParE is dispersed throughout the cell, ParE is required for ParC localization to the replisome (Wang et al., 2006). The FtsK C-terminus is also required for the localization of ParC, which explains, at least in part, the chromosome segregation defect of ftsK mutants (Wang et al., 2006).

4.3. Regulation of Cell Division Protein Expression In addition to the regulation of FtsZ polymerization and localization, coordination of cell division with other cell cycle and developmental events is achieved by regulating the synthesis of cell division proteins. Like other developmental regulators described in earlier sections, a combination of cell cycle dependent transcription and proteolysis is used to ensure that cell division proteins are present at the time they are required. 4.3.1. Transcriptional Regulation of Cell Division Genes In C. crescentus, ftsZ is part of a large cell division gene cluster whose overall organization is conserved in many bacteria. The regulation of the

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last three genes in the cluster, ftzQ-ftsA-ftsZ, has been studied in some detail (Kelly et al., 1998; Martin et al., 2004; Sackett et al., 1998; Wortinger et al., 2000). The bitopic inner membrane protein FtsQ is essential for cell division, but its precise role remains unknown, although it may be involved in peptidoglycan synthesis (Chen et al., 2002; Martin et al., 2004). FtsQ is part of a group of cell division proteins that localizes to the mid-cell late in cell division (Fig. 10; Buddelmeijer et al., 1998; Chen et al., 1999; Martin et al., 2004). FtsA, a peripheral membrane protein with ATPase activity, is a member of the actin family and, like FtsZ, is a highly conserved cell division protein (Bork et al., 1992; Rothfield et al., 1999; van den Ent and Lo¨we, 2000). In E. coli, ZipA, an inner membrane protein that binds FtsZ, and FtsA help stabilize the Z-ring and anchor it to the membrane (Addinall and Holland, 2002; Feucht et al., 2001; Pichoff and Lutkenhaus, 2002, 2005; Sa´nchez et al., 1994). Three promoters, PQA, PA, and PZ, drive transcription of ftsQ, ftsA, and ftsZ (Kelly et al., 1998; Sackett et al., 1998). The PQA promoter drives most of ftsQ and ftsA transcription (Sackett et al., 1998). PA is located between ftsQ and ftsA and is B10 times weaker than PQA. While ftsQ and ftsA are cotranscribed, a strong terminator is found between ftsA and ftsZ and prevents transcription from the ftsQ and ftsA genes from extending to ftsZ. ftsZ is transcribed from a strong promoter, PZ, which is located between ftsA and ftsZ (Kelly et al., 1998; Sackett et al., 1998). CtrA directly activates PQA transcription (Wortinger et al., 2000) and represses PZ transcription (Kelly et al., 1998), while DnaA activates the expression of ftsZ (Hottes et al., 2005). Transcription from all three promoters is low in swarmer cells. During the swarmer to stalked cell transition, the level of CtrA decreases and the level of DnaA increases, leading to the transcription of ftsZ concurrently with the start of DNA replication (Hottes et al., 2005; Kelly et al., 1998; Sackett et al., 1998). In the predivisional cell, CtrA is synthesized and activates the transcription of ftsQ and ftsA and represses the transcription of ftsZ (Kelly et al., 1998; Sackett et al., 1998; Wortinger et al., 2000). Following the completion of cell division, transcription of ftsQ and ftsA ceases through an unknown mechanism (Sackett et al., 1998). 4.3.2. The Proteolysis of Cell Division Proteins helps Establish a DNA Replication Checkpoint Coordination between cell division and DNA replication is ensured in part by a checkpoint that targets the expression of late cell division proteins. When DNA replication is blocked in C. crescentus, the transcription of ctrA from its P2 promoter is inhibited (Wortinger et al., 2000). In the absence of

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high levels of CtrA, ftsA and ftsQ are not transcribed (Wortinger et al., 2000). ctrA transcription from the P2 promoter depends on the presence of a low level of phosphorylated CtrA (see Section 2.1.1.1). ctrA transcription from the P1 promoter is activated by GcrA and supplies the initial CtrA, which is then phosphorylated (Holtzendorff et al., 2004). Therefore, the DNA replication checkpoint probably targets CtrA phosphorylation or GcrA activity, or both. One requirement for this checkpoint is that FtsA or FtsQ should be limiting for division in the next cell cycle. Indeed, the number of FtsA and FtsQ molecules fluctuates such that their concentration is low in swarmer and stalked cells, peaks in pre-divisional cells, and then dramatically decreases after cell division (Martin et al., 2004). Even when an inducible promoter drives ftsA and ftsQ transcription, FtsA and FtsQ levels vary during the cell cycle. The half-life of FtsA increases from 13 min in swarmer cells to 55 min in stalked cells, suggesting that FtsA is specifically degraded in certain cell types (Martin et al., 2004). Similarly, the half-life of FtsZ decreases from B80 min in stalked and early predivisional cells to B10 min in late predivisional cells (Kelly et al., 1998). Following cell division, degradation of FtsA and FtsQ reduces their concentration to 1 and 10% of their maximal level, respectively (Martin et al., 2004). These results strongly suggest that de novo synthesis of cell division proteins is required for each division cycle and sets the stage for checkpoint control through the regulation of their expression. FtsA and FtsQ are dynamically localized throughout the cell cycle (Fig. 10). After their synthesis in stalked cells, FtsA and FtsQ are distributed throughout cells in a spiral-like pattern. Both proteins are recruited to the mid-cell in late predivisional cells, consistent with their maximum level of expression and the requirement for FtsA in late stages of C. crescentus cell division (Martin et al., 2004).

4.4. Cell Shape Bacteria are characterized by a myriad of different shapes and sizes, including the large viviparous Epulopiscium fishelsoni (Angert et al., 1993; Fishelson et al., 1985), the star shaped bacterium, Stella strain IFAM1312 (Young, 2006), and the small, vibrioid, prosthecate bacterium, C. crescentus (Poindexter, 1964). Why do bacteria have different shapes and sizes? The specific shape of a bacterium is generally thought to confer a selective advantage in a certain environment (Young, 2006). For example, different bacterial shapes have varying abilities to take up nutrients or to protect the

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cell from predation (Young, 2007). The vibrioid shape of C. crescentus may contribute to its ability to disperse (Ausmees et al., 2003) and the stalk functions in nutrient uptake (Wagner et al., 2006). In addition, the shape of surviving C. crescentus cells changes dramatically during late stationary phase, indicating that changes in morphology are likely to confer additional advantages (Wortinger et al., 1998). What mechanisms govern the formation and maintenance of cell shapes? In C. crescentus, changes in cell shape occur throughout the cell cycle and are dictated by cell growth, cell division, and stalk elongation. This section will address how the shape of C. crescentus is generated, maintained, and changed throughout the cell cycle.

4.4.1. MreB, MreC, RodA, and PBP2 The mre operon (murein cluster e) contains the mreB, mreC, mreD, rodA (mrdB), and mrdA (PBP2) genes, which are involved in maintaining cell shape and are conserved in nearly all of the rod-shaped bacteria (Wachi et al., 1987). In C. crescentus and in rod-shaped bacteria, the proteins encoded by these genes determine and maintain bacterial cell shape by functioning as cytoskeletal proteins and by controlling peptidoglycan synthesis (Dye et al., 2005; Wachi et al., 1987; Wagner et al., 2005). MreB is an essential actin-like protein required for the maintenance of cell shape during growth. MreB depletion or treatment with A22, a drug that inhibits MreB polymerization, results in the formation of lemon-shaped cells (Fig. 11) (Figge et al., 2004; Gitai et al., 2004; Wagner et al., 2005). In C. crescentus, MreB also plays a role in stalk elongation (see Section 3.3.2) and chromosome segregation (see Section 4.2.5) (Gitai et al., 2004; Wagner et al., 2005). The transcript and protein levels of MreB remain constant throughout the cell cycle; however, MreB is dynamically localized during the cell cycle. MreB is primarily found as a cytoplasmic spiral that condenses to a ring at the mid-cell in a FtsZ and TipN-dependent manner during cell division (Fig. 11; Figge et al., 2004; Lam et al., 2006). Single MreB proteins have been shown to undergo treadmilling within the spiral structure in vivo (Kim et al., 2006). Treadmilling allows a protein polymer to remain stationary, despite the movement of individual proteins as new monomers are polymerized at one end of the polymer and old monomers are depolymerized from the other end (Kim et al., 2006). Addition of A22 results in a nearly immediate delocalization of MreB from the cytoplasmic spiral or mid-cell localization (Gitai et al., 2005).

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MreB B

CreS MreB MreC PBP2

MreC/PBP2

C CreS

mreB(-) A22 treated cells (MreB)

Wild-type

creS(-)

Mecillinam treated cells (PBP2)

mreC(-) rodA(-)

Figure 11 Cell shape proteins in C. crescentus. (A) Localization of cell shape proteins during the cell cycle. The proteins are represented in gray. MreB localizes as a spiral in swarmer cells, coalesces to a ring in the stalked and predivisional cells, and returns to a spiral at the end of cell division. MreC and PBP2 localize in a similar spiral-like pattern throughout the cell cycle. CreS aligns along the inner curve of the cell throughout the cell cycle. (B) Cell shape protein localization in swarmer cells. MreC and PBP2 co-align while the MreB spiral is offset from the MreC/PBP2 spirals. CreS lies along the inner curve of the bacterium. (C) Cell shapes resulting from the absence or inhibition of cytoskeletal proteins. Gene deletions and protein depletions are indicated by (). A22 targets MreB and cells grown in its presence have an identical phenotype to cells depleted for MreB. PBP2 is inhibited by the addition of mecillinam.

RodA, MreC, and PBP2 contribute to the maintenance of cell shape and control of cell width. Depletion of either MreC or RodA results in severe cell shape defects similar to that seen following MreB depletion (Fig. 11; Divakaruni et al., 2005, 2007; Wagner et al., 2005). Treatment of C. crescentus with mecillinam, an antibiotic that targets PBP2 in E. coli, generates wide filamentous cells (Fig. 11) (Koyasu et al., 1983; Seitz and Brun, 1998; Wagner et al., 2005). Throughout the entire cell cycle, MreC and PBP2 form longitudinal spirals, which encompass the whole cell

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(Fig. 11; Divakaruni et al., 2005; Dye et al., 2005; Figge et al., 2004; Gitai et al., 2004). The cytoplasmic spirals of MreC and PBP2 co-localize in vivo and affinity chromatography experiments have shown that MreC and PBP2 interact in vitro (Divakaruni et al., 2005; Dye et al., 2005; Figge et al., 2004). Notably, the spirals formed by MreC and PBP2 are offset from the spirals formed by MreB (Fig. 11). While the localization of MreB and MreC is independent of one another, proper localization of PBP2 requires the presence of both MreB and MreC (Divakaruni et al., 2005; Dye et al., 2005; Figge et al., 2004). In both E. coli and C. crescentus, cell elongation depends on RodA, which controls the transpeptidase activity of PBP2 (Shih and Rothfield, 2006; Wagner et al., 2005). Depleting either MreC or RodA disrupts peptidoglycan synthesis (Divakaruni et al., 2007; Ishino et al., 1986) but not the localization of PBP2. RodA and PBP2 are also required for stalk formation, indicating that stalk biosynthesis is a specialized form of cell elongation (Seitz and Brun, 1998; Wagner et al., 2005). The localization of RodA remains unknown due to the technical challenges associated with tracking the localization of inner membrane proteins. The function of MreD in maintaining the cell shape of C. crescentus has not been determined; however in E. coli, MreD is essential and its depletion results in spherical cells (Kruse et al., 2005). 4.4.2. CreS C. crescentus cells undergo morphological changes throughout the cell cycle, but the cells always remain curved. While depletion or disruption of mreB, mreC, rodA, or mrdA causes defects in cell morphology, they are not responsible for the characteristic curved shape of C. crescentus. A transposon mutagenesis screen for cell shape defects identified two straight mutants with mutations that mapped to the creS gene, which encodes the protein crescentin (Fig. 11; Ausmees et al., 2003). The straight phenotype was confirmed in deletion mutants of creS, suggesting that crescentin is responsible for imparting the curve shape to C. crescentus cells. Crescentin localizes along the concave side of the cell along the cytoplasmic side of the inner membrane (Fig. 11; Ausmees et al., 2003); however, CreS-GFP is nonfunctional and requires wild-type CreS for localization (Ausmees et al., 2003). A creS mutant that contains creS-gfp as its only copy of crescentin is straight and CreS-GFP localizes randomly in the cytoplasm as a curved filament (Ausmees et al., 2003). Crescentin assembles into curved polymers and is B25% identical and 40% similar and to cytoskeletal intermediate filaments, which play a role in controlling eukaryotic cell shape (Ausmees et al., 2003). The existence of intermediate filaments in bacterial cells

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completes the set of eukaryotic cytoskeletal homologs found within bacteria including the actin homologs, MreB/ParM/FtsA, and the tubulin homolog, FtsZ (Ayako et al., 1992; Bork et al., 1992; Erickson, 1995; Jensen and Gerdes, 1997; Lutkenhaus, 1993; Margolin, 2004; Sa´nchez et al., 1994; van den Ent et al., 2001). Crescentin is also required for the maintenance of cell shape during stationary phase. During late stationary phase, a small percentage of cells survive and adopt an elongated helical morphology (Wortinger et al., 1998). The elongated helical cells have a decreased level of FtsZ, explaining why the cells become elongated, but not why the cells become coiled. Crescentin is essential for the helical nature of stationary phase cells and is localized along the inner coil of the cell (Ausmees et al., 2003). This observation indicates that crescentin follows a helical pitch that is revealed only in elongated stationary cells. In exponential cells, the short filaments of crescentin appear only slight curved (Ausmees et al., 2003). 4.4.3. Peptidoglycan Synthesis While MreB, MreC, PBP2, and RodA maintain the proper cell shape throughout the cell cycle, these proteins are not directly responsible for mediating cell shape changes. How are changes in cell shape mediated? The generation and maintenance of C. crescentus cell shape requires temporal and spatial control of cell growth, cell division, and stalk elongation. These events require an expansion of the peptidoglycan cell wall, which occurs by insertion of new peptidoglycan into the existing peptidoglycan cell wall along the long axis of the cell (De Pedro et al., 1997). During cell division and stalk elongation, peptidoglycan precursor synthesis and peptidoglycan insertion is localized to the specific sites of elongation (Aaron et al., 2007; Divakaruni et al., 2007). In this section, the role of proteins involved in peptidoglycan synthesis and insertion at the sites of cell division and stalk synthesis is discussed. 4.4.3.1. FtsZ, MurG, and PBP2. Peptidoglycan synthesis at the site of cell division requires the presence of MurG, PBP2, and FtsZ. MurG, a glycosyltransferase involved in peptidoglycan precursor synthesis, contributes to the coordination of peptidoglycan synthesis along the length of the cell and PBPs are responsible for the insertion of new peptidoglycan to facilitate changes in cell shape (Aaron et al., 2007; Divakaruni et al., 2007). Experiments conducted in C. crescentus using D-cysteine or fluorescently labeled vancomycin, which incorporate into newly made peptidoglycan, determined that peptidoglycan synthesis

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occurs along the cell body and at the mid-cell (Aaron et al., 2007; De Pedro et al., 1997; Divakaruni et al., 2007). In cells depleted of FtsZ, no new synthesis was observed at the mid-cell, indicating that FtsZ directs peptidoglycan synthesis at that site (Aaron et al., 2007; Divakaruni et al., 2007). What is the role of FtsZ in directing peptidoglycan synthesis to the midcell? The transpeptidase protein, PBP2, which is likely to be involved in the insertion of new peptidoglycan into the cell wall, is mislocalized in the absence of FtsZ (Dye et al., 2005). When MreB is depleted, PBP2 mislocalizes from a spiral shape to the mid-cell; however, in cells depleted of FtsZ and treated with the MreB inhibitor A22, PBP2 remains localized as a spiral. This indicates that FtsZ plays an indirect role in PBP2 localization. While other bacteria form septa during cell division, C. crescentus must add cell wall material as it pinches and invaginates. To further elucidate the role of FtsZ in transverse peptidoglycan synthesis in C. crescentus, peptidoglycan synthesis was tracked using the localization of MurG, an enzyme that functions in a late step to produce a peptidoglycan precursor (Aaron et al., 2007; Divakaruni et al., 2007). Despite using similar strategies to localize MurG, conflicting data was obtained by the two research groups. Aaron et al. (2007) showed that MurG-GFP had a dynamic behavior in that it displayed diffuse localization in swarmer cells, mostly migrated to the mid-cell during cell division, and released from the mid-cell to diffuse localization again prior to completion of cell division. They determined that MurG-GFP localizes to the mid-cell in an FtsZ-dependent and MreB-independent manner (Aaron et al., 2007). This was demonstrated using a functional murG-mgfp fusion that provided the cell with its only copy of murG (Aaron et al., 2007). These observations are consistent with immunolocalization studies in E. coli, where MurG has also been shown to be localized to the mid-cell in an MreB-independent manner (Mohammadi et al., 2007). Conversly, Divakaruni et al. (2007) showed that MurG-mCherry displayed an MreB-dependent banded or punctate pattern along the cell with concentrations of localization at the poles. When the cells were treated with A22, MurG-mCherry became localized as single foci at the mid-cell or the poles, or both (Divakaruni et al., 2007). These observations were confirmed using immunolocalization (Divakaruni et al., 2007). How can the differences in MurG localization be reconciled? Both groups used C-terminal fluorescent protein fusions; however, murG-mCherry was expressed in the presence of wild-type murG in one case (Divakaruni et al., 2007), while murG-mgfp was the only copy of murG in the cell in the other case (Aaron et al., 2007). Thus, the differences in MurG localization

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observed by the two groups may be due to the differences in their respective MurG constructs. Despite the differences in the construction of the strains used to localize MurG, it is clear that mid-cell localization occurs and probably depends on FtsZ; however, it remains unclear if mid-cell localization of MurG depends on MreB. FtsZ is also required for transverse peptidoglycan synthesis to form the characteristic stalk crossbands (Divakaruni et al., 2007). The role of FtsZ in peptidoglycan synthesis of C. crescentus is consistent with results in E. coli, which show that FtsZ is required for peptidoglycan synthesis near the poles, indicating that the FtsZ function is not restricted to the mid-cell (Varma et al., 2007). 4.4.3.2. MreC, MltA, and MipA. In order for new peptidoglycan to be inserted into the cell wall at specific locations, localized cell wall degradation must occur. MltA, a lytic transglycoslase, and its interacting partner, MipA, are localized in a band-like pattern that is similar to that of PBP2, indicating that cell wall degradation and synthesis may be coupled (Divakaruni et al., 2007; Figge and Gober, 2003). In addition, both MltA and MipA are localized at the stalked cell pole where peptidoglycan synthesis is likely to occur (Divakaruni et al., 2007). Localization of MltA and MipA depends on the presence of MreC, but not MreB. In C. crescentus, MreC has been shown to interact with complexes of PBPs (Divakaruni et al., 2005). Taken together, these results suggest that MreC coordinates the localization of multiprotein complexes containing proteins for the degradation and synthesis of peptidoglycan. Efficient peptidoglycan synthesis leading to changes in cell shape is likely to require the coordination of a number of cell shape determining proteins, cell division proteins, and biosynthetic enzymes.

5. CONCLUDING REMARKS C. crescentus is a powerful model system for the study of cell cycle progression at the molecular level. Multiple levels of regulation, including both temporal and spatial regulation, are required to coordinate polar morphogenesis, chromosome replication, and cell division throughout the cell cycle. Cell cycle progression is mediated primarily by the activities of three global regulators, CtrA, DnaA, and GcrA, which are expressed in a specific temporal sequence. The actions of these three transcriptional regulators directly and indirectly impact on the expression of genes encoding both regulatory and structural proteins, at each phase of the cell cycle. The temporal expression of additional regulatory proteins, particularly

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location-specific two-component signal proteins, is necessary to fine-tune the processes that occur during each phase of the cell cycle. The ability of the cell to monitor progress in each phase of the cell cycle to direct the activation (or repression) of the regulators for subsequent steps in cell cycle progression has proven to be a critical means of feedback control. Regulators not only direct cell cycle progression, but are also subject to activation (or deactivation) based on the success of previous steps in the cell cycle. These regulatory feedback loops ensure that cell cycle progression does not continue until the previous phase is successfully completed and are critical for defining the physiology of the two distinct progeny cells. Despite significant progress in identifying and understanding the regulatory elements that direct cell cycle progression, a number of questions remain. Is cell cycle progression in C. crescentus truly hardwired? What signals, if any, are responsible for controlling cell cycle progression? Are multiple signals integrated to control global regulators? Recent studies indicate that c-di-GMP may act as an important signal in mediating cell cycle progression (for review see Jenal and Malone, 2006). Gain of motility, through flagellum assembly in the swarmer compartment of the predivisonal cell and flagellum activation in the new swarmer cell, requires a reduction in c-di-GMP levels. Conversely, loss of motility, holdfast formation, and stalk biogenesis during the swarmer to stalked cell transition requires an increase in c-di-GMP levels. The signal responsible for initiating production of c-di-GMP remains unknown. Given the large number of two-component signal transduction proteins involved in mediating cell cycle progression, it seems likely that additional signals remain to be discovered which impact cell cycle progression. Irrespective of the physiological and/or environmental signals which trigger the production, activation, and destruction of regulatory proteins, it is clear that successful progression through the cell cycle of C. crescentus requires multiple levels of regulation which must be precisely orchestrated in both time and space.

ACKNOWLEDGEMENTS We wish to thank members of the Brun laboratory for critical reading of this manuscript and providing unpublished data. Research in our laboratory is supported by grants GM077648 and GM51986 from the National Institutes of Health, MCB0731950 from the National Science Foundation, and by the Indiana METACyt Initiative of Indiana University, funded in part through a major grant from the Lilly Endowment, Inc. P.J.B.B. is supported by

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a postdoctoral National Institutes of Health National Research Service Award number F32AI072992 from the National Institute of Allergy and Infectious Diseases.

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Sulfur Metabolism in Phototrophic Sulfur Bacteria Niels-Ulrik Frigaard1 and Christiane Dahl2 1

Copenhagen Biocenter, Department of Biology, University of Copenhagen, Ole Maaløes Vej 5, 2200 Copenhagen N, Denmark 2 Institut fu¨r Mikrobiologie & Biotechnologie, Rheinische Friedrich-Wilhelms-Universita¨t Bonn, Meckenheimer Allee 168, D–53115 Bonn, Germany

ABSTRACT Phototrophic sulfur bacteria are characterized by oxidizing various inorganic sulfur compounds for use as electron donors in carbon dioxide fixation during anoxygenic photosynthetic growth. These bacteria are divided into the purple sulfur bacteria (PSB) and the green sulfur bacteria (GSB). They utilize various combinations of sulfide, elemental sulfur, and thiosulfate and sometimes also ferrous iron and hydrogen as electron donors. This review focuses on the dissimilatory and assimilatory metabolism of inorganic sulfur compounds in these bacteria and also briefly discusses these metabolisms in other types of anoxygenic phototrophic bacteria. The biochemistry and genetics of sulfur compound oxidation in PSB and GSB are described in detail. A variety of enzymes catalyzing sulfur oxidation reactions have been isolated from GSB and PSB (especially Allochromatium vinosum, a representative of the Chromatiaceae), and many are well characterized also on a molecular genetic level. Complete genome sequence data are currently available for 10 strains of GSB and for one strain of PSB. We present here a genome-based survey of the distribution and phylogenies of genes involved in oxidation of sulfur compounds in these strains. It is evident from biochemical and genetic analyses that the dissimilatory sulfur metabolism of these organisms is very complex and incompletely understood. This metabolism is modular in the sense that ADVANCES IN MICROBIAL PHYSIOLOGY, VOL. 54 ISBN 978-0-12-374323-7 DOI: 10.1016/S0065-2911(08)00002-7

Copyright r 2009 by Elsevier Ltd. All rights reserved

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individual steps in the metabolism may be performed by different enzymes in different organisms. Despite the distant evolutionary relationship between GSB and PSB, their photosynthetic nature and their dependency on oxidation of sulfur compounds resulted in similar ecological roles in the sulfur cycle as important anaerobic oxidizers of sulfur compounds.

Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anoxygenic phototrophic bacteria: physiology and taxonomy . . . . . . . 2.1. Purple Sulfur Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Green Sulfur Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Purple Non-Sulfur Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Aerobic Bacteriochlorophyll-Containing Bacteria . . . . . . . . . . . . . 2.5. Filamentous Anoxygenic Phototrophs . . . . . . . . . . . . . . . . . . . . 2.6. Heliobacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Anoxygenic phototrophic sulfur bacteria: ecology . . . . . . . . . . . . . . . 4. Transformations of sulfur compounds . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Purple Sulfur Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Green Sulfur Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. Purple Non-Sulfur Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4. Aerobic Bacteriochlorophyll-Containing Bacteria . . . . . . . . . . . . . 4.5. Filamentous Anoxygenic Phototrophs . . . . . . . . . . . . . . . . . . . . 4.6. Heliobacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Oxidative sulfur metabolism: enzymes and genes . . . . . . . . . . . . . . . 5.1. Oxidation of H2S . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2. Oxidation of Polysulfides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3. Uptake of External Sulfur . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.4. Properties of Sulfur Globules . . . . . . . . . . . . . . . . . . . . . . . . . . 5.5. Oxidation of Stored Sulfur to Sulfite . . . . . . . . . . . . . . . . . . . . . 5.6. Oxidation of Sulfite to Sulfate . . . . . . . . . . . . . . . . . . . . . . . . . . 5.7. Oxidation of Thiosulfate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.8. Other Enzymes Related to Sulfur Compound Oxidation . . . . . . . 6. Evolution of sulfur metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7. Sulfate assimilation in anoxygenic phototrophic bacteria . . . . . . . . . . 7.1. Historical Aspects and General Outline of the Pathway . . . . . . . . 7.2. Occurrence of Sulfate Assimilation in Anoxygenic Phototrophic Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.3. Uptake of Sulfate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.4. Activation of Sulfate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.5. Reduction to Sulfite . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.6. Reduction of Sulfite to Sulfide . . . . . . . . . . . . . . . . . . . . . . . . . . 7.7. Incorporation of Sulfide . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. 2.

105 106 107 109 112 113 114 114 115 116 118 118 123 123 124 125 126 126 130 134 135 138 143 148 155 162 165 167 167 169 174 174 175 176 177 177 179 179

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 105

ABBREVIATIONS ABC bacteria Alc. APAT APS Atb. BChl Cba. Chl. Cfl. Chp. DMSP Dsv. Ect. EPS FAP bacteria GSB Hlr. LGT LH complex MPT PAPS Pcs. PSB PSR Rba. RC RLP Rpl. rRNA SGP SQR Tca. Thb. Tms. Wol.

aerobic bacteriochlorophyll-containing bacteria Allochromatium Adenylylsulfate phospateadenylyl transferase adenosine-5u-phosphosulfate Acidithiobacillus bacteriochlorophyll Chlorobaculum Chlorobium Chloroflexus Chloroherpeton dimethylsulfoniopropionate Desulfovibrio Ectothiorhodospira extracellular polymeric substances filamentous anoxygenic phototrophic bacteria green sulfur bacteria Halorhodospira lateral gene transfer light-harvesting complex molybdopterin 3u-phosphoadenosine-5u-phosphosulfate Paracoccus purple sulfur bacteria polysulfide reductase Rhodobacter (photosynthetic) reaction center RuBisCO-like protein Rhodoplanes ribosomal RNA sulfur globule protein sulfide:quinone reductase Thiocapsa Thiobacillus Thiomicrospira Wolinella

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1. INTRODUCTION The most important common property of phototrophic prokaryotes is the possession of tetrapyrrole pigments and a photosynthetic apparatus enabling light-dependent electron transfer and energy conservation processes. Only a few groups, all within the domain of Bacteria, possess these properties. Fundamental physiological differences exist between these groups: (a) the oxygenic phototrophic bacteria (cyanobacteria) that use water as photosynthetic electron donor and thus produce molecular oxygen as a product; (b) the anoxygenic phototrophic bacteria (groups of purple and green bacteria and heliobacteria) that use electron donors other than water, such as reduced sulfur compounds, organic compounds, hydrogen or ferrous iron; and (c) the aerobic anoxygenic bacteriochlorophyll-containing (ABC) bacteria, a group of poorly characterized Proteobacteria with a predominantly chemoheterotrophic physiology but with the potential to produce photosynthetic pigment–protein complexes and to perform lightmediated electron transport (Imhoff, 2008). The utilization of reduced sulfur compounds, most notably sulfide, as photosynthetic electron donors is – although to a different extent – common among almost all groups of phototrophic bacteria. Classical in this respect are the purple sulfur bacteria (PSB; families Chromatiaceae and Ectothiorhodospiraceae) and green sulfur bacteria (GSB; family Chlorobiaceae) all of which utilize reduced sulfur compounds as electron donors and which collectively are known as phototrophic sulfur bacteria. A number of purple ‘‘non-sulfur’’ bacteria, some members of the filamentous anoxygenic phototrophs (FAP; relatives of Chloroflexus, previously known as green gliding bacteria or green non-sulfur bacteria), and a few representatives of the strictly anaerobic heliobacteria are also able to oxidize reduced sulfur compounds during phototrophic growth. Even certain species of cyanobacteria can perform anoxygenic photosynthesis at the expense of sulfide as an electron donor (Cohen et al., 1975). Photoautotrophic growth with sulfur compounds has so far not been described for any of the ABC bacteria. We will first introduce the physiology, taxonomy, and ecology of anoxygenic phototrophic bacteria. Then the sulfur-oxidizing capabilities of the various groups of anoxygenic phototrophic bacteria will be described. It should be emphasized that some older reviews still serve as a valuable source of information especially regarding sulfur oxidation patterns by phototrophic sulfur bacteria (Brune, 1989, 1995b). This review focuses on sulfur compound oxidation in the PSB (Chromatiaceae and Ectothiorhodospiraceae) and in the GSB (Chlorobiaceae) with special attention on the current

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 107

knowledge of the biochemical details of the metabolic pathways employed in these groups. Within the past few years, much useful information on this subject has been derived from comparative analyses of the various available genome sequences from GSB and purple bacteria (Frigaard and Bryant, 2008a; Sander and Dahl, 2008) and this will surely continue to be the case as more genome sequences become available. We also give a brief overview of the assimilatory sulfate reduction metabolism of anoxygenic phototrophic bacteria. Regarding the metabolism of organosulfur compounds in anoxygenic phototrophic bacteria, we refer the reader to recent articles by Cook and coworkers (Baldock et al., 2007; Denger et al., 2004, 2006).

2. ANOXYGENIC PHOTOTROPHIC BACTERIA: PHYSIOLOGY AND TAXONOMY The major groups of phototrophic bacteria are well-distinguished on the basis of fundamental structural, molecular, ecological, and physiological properties (Table 1). While the oxygenic cyanobacteria represent a separate large phylogenetic lineage not discussed further here, the anoxygenic phototrophic bacteria are found in only five different major phylogenetic lineages: (1) the FAP bacteria (Chloroflexus and relatives), (2) the GSB (comprising the family Chlorobiaceae), (3) the heliobacteria (comprising the family Heliobacteriaceae whose members are related to Gram-positive bacteria), (4) the purple bacteria (divided into purple sulfur and purple nonsulfur species, all of which belong to either the Alpha-, Beta- or Gammaproteobacteria and the ABC bacteria that also belong to the Alpha-, Beta- and Gammaproteobacteria, and (5) the phototrophic acidobacteria. The latter group is a recently discovered lineage of phototrophic bacteria of which only a single organism in enriched culture is known (Bryant et al., 2007). This bacterium, ‘‘Candidatus Chloroacidobacterium thermophilum,’’ belongs to the poorly characterized phylum Acidobacteria and was derived from microbial mats of alkaline siliceous hot springs. Like GSB, it synthesizes BChl a and c, chlorosomes, FMO protein, and type I reaction centers, but unlike GSB it grows under aerobic conditions. It is not yet available in pure culture and it is not known to oxidize sulfur compounds for growth. The variations in photosynthetic reaction centers (RCs), light-harvesting structures, and different CO2 fixation pathways among these groups are quite remarkable (Table 1). All known photosynthetic RCs can be divided into two main groups: type I, in which a Fe–S cluster is the terminal electron

108

Table 1 Major groups of anoxygenic phototrophic bacteria Purple sulfur bacteria (PSB)

Purple non-sulfur bacteria

Phylogenetic affiliation

Gammaproteobacteria Alpha- and Betaproteobacteria

Type of bacteriochlorophyll

BChl a, b

Aerobic bacteriochlorophyllcontaining (ABC) bacteria

Filamentous anoxygenic phototrophic (FAP) bacteria

Heliobacteria

Alpha- , Beta- and Chlorobiaceae Gammaproteobacteria (phylum Chlorobi)

Chloroflexaceae (phylum Chloroflexi)

BChl a, b

BChl a, Zn-BChl a

BChl a. Some also have BChl c, d

Heliobacteriaceae (phylum Firmicutes, order Clostridiales) BChl g

Pheophytin/quinone (type II) Antenna complexes LHI and LHII in intracytoplasmic membranes Preferred electron donor Reduced sulfur for phototrophic growth compounds, hydrogen, (organic compounds) Chemotrophic growth /þ Pathway of autotrophic Calvin cycle CO2 fixation

Pheophytin/quinone (type II) LHI and LHII in intracytoplasmic membranes Organic compounds, hydrogen, (sulfide, ferrous iron)

Pheophytin/ quinone (type II) LHI. Most have no LHII

þ Calvin cycle

þ If present: Calvin cycle. No photoautotrophic growth

Major habitat

Anoxic marine and freshwater

Eutrophic aquatic environments

Reaction center type

a

Anoxic marine and freshwater

None known

Green sulfur bacteria (GSB)

BChl c, d, e (a minor component in all) Fe-S (type I) FMO protein, chlorosomes Sulfide, hydrogen, (ferrous iron in some) – Reductive TCA cycle

Anoxic marine and freshwater

Pheophytin/ quinone (type II) LHI-like. Some have also chlorosomes Organic compounds, (sulfide)

Fe-S (type I) Nonea Organic compounds

/þ þ HydroxyproNone pionate pathway (Chloroflexus), Calvin cycle (Oscillochloris) Hot springs, Soil meta- and hypolimnion of stratified lakes, shallow freshwater pools

Heliobacteria contain no other antenna chlorophylls than those associated with the photosynthetic reaction center in the cytoplasmic membrane.

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Group

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 109

acceptor, and type II, in which a quinone is the terminal electron acceptor. Photosystem II of oxygenic phototrophic bacteria and eukaryotes and the purple bacterial RC are members of the type II group, whereas photosystem I of oxygenic phototrophic bacteria and eukaryotes and the homodimeric RCs of GSB and heliobacteria are members of the type I group. A combination of the two types of photosystems as found in oxygenic phototrophs (all cyanobacteria and all eukaryotic phototrophs) is required for the thermodynamically unfavorable utilization of water as an electron donor for photosynthesis. Due to the simpler architecture of their photosynthetic apparatus, all anoxygenic phototrophic bacteria depend on electron donors with standard potentials more negative than that of water (e.g., reduced sulfur compounds, hydrogen, and acetate). This results in comparatively narrower ecological niches for these bacteria than for cyanobacteria in extant ecosystems (Overmann, 2008). In the following sections, the properties of the various groups of anoxygenic phototrophic bacteria will briefly be described. Details on the transformations of sulfur compounds are described in Section 4.

2.1. Purple Sulfur Bacteria The PSB belong to the Gammaproteobacteria and fall into two families, the Chromatiaceae and the Ectothiorhodospiraceae. Both form coherent groups on the basis of their 16S ribosomal RNA (rRNA) sequences. The most important and easily recognized and distinguishing feature among these two families is the formation of sulfur globules within the confines of the cell wall in Chromatiaceae during growth on sulfide, polysulfides, thiosulfate, or elemental sulfur (Fig. 1A), while sulfur globules accumulate extracellularly in members of the Ectothiorhodospiraceae (Brune, 1995b; Dahl, 2008). The Ectothiorhodospiraceae member Thiorhodospira sibirica is an exception as it forms both extra- and intracellular sulfur globules (Bryantseva et al., 1999). All PSB fix CO2 using the Calvin cycle and their photosynthetic RC is type II. Very little genome sequence information is available for the phototrophic members of this group despite their diversity and ecological significance. The genome sequence of Halorhodospira halophila SL1 (DSMZ 244T) has been determined (www.jgi.doe.gov). The genome of Thermochromatium tepidum MC (DSMZ 3771T) has been sequenced by a commercial enterprise and is not publicly available. However, the genome of the important model organism Allochromatium vinosum DSMZ 180T is in the process of being sequenced by the JGI.

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Figure 1 Phototrophic sulfur bacteria with sulfur globules. (A) Thiocapsa sp. with intracellular sulfur globules, light microscopy, magnification  2000; (B) Chlorobium phaeovibrioides strain 9130 with extracellular sulfur globules, magnification  2000; (C) Thiocystis violaceae DSMZ 212 with intracellular sulfur globules. The vesicular intracytoplasmic membranes (‘‘chromatophores’’) are associated with the sulfur globules in a highly organized manner, magnification  22,000. (D) Part of a Thiocapsa roseopersicina (strain 8811) cell, the sulfur globule envelope is clearly visible, magnification  179,550. As a result of the preparation of the cells for electron microscopy, the localization of sulfur globules is visible as ‘‘holes’’ in the electron micrographs. All photos kindly provided by Hans G. Tru¨per, Bonn.

2.1.1. Chromatiaceae Most species have the vesicular type of intracytoplasmic membranes (Fig. 1C and D). Many species are motile by means of flagella, and about one-third of

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the species of the Chromatiaceae have gas vesicles. The gas-vesicle-bearing Chromatiaceae typically colonize low-light stratified environments. Two major physiological groups of Chromatiaceae can be distinguished: the metabolically versatile species and the metabolically specialized species. The specialized species (among them many large-celled species, e.g., Chromatium okenii, Allochromatium warmingii, and Isochromatium buderi) depend on strictly anaerobic conditions and are obligately phototrophic. Sulfide is required whereas thiosulfate and hydrogen are not used as electron donors. Only acetate and pyruvate are photoassimilated in the presence of CO2 and sulfide. Assimilatory sulfate reduction does not occur. The versatile species (among them many small-celled species, e.g., Allochromatium vinosum and Allochromatium minutissimum) use thiosulfate in addition to sulfide and elemental sulfur as electron donors (Imhoff, 2005a). Most of these species are capable of assimilatory sulfate reduction, can grow photoorganoheterotrophically in the absence of reduced sulfur compounds, and can grow as chemolithotrophs on reduced sulfur compounds (Gorlenko, 1974; Ka¨mpf and Pfennig, 1980; Kondrat’eva et al., 1976). Some species can even grow as chemoorganotrophs in which case the addition of sulfide or thiosulfate as a sulfur source is required because the assimilation of sulfate is repressed under aerobic conditions (Kondrat’eva et al., 1981). 2.1.2. Ectothiorhodospiraceae The Ectothiorhodospiraceae are distinguished from the Chromatiaceae by lamellar intracytoplasmic membrane structures, by significant differences in polar lipid composition, and by the dependence on saline and alkaline growth conditions. All species are motile by polar flagella. Ect. vacuolata harbors gas vesicles. Four phototrophic genera of Ectothiorhodospiraceae are validly described: Ectothiorhodospira, Halorhodospira, Thiorhodospira, and Ectothiorhodosinus. Formally, the genus Ectothiorhodosinus (Gorlenko et al., 2004) has no standing in the nomenclature. All species of the genus Ectothiorhodospira grow well under anoxic conditions in the light with reduced sulfur compounds as photosynthetic electron donors and in the presence of organic carbon sources and inorganic carbonate. Several species of the genus Ectothiorhodospira are also able to grow chemolithotrophically on sulfur compounds (e.g., Ectothiorhodospira haloalkaliphila and Ectothiorhodospira shaposhnikovii) (Imhoff, 2005b). Non-phototrophic genera of Ectothiorhodospiraceae include Alkalilimnicola (Yakimov et al., 2001), Alkalispirillum (Rijkenberg et al., 2001), and

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Arhodomonas (Adkins et al., 1993). Alkalilimnicola ehrlichii is capable of anaerobic chemoautotrophic growth using hydrogen, sulfide, or thiosulfate as electron donor and nitrate as electron acceptor (Hoeft et al., 2007). Other non-phototrophic members of Ectothiorhodospiraceae are also able to oxidize sulfur compounds, including species of Thioalkalivibrio and Thioalkalispira (Sorokin and Kuenen, 2005; Sorokin et al., 2001).

2.2. Green Sulfur Bacteria The GSB form a cluster of closely related genera united in a single family, Chlorobiaceae, which represents the only members of the phylum Chlorobi that have been isolated in pure culture (Garrity and Holt, 2001a). The taxonomy and nomenclature of GSB have recently been extensively modified to reflect recent progress in understanding their phylogeny (Imhoff, 2003). BChl c, d, and e are the major light-harvesting pigments. These pigments are organized into large organelles known as chlorosomes in all GSB characterized to date (Frigaard and Bryant, 2006). Chlorosomes allow a highly efficient capture of light energy and, as a result of this, growth at remarkably low light intensities. The only other organisms in which chlorosomes have been found are the ‘‘green FAPs’’ (see Section 2.5) and the phototrophic acidobacteria (see introduction to Section 2). BChl a is always present in the chlorosomes, the peripheral Fenna–Matthews–Olson (FMO) antenna protein, and the photosynthetic RC. In addition, GSB contain various carotenoids of the chlorobactene and isorenieratene series. Intracytoplasmic membranes are not formed. All characterized GSB have similar metabolic properties: they are strictly anaerobic, obligately phototrophic, and obligately autotrophic. They can grow with CO2 as sole carbon source but they will photoassimilate acetate if CO2 is present. In contrast to the purple bacteria, CO2 is fixed via the reductive tricarboxylic acid cycle (also known as the reverse citric acid cycle). Most strains use electrons derived from oxidation of sulfide, thiosulfate, elemental sulfur, and H2, but a few characterized strains can also oxidize Fe2þ (Heising et al., 1999). Sulfur is deposited outside of the cells (Fig. 1B). Some GSB form gas vesicles. One benthic species, Chloroherpeton thalassium, is motile by gliding. All known pelagic isolates of GSB are non-flagellated, although some have acquired motility by forming highly specific symbioses with motile chemoheterotrophic bacteria (Overmann and Schubert, 2002). GSB are commonly found in anoxic and sulfide-rich freshwater and estuarine environments, either in the water column, in sediments, or within microbial mats. They have also recently been found in the anoxic zone 100 m

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below the surface of the Black Sea (Manske et al., 2005; Overmann et al., 1992a), on deep-sea hydrothermal vents in the Pacific Ocean (Beatty et al., 2005), and in the microbial mats of Octopus and Mushroom Springs in Yellowstone National Park (Ward et al., 1998). Compared to other anoxygenic phototrophic bacteria, a large amount of genome sequence information is available for the GSB. Currently, the genome sequences of 10 strains of GSB are publicly available (www. ncbi.nlm.nih.gov): Cba. tepidum TLS (DSMZ 12025T), Chl. chlorochromatii CaD3, Chl. clathratiforme BU1 (DSMZ 5477T), Chl. ferrooxidans DSMZ 13031T, Chl. limicola DSMZ 245T, Chl. luteolum DSMZ 273T, Chl. phaeobacteroides DSMZ 266T, Chl. phaeovibrioides DSMZ 265, Prosthecochloris sp. BS1, Ptc. aestuarii SK413 (DSMZ 271T). The genomes of Cba. parvum NCIMB 8327 (DSMZ 263T) and Chp. thalassium ATCC 35110T are in the process of being sequenced.

2.3. Purple Non-Sulfur Bacteria The group of purple non-sulfur bacteria is by far the most diverse group of the photosynthetic purple bacteria. Representatives are found within the Alpha- and the Betaproteobacteria (Imhoff et al., 2005). Their diversity is reflected by highly variable cell morphology, internal membrane structure, carotenoid composition, utilization of carbon sources, and by the kind of electron donors used for photosynthesis. The intracytoplasmic membranes that harbor the photosynthetic apparatus can be small finger-like intrusions, vesicles, or various types of lamellae. In general, purple non-sulfur bacteria grow preferentially photoheterotrophically under anoxic conditions in the light. Mixotrophic growth is common during which fermentation products of other bacteria are used as the preferred carbon source. Some species even lack the ability to grow photolithoautotrophically (Imhoff et al., 2005). Most purple non-sulfur bacteria are metabolically very flexible and can switch between anoxygenic photosynthesis, fermentation, and aerobic or anaerobic respiration using nitrate, dimethyl sulfoxide, or trimethylamine-N-oxide as electron acceptor. Many species can grow photoautotrophically with hydrogen or sulfide as electron donor. Many of these organisms do not oxidize sulfide completely to sulfate but form sulfur globules as the terminal sulfur product. Purple non-sulfur bacteria are widely distributed in nature as indicated by their isolation from very diverse sources including pond water, marine coastal sediments, sewage, swine waste lagoons, and even earthworm droppings (Larimer et al., 2004). The photosynthetic RC of purple non-sulfur bacteria is type II and, if present, CO2 fixation occurs by the Calvin cycle. Due

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to interest in their physiological diversity and their use as model organisms, genome sequence information has been generated for several purple nonsulfur bacteria, including strains of Rhodobacter capsulatus (SB1003), Rhodobacter sphaeroides (2.4.1, ATCC 17025, ATCC 17029), Rhodospirillum rubrum (ATCC 11170T), Rubrivivax gelatinosus (IL144, PM1), and Rhodopseudomonas palustris (BisA53, BisB18, BisB5, CGA009, HaA2, TIE-1).

2.4. Aerobic Bacteriochlorophyll-Containing Bacteria The ABC bacteria are a relatively recently discovered group of phototrophic purple bacteria whose physiology and ecology is not completely understood (Hiraishi and Shimada, 2001). All ABC bacteria are capable of synthesizing BChl a under aerobic conditions and appear to be capable of photosynthetic growth only under aerobic conditions. The content of BChl in the cells is generally low and the BChl synthesis is strongly inhibited by light. The BChl found in most of these bacteria is BChl a esterified with phytol as in other purple bacteria. However, Acidiphilium species produce zinc-chelated BChl a as the major component. All ABC bacteria studied so far contain an active photosynthetic RC and LH1 light-harvesting complexes with BChl a. Peripheral LH2-like antennae are absent in most species. All ABC bacteria share aerobic chemoorganotrophy as the preferred mode of growth (Yurkov, 2006). While they are able to gain energy from their photosynthetic machinery, they do not grow solely at the expense of light energy. None of the characterized ABC bacteria are able to grow photolithotrophically with sulfur compounds as electron donors. Until recently, all known species of ABC bacteria were believed to belong to the Alphaproteobacteria, except Roseotales depolymerans which belongs to the Betaproteobacteria. ABC bacteria do not form a homogeneous cluster within the Alphaproteobacteria but are closely interspersed with phototrophic and non-phototrophic species (Imhoff and Hiraishi, 2005). The work of Fuchs et al. (2007) was the first to unequivocally prove an ABC bacterium (Congregibacter litoralis KT71) among the Gammaproteobacteria. A few strains of ABC bacteria belonging to the Roseobacter and Erythrobacter genera have had their genome sequenced. In addition, the genome sequence of Congregibacter litoralis KT71 is nearly complete (Fuchs et al., 2007). These sequences may help explain the physiology of this group of bacteria.

2.5. Filamentous Anoxygenic Phototrophs The FAP bacteria belong to one of the deepest branching bacterial phyla (Chloroflexi) of the Bacteria. This phylum contains both phototrophic and

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 115

non-phototrophic members, characterized by a filamentous morphology and gliding motility. Most but not all FAP bacteria are facultatively aerobic and preferentially utilize organic substrates in their phototrophic or chemotrophic metabolism. Chloroflexus, Chloronema, Oscillochloris, Chlorothrix, Roseiflexus, and Heliothrix are well-described genera, although no species of Chloronema, Chlorothrix, and Heliothrix have been characterized in pure culture (Hanada and Pierson, 2002). The FAP bacteria have taxonomically been organized in the family Chloroflexaceae (Garrity and Holt, 2001b) although it has been suggested that all the above-mentioned genera other than Chloroflexus should be organized into new families (Tourova et al., 2006). All FAP bacteria contain BChl a as a component of the light-harvesting antenna and a type II photosynthetic RC. In addition, four of the genera Chloroflexus, Chloronema, Oscillochloris, and Chlorothrix contain chlorosomes that often cause the cells to appear green (thus, the name green filamentous bacteria or ‘‘green FAPs’’ has been used to describe these organisms). Chlorosomes are large light-harvesting antenna structures that are attached to the cytoplasmic membrane and contain BChl c or d (Frigaard and Bryant, 2006). The only other organisms that contain chlorosomes are GSB and the phototrophic acidobacteria (see introduction to Section 2). FAP bacteria that do not contain chlorosomes have an orange, red, or pink coloration, and hence sometimes are called ‘‘red FAPs.’’ In Chloroflexus species, CO2 is fixed by the 3-hydroxypropionate cycle (Ivanovskii et al., 1993) while the Calvin cycle appears to be used in Oscillochloris trichoides DG-6 (Ivanovsky et al., 1999). The genome sequences of the following FAP strains have been, or are in the process of being, determined: Chloroflexus aggregans DSMZ 9485T, Chloroflexus aurantiacus J-10-fl (DSMZ 635T), Chloronema giganteum UdG9001, Oscillochloris sp. UdG9002, Roseiflexus castenholzii DSMZ 13941T, Roseiflexus sp. RS–1, Heliothrix oregonensis, and ‘‘Candidatus Chlorothrix halophila’’ (www.genomesonline.org).

2.6. Heliobacteria Heliobacteria occur in nature primarily in soil and in this regard they differ significantly in their ecology from purple and green phototrophs. Cells of heliobacteria stain Gram-negative, although the phylogenetic position of the group is together with typical Gram-positive bacteria. They belong to the Firmicutes phylum and group within the Bacillus/Clostridium lineage. Spores are formed, gas vesicles are not present, and their metabolism is strictly anaerobic. Heliobacteria can grow both photo- and chemotrophically. Photoheterotrophic growth occurs on a very limited number of organic

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compounds as carbon sources. Chemotrophic growth in the dark occurs by fermentation of pyruvate. Photoautotrophic growth on CO2/H2 or CO2/H2S has not been demonstrated with any species of heliobacteria. However, some strains oxidize sulfide to elemental sulfur during photoheterotrophic growth (Bryantseva et al., 2000). Heliobacteria contain a unique photosynthetic pigment called BChl g, which contains a vinyl group on ring I and an ethylidine group on ring II. This BChl distinguishes heliobacteria from all other phototrophic bacteria (Madigan, 2001) and gives them their characteristic brown color (Gest and Favinger, 1983). The pigment system of heliobacteria is simple: it consists of a single pigment–protein complex that is embedded in the cytoplasmic membrane and contains both the type I reaction center and all BChls that have an antenna function. Heliobacteria lack differentiated photosynthetic internal membranes, such as the membrane vesicles or lamellae of purple bacteria or the chlorosomes of green bacteria (Neerken and Amesz, 2001). The only heliobacterium for which genome sequence information is available is Heliobacterium modesticaldum Ice1 (ATCC 51547) (http:// genomes.tgen.org/helio.html).

3. ANOXYGENIC PHOTOTROPHIC SULFUR BACTERIA: ECOLOGY The various photosynthetic tetrapyrrole and carotenoid pigments in phototrophic bacteria lead to a distinct coloration of the cells ranging from green, brown, and red to purple. Older reports about colored waters or sediments occurring in various natural environments (Bavendamm, 1924; Engelmann, 1882, 1888; van Niel, 1931; Winogradsky, 1887) can nowadays be explained by massive blooms of anoxygenic phototrophic sulfur bacteria forming in the water column or in aquatic sediments under appropriate conditions when both sulfide and light are available and oxygen is deficient or lacking (Overmann, 2008). We know now that these phototrophic sulfur bacteria comprise the PSB and the GSB whose ecology and sulfur metabolism share many similarities (Brune, 1989, 1995b; Overmann, 2008; van Gemerden and Mas, 1995), although their phylogeny is rather different (see Section 2). In natural environments light and sulfide occur in opposing gradients. Therefore, growth of phototrophic sulfur bacteria is restricted to a narrow zone. In open waters, like lakes or lagoons, stratification of oxic and anoxic water layers is maintained by density differences. In benthic environments,

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gradients of sulfide are much steeper due to the higher rates of sulfate reduction and a lower frequency of turbulent mixing (Overmann, 2008). In some habitats of phototrophic sulfur bacteria, particularly in intertidal sediments, redox conditions change rapidly within hours. Certain species of PSB have adapted to these conditions and can switch to an aerobic chemolithotrophic growth mode and oxidize sulfide or thiosulfate or both with molecular oxygen. Under these conditions, the synthesis of pigments and pigment-binding proteins is repressed and the cells become colorless. In contrast, all GSB are obligately anaerobic and obligately phototrophic. These properties enforce a greater constraint on the possible niches available to GSB. Phototrophic sulfur bacteria typically occur in non-thermal aquatic ecosystems, although a few moderately thermophilic members of these groups have been described from hot spring mats, for example, the GSB Chlorobaculum tepidum and the PSB Thermochromatium tepidum (Castenholz et al., 1990; Madigan, 1986). Among the PSB, most members of the Chromatiaceae are typically found in freshwater and marine environments, whereas the Ectothiorhodospiraceae inhabit hypersaline waters. Some members of the GSB have also been isolated from a hypersaline athallassohaline lake in Spain (Vila et al., 2002). In coastal and most lacustrine waters, metabolically specialized Chromatiaceae dominate (van Gemerden and Mas, 1995). These organisms are obligately photolithotrophic, lack assimilatory sulfate reduction, cannot reduce nitrate, and assimilate only a few carbon sources. This is to be expected since metabolically versatile species of the Chromatiaceae have no strong selective advantage in most pelagic habitats (Overmann, 2008). In some lakes, the absorption of ultraviolet and blue light by humic substances is so significant that green-colored species of GSB have a selective advantage over their brown-colored counterparts and over PSB (Parkin and Brock, 1980). In most phototrophic organisms, the response to very low light intensities is an increase in the size of the photosynthetic unit (Drews and Golecki, 1995; Overmann, 2008): In purple bacteria, the size of the photosynthetic antenna is in the range of 20–2000 BChl a molecules per reaction center (Zuber and Cogdell, 1995). However, the specialized photosynthetic antenna organelles of GSB, the chlorosomes, are significantly larger than those of any other phototroph resulting in a ratio of about 5000–8000 BChl c molecules per RC (Frigaard et al., 2003). The theoretical quantum requirement for the CO2 fixation in PSB is 8–10.5 mol quanta/(mol CO2), but only 3.5–4.5 mol quanta/(mol CO2) for GSB (Brune, 1989). In general, GSB exhibit lower maintenance energy requirements and higher sulfide tolerance than PSB and are thereby especially well adapted to low-light habitats (Overmann, 2008).

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4. TRANSFORMATIONS OF SULFUR COMPOUNDS In the following section the sulfur oxidation capabilities of the various groups of anoxygenic phototrophic bacteria will be briefly described. PSB and GSB preferentially use reduced sulfur compounds as electron donors during photolithoautotrophic growth. Their sulfur-metabolizing capabilities are summarized in Table 2. Sulfur oxidation capabilities in the ABC bacteria, the FAP bacteria, and the heliobacteria are rather limited. Information about the enzymes involved in the bacteria is in most cases not available.

4.1. Purple Sulfur Bacteria Although both PSB and GSB photooxidize sulfide, PSB prefer lower sulfide concentrations than GSB in general. However, the tolerance towards high sulfide concentrations is strain-specific. For example, among the PSB the ability to tolerate sulfide extends to a concentration of up to 11 mM, as has been shown for Thiorhodococcus drewsii (Zaar et al., 2003). 4.1.1. Chromatiaceae Here we focus on those members of the Chromatiaceae that grow obligately or facultatively photoautotrophically. Closely related organisms may be non-phototrophic (e.g., Arsukibacterium ikkense (Schmidt et al., 2007) or members of the genus Rheinheimera (Brettar et al., 2002; Romanenko et al., 2003)). All phototrophic members of the Chromatiaceae use sulfide and sulfur of the oxidation state zero as photosynthetic electron donors (Table 2). Specialized species (e.g., Allochromatium warmingii, Isochromatium buderi, and Thiospirillum jenense) are unable to use thiosulfate, hydrogen, or organic compounds as electron donors (Imhoff et al., 1998). However, some of these species have never been tested for their ability to use thiosulfate and sulfite. A range of versatile species uses several different sulfur compounds including thiosulfate or sulfite. Those members of the Chromatiaceae that have been studied with respect to the utilization of externally added polysulfides (e.g., Alc. vinosum and Tca. roseopersicina) readily use these compounds as photosynthetic electron donors (Steudel et al., 1990; van Gemerden, 1987; Visscher et al., 1990). Utilization of polysulfides is not surprising because polysulfides are found in the growth medium of Alc. vinosum as intermediates in the oxidation of sulfide to internally stored sulfur (Prange et al., 2004).

Sulfur-metabolizing capabilities of phototrophic sulfur bacterial species or genera

Genus or species

Sulfur substrates used

Intermediates on sulfide

End product

Green Sulfur Bacteria (Chlorobiaceae) Ancalochloris perfilieviia,b Chlorobaculum chlorovibrioides Chlorobaculum limnaeum

Sulfide Sulfide, sulfur

Sulfur, EC Sulfur, EC

Sulfide, sulfur, thiosulfate in strain 1549 Sulfide, thiosulfate, sulfur, (tetrathionate in strain NCIB 8346) Sulfide, thiosulfate, sulfur Sulfide, thiosulfate, sulfur Sulfide Sulfide, thiosulfate in strain DSM 5477T, sulfur – Sulfide, thiosulfate only in strains 1630, 9330 and DSM 257, sulfur Sulfide, sulfur Sulfide, sulfur

Chlorobaculum parvum Chlorobaculum tepidum Chlorobaculum thiosulfatiphilum Chlorobium chlorochromatii Chlorobium clathratiforme Chlorobium ferrooxidans Chlorobium limicola Chlorobium luteolum Chlorobium phaeobacteroides Chlorobium phaeovibrioides Chloroherpeton thalassium Prosthecochloris aestuarii Prosthecochloris vibrioformis

Sulfide, sulfur, thiosulfate in strain DSM 265 Sulfide Sulfide, sulfur Sulfide, sulfur

Sulfate assimilation

Chemoautotrophic growth

Sulfate Sulfate



 

Sulfur, EC

Sulfate





Sulfur, EC

Sulfate





Sulfur, EC Sulfur, EC

Sulfate Sulfate

 

 

nd Sulfur, EC

nd Sulfate

 

 

– Sulfur, EC

Sulfate

þ 

 

Sulfur, EC Sulfur, EC

Sulfate Sulfate

c 

 

Sulfur, EC

Sulfate





Sulfur, EC Sulfur, EC Sulfur, EC

Sulfate Sulfate Sulfate

  

   (Continued )

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 119

Table 2

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Table 2 (Continued )

Purple Sulfur Bacteria Chromatiaceae Allochromatium Chromatium Halochromatium Isochromatium Lamprobactera Lamprocystis Marichromatium Rhabdochromatium Thermochromatium Thioalkalicoccus Thiobaca Thiocapsa Thiococcus Thiocystis

Sulfur substrates used

Intermediates on sulfide

End product

Sulfate assimilation

Chemoautotrophic growth

Sulfide, sulfur, thiosulfate, sulfite, (latter two not in Alc. warmingii) Sulfide, sulfur Sulfide, thiosulfate, sulfur, sulfite Sulfide, sulfur Sulfide, thiosulfate, sulfur Sulfide, thiosulfate, sulfur Sulfide, thiosulfate, sulfur, sulfite (only Mch. gracile) Sulfide, thiosulfate, sulfur Sulfide, sulfur Sulfide, sulfur Sulfide Sulfide, thiosulfate, sulfur, sulfite (only Tca. litoralis and Tca. pendens) Sulfide, sulfur Sulfide, thiosulfate (not in Tcs. gelatinosa), sulfur, sulfite in some strains

Sulfur, IC

Sulfate

þ (not in Alc. warmingii)

Some species

Sulfur, IC Sulfur, IC

Sulfate Sulfate

 

Sulfur, Sulfur, Sulfur, Sulfur,

IC IC IC IC

Sulfate S0 and sulfate Sulfate Sulfate

  /nd þ/

 þ (sulfide, thiosulfate)  þ þ/ þ/

Sulfur, Sulfur, Sulfur, Sulfur, Sulfur,

IC IC IC IC IC

Sulfate Sulfate Sulfate Sulfate Sulfate

nd nd nd nd þ/

   nd þ/

Sulfur, IC Sulfur, IC

Sulfate Sulfate

 þ in some strains

 þ

NIELS-ULRIK FRIGAARD AND CHRISTIANE DAHL

Genus or species

Thiolamprovum Thiopediaa Thiorhodococcus Thiorhodovibrio Thiospirilluma Ectothiorhodospiraceae Ectothiorhodosinusd Ectothiorhodospira Halorhodospira Thiorhodospira

Sulfide, Sulfide, Sulfide, sulfite Sulfide, Sulfide, Sulfide, Sulfide, Sulfide,

sulfur sulfur thiosulfate, sulfur,

Sulfur, IC Sulfur, IC Sulfur, IC

Sulfate Sulfate Sulfate

nd nd 

  þ

thiosulfate, sulfur sulfur thiosulfate, sulfur sulfur sulfur

Sulfur, Sulfur, Sulfur, Sulfur, Sulfur,

Sulfate Sulfate Sulfate Sulfate Sulfate

   nd nd

þ  þ/ þ 

Sulfur, EC Polysulfide, sulfur, EC

Sulfate Sulfate

nd þ in some species

nd þ in some species

Sulfur, EC

Sulfur or sulfate



Sulfur, EC & IC

Sulfate

þ in Hlr. halochloris nd

Sulfide, thiosulfate Sulfide, thiosulfate (not in Ect. marismortui), sulfur, sulfite (nd for some species) Sulfide, thiosulfate, sulfur only in Hlr. halophila Sulfide, sulfur

IC IC IC IC IC



The tabulated data were mostly taken from Overmann (2001); Imhoff (2003, 2005a, 2005b, 2008). Additional information was taken from Raymond and Sistrom (1969), Khanna and Nicholas (1982), Rees et al. (2002); Zaar et al. (2003); Gorlenko et al. (2004); Arunasri et al. (2005); Vogl et al. (2006). IC, intracellular; EC, extracellular; nd, not determined. a These genera have no clear phylogenetic standing because 16S rDNA sequences are not available. b Type strain not available. c Sulfate assimilation genes are present in strain DSMZ 273 but sulfate assimilation has not been investigated. d The genus Ectothiorhodosinus has no standing in nomenclature.

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Thiodictyona Thioflaviococcus Thiohalocapsa

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Some organic sulfur compounds can also serve as electron donors for photosynthetic growth of Chromatiaceae: Thiocapsa roseopersicina splits mercaptomalate to fumarate and sulfide and mercaptopropionate to acrylate and sulfide and then uses the formed sulfide as electron donor (Visscher and Taylor, 1993). This organism furthermore oxidizes dimethyl sulfide to dimethyl sulfoxide (Visscher and van Gemerden, 1991). During fermentative dark metabolism of Chromatiaceae, sulfur compounds (elemental sulfur) can serve as acceptors of electrons liberated by the oxidation of stored carbon compounds (polyhydroxyalcanoic acid). 4.1.2. Ectothiorhodospiraceae When PSB of the family Ectothiorhodospiraceae grow on sulfide as electron donor, extracellular sulfur globules are usually formed as intermediates (biphasic growth). Under the alkaline growth conditions which are optimal for Ectothiorhodospiraceae species, polysulfides are stable intermediates during oxidation of sulfide to sulfur (Then and Tru¨per, 1983, 1984). Ectothiorhodospira species are generally able to oxidize sulfide and sulfur (Table 2). When grown on elemental sulfur, Ect. halochloris does not oxidize this compound to sulfate, but reduces it to sulfide and polysulfide (Then and Tru¨per, 1984). With the exception of Ect. marismortui (Oren et al., 1989), thiosulfate is used by all species of the genus Ectothiorhodospira. On thiosulfate, sulfur globules are not detectable as an intermediate in all cases. In Ect. shaposhnikovii, for instance, sulfur is only formed from sulfide but not with thiosulfate as electron donor (Kusche, 1985). In other cases the ability to form sulfur globules from thiosulfate is not explicitly mentioned, but also not excluded (Tru¨per, 1968). In this context it is interesting to note that, in Ect. shaposhnikovii, thiosulfate oxidation does not stimulate growth, but is accompanied by the formation of tetrathionate and, later, higher polythionates (Gogtova and Vainstein, 1981). The ability to grow on sulfite has not been determined for all species, but appears to be widespread in the genus Ectothiorhodospira (Imhoff, 2005b). Members of the genus Halorhodospira oxidize sulfide to sulfur which is further oxidized to sulfate by some species. Thiosulfate is only used by Hlr. halophila (Raymond and Sistrom, 1969) and poorly by Halorhodospira neutriphila (Hirschler-Rea et al., 2003). Sulfur can also be used by some species (e.g., Hlr. halophila). The most restricted sulfur-oxidation capacities are observed in Hlr. halochloris and Hlr. abdelmalekii. Both species do not appear to be able to oxidize sulfur to sulfate (Then and Tru¨per, 1983, 1984). Thiorhodospira sibirica uses sulfide and sulfur, but not thiosulfate as electron donors for photoautotrophic growth (Bryantseva et al., 1999). The final

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 123

oxidation product is sulfate (Table 2). Ectothiorhodosinus mongolicum grows preferentially as a photoorganoheterotroph, but is able to grow photolithotrophically with sulfide. Thiosulfate is only used in the presence of organic matter. The final oxidation product is sulfate (Gorlenko et al., 2004).

4.2. Green Sulfur Bacteria Five genera with a total of 16 species of GSB are currently recognized (Imhoff, 2003, 2008; Vogl et al., 2006). GSB exhibit little variation in their ability to oxidize sulfur compounds and almost all GSB are capable of oxidizing sulfide and elemental sulfur to sulfate (Table 2). One exception is Chlorobium ferrooxidans for which only Fe2þ and H2 have been described as photosynthetic electron donors (Heising et al., 1999). Chl. ferrooxidans does not grow on sulfide, thiosulfate, or elemental sulfur, and is even inhibited by 1 mM sulfide when grown on H2. However, in general, GSB have a high affinity for sulfide, and sulfide is usually the preferred substrate even if other sulfur substrates are available (Brune, 1989, 1995b). Initially, sulfide is typically incompletely oxidized to elemental sulfur, which is deposited extracellularly as sulfur globules (Fig. 1B). These sulfur globules are oxidized completely to sulfate when the sulfide has been consumed. Some strains belonging to the genera Chlorobium and Chlorobaculum oxidize thiosulfate for growth (Imhoff, 2003). No Prosthecochloris species have been reported to utilize thiosulfate. Chlorobaculum parvum NCIB 8346 (formerly Chlorobium vibrioforme subsp. thiosulfatophilum; Imhoff, 2003) and a strain described by Helge Larsen as Chlorobium thiosulfatophilum can use tetrathionate as electron donor (Khanna and Nicholas, 1982; Larsen, 1952). Sulfite utilization has not yet been described for any GSB.

4.3. Purple Non-Sulfur Bacteria In general, purple non-sulfur bacteria are to a much lesser extent capable of tolerating and using toxic sulfur compounds such as sulfide than the PSB. However, a number of purple non-sulfur bacteria are able to grow photolithoautotrophically with reduced sulfur compounds (Brune, 1995b; Hansen and van Gemerden, 1972; Sander and Dahl, 2008). Phototrophic Betaproteobacteria (found among the Rhodocyclales and the Burkholderiales) have not been reported to use reduced sulfur compounds as electron donors and sulfide inhibits growth even at low concentrations

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(Brune, 1995b; Imhoff et al., 2005). Nevertheless, sox genes are present in the genome of Rubrivivax gelatinosus (Sander and Dahl, 2008) indicating a potential to oxidize thiosulfate to sulfate. Within all three taxonomic groups of the alphaproteobacterial purple non-sulfur bacteria (Rhodospirillales, Rhizobales, Rhodobacterales), species are found that are able to use reduced sulfur compounds. These organisms vary considerably with respect to intermediates and final products of sulfur compound oxidation. In many species (e.g., Rhodobacter capsulatus, Rhodospirillum rubrum) sulfur formed outside the cells is the end product of sulfide oxidation. However, in others, among them the species of the genera Rhodovulum, Rhodopseudomonas palustris, or Blastochloris sulfoviridis, sulfate is the end product of sulfide oxidation (reviewed in Brune, 1995b; Imhoff et al., 2005; Sander and Dahl, 2008). Rhodomicrobium vanniellii oxidizes sulfide to tetrathionate while Rhodobium marinum oxidizes it to sulfur and thiosulfate when grown photomixotrophically (Imhoff, 1983). Thiosulfate is also used by many species and oxidized either to tetrathionate (Rhodopila globiformis; Then and Tru¨per, 1981) or completely to sulfate (e.g. Rhodovulum species; Appia-Ayme et al., 2001; Brune, 1995b; Imhoff et al., 2005). Some species, for example, Rhodoplanes roseus and Rpl. elegans, use only thiosulfate but not sulfide as electron donors (Hiraishi and Ueda, 1994). Sulfur is also used as a substrate by some species, for example, Rhodopseudomonas julia and most species of the genus Rhodovulum (Imhoff et al., 2005). None of the purple non-sulfur bacteria are able to use sulfite as the sole electron donor. For more detailed information on sulfur metabolism in purple non-sulfur bacteria, the reader is referred to a recent review on sulfur oxidation in purple bacteria (Sander and Dahl, 2008).

4.4. Aerobic Bacteriochlorophyll-Containing Bacteria In general, ABC bacteria cannot grow photolithoautotrophically on reduced sulfur compounds. However, several representatives of this group can oxidize inorganic sulfur compounds. One example is Roseinatronobacter thiooxidans, a strictly aerobic, obligately heterotrophic alkaliphile that can oxidize sulfide, thiosulfate, sulfite, and elemental sulfur to sulfate in the presence of organic compounds. This bacterium appears to be a sulfuroxidizing lithoheterotroph as the efficiency of organic carbon utilization increases notably in the presence of thiosulfate (Sorokin et al., 2000). Thiosulfate-oxidizing activity was also shown in Erythromicrobium hydrolyticum, strain E4(1), and Roseococcus thiosulfatophilus, strain RB-7, in the presence of organic carbon substrate and aeration (Yurkov et al., 1994). The

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 125

genomes of all currently genome-sequenced ABC bacteria contain the genes soxB, soxAX, soxYZ, and soxCD encoding for a multienzyme complex that has been well characterized biochemically from Paracoccus pantotrophus (Friedrich et al., 2005). In Pcs. pantotrophus the encoded Sox proteins catalyze the oxidation of thiosulfate to sulfate. Taken together these findings indicate that many ABC bacteria are able to oxidize thiosulfate (Sander and Dahl, 2008). While virtually nothing is known about metabolism of inorganic sulfur compounds in these organisms, it is well established that members of the Roseobacter group are able to degrade dimethylsulfoniopropionate (DMSP), an organic sulfur compound produced in abundance by marine algae. Some isolates have been reported to carry out the major DMSP transformations that have been observed in natural bacterial communities. Among these are two competing pathways for DMSP degradation and a pathway for incorporation of the sulfur moiety of DMSP into bacterial protein (Gonzalez et al., 2003; Moran et al., 2003). In contrast to sulfide and thiosulfate, DMSP and dimethyl sulfide are abundant in aerobic marine environments (Liss, 1999). On a biochemical level, sulfur-dependent chemolithotrophy and oxidative sulfur metabolism have best been described in Acidiphilium acidophilum (formerly Thiobacillus acidophilus) (Hiraishi et al., 1995; Meulenberg et al., 1992b; Pronk et al., 1990). In this organism, the utilization of thiosulfate is initiated by the oxidative condensation of two molecules of thiosulfate yielding tetrathionate. This step is catalyzed by the periplasmic thiosulfate:cytochrome c oxidoreductase (Meulenberg et al., 1993). Furthermore, evidence was obtained that tetrathionate oxidation also takes place in the periplasm (Meulenberg et al., 1993). In addition, a tetrathionate hydrolase (de Jong et al., 1997), a trithionate hydrolase (Meulenberg et al., 1992a) and a sulfite:cytochrome c oxidoreductase (de Jong et al., 2000) have been characterized from the organism.

4.5. Filamentous Anoxygenic Phototrophs Slow photoautotrophic growth with hydrogen or sulfide as electron donor has been observed in some strains of Chloroflexus aurantiacus (Holo and Sirevag, 1986; Madigan and Brock, 1975, 1977) and Oscillochloris trichoides (Keppen et al., 2000), but not in Chloroflexus aggregans (Hanada et al., 1995). When sulfide is oxidized by Chloroflexus or Oscillochloris cultures, it is deposited as elemental sulfur outside the cells, often affixed to the cells. Sulfide-dependent photoautotrophic growth has also been observed in natural populations of Chloroflexus (Giovannoni et al., 1987; Ward et al.,

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1998). A survey of four strains of Oscillochloris trichoides showed that the preferred mode of growth was photolithoheterotrophic, although they also grew photoautotrophically with sulfide as electron donor (Keppen et al., 2000). All of the Oscillochloris trichoides stains tolerated sulfide up to about 4 mM and oxidized sulfide only to elemental sulfur. Thus, although sulfide is not the preferred substrate for photosynthetic cultivation of FAPs under laboratory conditions (Garrity and Holt, 2001b), sulfide oxidation could be important in certain natural environments. The natural habitats of Chloronema, Heliothrix, and Roseiflexus species contain no or very low concentrations of sulfide (Garrity and Holt, 2001b; Pierson et al., 1985).

4.6. Heliobacteria Heliobacteria cannot grow phototrophically using sulfide as the sole electron donor, but if added, sulfide is usually oxidized to elemental sulfur (Bryantseva et al., 2000; Madigan, 2001). Heliobacterium sulfidophilum and Heliobacterium undosum are especially tolerant to sulfide (up to 2 mM at pH 7.5) (Bryantseva et al., 2000), whereas the sulfate-fixing Heliophilum fasciatum is inhibited by sulfide concentrations above 0.1 mM (Ormerod et al., 1996). Some strains perform assimilatory sulfate reduction, whereas others require a reduced sulfur compound (e.g., thiosulfate, sulfide, methionine, or cysteine) for biosynthetic purposes. The only heliobacterium for which genome sequence information is available, Heliobacterium modesticaldum Ice1 (ATCC 51547T) (http://genomes.tgen.org/helio.html), cannot grow on sulfate as the sole sulfur source but requires thiosulfate, sulfide, methionine, or cysteine (Kimble et al., 1995).

5. OXIDATIVE SULFUR METABOLISM: ENZYMES AND GENES Here, we attempt to describe the enzymes or multienzyme systems involved in sulfur compound oxidation in phototrophic sulfur bacteria and summarize information on their occurrence. On a molecular genetic and biochemical level, sulfur oxidation is best characterized in the PSB Alc. vinosum and in the GSB Cba. tepidum. However, our knowledge of these processes is still rather incomplete. An overview of the currently proposed metabolic network is shown in Fig. 2. This figure is based on a combination of biochemical evidence and genome sequence analyses.

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 127

Figure 2 Overview of proposed pathways of thiotrophic sulfur transformations in phototrophic sulfur bacteria (PSB and GSB). Probably no single organism has all the reactions shown here. In the periplasm, the polysulfur chains are probably very short (n probably around 3 or 4), whereas the polysulfur chains in the sulfur globules can be very long (n W 3 and possibly up to n W 105 as for polymeric sulfur (Prange et al., 2002a; Dahl and Prange, 2006)). Transport into the periplasm of sulfite formed by cytoplasmic Dsr proteins and oxidation of sulfite to sulfate in the periplasm as previously suggested (Dahl and Prange, 2006; Dahl, 2008) cannot be completely excluded.

Many enzymes potentially involved in sulfur metabolism can readily be identified in the genome sequences by sequence homology with known enzymes. Tables 3 and 4 show a detailed survey of genes known to be involved in or potentially involved in the oxidative sulfur metabolism in the genome-sequenced strains of phototrophic sulfur bacteria. These tables show that PSB and GSB share a number of genes potentially involved in the oxidation of reduced sulfur compounds: For example, genes for the sulfideoxidizing enzyme flavocytochrome c (FccAB) and sulfide:quinone oxidoreductase (SQR) occur in most PSB and GSB. With only two exceptions, the dsr genes including dsrAB for dissimilatory sulfite reductase are present. The Sox system is involved in oxidation of thiosulfate and can account for this activity observed in all strains capable of oxidizing thiosulfate. The

128

Table 3 Occurrence of genes related to dissimilatory sulfur metabolism in genome-sequenced phototrophic sulfur bacteriaa sqr

fccAB

sox

soy

sgp

dsr

sat

aprBA

aprM

qmo

sorAB

Green Sulfur Bacteria Chlorobaculum parvum DSMZ 263T Chlorobaculum tepidum TLS (DSMZ 12025T) Chlorobium chlorochromatii CaD3 Chlorobium clathratiforme BU-1 (DSMZ 5477T) Chlorobium ferrooxidans DSM 13031T Chlorobium limicola DSMZ 245T Chlorobium luteolum DSMZ 273T Chlorobium phaeobacteroides DSMZ 266T Chlorobium phaeovibrioides DSMZ 265 Chloroherpeton thalassium ATCC 35110T Prosthecochloris aestuarii DSMZ 271T Prosthecochloris sp. BS1

þ þ þ þ þ þ þ þ þ þ þ þ

þ þ þ þ  þ  þ þ þ þ þ

þ þ þ þ     þ   

   þ  þ  þ  þ þ 

           

þ þ þ þ  þ þ þ þ  þ þ

 þ þ þ        þ

 þ þ þ        þ

           

 þ þ þ        þ

           

Purple Sulfur Bacteria Ectothiorhodospiraceae Halorhodospira halophila SL1 (DSMZ 244T) Chromatiaceae Allochromatium vinosum DSMZ 180T

 ?

3 copiesb

þ





þ











Enzyme activity

þ

þ

?

þ

þ

þ

þ

þ

?



The following abbreviations designate occurrence of a core set of several genes: dsr: dsrABCEFHLNMKJOP, qmo: qmoABC, sgp: sgpABC, sox: soxBXAYZ. a This survey is based on Frigaard and Bryant (2008) and additional genome sequence analyses, except for Alc. vinosum for which only limited sequence information is available. Genomes were analyzed by BLAST searches using the resources provided by Integrated Microbial Genomes (DOE Joint Genomes Institute, http://img.jgi.doe.gov) and GenBank (http://www.ncbi.nlm.nih.gov). Baits: FccAB from Alc. vinosum (AAA23316 and AAB86576); SQR from Rhodobacter capsulatus (CAA66112); SorAB from Starkeya novella (AAF64400 and AAF64401); Dsr proteins from Alc. vinosum (U84760), Sox proteins from Paracoccus denitrificans (CAA55824: SoxB, CAA55829: SoxC, CAB94380: SoxY, CAB94381: SoxZ, CAA55827: SoxA, CAB94379: SoxX); APS reductase (AprMBA) from Alc. vinosum (U84759), Qmo proteins from Cba. tepidum (QmoA: CT0866, QmoB: CT0867, QmoC: CT0868). b Relationships of the encoded proteins to FccAB higher in all cases than that to SoxEF, however similarity to SoxEF is still significant.

NIELS-ULRIK FRIGAARD AND CHRISTIANE DAHL

Organism

Organism

PSRLC1

PSRLC2

PSRLC3

hdr-hyd

rlp

hup

Green Sulfur Bacteria Chlorobaculum parvum DSMZ 263T Chlorobaculum tepidum TLS (DSMZ 12025T) Chlorobium chlorochromatii CaD3 Chlorobium clathratiforme BU-1(DSMZ 5477T) Chlorobium ferrooxidans DSMZ 13031T Chlorobium limicola DSMZ 245T Chlorobium luteolum DSMZ 273T Chlorobium phaeobacteroides DSMZ 266T Chlorobium phaeovibrioides DSMZ 265 Chloroherpeton thalassium ATCC 35110T Prosthecochloris aestuarii DSMZ 271T Prosthecochloris sp. BS1

þ þ þ þ   þ  þ þ  þ

  þ þ  þ  þ   þ þ

  þ   þ þ þ þ þ þ 

þ þ þ þ þ þ þ þ þ   

þ þ þ þ þ þ þ þ þ þ þ þ

þ   þ þ þ þ þ  þ þ þ

Purple Sulfur Bacteria Ectothiorhodospiracea Halorhodospira halophila SL1 (DSMZ 244T)









a

b



b



b

The following abbreviations designate occurrence of a core set of several genes: hdr-hyd: hdrDA-orf1247-orf1248-hydB2G2, hup, hupSLCD. Hlr. halophila has three polysulfide-reductase-like complexes that are rather distantly related to the PSRLC of GSB and of which one (encoded by Hhal_1934, 1935 and 1936) presumably is cytoplasmic as is the PSRLC3 of GSB. b

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 129

Table 4 Occurrence of genes possibly related to transformation of sulfur compounds and hydrogen in genome-sequenced phototrophic sulfur bacteriaa

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adenosine-5u-phosphosulfate reductase (Apr), sulfate adenylyltransferase (Sat), and either the quinone-interacting membrane-bound oxidoreductase (Qmo) complex or AprM probably constitute a sulfite-oxidizing system. A putative alternative sulfite-oxidizing system has been identified in the GSB strains that do not have the Apr, Sat, and Qmo (or AprM) enzymes. Homologs of sulfhydrogenase and heterodisulfide reductase are also found in GSB (Table 4). Homologs of polysulfide reductase are present in GSB as well as in the genome-sequenced PSB Hlr. halophila (Table 4; Frigaard and Bryant, 2008a). However, functions cannot easily be assigned to these proteins in phototrophic sulfur bacteria because (1) they are too distantly related to characterized enzymes; (2) they are not distributed among the organisms in a manner that obviously correlates with known physiological traits; and (3) the sulfur compound and hydrogen oxidation properties of analyzed strains can be accounted for by other enzymes. We will now first describe the oxidation of sulfide. The following sections are dedicated to our current knowledge on the properties of sulfur deposited by PSB and GSB and to the proteins involved in its further oxidation to sulfite. We will then describe the current state of affairs regarding sulfite oxidation to sulfate and finally describe the pathway of thiosulfate oxidation.

5.1. Oxidation of H2S Different enzymes are candidates for sulfide oxidation: sulfide:quinone oxidoreductase (SQR) (Schu¨tz et al., 1997), a flavocytochrome c sulfide dehydrogenase (FccAB) (Brune, 1995b; Chen et al., 1994; Kostanjevecki et al., 2000; Meyer and Cusanovich, 2003), and also the Sox system (AppiaAyme et al., 2001). The occurrence of the respective genes in GSB and PSB genomes is summarized in Table 3. 5.1.1. Flavocytochrome c Flavocytochrome c is usually a soluble, periplasmic enzyme consisting of a large sulfide-binding FccB flavoprotein subunit (41–47 kDa) and a smaller FccA cytochrome c subunit (Brune, 1995b). FccA from Alc. vinosum is 21 kDa and binds two heme c (van Beeumen et al., 1991) while in the GSB FccA is only 10 kDa and binds a single heme. In GSB, the FccB subunit has high sequence similarity (approximately 50% amino acid sequence identity) to a flavoprotein (SoxJ) encoded in the sox gene cluster (Section 5.7.1). In Ect. valuolata and also in Cba. thiosulfatiphilum DSMZ 249 (formerly Chl.

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limicola; Imhoff, 2003) the protein is membrane-bound (Kostanjevecki et al., 2000; Verte´ et al., 2002). In the latter strain, membrane attachment is possibly due to an unusual signal peptide in the FccA subunit that is not cleaved but supposedly anchors the protein in the cytoplasmic membrane. The predicted signal peptide in FccA from GSB is followed by a highly variable, 15- to 25-residue sequence, which is rich in alanine and proline and which is suggested to act as a flexible arm (Verte´ et al., 2002). FccAB is constitutively expressed in Alc. vinosum (Bartsch, 1978) and also in Cba. thiosulfatiphilum DSMZ 249 (Verte´ et al., 2002). In vitro, flavocytochromes can efficiently catalyze electron transfer from sulfide to a variety of small c-type cytochromes (e.g., cytochrome c550 from Alc. vinosum; Bosshard et al., 1986; Davidson et al., 1985) that may then donate electrons to the photosynthetic reaction center. However, the in vivo role of flavocytochrome c is unclear for a number of reasons. (1) Although many sulfide-utilizing organisms produce flavocytochrome c, some sulfideutilizing GSB and PSB do not, which clearly demonstrates that flavocytochrome c is not essential for sulfide oxidation (Brune, 1995b). (2) Mutants of Alc. vinosum (Reinartz et al., 1998) and Cba. tepidum (S.O. Barbas and N.-U. Frigaard, unpublished) deficient in flavocytochrome c exhibit sulfide oxidation rates similar to those of the wild type. (3) The pattern of sulfide oxidation and the concomitant formation of elemental sulfur that is subsequently oxidized to sulfate upon sulfide depletion, is similar in Chl. luteolum DSMZ 273T, which does not contain flavocytochrome c, and in Chl. limicola DSMZ 245T, which does contain flavocytochrome c (Steinmetz and Fischer, 1982). If indeed the FccAB flavocytochrome c oxidizes sulfide in vivo, both GSB and PSB apparently have alternative sulfide-oxidizing enzyme systems, possibly sulfide:quinone reductase (Section 5.1.2) that may be quantitatively more important. However, it is also possible that flavocytochrome c is advantageous under certain growth conditions and that such conditions have not yet been identified in these bacteria. Based on the large difference of redox potential between flavocytochrome c and the photosynthetic reaction center, Brune (1995b) suggested that flavocytochrome c may represent a high affinity system for sulfide oxidation that might be of advantage for the cells especially at very low sulfide concentrations. 5.1.2. Sulfide:Quinone Oxidoreductase As an alternative to sulfide oxidation via flavocytochrome c, the transfer of electrons from sulfide primarily into the quinone pool was proposed based on energetic considerations as well as on the inhibitory effect of rotenone,

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CCCP, and antimycin A on NAD photoreduction by sulfide (Brune, 1989; Brune and Tru¨per, 1986). Indeed, sulfide:quinone reductase (SQR; EC 1.8.5.-) catalyzing oxidation of sulfide with an isoprenoid quinone as the electron acceptor has been found in both chemotrophic and phototrophic prokaryotes as well as in some mitochondria (Griesbeck et al., 2000; Theissen et al., 2003). Membrane-bound SQR activity has been biochemically demonstrated in GSB and also PSB and presumably feeds electrons into the photosynthetic electron transfer chain via a quinol-oxidizing Rieske iron-sulfur protein/ cytochrome b complex (Reinartz et al., 1998; Shahak et al., 1992). The genome sequences of all GSB strains encode one (CT0117 in Cba. tepidum TLS) or two homologs of the biochemically characterized SQRs from Rhodobacter capsulatus (CAA66112) and Oscillatoria limnetica (AAF72962) (Table 3). This includes Chl. ferrooxidans (ZP_01385816), which cannot grow on sulfide as the sole electron donor (Heising et al., 1999). This organism may benefit from SQR activity as a supplement to its energy metabolism. It could also use SQR as a protective mechanism to remove sulfide, which prevents growth when present in high concentrations. The SQR homologs of GSB are flavoproteins with predicted masses of about 53 kDa, and each contains all three conserved cysteine residues that are essential for sulfide oxidation in Rhodobacter capsulatus SQR (Griesbeck et al., 2002). The significance of SQR in GSB is not yet clear. An sqr mutant of Cba. tepidum exhibits a lower sulfide oxidation rate than the wild type, which demonstrates that although SQR participates in sulfide oxidation in this organism, an alternative sulfide oxidation mechanism exists (Oco´n Barbas and Frigaard, unpublished). Although Alc. vinosum membranes exhibit SQR activity, it has so far not been possible to detect a sqr-related gene via Southern hybridization with heterologous probes or heterologous PCR nor could the protein be detected with antibodies directed against the Rba. capsulatus protein (M. Reinartz and C. Dahl, unpublished). The enzyme from Alc. vinosum and possibly other PSB may have properties distinct from those of characterized SQRs. In general, the sequences of SQR proteins are not well conserved and it is possible that a high degree of sequence variation could account for the lack of detection. In accordance, the Hlr. halophila genome contains one only distantly related homolog (Hhal_1665) of the biochemically well-characterized SQR from Rhodobacter capsulatus (Griesbeck et al., 2002; Schu¨tz et al., 1999). In Rba. capsulatus, SQR is absolutely essential for the oxidation of sulfide (Schu¨tz et al., 1999). The enzyme is a peripherally membrane-bound flavoprotein with its active site located in the periplasm (Schu¨tz et al., 1997, 1999). Interestingly, a classical N-terminal signal peptide is missing and

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 133

co-transport with the products of neighboring genes was excluded by gene deletion. However, nuclease treatment showed that 38 carboxy-terminal amino acids are necessary for the translocation, indicating a hitherto unknown transport mechanism (Schu¨tz et al., 1999). The primary product of the SQR reaction is soluble polysulfide whereas elemental sulfur does not appear to be formed in vitro (Griesbeck et al., 2002). Very probably, disulfide (or possibly a longer chain polysulfide) is the initial product of sulfide oxidation, which is released from the enzyme. Polysulfide anions of different chain lengths are in equilibrium with each other and longer-chain polysulfides can be formed by disproportionation reactions from the initial disulfide (Steudel, 1996b). When whole cells of Rba. capsulatus grow with sulfide, elemental sulfur is formed as the final product. In principle, elemental sulfur can form spontaneously from polysulfides (Steudel, 1996b). In experiments using isolated spheroplasts from Chl. vibrioforme and Alc. minutissimum, soluble polysulfides have also been detected as the product of sulfide oxidation (Blo¨the and Fischer, 2000). Polysulfides were also detected as primary products of sulfide oxidation by whole cells of Alc. vinosum (Prange et al., 2004) and have been reported as intermediates of the oxidation of sulfide to extracellular sulfur by species of the PSB family Ectothiorhodospiraceae (Then and Tru¨per, 1983; Tru¨per, 1978). While transient formation of polysulfide by the latter organism species had originally been attributed to chemical reaction between H2S and elemental sulfur promoted by the alkaline culture medium (Tru¨per, 1978), it now appears more likely that they present enzymatically generated intermediates. Some GSB additionally contain distantly related homologs of SQR with no assigned or obvious function (here denoted SQRLP1 and SQRLP2 for SQR-like proteins type 1 and type 2). SQRLP1 is present in Cba. tepidum TLS (CT1087) and in three other GSB. Among the GSB, SQRLP2 is only present in Cba. tepidum TLS (CT0876). Interestingly, SQRLP2 clusters in phylogenetic analyses with proteins of unknown function from various archaea. In addition, SQRLP2 from Cba. tepidum TLS shares 54% amino acid sequence identity with a protein from Sulfitobacter strain NAS14.1, which is a marine Roseobacter-like strain that grows aerobically on dimethylsulfoniopropionate (https://research.venterinstitute.org/moore/). Otherwise, SQRLP2 has few homologs in the databases. 5.1.3. Sox Proteins and Sulfide Oxidation In Rhodovulum sulfidophilum, a member of the purple non-sulfur bacterial family Rhodobacteraceae, the Sox enzyme system catalyzing the oxidation of

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thiosulfate to sulfate (see Section 5.7.1), is also indispensable for the oxidation of sulfide in vivo (Appia-Ayme et al., 2001). The same might well be the case for other purple non-sulfur bacteria containing sox genes. However, in Alc. vinosum mutants deficient of either flavocytochrome c (Reinartz et al., 1998), sox genes or both (D. Hensen, B. Franz and C. Dahl, unpublished), sulfide oxidation proceeds with wild-type rates indicating that SQR plays the main role in sulfide oxidation in this organism. Occurrence of sox genes in GSB and PSB indicates a tight correlation with the ability to oxidize thiosulfate (Table 3). 5.1.4. Other Proposed Sulfide-Oxidizing Enzymes In Alc. vinosum, a dissimilatory sulfite reductase (DsrAB) operating in reverse, that is, in the direction of sulfite formation, has also been discussed to be involved in sulfide oxidation (Schedel et al., 1979). However, this protein is clearly not essential for sulfide oxidation but rather absolutely required for oxidation of intracellularly stored sulfur (Pott and Dahl, 1998). This finding is consistent with the occurrence of dsr genes exclusively in sulfur-storing PSB but not in purple non-sulfur bacteria (Sander and Dahl, 2008). It should be noted that cytochromes without flavin groups have also been proposed to mediate electron transfer from sulfide to the reaction center in some PSB (Brune, 1989; Fischer, 1984; Leguijt, 1993).

5.2. Oxidation of Polysulfides As outlined above, polysulfides appear to be the primary product of the oxidation of sulfide in a number of purple and green bacteria. Utilization of externally added polysulfides has been studied in Alc. vinosum and Tca. roseopersicina. Both organisms readily used these compounds as photosynthetic electron donors (Steudel et al., 1990; van Gemerden, 1987; Visscher et al., 1990). It is currently unknown how polysulfides are converted into sulfur globules. Theoretically this could be a purely chemical, spontaneous process as longer polysulfides are in equilibrium with elemental sulfur (Steudel et al., 1990). However, we have shown that Alc. vinosum sulfur globules do not contain major amounts of sulfur rings but probably consist of long-chains of sulfur with organic residues at one or both ends (Prange et al., 1999, 2002a). Such organylsulfanes must eventually be formed by an unknown (enzymatic) mechanism.

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5.3. Uptake of External Sulfur Very many GSB and PSB including Alc. vinosum are able to oxidize externally supplied solid, virtually insoluble, elemental sulfur. This is a challenging biochemical problem in that elemental sulfur is extremely hydrophobic and virtually insoluble in water. In addition, elemental sulfur cannot be attacked via oxygenation reactions in the absence of molecular oxygen under anoxic conditions. In PSB like Alc. vinosum, elemental sulfur is first taken up by the cells and intracellular sulfur globules are formed as an obligate intermediate before the sulfur stored in the globules is oxidized to the final product sulfate (Brune, 1995b; Dahl, 2008; Franz et al., 2007). In contrast, GSB and PSB of the family Ectothiorhodospiraceae do not form intracellular sulfur globules. Very little is known on the attachment and utilization of solid elemental sulfur by phototrophic sulfur bacteria. Neither uptake of elemental sulfur nor the transformations necessary for its oxidation under anoxic conditions are well understood. Enzymes catalyzing the uptake and oxidation of externally added S0 have not yet been isolated from any phototrophs. Utilization of solid elemental sulfur must include binding and/or activation of the sulfur as well as transport inside the cells. As pointed out by Chan et al. (2008), two different strategies would in principle be possible, analogous to the reduction of insoluble electron acceptors (e.g., metals) (Lloyd, 2003): physical contact of the cells to their insoluble substrate and direct electron transfer from the cell envelope to the substrate via outer membrane proteins (Myers and Myers, 2001) or excretion of reducing substances that can act on substrate distant from the cells (Hernandez et al., 2004). Conclusive evidence for one of these models has not yet been obtained for phototrophic sulfur bacteria.

5.3.1. Properties of Elemental Sulfur ‘‘Elemental sulfur’’ (‘‘S0’’) has the formal valency zero. It tends to catenate and form chains with various lengths (SN or Sm) and ring sizes (Sn) (Steudel, 2000; Steudel and Eckert, 2003). All sulfur allotropes are hydrophobic, not wetted by water and they hardly dissolve in water. Cyclic, orthorhombic a-sulfur (a-S8) is the most stable form of elemental sulfur at ambient pressure and temperature (Roy and Trudinger, 1970; Steudel, 2000). Polymeric sulfur consists mainly of chain-like macromolecules (Steudel and Eckert, 2003). Customary in trade, typical elemental sulfur (‘‘flowers of sulfur’’) mainly consists of S8 rings and traces of S7 rings which are responsible for the yellow color (Steudel and

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Holz, 1988). In addition, elemental sulfur sublimed at ambient pressure always contains some polymeric sulfur (Steudel and Eckert, 2003).

5.3.2. Adhesion to Elemental Sulfur Some information is available about adhesion to extracellular sulfur by chemotrophic sulfur oxidizers and by GSB. It is well established that acidophilic chemotrophic sulfur-oxidizing bacteria like Acidithiobacillus thiooxidans or Acidithiobacillus ferrooxidans attach to sulfur by glycocalyxlike extracellular polymeric substances (EPS) (Espejo and Romero, 1987; Vogler and Umbreit, 1941), specifically lipopolysaccharides (Bryant et al., 1984; Gehrke et al., 1998). In general, EPS are a complex mixture of high molecular weight polymers (MWW10,000). They significantly influence the surface physicochemical properties, which are of considerable importance in governing bacterial adhesion. The origin of EPS is very complex, and their components and content heavily depend on many factors such as the substrate. In phototrophic sulfur bacteria, the presence, compositions, and structures of EPS have not been systematically elucidated. Structures attached to the cell wall (the so-called ‘‘spinae’’) have been postulated to mediate adhesion of the GSB Chlorobaculum limnaeum UdG 6038 (formerly Chlorobium limicola; Imhoff, 2003) to extracellularly deposited sulfur (Pibernat and Abella, 1996). The presence of a large capsule in Cba. limnaeum UdG 6038 concomitant with a high number of spinae was interpreted as the spinae acting as the frame for capsule stabilization. Concomitant with the frequent observation of large numbers of sulfur globules per cell by phase-contrast microscopy a role for both spinae and capsule in the metabolism of external sulfur globules has been suggested (Pibernat and Abella, 1996). In Atb. ferrooxidans, Rojas et al. (1995) have demonstrated the role of a capsule in retaining sulfur colloids during pyrite oxidation as temporary energy reservoirs. In spined GSB, the presence of spinae together with the capsule favors the retention of the external sulfur globules physically attached to the outer membrane of the cell wall. Moreover, it has recently been shown for the PSB Alc. vinosum that intimate physical cell-sulfur contact is a prerequisite for uptake of elemental sulfur (Franz et al., 2007). The surface properties of bacteria that have been found to affect adhesion to insoluble substrates like sulfur are cell surface hydrophobicity (Devasia et al., 1993; Gehrke et al., 1998; Takeuchi and Suzuki, 1997) and

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electrokinetic potential (Blake et al., 1994). Atb. ferrooxidans cells grown on sulfur or pyrite exhibited greater hydrophobicity than cells grown on soluble ferrous iron (Devasia et al., 1993). In a different study, sulfur-grown cells of Atb. ferrooxidans were found to exhibit purely hydrophobic surface properties and did not attach to charged particles such as pyrite. EPS from sulfur-grown cells contained more lipids, more free fatty acids, and a much higher ratio of uncharged (glucose) versus charged sugars (glucuronic acid) than EPS from pyrite-grown cells. It was concluded that attachment to hydrophobic substrates such as sulfur is dominated by van der Waal’s attraction forces (Gehrke et al., 1998). Measurements of zeta potentials of Atb. ferrooxidans cells showed that net surface charge was such as to minimize charge repulsive forces in the organism’s interaction with any substrate surface. When grown on elemental sulfur, washed cells were close to their isoelectric point (with a slight tendency to the negative), while they were negatively charged when grown on pyrite or ferrous iron (Blake et al., 1994). Besides hydrophobic interactions, hydrophilic interactions have also been implicated in cell-sulfur contact of Atb. ferrooxidans (Ohmura et al., 1993; Takeuchi and Suzuki, 1997). 5.3.3. Activation and Uptake of Elemental Sulfur In all cases so far, a reaction activating elemental sulfur prior to its oxidation is postulated, due to the stability and low water solubility of the substrate. In the case of cyclo-octasulfur, this activation reaction could be an opening of the S8 ring by nucleophilic reagents, resulting in the formation of linear inorganic or organic polysulfanes. In addition, the reduction of elemental sulfur to water-soluble sulfide has been discussed. Both reactions could be carried out by thiol groups of cysteine residues. Along this line, it was proposed for Acidithiobacillus and Acidiphilium that extracellular elemental sulfur is mobilized by thiol groups of special outer membrane proteins and transported into the periplasmic space as persulfide sulfur (Rohwerder and Sand, 2003). Experimental evidence for the existence of two different outer membrane proteins (OMPs) involved in cell-sulfur adhesion and sulfur uptake in Acidithiobacillus has been obtained by Buonfiglio et al. (1999) and Ramı´ rez et al. (2004). In this respect it might be interesting to note that the complete genome sequences of 10 GSB able to oxidize externally added elemental sulfur contain a gene related to that for the sulfur-induced OMP of Atb. ferrooxidans identified by Ramı´ rez et al. (2004) (CT2230 in Cba. tepidum). In several of these GSB the gene is located in the

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vicinity of other genes encoding enzymes of sulfur metabolism (subunits of potential polysulfide reductases, dissimilatory sulfite reductase (dsr) operon). In Chlorobium clathratiforme BU1 (DSMZ 5477T) the gene is even part of the dsr operon. Most conspicuously, the gene is absent in Chlorobium ferrooxidans that cannot oxidize reduced sulfur compounds. A closely related gene is not present in the genome of Hlr. halophila. However, a different gene encoding a potential outer membrane porin (Hhal_1947) is found in the sulfur gene cluster of Hlr. halophila (Dahl, 2008) where it is situated immediately upstream of genes encoding a potential flavocytochrome c. 5.3.4. Which Species of Elemental Sulfur are Metabolized? The bonding energy between S–S bonds in polymeric sulfur is 2.4 kJ/mol weaker than in cyclo-octasulfur (Steudel, 1985, 1996a; Steudel and Eckert, 2003). Therefore, chain-like sulfur (polymeric sulfur) might be the more easily accessible species of ‘‘elemental sulfur’’ for microorganisms. Recently, it was demonstrated that Alc. vinosum indeed uses only, or at least strongly prefers, the polymeric sulfur (sulfur chains) fraction of commercially available elemental sulfur and is probably unable to take up and form sulfur globules from cyclo-octasulfur (Franz et al., 2007). Evidence for the formation of intermediates like sulfide or polysulfides during uptake of elemental sulfur was not obtained. It may be speculated that ‘‘sulfur chains’’ rather than the more stable ‘‘sulfur rings’’ are the microbiologically preferred form of elemental sulfur also for other sulfur-oxidizing bacteria. This hypothesis gains some support in the recent study of Urich et al. (2006) who investigated the influence of different sulfur species on enzyme functions in the sulfur oxygenase reductase from Aquifex aeolicus. Theoretical considerations on the basis of the crystal structure of this enzyme led to the hypothesis that linear sulfur but not cyclic sulfur species can serve as a substrate for this enzyme.

5.4. Properties of Sulfur Globules In anoxygenic phototrophic sulfur bacteria, sulfur appears to be generally deposited outside the cytoplasm. GSB and PSB of the family Ectothiorhodospiraceae form extracellular sulfur globules, while the globules are located in the periplasmic space in members of the family Chromatiaceae (Pattaragulwanit et al., 1998).

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5.4.1. Intracellular versus Extracellular Sulfur Deposition Sulfur deposited inside the cell is unattainable for other individuals. It might be concluded that deposition of sulfur outside the cells would make this sulfur available to all individuals in the culture. However, using sulfidelimited continuous cultures of Cba. thiosulfatiphilum DSMZ 249, it was shown that elemental sulfur produced by the GSB, although being deposited extracellularly, is not easily available for other individuals, and apparently remains (in part) attached to the cells (van Gemerden, 1986). It was even concluded that functionally the elemental sulfur formed by GSB is not extracellular and that, in essence, not even a GSB cell can profit from the extracellular sulfur produced by another GSB cell. Microscopic observations indeed show that most sulfur is attached to the cells and very little is free floating in the medium (Tru¨per, 1984; Tru¨per and Genovese, 1968). The mechanistic explanation of this phenomenon could be the production of extracellular capsules of, for example, polysaccharides or the presence of tubes of pili outside the cell (see Section 5.3.2). The latter were observed in cultures of Chl. limicola strain DSMZ 257 and Cba. thiosulfatiphilum DSMZ 249 (Cohen-Bazire, 1963) and also Cba. limnaeum UdG 6038 (Pibernat and Abella, 1996). 5.4.2. Speciation of Sulfur in Sulfur Deposits Various techniques and approaches have been used to determine the exact chemical nature of the ‘‘elemental sulfur’’ in phototrophic sulfur bacteria. Different modifications of sulfur were detected or claimed and several models were proposed. Originally, orthorhombic sulfur crystals in Thiocystis violaceae and other phototrophic bacteria were proposed by Tru¨per and Hathaway (1967). However, microscopic investigations (polarized microscopy; freeze etch microscopy) yielded first evidence that the elemental sulfur in the globules is not crystalline orthorhombic sulfur (Hageage et al., 1970; Remsen, 1978; Remsen and Tru¨per, 1973). Using X-ray diffraction, Hageage et al. (1970) determined the intracellular sulfur of Chromatium okenii as ‘‘spherically symmetrical aggregates of radially arranged arrays of S8 molecules that were in a metastable, liquid modification.’’ By determining and calculating the density of non-sulfur-containing bacteria and two sulfurglobule-containing Chromatium spp. in comparison with the density of elemental sulfur, Guerrero et al. (1984) postulated that the intracellularly deposited sulfur is complexed with another low-dense component and might be ‘‘hydrated sulfur.’’ Raman spectroscopic investigations showed that the sulfur globules of Alc. vinosum and Hlr. abedelmalekii contain only small

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amounts of S8 rings (Steudel, 1985; Then, 1984). Steudel and coworkers suggested that the sulfur in the sulfur globules consists of a nucleus of S8 rings surrounded by water and long-chain sulfur species like polysulfides act as amphiphilic interface (Steudel, 1989; Steudel et al., 1990). As another model for the modification of sulfur in sulfur bacteria, sulfur-sols (small milky-colored emulsions of elemental sulfur (sulfur rings) in water) were proposed (Steudel, 1996b, 2003; Steudel and Albertsen, 1999). Finally, X-ray absorption near-edge structure (XANES) spectroscopy at the sulfur K-edge using synchrotron radiation was used as an in situ approach to investigate the sulfur speciation in intact phototrophic bacterial cells (Pickering et al., 2001; Prange et al., 1999, 2002a). In almost all earlier investigations, the sulfur globules were extracted from the cells prior to analysis (e.g., X-ray diffraction) causing changes in the chemical structure of the sulfur (Prange et al., 2002a). In contrast, XANES is not only a sensitive but also a non-destructive method (Prange and Modrow, 2002). XANES revealed only one main form of sulfur in sulfur globules of GSB and PSB (Prange et al., 2002a). Despite the different site of deposition (outside or inside the confines of the cell) the sulfur mainly consists of long sulfur chains very probably terminated by organic residues (mono-/bis-organyl polysulfanes) in all investigated members of the Chromatiaceae, Ectothiorhodospiraceae, and Chlorobiaceae. The organic residue present at the end of the sulfur chains appears to be glutathione or very similar to glutathione (Prange et al., 2002a) and could be responsible for keeping the sulfur in a ‘‘liquid’’ state at ambient pressure and temperature conditions. Furthermore, this finding supports earlier speculations that reduced glutathione (probably in its amidated form) could act as a carrier molecule of sulfur to and from the globules (Bartsch et al., 1996; Pott and Dahl, 1998) (see also Section 5.5.2). XANES spectroscopy also yielded evidence that the sulfur chains in globules of Alc. vinosum are gradually shortened during oxidation of intracellularly stored sulfur to sulfate (Prange et al., 2002a). It should be mentioned that some controversy has arisen about the interpretation of data acquired by XANES spectroscopy. Investigations of phototrophic sulfur bacteria by two different groups (Pickering et al., 2001; Prange et al., 2002a) yielded partly comparable experimental data but were interpreted in quite a different way. Pickering et al. (2001) concluded on the basis of theoretical considerations that the sulfur is ‘‘simply solid S8.’’ The discrepancies are mainly based on the measurement mode (George et al., 2002; Prange et al., 2002b). The model for the sulfur globules of Alc. vinosum that corresponds best with the available experimental data consists of long sulfur chains terminated by organic groups as was suggested by Kleinjan et al. (2003).

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Sulfur from sulfur globules isolated in the presence of oxygen from anaerobically grown Alc. vinosum was found as S8 rings (Prange et al., 2002a), indicating the influence of oxygen and the necessity of in situ methods like XANES spectroscopy that can be applied without destruction of the original sulfur environment. 5.4.3. Intracellular Localization of Sulfur Deposits As already pointed out, the intracellular sulfur globules of Chromatiaceae reside in the periplasm (Pattaragulwanit et al., 1998). While these sulfur globules appear to be more or less evenly distributed in many species (see also Fig. 1C), they can have very special and conspicuous localizations in other species. In Allochromatium warmigii, for example, globules are predominantly located at the two poles of the cell. Dividing cells form additional sulfur globules near the central division plane. In Lamprobacter modestohalophilus the sulfur globules appear in the center of cells, while they are found in the peripheral part of the cells that is free of gas vesicles in species of the genera Lamprocystis and Thiodictyon. Sulfur globules are also found in the cell periphery in Thiopedia rosea (Imhoff, 2005a). For Thiorhodovibrio winogradsky a formation of up to 10 small sulfur globules in a row along the long cell axis has been reported (Overmann et al., 1992b). The specialized arrangement of sulfur inclusions suggests an important structure function relationship. 5.4.4. Sulfur Globule Envelopes in Chromatiaceae Like many chemotrophic sulfur-oxidizing bacteria that form intracellular sulfur globules, the sulfur globules in the Chromatiaceae are enclosed by a protein envelope (Fig. 1D) (Brune, 1995a; Dahl, 1999; Dahl and Prange, 2006). In Alc. vinosum, this envelope is a monolayer of 2–5 nm consisting of three different hydrophobic ‘‘sulfur globule proteins’’ (SGPs) of 10.5 kDa, 10.6 kDa (SgpA and SgpB), and 8.5 kDa (SgpC). In Tca. roseopersicina strain 6311 (DSM 219) the envelope contains only two proteins of 10.7 kDa and 8.7 kDa (Brune, 1995a). The two larger sulfur globule proteins (SgpA and SgpB) of Alc. vinosum are homologous to each other and to the larger protein of Tca. roseopersicina. The smaller sulfur globule proteins (SgpC) in Alc. vinosum and Tca. roseopersicina are also homologous, indicating that these proteins are highly conserved between different species of the family Chromatiaceae. The proteins are targeted to the bacterial periplasm by Secdependent signal peptides (Pattaragulwanit et al., 1998). The targeting process was experimentally verified with phoA fusions in E. coli

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(Pattaragulwanit et al., 1998) and also in Alc. vinosum (Prange et al., 2004). Electron micrographs of two other species of the family Chromatiaceae (Thiocystis violaceae and Tca. roseopersicina) also demonstrated an extracytoplasmic localization of the sulfur globules (Pattaragulwanit et al., 1998). All three sulfur globule proteins are rich in glycine and aromatic amino acids, particularly tyrosine. The amino acid sequences contain tandem repeats typically found in cytoskeletal keratin or plant cell wall proteins suggesting that they are structural proteins rather than enzymes involved in sulfur metabolism (Brune, 1995a). A direct/covalent attachment of chains of stored sulfur to the proteins enclosing the globules is unlikely as none of the Sgp proteins sequenced so far contains cysteine residues. In Alc. vinosum SgpC appears to play an important role in sulfur globule expansion (Prange et al., 2004). SgpA and SgpB are in part, but not fully, competent to replace each other (Pattaragulwanit et al., 1998; Prange et al., 2004). The construction of Alc. vinosum mutants lacking SgpA and SgpC or all three Sgp proteins was not possible, leading to the conclusion that a basic level of sulfur globule proteins is obligatory for cell survival even under conditions that do not allow sulfur globule formation (Prange et al., 2004). Experiments with a sgpBC double mutant clearly showed that an envelope is indispensable for the formation and deposition of intracellular sulfur (Prange et al. 2004). Each of the three sgp genes of A. vinosum forms a separate transcriptional unit (Pattaragulwanit et al., 1998). All are constitutively expressed; however, the expression of sgpB and sgpC is significantly enhanced under photolithoautotrophic compared to photoorganoheterotrophic conditions. Interestingly, sgpB is expressed 10 times less than sgpA and sgpC implying that SgpA and SgpC are the ‘‘main proteins’’ of the sulfur globule envelope (Prange et al., 2004). Proteinaceous envelopes have never been reported for extracellular sulfur globules. Neither the complete genome sequence of Hlr. halophila nor those of GSB (Frigaard and Bryant, 2008a) contain potential sgp genes (Table 3). As outlined above, the sulfur speciation in sulfur globules of anoxygenic phototrophic bacteria is nearly identical, irrespective of whether it is accumulated in globules inside or outside the cells. It therefore appears that the Sgp proteins themselves are not responsible for keeping the sulfur in a certain chemical structure. 5.4.5. Reduction of Sulfur Deposits Sulfur globules can also serve as an electron-acceptor reserve that allows a rudimentary anaerobic respiration with sulfur. Under anoxic conditions in

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the absence of light, both PSB and GSB can reduce stored sulfur back to sulfide (Kondrat’eva, 1979; Paschinger et al., 1974; Tru¨per, 1978; van Gemerden, 1968). Some GSB strains photochemically disproportionate elemental sulfur into sulfide and thiosulfate but in the absence of CO2 and growth (Tru¨per et al., 1988). Sulfide production from sulfur has also been observed in heliobacteria that are transferred from the light to the dark (Starynin and Gorlenko, 1993). Van Gemerden (1968) showed that the dark reduction of stored sulfur to H2S by Alc. vinosum was coupled to oxidation of glycogen to poly-b-hydroxybutyrate and suggested that sulfur reduction was used to dispose of excess reducing equivalents formed during anaerobic breakdown of glucose. Given that the sulfur globules are not directly accessible to cytoplasmic enzymes, the involvement of quinonereactive membrane-bound enzymes resembling polysulfide reductase PsrABC from Wolinella succinogenes (Krafft et al., 1992, 1995) would be a likely possibility. The molybdopterin-containing active site subunit PsrA of this enzyme is located in the periplasm. Genes encoding homologous proteins that are predicted to reside in the periplasm are found in the genomes of many GSB and also in Hlr. halophila (see Section 5.8.2 and Table 4).

5.5. Oxidation of Stored Sulfur to Sulfite The oxidative degradation of sulfur deposits in PSB and GSB is a main subject of current research on sulfur metabolism in phototrophic bacteria but is still not completely understood. In the case of extracellularly deposited sulfur, this process does not only involve oxidation of the sulfur but must include binding, activation and transport into the cells (see above Section 5.3). 5.5.1. Occurrence and Arrangement of dsr Genes The only gene region known so far to be essential for oxidation of stored sulfur was localized by interposon mutagenesis in Alc. vinosum (Dahl et al., 2005; Pott and Dahl, 1998). A total of 15 open reading frames, designated dsrABEFHCMKLJOPNRS, were identified (Fig. 3). The first two of these genes encode the reverse dissimilatory sulfite reductase (DsrAB) of Alc. vinosum (Hipp et al., 1997; Schedel et al., 1979). Very similar gene clusters are also found in Hlr. halophila and GSB (Table 3 and Fig. 3). In Hlr. halophila the dsr gene cluster in addition contains genes encoding putative regulatory proteins and proteins possibly involved in sulfate

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Figure 3 Schematic overview of the gene organization around the dsr genes in the PSB Alc. vinosum DSMZ 180T and Hlr. halophila SL1 and the GSB Chl. phaeovibrioides DSMZ 265 and Cba. tepidum TLS. The numbers indicate genes of unknown function; genes with the same number are homologous.

transport downstream of dsrN (Dahl, 2008). GSB contain a cluster, dsrNCABLEFHTMKJOP, the only difference to Alc. vinosum being the absence of dsrRS and the presence of dsrT. This cluster is present in all GSB, except Chl. ferrooxidans and Chp. thalassium, and it most likely encodes the same function as in Alc. vinosum. In Cba. tepidum TLS the dsr genes are split into two clusters, and three functional dsr genes are duplicated (dsrA, dsrC, and dsrL) (Fig. 3). This may be due to a frameshift mutation in the dsrB gene in a recent ancestor of the TLS strain that rendered the gene nonfunctional. This could have been selected for a duplication, rearrangement, and subsequent frameshift mutation of a small segment of the genome, which restored a functional dsrB gene but also resulted in a duplication of the dsrCABL gene cluster. The two regions that contain a dsrCABL cluster in Cba. tepidum TLS are 99.4% identical at the nucleotide level. From the currently available data, it appears that the dsr genes only occur as a single cluster in all other genome-sequenced GSB. The absence of dsr genes in Chl. ferrooxidans is consistent with the observation that this bacterium is incapable of growth on elemental sulfur and sulfide. Because this strain appears to have ancestors that are sulfurand sulfide-oxidizing GSB that contain the dsr genes (Frigaard and Bryant, 2008a, b), it seems highly likely that Chl. ferrooxidans has lost the dsr genes as a consequence of adapting to growth on Fe2þ. The absence of dsr genes in Chp. thalassium is especially interesting for two reasons: first, this organism is a very early diverging GSB, and second, this organism grows poorly on elemental sulfur. Like other GSB, Chp. thalassium grows well on sulfide and forms extracellular sulfur globules as an oxidation product (Gibson et al., 1984). However, this elemental sulfur is only very slowly oxidized; and this behavior could be due to the absence of the Dsr system. It is at present

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unclear what might constitute an alternative sulfur-oxidizing system in Chp. thalassium (Table 4). Such a system might somehow involve the RuBisCOlike protein (RLP), which is present in all GSB including Chp. thalassium and which has been shown to be involved in growth on elemental sulfur in Cba. tepidum TLS (Hanson and Tabita, 2001, 2003). It is an interesting possibility that it might have been the acquisition of the Dsr-dependent system, which seems to be involved in efficient and complete oxidation of elemental sulfur, that led to the relatively recent, explosive radiation of the lineages of GSB that are not closely related to Chloroherpeton (Frigaard and Bryant, 2008a, b). Most of the dsr genes are widespread not only in phototrophic but also in chemotrophic sulfur oxidizers; in addition they occur in sulfate-reducing bacteria (Grimm et al., 2008; Mussmann et al., 2005; Sander et al., 2006). In sulfate-reducing prokaryotes, sulfite reductase catalyzes the reduction of sulfite to sulfide as the final step of sulfate respiration (Matias et al., 2005). The genes dsrABCNMKJOP represent a core unit occurring in both sulfuroxidizing and sulfate-reducing organisms. In contrast, other dsr genes appear to be specific for one of the physiological groups. The gene dsrD has so far only been detected in sulfate-/sulfite-reducing prokaryotes, whereas the genes dsrEFH appear to be typical for sulfur oxidizers (Sander et al., 2006). DsrL homologs were not only detected in sulfur oxidizers, but also in a few sulfite-respiring bacteria such as Desulfitobacterium hafniense and Moorella thermoacetica (Mussmann et al., 2005). In several cases phylogenetic analysis of the common Dsr proteins yielded two separate clusters consisting of proteins from sulfate reducers on the one hand and of proteins from sulfur oxidizers on the other (Sander et al., 2006). Within the GSB, DsrA and other Dsr proteins constitute a monophyletic group. However, the dsr genes have experienced lateral gene transfer (LGT) within the GSB phylum; for example, DsrA from Prosthecochloris aestuarii DSMZ 271T is located within the Chlorobium/Chlorobaculum cluster (Frigaard and Bryant, 2008a). In contrast to DsrAB sulfite reductase and other cytoplasmic Dsr proteins, the components of the membrane-bound DsrMKJOP complex of GSB do not cluster with the proteins of other sulfur oxidizers but affiliate with the sulfate-/sulfite-reducing prokaryotes. This phenomenon suggests a horizontal gene transfer, which is also supported by the presence of dsrT (Mussmann et al., 2005) in GSB, a gene otherwise only found in sulfate-/sulfite-reducing prokaryotes (Sander et al., 2006). The Dsr system in GSB is therefore considered to have an intriguing chimeric nature possibly generated by lateral gene transfer of dsrTMKJOP from a sulfatereducing prokaryote to a common ancestor of GSB.

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5.5.2. Dsr Gene Products and their Proposed Functions In Alc. vinosum, the dsrAB products form the cytoplasmic a2b2-structured sulfite reductase. This protein is closely related to the dissimilatory sulfite reductases from sulfate-reducing bacteria and archaea (Hipp et al., 1997). The prosthetic group of DsrAB is siroamide-[Fe4S4] with siroamide being an amidated form of the classical siroheme. The dsrN-encoded protein resembles cobyrinic acid a,c diamide synthases and probably catalyzes the glutamine-dependent amidation of siroheme (Lu¨bbe et al., 2006). An involvement of DsrR in biosynthesis of siroamide is possible based on the finding that the genes dsrN and dsrR are fused in the chemotrophic sulfuroxidizing bacterial endosymbiont (Candidatus Ruthenia magnifica) of the bivalve Calyptogena magnifica (Newton et al., 2007). An Alc. vinosum DdsrN mutant showed a reduced sulfur oxidation rate. Alc. vinosum is apparently able to incorporate siroheme instead of siroamide into sulfite reductase, thereby retaining some function of the enzyme (Lu¨bbe et al., 2006). Adjacent to dsrAB, the dsrEFH genes are located. The products of these three genes show significant similarity to each other (Pott and Dahl, 1998) and in Alc. vinosum they form a soluble cytoplasmic a2b2g2-structured 75 kDa holoprotein (Dahl et al., 2005). DsrC is a small soluble cytoplasmic protein with a highly conserved C-terminus including two conserved cysteine residues. Proteins closely related to DsrEFH and DsrC have recently been shown to act as parts of a sulfur relay system involved in thiouridine biosynthesis at tRNA wobble positions in E. coli (Ikeuchi et al., 2006; Numata et al., 2006). The dsrM-encoded protein is predicted to be a membrane-bound b-type cytochrome and shows similarities to a subunit of heterodisulfide reductases from methanogenic archaea. The iron–sulfur protein DsrK exhibits relevant similarity to the catalytic subunit of heterodisulfide reductases (Dahl et al., 1999; Pott and Dahl, 1998). DsrK is predicted to reside in the cytoplasm. DsrP is another integral membrane protein. The periplasmic proteins DsrJ and DsrO are a triheme c-type cytochrome and an iron–sulfur protein, respectively. DsrKJO were copurified from membranes of Alc. vinosum indicating the presence of a transmembrane electron-transporting complex consisting of DsrKMJOP (Dahl et al., 2005). In Alc. vinosum, individual in-frame deletions of the dsrMKJOP genes lead to the complete inability of the mutants to oxidize stored sulfur (Sander et al., 2006). DsrL is a cytoplasmic iron–sulfur flavoprotein with NADH:acceptor oxidoreductase activity (Y. Lu¨bbe and C. Dahl, unpublished). In frame deletion of dsrL completely inhibited the oxidation of stored sulfur (Lu¨bbe et al., 2006). DsrS is a soluble cytoplasmic protein of unknown function.

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In Alc. vinosum, the dsr genes, with the exception of the constitutively expressed dsrC, are expressed and the encoded proteins are formed at a low basic level even in the absence of sulfur compounds. An increased production of all Dsr proteins is induced by sulfide and/or stored sulfur (Dahl et al., 2005). Since the proteins encoded at the dsr locus are either cytoplasmic or membrane-bound they cannot act directly on the extracytoplasmic sulfur globules. As already mentioned above, dissimilatory sulfite reductase catalyzes the six electron reduction of sulfite to sulfide in sulfate-reducing bacteria. Along this line, it has been proposed that sulfur stored in periplasmic or extracellular sulfur globules in PSB and GSB is reductively activated, transported to, and further oxidized in the cytoplasm by sulfite reductase operating in reverse. Different models have been suggested to explain the roles of the dsr-encoded proteins in such a scenario (Dahl et al., 2005; Pott and Dahl, 1998; Cort et al., 2008). In the model shown in Fig. 2 the NADH:acceptor oxidoreductase activity of DsrL is taken into account. The protein carries a thioredoxin motif CysXXCys immediately preceding the carboxy-terminal iron–sulfur cluster-binding sites. This indicates a potential disulfide reductase activity. Therefore, the possibility exists that DsrL uses NADH as electron donor for reduction of a di- or persulfidic compound. It is suggested that DsrL is involved in the reductive release of sulfide from a perthiolic organic carrier molecule transporting sulfur from the periplasmic sulfur globules to the cytoplasm (Dahl et al., 2005). Glutathione amide is a likely candidate for the carrier molecule, as it bears an amide group at the glycyl moiety of glutathione and is especially resistant to autoxidation. It was found mainly as a persulfide in cells of Alc. vinosum grown photoautotrophically on sulfide (Bartsch et al., 1996). Recently, transporters have been characterized in E. coli mediating export (Pittman et al., 2005) and import (Suzuki et al., 2005) of glutathione. Shuttling of glutathione amide between cytoplasm and periplasm in phototrophic sulfur bacteria like Alc. vinosum, therefore, also appears feasible. From Alc. vinosum, DsrL is copurified with sulfite reductase (Y. Lu¨bbe and C. Dahl, unpublished). Sulfide released from the perthiol by the action of DsrL is possibly directly passed to the dsrAB-encoded sulfite reductase thereby reducing losses caused by evaporation of gaseous H2S. On the other hand, sulfite reductase specifically interacts also with the membrane-bound Dsr proteins and DsrEFHC (Dahl et al., 2005). Electrons released from the oxidation of sulfide by sulfite reductase may therefore be fed into photosynthetic electron transport via DsrC and DsrKMJOP, which would be analogous to the pathway postulated for sulfate reducers (Pires et al., 2006), operating in the reverse direction. DsrM could operate as a quinone reductase, DsrP as a quinol oxidase and

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finally the c-type cytochrome DsrJ would be reduced (Dahl et al., 2005). From here, electrons could be transferred to HiPIP, the primary electron donor to the photosynthetic reaction center (Vermeglio et al., 2002). The function of DsrEFH remains unclear but, as it occurs exclusively in sulfur oxidizers and shows interaction with DsrC (Cort et al., 2008), it may be important for the pathway to operate in the sulfide oxidizing direction. On the other hand, sulfur-transfer reactions as performed by the related TusBCD and TusE proteins in E. coli could also be important for the Dsr-catalyzed sulfiteformation pathway (Ikeuchi et al., 2006). Indeed, the similarity of DsrEFH to TusBCD not only on the amino acid sequence but also on a structural level points to the possibility that DsrEFH also has sulfur transferase activity (Dahl et al., 2007; C. Dahl, A. Schulte, F. Grimm, J. Sander, U. Selan, C. Hong and D.H. Shin, unpublished).

5.6. Oxidation of Sulfite to Sulfate While GSB are generally unable to oxidze externally supplied sulfite, some PSB can use sulfite as photosynthetic electron donor. In addition, sulfite appears to be generated in the cytoplasm of all phototrophic sulfur bacteria as the product of the Dsr system (Fig. 2). Currently, two different pathways for sulfite oxidation are known in chemotrophic and phototrophic sulfur oxidizers (Kappler and Dahl, 2001): (a) direct oxidation by sulfite dehydrogenase (EC 1.8.2.1), typically a molybdenum-containing enzyme; and (b) indirect, AMP-dependent oxidation via adenosine-5u-phosphosulfate (adenylylsulfate, APS). While the APS reductase pathway has been shown in several but not all GSB and Chromatiaceae, it has never been detected in PSB of the family Ectothiorhodospiraceae. In accordance, potential APS reductase genes (aprBA, see below) are not found in the genome of Hlr. halophila (Table 3). Enzymatic evidence for the simultaneous occurrence of both pathways in the same organism has been provided for several PSB and GSB (Brune, 1995b; Kappler and Dahl, 2001; Tru¨per and Fischer, 1982). The occurrence of one or both sulfite-oxidation pathways can vary between genera of the same family and even between different strains of the same genus (Frigaard and Bryant, 2008a; Kappler and Dahl, 2001). 5.6.1. Indirect Pathway via Adenylylsulfate (APS) When the indirect pathway is employed, sulfite is oxidized by adenosine-5uphosphosulfate (APS) reductase (also called adenylylsulfate reductase, EC

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 149

1.8.99.2) in a reaction that consumes sulfite and AMP and generates APS and reducing equivalents. Theoretically, APS could then be hydrolyzed to AMP and sulfate by adenylylsulfatase (EC 3.6.2.1) but there is no evidence for this enzyme in GSB or PSB. Alternatively, the energy of the phosphosulfate anhydride bond in APS can be conserved by the action of ATP:sulfate adenylyltransferase (also called ATP sulfurylase; EC 2.7.7.4). ATP sulfurylase generates ATP and sulfate from APS and pyrophosphate. Alternatively, adenylylsulfate:phosphate adenylyltransferase (APAT, formerly ADP sulfurylase (EC 2.7.7.5) (Bru¨ser et al., 2000)) generates ADP and sulfate from APS and phosphate. Since ADP can be converted to ATP and AMP by adenylate kinase, both sulfate-liberating enzymes catalyze substrate phosphorylations, which have been proposed to be of energetic importance, especially in chemolithoautotrophic bacteria (Peck, 1968). The APS pathway is also known from sulfate reduction, where it operates in the reverse direction serving assimilatory and dissimilatory purposes. The dissimilatory APS reductases are highly conserved among sulfur-oxidizing and sulfate-reducing prokaryotes (Hipp et al., 1997; Meyer and Kuever, 2007). APS reductases functioning in assimilatory sulfate reduction are completely different enzymes related to 3u-phosphoadenosine-5u-phosphosulfate (PAPS) reductases (Bick et al., 2000; Kopriva et al., 2001) (see Section 7). Indirect AMP-dependent oxidation of sulfite to sulfate via APS (Fig. 2) occurs in the bacterial cytoplasm with APS reductase being membranebound (e.g., in many Chromatiaceae) or soluble, and ATP sulfurylase and APAT being soluble enzymes (Brune, 1995a; Bru¨ser et al., 2000). In Alc. vinosum, the genes for ATP sulfurylase (sat) and APS reductase (aprMBA, with aprM encoding a putative membrane anchor) form an operon (Hipp et al., 1997; A. Wynen, H.G. Tru¨per and C. Dahl, unpublished, GenBank No. U84759). In the genomes of four GSB the genes for ATP sulfurylase and APS reductase are located directly adjacent to each other (Frigaard and Bryant, 2008a). Genes related to aprM are not present. Instead, the APS reductase and ATP sulfurylase genes in GSB are always clustered with genes encoding a so-called Qmo complex (qmoABC). Despite the absence of a recognizable APS reductase in its genome sequence, an APS reductase activity has been purified from Chl. limicola DSMZ 245T and biochemically characterized although not sequenced (Kirchhoff and Tru¨per, 1974). This APS reductase from Chl. limicola DSMZ 245T was reported to have a molecular mass of about 200 kDa and to contain one flavin per molecule and non-heme iron. Apart from potential uncharacterized membrane subunits (e.g., AprM in Alc. vinosum), all known dissimilatory APS reductases form heterodimers

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with one a-subunit (AprA) of 70–80 kDa (1 FAD) and one b-subunit (AprB) of 18–23 kDa (2 [4Fe-4S] centers) (Fritz et al., 2000; Hipp et al., 1997; Lampreia et al., 1994; Molitor et al., 1998; Speich et al., 1994). A catalytic mechanism has been proposed in which sulfite initially forms a complex with the flavin (Brune, 1995b, and references therein). This then reacts with AMP to yield APS, releasing two electrons that are transferred via the flavin to the iron–sulfur centers. The proteins that mediate the electron transport between the cytoplasmic AprAB APS reductase and the quinol/quinone pool in the membrane are still poorly characterized. However, experimental evidence is accumulating that the membrane-bound redox complex QmoABC (quinone-interacting membrane-bound oxidoreductase) is involved (Pires et al., 2003). This complex shares homology with subunits of heterodisulfide reductases. In Cba. tepidum TLS and three other GSB strains as well as in the sulfate reducers Desulfovibrio desulfuricans and Archaeoglobus fulgidus, QmoA and QmoB are cytoplasmic, nucleotide- and iron–sulfur-cluster-binding subunits and QmoC is a membrane-bound, heme-b-binding subunit that exchanges electrons with the isoprenoid quinone pool (Frigaard and Bryant, 2008a; Pires et al., 2003). The Qmo complex from Desulfovibrio desulfuricans was biochemically characterized and shown to have quinol-oxidizing activity (Pires et al., 2003). As in the four GSB, the genes aprBA-qmoABC are clustered in Dsv. desulfuricans and other Desulfovibrio-like strains. In summary, the enzymes encoded by the sat-aprBA-qmoABC operon in GSB, in principle, allow oxidation of sulfite to sulfate via an APS intermediate with concomitant reduction of membrane-bound quinones. We propose that the membrane protein AprM serves a function analogous to that of QmoABC in Alc. vinosum. Interestingly, the Apr proteins of sulfur-oxidizing phototrophic bacteria and sulfate-reducing prokaryotes diverged into two phylogenetic lineages, with the GSB affiliated with the sulfate reducers while Alc. vinosum-related sequences form a distinct group. This phylogenetic separation is exactly reflected in the differing presence of the putative proteins functionally associated with Apr, that is, the QmoABC complex and AprM (Meyer and Kuever, 2007). With regard to release of sulfate from APS via ATP sulfurylase and/or APAT, activity in crude cell extracts for both enzymes has been described for some strains of PSB (Dahl and Tru¨per, 1989; Tru¨per and Fischer, 1982). In other strains, one or the other has been reported. Likewise, some GSB strains have been reported biochemically to contain either ATP sulfurylase activity (strains DSMZ 249 and DSMZ 257) or ADP sulfurylase activity (strains DSMZ 263T and NCIMB 8346) but not both activities (Bias and Tru¨per, 1987; Khanna and Nicholas, 1983). The genome of none of these

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 151

strains has been sequenced. It should be noted that specific ‘‘ADP sulfurylase’’ activities in many reported cases were below 100 mU/(mg protein) and that such activites may be due to other enzymes not specifically involved in oxidative sulfur metabolism (Bru¨ser et al., 2000). The best characterized ATP sulfurylase (Sat) from any sulfur-oxidizing bacterium is the enzyme from the endosymbiont of the hydrothermal vent worm Riftia pachyptila (Beynon et al., 2001; Renosto et al., 1991). Like all other ATP sulfurylases, the enzyme is strictly Mg2þ-dependent. The Vmax of ATP synthesis is seven times higher than that of molybdolysis, the assay used for measuring the APS-producing reaction. The Riftia symbiont enzyme also has a higher kcat for the ATP synthesis direction (257/s compared to 64/s for the assimilatory enzyme from Penicillium chrysogenum that works in the sulfate activating direction (Renosto et al., 1991)). The native enzyme appears to be a dimer (MW 90 kDa) composed of identical size subunits (396 residues). The ATP sulfurylase from Alc. vinosum is isolated as a monomer with an apparent molecular mass of 45 kDa (A. Wynen, C. Dahl and H.G. Tru¨per, unpublished). Four GSB genomes encode highly similar, sat-derived ATP-sulfurylases (Table 3; CT0862 in Cba. tepidum TLS). More information is available for ATP sulfurylases from sulfateassimilating or sulfate-reducing organisms in which the activation of the chemically inert sulfate by adenylylation is the relevant reaction. Two completely different, unrelated types of ATP sulfurylase can be distinguished: the heterodimeric CysDN type occurs exclusively in sulfateassimilating prokaryotes, for example, E. coli (Leyh, 1993). The other ATP sulfurylases characterized in sufficient detail are monomers or homooligomers of 41–69 kDa (Gavel et al., 1998; Sperling et al., 1998; Yu et al., 2006). Size variations are due to APS kinase or PAPS-binding allosteric domains residing on the same polypeptide in some cases. Five highly conserved regions are present, two of which are rich in basic amino acids, suggesting that they may participate in binding of MgATP2and SO2 4 . The homo-oligomeric ATP sulfurylases are related with the sat-encoded dissimilatory ATP sulfurylases from sulfur oxidizers on the amino acid sequence level. The in vivo role of APAT is especially difficult to assign because all phototrophic bacteria with significant APAT activity (W100 mU/mg in crude extracts, see above and cf. Bru¨ser et al., 2000) also contain ATP sulfurylase. It has been hypothesized that APAT may serve to ensure a high turnover of APS under pyrophosphate-limiting conditions as this enzyme is independent of the energy-rich pyrophosphate molecule (Bru¨ser et al., 2000). In fact, the existence of APAT as an independent entity has been questioned

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for a long time. Only in 2000 the enzyme was purified from the chemotrophic sulfur oxidizer Thiobacillus denitrificans (Bru¨ser et al., 2000). This enzyme is a homodimer of 41.4 kDa subunits. The KM values for APS and phosphate are 300 mM and 12 mM, respectively. The pH optimum is 8.5–9.0. Catalysis is strictly unidirectional and occurs by a PingPong mechanism with a covalently bound AMP as intermediate. Histidine modification suggested a histidine as the nucleotide-binding residue. APAT from Thb. denitrificans belongs to the histidine triad (HIT) superfamily of nucleotide hydrolases and transferases and among this to the GalT family which contains three known subfamilies of enzymes (Brenner, 2002; Mccoy et al., 2006): the GalT-like UDP-hexose:hexose-1-P-uridylyltransferases, APAT and diadenosine 5u,5uuu-P1,P4-tetraphosphate (Ap4A) phosphorylase (Booth and Guidotti, 1995). Ap4A phosphorylase from yeast also has APAT activity while APAT from Thb. denitrificans does not exhibit Ap4A phosphorylase activity. The in vivo function of the latter enzyme may therefore indeed be the formation of ADP and sulfate from phosphate and APS. However, genetic evidence for this assumption is currently missing. 5.6.2. Direct Pathway While sulfite oxidases can transfer electrons to oxygen, ferricyanide, and sometime cytochrome c, the sulfite dehydrogenases can use one or both of the latter electron acceptors but not oxygen (Kappler, 2008; Kappler and Dahl, 2001). The oxygen-dependent sulfite oxidases are not relevant in anoxygenic phototrophic bacteria. The sulfite dehydrogenases belong to the sulfite oxidase family of molybdoenzymes. This family comprises established sulfite-oxidizing enzymes and proteins related to these as well as assimilatory nitrate reductases from plants (Hille, 1996). All these enzymes contain a single molydopterin cofactor; other redox active centers may be present in addition. The best characterized sulfite-oxidizing enzymes from the sulfite oxidase family are those from avian and mammalian sources (Kisker et al., 1997). These proteins are homodimers containing heme b and molybdenum coordinated via an MPT-type molybdenum pterin cofactor. The periplasmic SorAB protein from Starkeya novella is currently the best characterized bacterial sulfite-oxidizing enzyme (Doonan et al., 2006; Feng et al., 2003; Kappler and Bailey, 2005; Kappler et al., 2000; Raitsimring et al., 2005). It is a heterodimer of a large MoCo-dimer domain (40.2 kDa) and a small cytochrome c subunit (8.8 kDa). Its molybdenum pterin cofactor is of the MPT-type with a 1:1 ratio between Mo and MPT. During catalysis, electrons are sequentially transferred to a single heme c552 (Em8.0 ¼ þ280 mV) located

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 153

on the smaller subunit and passed on from there to a cytochrome c550 from the same organism, thought to be the enzyme’s natural electron acceptor. Related proteins also appear to be localized in the periplasm and to contain a heme c-binding subunit (Myers and Kelly, 2005). Genes closely related to sorAB do not occur in the currently available genomes of PSB and GSB (Table 3). Biochemical studies and, most importantly, the sequencing of a large number of bacterial genomes in the past few years revealed that many bacterial genes exist that encode proteins belonging to the sulfite oxidase family (Kappler, 2008). While the well-characterized bacterial sulfite dehydrogenases are soluble proteins, membrane-bound bacterial sulfiteoxidizing enzymes have also been reported in the literature (reviewed in Kappler, 2008; Kappler and Dahl, 2001). Most of the established or predicted soluble members of the sulfite oxidase family are periplasmic enzymes; however, some of the proteins belonging to this group (however, without a biochemically characterized function) are predicted to reside in the bacterial cytoplasm (Kappler, 2008). Direct oxidation of sulfite to sulfate in the bacterial cytoplasm can, therefore, not generally be excluded.

5.6.3. Sulfite Oxidation in Phototrophic Sulfur Bacteria: Unsolved Questions Although enzymes participating in the indirect sulfite oxidation pathway in PSB have been studied for more than 30 years (Tru¨per and Rogers, 1971), their role in vivo is still questionable. As outlined above, the oxidative APS reductase pathway occurs exclusively in members of the Chromatiaceae and in some but not all GSB. In addition, APS reductase is clearly dispensable in Alc. vinosum (Dahl, 1996). The growth rates of the wild type and an APSreductase-deficient Alc. vinosum mutant show little differences under lightlimiting conditions. A difference is observed only at saturating irradiances. Under these conditions, the wild type grows considerably faster, indicating that the presence of a second pathway of sulfite oxidation allows a higher rate of supply of reducing power (Sanchez et al., 2001). Further experiments with Alc. vinosum indicated the involvement of a molybdoenzyme in sulfite oxidation. However, so far, all attempts have failed to prove the existence of sorAB-related genes or the respective protein (U. Kappler and C. Dahl, unpublished). As already pointed out above, none of the sequenced PSB or GSB contain genes homologous to those encoding proteins of the sulfite oxidase family. As Hlr. halophila and several of the GSB do not possess genes encoding for the APS pathway, different means for sulfite oxidation must be present.

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As is apparent from Table 3, only four of 12 genome-sequenced GSB strains contain a Sat-AprBA-QmoABC enzyme system that potentially oxidizes sulfite. The sat-aprBA-qmoABC genes in Cba. tepidum TLS are immediately downstream of the dsr gene cluster, which corroborates the suggestion that they are involved in the biochemistry of the DSR system. An alternative enzyme that could oxidize the sulfite produced by the DSR system is the so-called PSRLC3 complex (see Section 5.8.2). This system is homologous with the polysulfide reductase (PSR) from Wolinella succinogenes (Krafft et al., 1992). In Wol. succinogenes, PSR and a hydrogenase allow respiration on polysulfide using H2 as the electron donor. However, homologs of PSR are also involved in metabolizing thiosulfate, tetrathionate, and other inorganic and organic compounds. PSRLC3 in GSB is encoded by a homolog of the Wol. succinogenes psrA gene (ABL64190 in Chl. phaeobacteroides DSMZ 266) and fusion of psrB- and psrC-homologous genes (ABL64189 in Chl. phaeobacteroides DSMZ 266). Sequence analysis of the PsrBC-like subunit of PSRLC3 suggests that the PsrC-like domain has an orientation in the cytoplasmic membrane that is opposite that of the PsrC subunit of Wol. succinogenes PSR, such that the PsrB-like domain of PSRLC3 is in the cytoplasm. In addition, the PsrA-like subunit of PSRLC3 does not have any obvious signal sequence. Thus, the catalytic PsrA-like catalytic subunit and the PsrB-like domain of PSRLC3 are probably located in the cytoplasm. In contrast, PsrA and PsrB are established periplasmic proteins in Wol. succinogenes. Interestingly, the genes in GSB encoding PSRLC3 are immediately upstream of the dsr gene cluster (Fig. 3). In addition, three related genes (Hhal_1934, 1935, and 1936) are found in the sulfur gene cluster of Hlr. halophila. In accordance with the situation in the GSB, the molydopterin-binding putative active-site-bearing subunit (PsrA) as well as PsrB are predicted to be localized in the cytoplasm. In summary, it is an attractive possibility that PSRLC3 in GSB and a similar PSR-like enzyme in PSB are involved in cytoplasmic oxidation of the sulfite produced by the Dsr system (Fig. 2) (Section 5.5). If so, these cytoplasmic PSR-like enzymes could provide all of the Dsr-containing green and purple bacterial organisms that lack the putative Sat-Apr sulfite oxidation system (Table 3) with a means to oxidize sulfite (except Chl. parvum DSMZ 263T for which no putative sulfite-oxidizing system is known). PSRLC3 is not widespread among other organisms, but a homologous complex with an overall amino acid sequence identity of approximate 50% is found in Chloroflexus aurantiacus, Roseiflexus sp. RS-1, and a few members of the high-GC Firmicutes. In view of the current lack of experimental evidence for the presented suggestion, we would like to point out one further possibility. Analysis of a

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 155

soxY-deficient mutant of Alc. vinosum showed that this strain is severely impaired in the oxidation of sulfite (Hensen, Franz, and Dahl, unpublished). The Sox system is also a likely candidate for sulfite oxidation in purple Alpha- and Betaproteobacteria as sulfite is accepted in vitro as a substrate of the reconstituted Sox system of Pcs. denitrificans (Friedrich et al., 2001; Sander and Dahl, 2008). Clearly, the question of sulfite oxidation in phototrophic sulfur bacteria will require special attention in the future.

5.7. Oxidation of Thiosulfate Thiosulfate (S2O2 3 ) is a rather stable and environmentally abundant sulfur compound of intermediate oxidation state. It fulfills an important role in the natural sulfur cycle and is used by many phototrophic and chemotrophic sulfur oxidizers (Jørgensen, 1990; Sorokin et al., 1999). Two completely different pathways of thiosulfate oxidation appear to exist in PSB. In the first, thiosulfate is completely oxidized to sulfate via multiple steps. In the second, tetrathionate is produced by oxidation of two thiosulfate anions via thiosulfate dehydrogenase (thiosulfate:acceptor oxidoreductase, EC 1.8.2.2). 5.7.1. Oxidation of Thiosulfate to Sulfate: The Sox Multienzyme System Many PSB and GSB can oxidize thiosulfate completely to sulfate (Table 2). In batch cultures of PSB growing on thiosulfate, the formation of sulfur globules is sometimes – but not always – observed. Sulfur globule formation on thiosulfate-containing medium has also been observed for at least some GSB (Brune, 1989; Steinmetz and Fischer, 1982) including Cba. tepidum TLS and Cba. parvum DSMZ 263T (Steinmetz and Fischer, 1982). Experiments with thiosulfate labeled with 35S either in the sulfane (35SSO2 3 ) or sulfone (S35SO2 ) position furthermore showed that Cba. thiosulfatiphi3 lum rapidly released the sulfone sulfur as sulfate, while the sulfane sulfur was retained (probably as elemental sulfur) with the bacterial cells upon filtration (Khanna and Nicholas, 1982). Elemental sulfur may be an intermediate of thiosulfate oxidation in other GSB too, although it may not always be detectable due to a high turnover rate (Brune, 1989; Dahl, 2008). The kinetics of elemental sulfur formation and degradation are probably complex and depend on growth rate; for example, rapidly growing batch cultures of Cba. tepidum TLS under high light form much higher transient concentrations of elemental sulfur from thiosulfate than slowly growing cultures under low light (N.-U. Frigaard, unpublished). Polysulfides have also been suggested as intermediates occurring in the periplasm of GSB

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during thiosulfate oxidation (Frigaard and Bryant, 2008a). Despite the obvious lack of sulfur globule accumulation under certain growth conditions or in certain strains, the formation of sulfur globules is an obligatory step during the oxidation of thiosulfate to sulfate in Alc. vinosum and probably also in other PSB. Two independent lines of evidence lead to this conclusion: (1) an Alc. vinosum mutant unable to form sulfur globules due to the lack of sulfur globule proteins cannot grow on thiosulfate (Prange et al., 2004) and (2) Alc. vinosum dsr mutants blocked in sulfur oxidation form intracellular sulfur globules from thiosulfate as a dead end product (Pott and Dahl, 1998). In addition, studies with radioactively labeled thiosulfate demonstrated very clearly that the more reduced sulfane and the more oxidized sulfone sulfur atoms are processed differently in PSB (Smith and Lascelles, 1966; Tru¨per and Pfennig, 1966). Only the sulfane sulfur accumulates as stored sulfur [S0] before further oxidation, whereas the sulfone sulfur is rapidly converted into sulfate and excreted. The formation of sulfur as an intermediate in PSB is different from the thiosulfate-oxidizing pathway (Sox pathway) that occurs in a wide range of facultatively chemoor photolithotrophic bacteria like Pcs. pantotrophus or Rhodovulum sulfidophilum (Appia-Ayme et al., 2001; Friedrich et al., 2001). In the latter, both sulfur atoms of thiosulfate are oxidized to sulfate without the appearance of sulfur deposits as intermediates. The Sox complex, a periplasmic thiosulfate-oxidizing multienzyme complex was first found and characterized in Pcs. versutus (Lu et al., 1985) and Pcs. pantotrophus (Friedrich et al., 2001; Rother et al., 2001). In Pcs. pantotrophus the Sox complex is essential for thiosulfate oxidation in vivo and catalyzes reduction of cytochrome c coupled to the oxidation of thiosulfate, sulfide, sulfite, and elemental sulfur in vitro. The sox gene cluster of Pcs. pantotrophus comprises 15 genes (soxRSVWXYZABCDEFGH). SoxR is a repressor protein of the ArsR family. SoxS is a periplasmic thioredoxin and essential for the full expression of the sox gene cluster (Rother et al., 2005). SoxV is a membrane protein and SoxW is another periplasmic thioredoxin. SoxF is a monomeric flavoprotein with sulfide dehydrogenase activity (Bardischewsky et al., 2006). The following seven genes (soxX to soxD) encode four periplasmic proteins (SoxXA, SoxYZ, SoxB, and Sox(CD)2) which constitute the core Sox system (Fig. 4A): SoxXA is composed of two c-type cytochromes, the diheme SoxA and the monoheme SoxX. The heme group in the SoxX subunit has a His/Met axial ligation. One of the hemes in SoxA (heme 1) has a classical His/Cys coordination. This heme is redox inactive. Heme 2 of SoxA is also His/Cys coordinated, albeit the cysteine is modified into a cysteine persulfide ligand resulting in an unusually low midpoint redox potential of 432 7 15 mV

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 157

A

B -

2e + H

S2O32SoxYZ -SH

+

SoxAX

SO42-+ H+

SoxAX

SoxYZ -SH

SoxYZ -S-S-SO32-

2-

SoxYZ -S-S-SO3

SoxB H2O

2e- + H+

S2O32-

H2O

H2O

SoxYZ -S-SO326e- + 7H+ SoxCD

SoxB SoxYZ -S-SH

3 H2O

SO42-+ H+

R-Sn+1H or H-Sn+1

SoxB ?

SoxYZ -S-SH

SO42-+ H+

R-SnH or H-Sn-

Figure 4 Proposed pathway of thiosulfate oxidation catalyzed by Sox systems in (A) organisms that do not form sulfur globules en route to sulfate and (B) organisms that form sulfur globules as intermediates. All reactions take place in the periplasm. Cycle A reflects the model as proposed by Friedrich et al. (2001, 2005) for the aerobic chemolithotrophic Paracoccus denitrificans. It involves SoxCD acting as a sulfur dehydrogenase. Cycle B is suggested to operate in PSB and GSB lacking SoxCD. The boxed names indicate Sox proteins catalyzing transformations of sulfur compounds bound to SoxYZ. The sulfur compound binding protein SoxYZ is circled. Sulfur compounds are bound by a conserved SoxY cysteine residue (Cys138 in Pcs. pantotrophus). The sulfur atom of this residue is indicated by a capital letter S. Two proposed sulfur acceptor substrates are shown as ‘‘RSnH’’ and ‘‘HS n .’’ Final deposition of sulfur in sulfur globules is proposed to proceed as indicated in Fig. 2. This figure is adapted from Sander and Dahl, 2008 with permission.

(Bamford et al., 2002; Cheesman et al., 2001; Dambe et al., 2005; Kappler et al., 2005; Reijerse et al., 2007). The latter heme is the one believed to participate in catalysis. SoxYZ is free of cofactors and able to covalently bind sulfur compounds of various oxidation states to a conserved cysteine residue (Quentmeier and Friedrich, 2001). The monomeric SoxB has been shown to interact with the SoxYZ complex (Quentmeier et al., 2003) and contains a dinuclear manganese cluster (Epel et al., 2005). The protein is proposed to function as a sulfate thiohydrolase. Sox(CD)2 is composed of the molybdoprotein SoxC and the diheme c-type cytochrome SoxD (Quentmeier et al., 2000). The proposed mechanism for thiosulfate oxidation requires four different proteins: SoxB, SoxXA, SoxYZ, and SoxCD (Friedrich et al., 2001) (Fig. 4A). SoxXA is proposed to fuse the sulfur substrate (thiosulfate) to the sulfhydryl Cys138 near the carboxyterminus of SoxY to initiate the reaction cycle. SoxB is believed to act as a sulfate thiol esterase and to be responsible for hydrolytic cleavage of a sulfate group from the bound sulfur substrate. Sox(CD)2 then oxidizes the remaining sulfane sulfur, acting as a sulfur dehydrogenase. Further action of

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SoxB releases a second sulfate molecule and thereby restores SoxYZ. As pointed out by Sauve´ et al. (2007), SoxYZ is the pivotal component of the Sox system since it participates in every reaction of the pathway. The crystal structure of Pcs. denitrificans SoxYZ revealed that the conserved carrier cysteine is located on a peptide swinging arm probably allowing the cysteinebound intermediates to access and orient themselves within the active sites of multiple reaction partners (Sauve´ et al., 2007). Since the identification of sox genes in Paracoccus, sox gene clusters have been found by cloning or genome-sequencing approaches not only in a great number of chemotrophic but also in many phototrophic sulfur-oxidizing bacteria, including GSB as well as purple sulfur and non-sulfur bacteria (Friedrich et al., 2005; Frigaard and Bryant, 2008a; Hensen et al., 2006; Sander and Dahl, 2008) (see also Table 3). In all GSB that have sox genes, they occur in a conserved cluster of eight genes, soxJXYZAKBW (Frigaard and Bryant, 2008a). Only one GSB strain that contains this sox cluster has not been demonstrated to grow on thiosulfate (Chl. chlorochromatii CaD3), whereas all strains that grow on thiosulfate and that have been examined for sox genes indeed contain the sox cluster. This strongly indicates that oxidation of thiosulfate in GSB is dependent on sox genes. The gene soxJ (CT1015 in Cba. tepidum TLS) encodes a putative FAD-containing dehydrogenase related to the sulfidebinding FccB flavoprotein subunit of flavocytochrome c. The gene soxK (CT1020 in Cba. tepidum TLS) encodes a hypothetical 11 kDa protein with a signal peptide and with a homolog encoded in the sox cluster of the PSB Alc. vinosum (ORF9/ABE01362) (see below). However, a homolog is not present in the Hlr. halophila sulfur gene cluster (Dahl, 2008). SoxA in GSB and some other bacteria such as Alc. vinosum and Starkeya novella have only one heme-binding motif corresponding to the heme 2 in Pcs. pantotrophus. This difference is reflected in a phylogenetic sequence analysis, in which SoxA sequences from GSB and Alc. vinosum group separately from the SoxA sequences of Pcs. denitrificans and Rhodobacter sphaeroides that have two heme-binding motifs. The soxCD genes, which are essential components of the Sox system in Pcs. pantotrophus, do not occur in the genome sequences of GSB. This observation suggests (1) that the persulfide-form of the SoxYZ carrier protein (SoxYZ-S-SH) is transformed back to the unmodified form (SoxYZ-SH) differently in GSB and in Pcs. pantotrophus and (2) that the sulfane moiety from the thiosulfate molecules that become attached to the SoxYZ carrier protein is not completely oxidized to sulfate in the GSB Sox system as they are in the Pcs. pantotrophus Sox system. In this process, a net electron gain in the GSB Sox system is only accomplished by the SoxAXdependent oxidation. The conservation of the soxJ and soxK genes in the

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GSB sox gene cluster suggests that SoxJ and SoxK may substitute for SoxCD in GSB in the process that regenerates the SoxYZ complex (Fig. 4). Alc. vinosum also contains sox genes.Gene inactivation and complementation studies clearly showed that the soxBXA and soxYZ genes, located in two independent gene regions, are essential for thiosulfate oxidation in Alc. vinosum (Hensen et al., 2006). Three periplasmic Sox proteins were purified from Alc. vinosum: the heterodimeric c-type cytochrome SoxXA (SoxX 11 kDa, SoxA 29 kDa; one covalently bound heme is present in each subunit), the heterodimeric SoxYZ (SoxY 12.7 kDa, SoxZ 11.2 kDa), and the monomeric SoxB (62 kDa, predicted to bind two manganese atoms) (Hensen et al., 2006). In Alc. vinosum the genes soxB and soxXA are transcribed divergently. Upstream of soxB a gene encoding a potential regulator protein is located and immediately downstream of soxA two further interesting genes are found. The first (ORF9) is homologous to soxK and encodes a hypothetical 12.2 kDa (9.2 kDa after processing) protein with a signal peptide. The second (rhd) encodes a putative periplasmic protein (22.2 kDa after processing) containing a conserved domain typical for rhodaneses. A homologous gene is not found close to either sox genes of GSB or in Hlr. halophila. In vitro, rhodaneses (thiosulfate sulfur transferases) can catalyze the transfer of the sulfane sulfur atom of thiosulfate to cyanide yielding thiocyanate (rhodanide, SCN) and sulfite. This is, however, not the physiological role in most cases. In the past, the detection of rhodanese and thiosulfate reductase activity in phototrophic sulfur bacteria led to the assumption that thiosulfate would be cleaved into sulfite and sulfide in the presence of suitable reduced thiol acceptors like glutathione and dihydrolipoic acid, and that the H2S formed during the proposed reaction would be immediately oxidized to sulfur stored in sulfur globules (Brune, 1989, 1995b; Dahl, 1999). However, gene inactivation showed that the Alc. vinosum rhd product does not play such a vital role and is dispensable for thiosulfate oxidation (Hensen et al., 2006). The physiological role of the rhd-encoded protein remains to be elucidated. The deduced properties of other genes encoded in immediate vicinity of the Alc. vinosum sox genes were described in detail by Hensen et al. (2006). A function in oxidative sulfur metabolism for these hypothetical proteins is not obvious. In Hlr. halophila putative sox genes are clustered but not organized in a single operon (Dahl, 2008). Hlr. halophila contains a gene homologous to soxH. A related gene is neither present in GSB nor located close to the sequenced sox genes in Alc. vinosum. The soxH gene is not required for lithotrophic growth on thiosulfate in Pcs. pantotrophus (Rother et al., 2001). In Hlr. halophila, soxBHYZ appear to be co-transcribed. They are separated

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from a gene encoding a fusion of SoxXA by a divergently oriented cluster of four genes, among them fccAB possibly encoding a flavocytochrome c (sulfide dehydrogenase). The derived FccB polypeptide also shows similarity to SoxF, an important though not essential component of the Pcs. pantotrophus Sox system (Bardischewsky et al., 2006). However, the similarity is significantly lower than that to the flavoprotein subunit FccB of Alc. vinosum flavocytochrome c (Dolata et al., 1993; Reinartz et al., 1998). The gene immediately upstream of fccB in Hlr. halophila is clearly related to fccA encoding the cytochrome c subunit of Alc. vinosum flavocytochrome c while similarity to soxE from Pcs. pantotrophus is below detection limits in searches using the BLAST algorithm (Altschul et al., 1990). On the basis of these results and the model suggested by Friedrich et al. (2001) for non-sulfur-storing bacteria (Fig. 4A), a model for thiosulfate oxidation in sulfur-storing organisms is proposed (Fig. 4B). The initial oxidation and covalent binding of thiosulfate to SoxYZ is catalyzed by SoxXA and sulfate is then hydrolytically released by SoxB. Due to the lack of the ‘‘sulfur dehydrogenase’’ SoxCD, the sulfane sulfur atom still hooked up to SoxY cannot be directly further oxidized in organisms like Alc. vinosum. Probably, the sulfur is instead transferred to growing sulfur globules. Such a suggestion is feasible as the sulfur globules in Alc. vinosum and in many if not all other organisms forming intracellular sulfur deposits reside in the bacterial periplasm (Dahl and Prange, 2006; Pattaragulwanit et al., 1998) and therefore in the same cellular compartment as the Sox proteins. How the transfer of SoxY-bound sulfur to the sulfur globules is achieved is currently unclear as the lack of the potential sulfur transferase encoded immediately adjacent to soxXA in Alc. vinosum did not lead to a detectable phenotype. Possibly, other sulfur transferases present in the cells function as a back up system (Hensen et al., 2006). In GSB and members of the Ectothiorhodospiraceae like Hlr. halophila, additional steps may be necessary for formation of extracellular sulfur globules, that is, sulfur deposited beyond the outer membrane. As outlined above, the Sox system appears to occur in two different versions in two physiologically different groups of bacteria that oxidize thiosulfate completely to sulfate: those that form sulfur as an intermediate and those that perform a direct oxidation without accumulation of sulfur (Hensen et al., 2006). A survey of currently sequenced purple bacterial genomes supported the presence of two subgroups of organisms containing sox gene clusters (Sander and Dahl, 2008): those containing genes for the potential sulfur dehydrogenase SoxCD and those lacking these genes. This conspicuous difference is well related to the intermediates formed in the organisms. A complete sox gene cluster occurs in organisms not forming

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deposits of elemental sulfur. These include the thiosulfate-oxidizing purple non-sulfur bacteria. The soxCD genes are not present in most organisms forming sulfur deposits. These include the GSB and PSB. However, it should be pointed out that the chemotrophic sulfur-oxidizing epsilonproteobacteria Thiomicrospira crunogena and Sulfurimonas denitrificans (Scott et al., 2006; Sievert et al., 2008) and the marine gammaproteobacterium Neptuniibacter caesariensis MED92 (Arahal et al., 2007) have soxCD, although their other sox genes are more closely related to those of Cba. tepidum, Alc. vinosum, and Thb. denitrificans, which do not have soxCD and produce sulfur globules. Interestingly, Tms. crunogena is able to form sulfur from thiosulfate (Javor et al., 1990) while elemental sulfur formation by S. denitrificans has not been reported. Sievert et al. (2008) speculated that Tms. crunogena soxCD is regulated and turned on and off depending on environmental conditions. Furthermore, these authors proposed that SoxCD may even function differently in S. denitrificans and Tms. crunogena and that it might exhibit sulfite dehydrogenase activity in these organisms. 5.7.2. Oxidation of Thiosulfate to Tetrathionate: Thiosulfate Dehydrogenase The formation of tetrathionate from thiosulfate has been mainly studied in chemoorganotrophic bacteria that use thiosulfate as a supplemental but not as the sole energy source (Jørgensen, 1990; Podgorsek and Imhoff, 1999; Sorokin et al., 1999). The pathway occurs only in a few PSB including Alc. vinosum (Hensen et al., 2006; Smith and Lascelles, 1966). In Alc. vinosum the ratio between tetrathionate and sulfate formed from thiosulfate is strongly pH-dependent with more tetrathionate as the product under slightly acidic conditions (Smith, 1966). In Alc. vinosum thiosulfate dehydrogenase is a periplasmic 30 kDa monomer with an isoelectric point of 4.2. The enzyme contains heme c and is reduced by thiosulfate at pH 5.0 but not at pH 7.0. In accordance, the pH optimum of the enzyme was determined to be 4.25 (Hensen et al., 2006). An examination of the kinetic properties of Alc. vinosum thiosulfate dehydrogenase with ferricyanide as artificial electron acceptor was initiated but interpretation of experimental results is complicated by the fact that enzymes that use two molecules of the same substrate do not follow regular Michaelis–Menten kinetics. However, some important constants could be estimated: the limiting Vmax is about 34,000 units/(mg protein) (corresponding to a kcat of 1.7  104/s) and the [S]0.5 for ferricyanide is about 0.5 mM. [S]0.5 is the substrate concentration that yields half maximal velocity. It is important to note that it is not identical to Km as a Km cannot be given for reactions not following Michaelis–Menten kinetics (Segel, 1993). While thiosulfate did not display

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strong substrate inhibition at any of the experimental ferricyanide levels, ferricyanide did show substrate inhibition on Alc. vinosum thiosulfate dehydrogenase (Hensen et al., 2006). Furthermore, the enzyme was significantly inhibited by sulfite (50% inhibition at 80 mM of sulfite). Under optimized assay conditions cytochrome c from yeast is used as electron acceptor instead of ferricyanide by the enzyme, whereas horse heart cytochrome c is not accepted. The properties of Alc. vinosum thiosulfate dehydrogenase described by Hensen et al. (2006) are compatible with older data presented by Smith (1966) and Fukumori and Yamanaka (1979). In both reports a tetrathionate-forming activity with a pH optimum in the acidic range was described. The presence of a tetrathionate-forming enzyme operating at pH 8.0 in Alc. vinosum (Knobloch et al., 1981; Schmitt et al., 1981) could not be confirmed by Hensen et al. (2006). Thiosulfate dehydrogenases from other bacterial sources show remarkable heterogeneity with respect to structural properties and catalytic characteristics (Brune, 1989; Kusai and Yamanaka, 1973; Then and Tru¨per, 1981; Visser et al., 1996) which has been interpreted as indicating convergent rather than divergent evolution (Visser et al., 1996). A gene sequence encoding a heme-containing thiosulfate dehydrogenase has not yet been reported. A Blast search with the amino-terminal sequence of the enzyme from Alc. vinosum yielded only one significantly related sequence, a hypothetical c-type cytochrome from Cupriavidus (Ralstonia, Wautersia) metallidurans (Hensen et al., 2006).

5.8. Other Enzymes Related to Sulfur Compound Oxidation 5.8.1. RuBisCO-Like Protein The enzyme RuBisCO (ribulose 1,5-bisphosphate carboxylase/oxygenase) catalyzes the key step in the Calvin–Benson–Bassham CO2 fixation pathway in many phototrophic and chemotrophic organisms (Tabita, 1999). Genome sequencing revealed that some bacteria and archaea contain homologs of RuBisCO, called RuBisCO-like proteins (RLPs), which do not have the same enzymatic activity as bonafide RuBisCO. For example, in Bacillus subtilis RLP functions as a 2,3-diketomethythiopentyl-1-phosphate enolase in the methionine salvage pathway of this organism (Ashida et al., 2003). All GSB genomes sequenced to date contain an RLP, as does Alc. vinosum (Q93UZ0) but not Hlr. halophila. GSB do not have other recognizable genes

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for a methionine salvage pathway and thus the RLP probably does not function in methionine metabolism. A mutant of Cba. tepidum TLS lacking RLP (CT1772) has a pleiotropic phenotype with increased levels of oxidative stress proteins and defects in photopigment content, photoautotrophic growth rate, carbon fixation rates, and the ability to oxidize thiosulfate and elemental sulfur (Hanson and Tabita, 2001, 2003). Notably, sulfide oxidation is not affected in the rlp mutant of Cba. tepidum TLS. Hanson and Tabita subsequently suggested that RLP is involved in the biosynthesis of a low-molecular-weight thiol, which is essential for oxidation of thiosulfate and elemental sulfur. The possible role of such a hypothetical thiol as a carrier of sulfane sulfur is illustrated in Fig. 2. However, the function of GSB RLP is probably not limited to oxidation of inorganic sulfur compounds because Chl. ferrooxidans contains an RLP very similar to the RLP in other GSB, even though this organism cannot grow on inorganic sulfur compounds and does not contain genes thought to be involved in oxidation of thiosulfate (sox genes) and elemental sulfur (dsr genes) (Table 4). 5.8.2. Polysulfide-Reductase-Like Complexes The well-characterized polysulfide reductase (PSR) in the chemotrophic Wolinella succinogenes allows respiration on polysulfide by producing sulfide (Krafft et al., 1992). The Wol. succinogenes PSR is encoded by the psrABC genes and consists of two periplasmic subunits, a molybdopterin-containing PsrA subunit and a [4Fe-4S]-cluster-binding PsrB subunit, and a membraneanchoring PsrC subunit that binds an isoprenoid quinone and exchanges electrons with PsrB. The PsrA subunit is translocated to the periplasm by the twin-arginine transport (Tat) system. Several homologs of PSR are found in the genome sequences of phototrophic sulfur bacteria but because PSR homologs have been implicated in metabolism of thiosulfate, tetrathionate, and other inorganic and organic compounds, the function of the PSR-like complexes in GSB and PSB cannot easily be established from sequence analysis alone. Three types of complexes, here denoted polysulfide-reductase-like complex 1, 2, and 3 (PSRLC1, PSRLC2, and PSRLC3), with sequence similarity to the characterized PSR in Wol. succinogenes, are found in the genome sequences of GSB (Table 4). Similar to the case of Wol. succinogenes PSR, PSRLC1 (comprising CT0494, CT0495, and CT0496 in Cba. tepidum TLS) and PSRLC2 (comprising Cpha266_2562, 2563, and 2564 in Chl. phaeobacteroides DSMZ 266) are encoded by three genes. For both PSRLC1 and PSRLC2, the PsrA-like subunits with the catalytic site have a Tat signal

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sequence and thus should be translocated into the periplasm. PSRLC3 is encoded by two genes similar to psrA and a fusion of psrBC. The PsrA-like subunit of this complex does not have a Tat signal and presumably is located in the cytoplasm. The PSRLC3 has been implicated in sulfite oxidation as discussed in Section 5.6.3. Homologs of PSRLC1 and PSRLC2 are found in many other organisms; for example, Carboxydothermus hydrogenoformans has a PSRLC1 homolog that has an overall amino acid sequence identity of approximate 50% with the PSRLC1 of Cba. tepidum TLS. Hlr. halophila also has three types of PSR-like complexes with unknown function, each of which is encoded by three genes. The genes of one of the PSRLC (Hhal_1164, 1165, 1166) are located immediately adjacent, but divergently transcribed, to genes encoding an FccAB flavocytochrome c homolog (Hhal_1162, 1163). The PsrA-like subunit of this PSRLC and one of the other PSRLC (Hhal_0353, 0354, 0355) contain a Tat signal sequence. However, the PsrA-like subunit of the third PSRLC in Hlr. halophila (Hhal_1936, 1935, 1934) does not have a Tat signal sequence and the genes encoding this complex cluster with genes related to sulfur metabolism. Thus, it is possible that a PSR-like complex in the GSB and Hlr. halophila that do not have an APS reductase (AprBA) functions in sulfite oxidation (see Section 5.6.3). 5.8.3. Sulfhydrogenase-Like and Heterodisulfide-Reductase-Like Complexes A putative cytoplasmic abgd-heterotetrameric, bi-directional hydrogenase, which resembles Pyrococcus furiosus hydrogenase II that catalyzes H2 production, H2 oxidation, as well as the reduction of elemental sulfur and polysulfide to sulfide (Ma et al., 2000), is present in all sequenced GSB genomes, except those of Chl. phaeovibrioides DSMZ 265 and Chl. luteolum DSMZ 273T (Table 4). The respective genes are not present in the PSB Hlr. halophila. The genes encoding this putative sulfhydrogenase form a conserved hyd1 cluster, hydB1G1DA (CT1891–CT1894 in Cba. tepidum TLS), except in Chl. chlorochromatii CaD3 in which the genes are split into two clusters, hydB1G1 and hydDA. Since Cba. tepidum TLS is unable to grow on H2, these genes apparently do not confer the ability to oxidize large amounts of H2. Likewise, the presence of this enzyme in Chl. ferrooxidans DSMZ 13031T and its absence from Chl. phaeovibrioides DSMZ 265, Chl. luteolum DSMZ 273T, and Hlr. halophila suggests that its primary role is not related to elemental sulfur or polysulfide metabolism. There are two types of complexes with sequence homology to heterodisulfide reductases encoded in the genomes of the sequenced GSB. One is the Qmo complex, which is probably involved in intracellular sulfite oxidation as

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discussed above (Section 5.6.1). The other is encoded by genes that form a conserved cluster with genes encoding a putative hydrogenase. This hdr-hyd2 gene cluster, hdrD-hdrA-orf1247-orf1248-hydB2-hydG2, is conserved in seven of the sequenced strains. The hdrD gene in GSB is a fusion of the hdrC and hdrB genes found in other organisms. As with the Hyd1 complex mentioned above, the presence of Hdr-Hyd2 in Chl. ferrooxidans DSMZ 13031T and its absence from two other GSB strains and also from Hlr. halophila, suggests that its presence is not essential in elemental sulfur or polysulfide metabolism.

6. EVOLUTION OF SULFUR METABOLISM The dissimilatory oxidation of sulfur compounds as a means to supply phototrophic sulfur bacteria with electrons for thiotrophic growth clearly is a complex metabolism. Nevertheless, increasing biochemical and genetic information reveal that many similarities exist in the enzyme systems that transform sulfur compounds in thiotrophic organisms and that these similarities in many cases clearly are due to lateral gene transfer events. However, it is also apparent that the exact genetic composition in a given organism is optimized for the particular lifestyle of the organism. As described above (Section 5.5), the Dsr system appears to allow efficient oxidation of elemental sulfur in both PSB and GSB. However, the genetic history of the Dsr system is somewhat different in these two types of bacteria, since parts of the Dsr system in GSB (DsrAB and other proteins) are related to the Dsr system in PSB, whereas other parts (DsrMKJOPT) are related to the Dsr system in sulfate-reducing bacteria (Sander et al., 2006). In fact, the observation that the earliest diverging GSB, Chp. thalassium, does not possess the Dsr system, suggests that the common ancestor of all currently known GSB did not posses the Dsr system. Thus, it is an interesting possibility that it might have been the acquisition of a (chimeric) Dsr system that has led to the relatively recent, explosive radiation of the lineages of GSB that are not closely related to Chloroherpeton. Genetic evidence also suggests that the GSB that contain the Dsr system may oxidize the sulfite generated by the Dsr enzymes in different manners, some using the Sat-Apr-Qmo system (Section 5.6.1), while others may use the putative PSRLC3 complex (Sections 5.6.3 and 5.8.2). The former system may present an advantage over the latter system by apparently conserving a high-energy phosphate anhydride bond (Fig. 2). One GSB strain, Chl. ferrooxidans, that grows on ferrous iron has lost the Dsr system and other sulfur-oxidizing enzymes and consequently the ability to grow on sulfur compounds altogether.

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Thiosulfate oxidation is found only in certain GSB strains and is dependent on a modified Sox system (Section 5.7.1). It differs conspicuously from the well-characterized Sox system of certain chemotrophic bacteria by not including the SoxCD enzyme (Fig. 4). The absence of SoxCD disables GSB from oxidizing thiosulfate completely to sulfate in the periplasm using the Sox system. As a consequence of not having SoxCD, these organisms probably transiently accumulate polysulfide species, and possibly elemental sulfur, which subsequently is oxidized by the Dsr system. The same is true for the PSB Alc. vinosum that has a lifestyle similar to GSB. The question then arises as to why these bacteria do not oxidize thiosulfate (and possibly other sulfur compounds) completely to sulfate using the Sox system? One possible explanation is that the absence of SoxCD forces the oxidation of sulfur to occur via the Dsr system, thus possibly conserving energy by generating a transmembrane proton gradient and high-energy phosphate anhydride bonds (Fig. 2), which do not occur if the sulfur is oxidized completely to sulfate in the periplasm (L.H. Gregersen and N.-U. Frigaard, unpublished). This metabolic ‘‘trick’’ could be a hallmark of phototrophic sulfur bacteria since all known Sox-containing PSB and GSB contain the Dsr system and not SoxCD, whereas other bacteria that contain a closely related Sox system (e.g., the marine gammaproteobacteria Neptuniibacter caesariensis MED92 (Arahal et al., 2007) and Congregibacter litoralis KT71 (Fuchs et al., 2007)) contain SoxCD but not the Dsr system. Thiosulfate utilization by the Sox system in GSB presents an interesting case study of lateral transfer. The Sox proteins are only present in some GSB strains and have phylogenies that are incongruent with that for ribosomal RNAs (Frigaard and Bryant, 2008a). This indicates that the sox genes in GSB were not inherited vertically from a common ancestor. However, the Sox proteins from GSB form a monophyletic cluster in phylogenetic analyses, which strongly implies that the currently known sox genes in GSB have only been laterally exchanged within the GSB. In addition, all eight genes in the sox gene cluster of GSB have congruent phylogenies. This suggests that all eight sox genes were transferred simultaneously as one conserved cluster to each recipient strain. How might such a transfer have occurred? Two GSB strains, DSMZ 273T and DSMZ 265, which are very closely related in terms of ribosomal RNA phylogeny and genome organization, differ in one important respect: the latter strain contains the sox gene cluster whereas the former strain does not (Table 3). Analysis of the genome sequences reveals that the sox cluster in strain DSMZ 265 resides on an 11 kbp island that contains four additional genes. This island appears to have been inserted into a region of the genome in a recent

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ancestor that was not involved in sulfur metabolism. This ancestor was likely to be similar to strain DSMZ 273T and unable to use thiosulfate. The genes on the island include a transposase, an integrase, and an RNA-directed DNA polymerase, all of which are indicative of a mobile element. Such a mobile element may be carried by a plasmid or a virus. The observation that the sox genes only occur in the Chlorobium/Chlorobaculum lineage, and not other GSB lineages, suggests that the putative mobile element only transfers within this lineage of closely related organisms – a trait observed with many plasmids and viruses. To our knowledge, no virus of any kind that infects GSB has yet been isolated but there is no reason to believe such viruses do not exist.

7. SULFATE ASSIMILATION IN ANOXYGENIC PHOTOTROPHIC BACTERIA Assimilatory metabolism of sulfate in anoxygenic phototrophic bacteria has not been covered in the reviews by Brune (1989, 1995b). We therefore take this opportunity to summarize the currently available information for all groups of these organisms and include not only GSB and PSB but also purple non-sulfur bacteria, FAP bacteria, aerobic anoxygenic bacteria, and one representative of the Heliobacteria.

7.1. Historical Aspects and General Outline of the Pathway Assimilatory reduction of sulfate in phototrophic organisms (mostly plants and algae) has been investigated mostly from a nutritional point of view. Sulfate was believed generally not to be limiting for growth (Mothes and Specht, 1934). For Enterobacteria, a pathway (Fig. 5) was described (reviewed in Kredich, 1996) which was believed to be operating in phototrophic bacteria as well: (1) activation of sulfate to form a sulfonucleotide (adenosine-5uphosphosulfate); (2) followed by a second activation step with ATP to form 3u-phospho-adenosine-5u-phosphosulfate (PAPS); (3) a thioredoxin-dependent PAPS reductase then forms free sulfite; (4) sulfite reductase uses electrons from NAD(P)H or ferredoxin, respectively, and free sulfide is released that is finally (5) integrated into cysteine. It should be noted that activated sulfate in the form of PAPS is generally very important for sulfate transfer reactions to numerous substrates yielding an array of sulfonated

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Sulfateout cysPTWA or sul1

Sulfatein ATP ATP sulfurylase cysDN or sat PPi APS cysC

APS kinase 2 e-

Sulfotransferases

APS reductase PAPS

cysH

2 AMP

PAPS reductase

e-

Sulfonated products e.g. sulfolipids

cysH PAP

Sulfite

6 e- + 6 H+ Sulfite reductase

Serine + Acetyl-CoA Serine acetyltransferase

cysI or cysIJ

cysE 3 H2O Sulfide

O-Acetylserine O-acetylserine (thiol)-lyase A or B cysK or cysM

Cysteine

Figure 5 Assimilatory sulfate reduction and cysteine biosynthesis. Alternative pathways and enzymes occurring in anoxygenic phototrophic bacteria are shown. Direct reduction of APS appears to be more common than direct reduction of PAPS (cf. Table 5). Gene names are boxed. Except for the membrane-bound transporters, all enzymes involved are localized in the cytoplasm.

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 169

compounds, for example, sulfolipids (Negishi et al., 2001). Such biosyntheses can also occur in organisms that are not able to perform the reductive steps, that is, the capacity for activation of sulfate is not necessarily combined with the capacity for assimilatory sulfate reduction. In the 1970s, the simple transfer of the enterobacterial pathway was challenged for cyanobacteria, unicellular green algae, as well as for Euglena (reviewed by Schmidt and Ja¨ger, 1992): It was especially questioned whether PAPS was indeed the substrate for the first reduction step or whether APS was reduced without further activation. Evidence was presented for both alternatives (Abrams and Schiff, 1973; Brunold and Schiff, 1976; Kanno et al., 1996; Li and Schiff, 1991; Schmidt, 1973; Schwenn, 1989). The true nature of the reaction mechanism remained elusive (reviewed in Hell, 1997). For plants, the final solution was approached with the proof of APS reductase in Arabidopsis thaliana and biochemical identification of the enzyme from Lemna (Gutierrez-Marcos et al., 1996; Setya et al., 1996; Suter et al., 2000). PAPS turned out to be the primary substrate for reduction in diverse eubacteria (including enterobacteria), archaea, and fungi, but not in plants, algae, and cyanobacteria. In the latter groups, APS is the major substrate for reduction and this also appears to hold true for most anoxygenic phototrophic bacteria (Sander and Dahl, 2008). In bacteria including anoxygenic phototrophic bacteria, sulfate reduction pathways are clearly induced under sulfur-limited conditions. Today it is generally accepted that sulfate uptake, reduction and fixation of sulfide are the most regulated steps in this process (Haverkamp and Schwenn, 1999; Kredich, 1996; Neumann et al., 2000; Sander and Dahl, 2008).

7.2. Occurrence of Sulfate Assimilation in Anoxygenic Phototrophic Bacteria The genes related to assimilatory sulfate reduction in the genomes of anoxygenic phototrophic bacteria currently available are tabulated in Table 5. It appears that the enzymes involved in sulfate assimilation in these organisms are not uniform but may vary even within the same family (compare Alc. vinosum and Tcs. roseopersicina). Some anoxygenic phototrophic bacteria are very much specialized for living in habitats with reduced sulfur compounds and such bacteria may completely lack a sulfate reduction pathway. The GSB are a good example for this specialization. Most members of the Chlorobiaceae lack genes for the enzymes catalyzing sulfate assimilation (in Table 5 Cba. tepidum is given as an example). Within the Chromatiaceae and Ectothiorhodospiraceae the

170

Table 5

Occurrence of genes related to assimilatory sulfate reduction in anoxygenic phototrophic bacteria cysPTWA or sul1

sat

cysDN

cysH [4Fe-4S]2þ

cysH cysJ no [4Fe-4S]2þ

cysI

Alphaproteobacteria Bradyrhizobiaceae Rhodopseudomonas palustris BisA53

cysPTWA



þ





þ

Rhodopseudomonas palustris BisB18

cysPTWA



þ





þ

Rhodopseudomonas palustris BisB5

cysPTWA



þ





þ

cysPTWA



þ





þ

cysPTWA



2 copies (bothþAPS kinase) 2 copies (bothþAPS kinase) þ (þ APS kinase) þ (þ APS kinase) þ (þ APS kinase)

þ





þ

sul1 no cysP, no sul1 sul1 sul1

þ (þ APS kinase) þ (þ APS kinase)

 

þ þ

 

 

þ þ

þ (þ APS kinase) 

 

þ þ

 

 

þ þ

3 copies cysPTWA þ (þ APS kinase)



þ





þ

2 copies cysPTWA þ (þ APS kinase)









þ

cysPTWA

þ (þ APS kinase)



þ





þ

sul1 sul1

þ (þ APS kinase) þ

 

þ þ

 

 

þ þ

Rhodopseudomonas palustris CGA009 (ATCC BAA-98) Rhodopseudomonas palustris HaA2 (ATCC BAA-1122) Rhodobacteraceae Roseovarius sp. 217a Roseovarius nubinhibens ISM (DSMZ 15170T)a Roseobacter sp. MED193a Roseobacter denitrificans OCh 114 Rhodobacter sphaeroides ATCC 17025a Rhodobacter sphaeroides ATCC 17029a Rhodobacter sphaeroides 2.4.1 (ATCC 55304) Rhodobacterales bacterium HTCC2654a Dinoroseobacter shibae DFL_12a

NIELS-ULRIK FRIGAARD AND CHRISTIANE DAHL

Organism

–



þ (þ APS kinase)

þ



þ

þ

Rhodospirillaceae Rhodospirillum rubrum ATCC 11170T

cysPTWA



þ





þ

cysPTWA



þ (þ APS kinase) þ

þ



(þ)

þ

cysPTWA cysPTWA

 

þ þ

þ þ

 

 

þ þ

sul1

þ



þ



þ

2 copies sul1



þ



þ

sul1



2 copies (one þ copyþAPS kinase) þ (þ APS þ kinase)







unknown unknown

þ (dissimilatory) unknown

þ unknown

þ 

 þ

 þ

þ þ

cysPTWA



þ





þ

cysPTWA 

 þ (dissimilatory)

þ (þ APS kinase) þ 

þ 

 

 

þ 

sul1

2 copies (oneþAPS  kinase)









Rhodocista centenaria ATCC 51521a Betaproteobacteria Comamonadaceaee Rubrivivax gelatinosus PM1 Rhodoferax ferrireducens T118 (DSMZ 15236T) Gammaproteobacteria Congregibacter litoralis KT71a Ectothiorhodospiracea e Alkalilimnicola ehrlichei MLHE-1 (DSMZ 17681T) Halorhodospira halophila SL1 (DSMZ 244T) Chromatiaceae Allochromatium vinosum DSMZ 180Tb Thiocapsa roseopersicina M1c Green Sulfur Bacteria (Chlorobiaceae) Chlorobium ferrooxidans DSMZ 13031T Chlorobium luteolum DSMZ 273T Chlorobaculum tepidum TLS (DSMZ 12025T) FAP Bacteria (Chloroflexaceae) Chloroflexus aggregans DSMZ 9485T

(Continued )

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 171

Acetobacteraceae Acidiphilium cryptum JF-5a

172

Table 5 (Continued ) cysPTWA or sul1

sat

cysDN

cysH [4Fe-4S]2þ

cysH cysJ no [4Fe-4S]2þ

cysI

Chloroflexus aurantiacus J-10-fl (DSMZ 635T) Roseiflexus castenholzii DSMZ 13941Ta Roseiflexus sp. RS-1 Heliobacteria Heliobacterium modesticaldum ATCC51547Ta

sul1

þ (þ APS kinase)



þ





þ

sul1 sul1

þ (þ APS kinase) þ (þ APS kinase)

 

þ þ

 

 

 

cysPTWA



þ









This survey is based on complete or draft genome sequence data, except for Alc. vinosum and Tca. roseopersicina for which only limited sequence information is available. The following purple bacterial genera are ABC bacteria: Roseovarius, Roseobacter, Dinoroseobacter, Acidiphilium, and Congregibacter. Four non-phototrophic species (underlined) that do not contain genes encoding reaction center polypeptides are included for comparative reasons. Cba. tepdium is shown as an example of those GSB that are incapable of assimilatory sulfate reduction. a draft genomes at the time of compilation of this table, Genomes were analyzed by BLAST searches using the resources provided by Integrated Microbial Genomes (DOE Joint Genomes Institute, http://img.jgi.doe.gov) or by the Phototrophic Prokaryotes Sequencing Project (http:// genomes.tgen.org). b Neumann et al. (2000). c Haverkamp and Schwenn (1999).

NIELS-ULRIK FRIGAARD AND CHRISTIANE DAHL

Organism

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 173

specialized species are also unable to grow photoorganotrophically and accordingly none of these species assimilates sulfate as a sulfur source (Imhoff, 2005a, 2005b). On the other hand, very many versatile purple and even a few green bacteria are able to assimilate and reduce sulfate in the absence of a source of reduced sulfur. Chl. ferrooxidans DSMZ 13031T is one example for a GSB that grows with sulfate as the sole sulfur source and cannot utilize sulfide, thiosulfate, or elemental sulfur as electron donors (Heising et al., 1999). In agreement with this observation, the Chl. ferrooxidans genome encodes a single gene cluster that includes the assimilatory sulfate reduction genes cysIHDNCG and the sulfate permease genes cysPTWA, which are transcribed in opposite directions. These assimilatory sulfate reduction genes share a high degree of sequence similarity with those of the clostridia Clostridium thermocellum and Desulfitobacterium hafniense. An identical cys gene cluster is observed in Chl. luteolum DSMZ 273T raising the possibility that this strain also is capable of assimilatory sulfate reduction and growth in the absence of reduced sulfur compounds using electron donors such as H2 and Fe2þ. The GSB Chl. phaeovibrioides DSMZ 265 and Ptc. aestuarii DSMZ 271T also contain cysC and cysN homologs. However, in these two organisms the genes occur in a cluster that appears to be involved in another aspect of sulfur metabolism. This cluster also contains a cysQ homolog that may encode a phosphatase acting on APS or PAPS (Neuwald et al., 1992) and a homolog of the ArsB/NhaD superfamily of permeases that translocates Naþ and various anions such as sulfate across the cytoplasmic membrane (500231320 and 500231330, respectively, in Chl. phaeovibrioides DSMZ 265). It is therefore possible that Chl. phaeovibrioides DSMZ 265 and Ptc. aestuarii DSMZ 271T possess a system that processes APS or sulfite (or both) differently than in other GSB and that actively excretes sulfate. Among the FAP bacteria, the ability to assimilate sulfate may or may not be present, exemplified by the sequenced Chloroflexus aggregans, Chloroflexus aurantiacus, and Roseiflexus strains. While Cfl. aurantiacus harbors a set of genes the products of which can catalyze the complete activation and reduction of sulfate, Cfl. aggregans and Roseiflexus strains are predicted to be able to take up and activate, but not to be capable of reducing sulfate (Table 5). There is currently only one genome-sequenced representative of the Heliobacteria, Heliobacterium modesticaldum Ice1 (ATCC 51547T) (http://genomes.tgen.org/helio.html). This organism cannot grow on sulfate as the sole sulfur source. Accordingly, genes for proteins involved in reduction of (P)APS and sulfite are not present.

174

NIELS-ULRIK FRIGAARD AND CHRISTIANE DAHL

7.3. Uptake of Sulfate Assimilatory sulfate reduction commences with the uptake of extracellular sulfate (Fig. 5). In Escherichia coli the genes cysTWA encode three membrane-bound components of a periplasmic substrate-binding transport system (Hryniewicz and Kredich, 1991). The preceding gene cysP has been shown to encode a periplasmic thiosulfate binding protein distinct from the sulfate binding protein of Salmonella typhimurium. In plants as well as many lower eukaryotes, sulfate transport is mediated by both high- and low-affinity systems. Both act as thermodynamic active processes which are thought to be driven by proton motive force, through a Hþ/SO2– 4 co-transport (MendozaCo´zatl et al., 2005). The characterized high-affinity transporters Sul1 from Saccharomyces cerevisiae (Cherest et al., 1997) and Hst1At from Arabidopsis thaliana (Vidmar et al., 2000) are prominent members of the MFS superfamily of sulfate permeases and related transporters (COG0659). This family also comprises a number of bacterial proteins. Most anoxygenic phototrophic bacteria encode genes homologous to cysPTWA indicating that they use a sulfate transport system related to that of Enterobacteria (Table 5). In some anoxygenic phototrophic bacteria, including Hlr. halophila, related genes do not appear to be present. Instead, a gene is found encoding for a potential sulfate permease related to the highaffinity sulfate transporter of Saccharomyces crevisiae. However, in Hlr. halophila, Rhodobacterales bacterium HTCC2654, and Rosoevarius sp. 217, this gene is located in immediate vicinity of clusters of genes involved in oxidative sulfur metabolism (Sander and Dahl, 2008). Therefore, the encoded transporter may also be involved in export of sulfate produced in the cytoplasm as an end product of the oxidation of reduced sulfur compounds.

7.4. Activation of Sulfate Once inside of the cell, the low reactivity of sulfate must first be overcome by the formation of a phosphate–sulfate anhydride bond in the compound adenosine-5u-phosphosulfate (adenylylsulfate, APS). This reaction is catalyzed by the enzyme ATP sulfurylase (EC 2.7.7.4) (Leustek and Saito, 1999). Due to instability of APS and the unfavourable reaction equilibrium (DGu ¼ þ45 kJ/mol) the backward reaction was believed to be prevented by cleavage of pyrophosphate by abundant pyrophosphatase activity and further activation by APS kinase to PAPS (Leustek et al., 2000). As

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 175

elaborated above, PAPS is indeed very important for sulfate transfer reactions to numerous substrates (Negishi et al., 2001). Assimilatory ATP sulfurylases occur in two different forms: a heterodimeric CysDN type from E. coli (Leyh, 1993) and an unrelated homooligomeric type found in bacteria, plants, and fungi (Foster et al., 1994; MacRae et al., 2001). The latter polypeptides are related to the sat-encoded dissimilatory ATP sulfurylases of sulfate-reducing and chemotrophic sulfuroxidizing prokaryotes (Beynon et al., 2001; Sperling et al., 1998). In both, heterodimeric and homo-oligomeric, ATP sulfurylases carboxy-terminal APS kinase domains may be present. In anoxygenic phototrophic bacteria both types of ATP sulfurylases can be present. Genes encoding the Sat-type of ATP sulfurylase are found in Rhodobacteraceae and FAP bacteria while the other analyzed genomes contain cysDN genes. In most of these cases, cysN is fused with an APS kinase gene.

7.5. Reduction to Sulfite For incorporation of reduced sulfur into biomolecules, for example amino acids, sulfate in APS is reduced to sulfite and finally into sulfide. This process may occur differently, depending on the organism (Fig. 5). Similar to the situation in E. coli APS may first be phosphorylated by an APS kinase (EC 2.7.1.25, gene cysC) using ATP to produce PAPS and ADP. In the following reaction, PAPS reductase (EC 1.8.99, gene cysH) generates free sulfite using reduced thioredoxin as electron donor. The other pathway involves the direct reduction of APS by APS reductase (EC 1.8.99.x, also cysH) (Bick et al., 2000). Assimilatory APS reductases and PAPS reductases are related on a sequence level. The signature of the plant, algae and cyanobacteria type APS reductase was originally seen in a two domain structure consisting of a reductase domain with an [4Fe-4S]2þ cluster and a C-terminal thioredoxin/ glutaredoxin domain, using glutathione as electron donor. In contrast, PAPS reductase was believed to have only the reductase domain without an iron–sulfur cluster and to use thioredoxin as electron donor (Kopriva et al., 2002). Today, the situation indicates even more variants of assimilatory sulfonucleotide reductases: (1) the E. coli CysH-like enzymes reduce PAPS using thioredoxin or glutaredoxin as electron donor, have no [4Fe-4S]2þ cluster, and are found in many eubacteria, archaea, and fungi (Berendt et al., 1995); (2) another CysH-like group reduces APS rather than PAPS, uses thioredoxin as electron donor, has an [4Fe-4S]2þ cluster, and was found

176

NIELS-ULRIK FRIGAARD AND CHRISTIANE DAHL

in Pseudomonas aeruginosa (Bick et al., 2000); (3) yet another protein with strong homology to CysH is from Bacillus subtilis and is able to reduce APS and PAPS (Berndt et al., 2004); (4) the plant-type sulfonucleotide reductase (e.g. Arabidopsis thaliana, Enteromorpha intestinalis) uses APS, carries an [4Fe-4S]2þ cluster, and contains a C-terminal glutaredoxin-like domain (Bick et al., 1998; Gao et al., 2000); (5) a reductase from the moss Physcomitrella patens that shares high similarity with CysH-like proteins, has no iron–sulfur cluster but prefers APS rather than PAPS (Kopriva et al., 2007). This functional variability seems independent of oxygenic or phototrophic lifestyles and reflects the complex evolutionary development of assimilatory sulfate reduction (Kopriva, 2008). Table 5 shows that almost all of the cysH genes present in anoxygenic phototrophs encode proteins predicted to bind an iron–sulfur cluster. Based on the above considerations it appears most likely that these proteins reduce APS rather than PAPS. Alkalimnicola ehrlichii is an exception as it contains a bonafide PAPS reductase gene albeit in addition to a gene encoding an assimilatory APS reductase. For Tcs. roseopersicina strain M1 the presence of a PAPS reductase gene and a product dedicated to the reduction of PAPS has been shown biochemically (Haverkamp and Schwenn, 1999).

7.6. Reduction of Sulfite to Sulfide Sulfite is finally reduced to sulfide by an assimilatory sulfite reductase (EC 1.8.7.1). Again, different types can be differentiated: The NADPHdependent assimilatory sulfite reductases of enterobacteria are composed of two different subunits: a siroheme-[4Fe-4S]-containing protein (CysI) and a flavoprotein (CysJ) (Murphy et al., 1973; Ostrowski et al., 1989; Siegel et al., 1973). In contrast the ferredoxin-dependent sulfite reductases from cyanobacteria, algae, and higher plants are much simpler homooligomers of just a siroheme-[4Fe-4S]-containing protein. These enzymes are only very distantly related to their dissimilatory counterparts (Dhillon et al., 2005). The data presented in Table 5 indicate that the ferredoxin-dependent sulfite reductase is more common in purple bacteria than the NADPHdependent enzyme consisting of two different polypeptides. Tca. roseopersicina strain M1 is currently the only purple bacterium for which the existence of a cysIJ-encoded sulfite reductase has been firmly established (Haverkamp and Schwenn, 1999).

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 177

7.7. Incorporation of Sulfide The final step in cysteine biosynthesis is catalyzed by either O-acetylserine (thiol)-lyase A or O-acetylserine (thiol)-lyase B, encoded by the genes cysK and cysM, respectively (Kredich, 1992, 1996). The CysK and CysM proteins from E. coli are 43% identical. CysK synthesizes cysteine from O-acetylserine and sulfide, while the CysM protein differs in that it can utilize thiosulfate in addition to sulfide. The reaction between O-acetylserine and thiosulfate produces S-sulfocysteine, which is converted into cysteine by an as yet uncharacterized mechanism. On a biochemical level, cysteine and S-sulfocysteine biosynthesis in anoxygenic phototrophic bacteria were studied by Hensel and Tru¨per (1976, 1981). Both activities were found in several species of Chromatiaceae and Ectothiorhodospiraceae, and in Rhodocyclus tenuis and Rubrivivax gelatinosus, both purple non-sulfur bacteria belonging to the Betaproteobacteria. The tested alphaproteobacterial purple non-sulfur bacteria exhibited only O-acetylserine (thiol)-lyase A activity, that is, they did not catalyze reaction of O-acetylserine with thiosulfate. The two different O-acetylserine (thiol)-lyases were purified from Alc. vinosum (Hensel and Tru¨per, 1981). Sulfide for cysteine biosynthesis cannot only be obtained from the transport and reduction of inorganic sulfate as described but a number of bacteria can also derive it from organic sulfonate compounds such as taurine (van der Ploeg et al., 1996, 1998, 2001).

8. CONCLUSIONS The combination of available biochemical information and the survey of genome sequence data presented here allows some general conclusions to be made about the sulfur compound oxidation enzymes in GSB and PSB. Sulfide:quinone oxidoreductase (encoded by sqr) appears to be especially important for the oxidation of sulfide. Flavocytochrome c is less widespread. Sox genes (SoxXABYZ) occur in both GSB and PSB and are responsible for the oxidation of thiosulfate. In many cases, elemental sulfur appears as an intermediate of sulfide and thiosulfate oxidation. All thiosulfate-utilizing GSB strains have an identical sox gene cluster (soxJXYZAKBW). The soxCD genes found in certain other thiosulfate-utilizing organisms like Pcs. pantotrophus are absent from all investigated GSB and PSB. A putative complex denoted SoyYZ, related to the thiosulfate-binding SoxYZ complex, could be involved in the processing of an as yet unidentified sulfur compound in GSB (Frigaard and Bryant, 2008a).

178

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Siroamide-sulfite reductase (DsrAB) and proteins encoded by several other dsr genes are essential for the oxidation of sulfur formed as an intermediate during sulfide or thiosulfate oxidation. Most GSB and PSB utilizing sulfide or elemental sulfur contain the dissimilatory sulfite reductase dsrNCABLEFHTMKJOP genes. Although Chp. thalassium appears to have an (probably less efficient) alternative enzyme system for elemental sulfur oxidation, the dsr genes appear to be involved in elemental sulfur utilization in all other cases. Sulfite is the product of this process. The question of sulfite oxidation in purple bacteria will require special attention in the future, as the APS reductase pathway is restricted to only some members of the Chromatiaceae and Chlorobiaceae. It has even been shown that this pathway is not essential in Alc. vinosum. In addition, genes for a classical sulfite:acceptor oxidoreductase are present neither in GSB nor in PSB. Analyses indicate that, although the phylogenies of some enzymes (e.g., the DsrA protein) are congruent with the organismal phylogeny at the phylum level, the phylogenies of other enzymes are not (e.g., the SQR protein). Some enzyme systems that are only present in some strains exhibit a phylogeny incongruent with the cellular core phylogeny (e.g., the Sox and the Sat-Apr-Qmo systems). Thus, these systems appear to result from LGT rather than gene elimination. In the case of the sox gene cluster, evidence for phage-mediated LGT by a mobile genetic element was identified in GSB. As a final point, although the GSB are closely related, with the exception of Chp. thalassium, genomic analyses clearly show that gene elimination and LGT substantially influence the distribution of sulfur metabolism genes both within the GSB and among prokaryotes from other phyla. These observations illustrate the dynamic structures of prokaryotic genomes and in addition demonstrate that even organisms that superficially appear to be very closely related on the basis of their cellular core machinery nevertheless can have unexpected differences in physiology and life style. Assimilatory sulfate reduction occurs in many anoxygenic phototrophic bacteria. Two different types of sulfate transporters appear to be used. While sulfate is activated to adenosine-5u-phosphosulfate via a heterodimeric cysDN-encoded ATP sulfurylase in members of the Bradyrhizobiaceae, Acetobacteraceae, Rhodospirillaceae, Comamonadaceae, a few GSB, and Heliobacterium modesticaldum, the homo-oligomeric sat-encoded type of this enzyme occurs in Rhodobacteraceae and is also present in the PSB Alc. vinosum as well as in FAP bacteria. In most sulfate-assimilating anoxygenic phototrophic bacteria, APS appears to be immediately reduced without further phosphorylation to PAPS. The generated sulfite is reduced further to sulfide by assimilatory sulfite reductase. This enzyme appears to be ferredoxin-dependent in almost all cases, a flavoprotein subunit able to

SULFUR METABOLISM IN PHOTOTROPHIC SULFUR BACTERIA 179

use NADPH as an electron donor – as in E. coli – is not encoded in the sequenced genomes of anoxygenic phototrophic bacteria.

ACKNOWLEDGEMENTS Support by the Deutsche Forschungsgemeinschaft to C.D. is gratefully acknowledged. Birgitt Hu¨ttig provided excellent technical assistance. N.-U.F. gratefully acknowledges support from the Danish Natural Science Research Council (grant 21-04-0463). We thank Hans G. Tru¨per for the light and electron micrographs presented in Fig. 1.

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Visscher, P.T., Nijburg, J.W. and van Gemerden, H. (1990) Polysulfide utilization by Thiocapsa roseopersicina. Arch. Microbiol. 155, 75–81. Visscher, P.T. and van Gemerden, H. (1991) Photoautotrophic growth of Thiocapsa roseopersicina on dimethyl sulfide. FEMS Microbiol. Lett. 81, 247–250. Visser, J.M., de Jong, G.A.H., Robertson, L.A. and Kuenen, J.G. (1996) Purification and characterization of a periplasmic thiosulfate dehydrogenase from the obligately autotrophic Thiobacillus sp. W5. Arch. Microbiol. 166, 372–378. Vogl, K., Glaeser, J., Pfannes, K.R., Wanner, G. and Overmann, R. (2006) Chlorobium chlorochromatii sp. nov., a symbiotic green sulfur bacterium isolated from the phototrophic consortium ‘‘Chlorochromatium aggregatum’’. Arch. Microbiol. 185, 363–372. Vogler, K.G. and Umbreit, W.W. (1941) The necessity for direct contact in sulphur oxidation by Thiobacillus thiooxidans. Soil Sci. 51, 331–337. Ward, D.M., Ferris, M.J., Nold, S.C. and Bateson, M.M. (1998) A natural view of microbial biodiversity within hot spring cyanobacterial mat communities. Microbiol. Mol. Biol. Rev. 62, 1353–1370. Winogradsky, S.N. (1887) U¨ber Schwefelbakterien. Bot. Ztg. 45, 489–508. Wynen, A., Dahl, C. and Tru¨per, H.G. (Unpublished). Wynen, A., Tru¨per, H.G. and Dahl, C. (Unpublished). Yakimov, M.M., Guiliano, L., Chernikova, T.N., Gentile, G., Abraham, W.R., Lunsdorf, H., Timmis, K.N. and Golyshin, P.N. (2001) Alcalilimnicola halodurans gen. nov., sp. nov., an alkaliphilic, moderately halophilic and extremely halotolerant bacterium, isolated from sediments of soda-depositing Lake Natron, East Africa Rift Valley. Int. J. Syst. Evol. Microbiol. 51, 2133–2143. Yu, Z., Lansdon, E.B., Segel, I.H. and Fisher, A.J. (2006) Crystal structure of the bifunctional ATP sulfurylase-APS kinase from the chemolithotrophic thermophile Aquifex aeolicus. J. Mol. Biol. 365, 732–743. Yurkov, V.V. (2006) Aerobic phototrophic proteobacteria. In: The Prokaryotes (M. Dworkin, S. Falkow, E. Rosenberg, K.-H. Schleifer and E. Stackebrandt, eds), Vol. 5, pp. 562–584. Springer, New York. Yurkov, V.V., Krasil’nikova, E.N. and Gorlenko, V.M. (1994) Thiosulfate metabolism in the aerobic bacteriochlorophyll-a-containing bacteria Erythromicrobium hydrolyticum and Roseococcus thiosulfatophilus. Microbiology 63, 91–94. Zaar, A., Fuchs, G., Golecki, J.R. and Overmann, J. (2003) A new purple sulfur bacterium isolated from a littoral microbial mat, Thiorhodococcus drewsii sp. nov. Arch. Microbiol. 179, 174–183. Zuber, H. and Cogdell, R.J. (1995) Structure and organization of purple bacterial antenna complexes. In: Anoxygenic Photosynthetic Bacteria (R.E. Blankenship, M.T. Madigan and C.E. Bauer, eds), Advances in Photosynthesis, Vol. 2, pp. 315–348. Kluwer Academic Publishers, Dordrecht.

Carbon, Iron and Sulfur Metabolism in Acidophilic Micro-Organisms D. Barrie Johnson and Kevin B. Hallberg School of Biological Sciences, Bangor University, Bangor, U.K.

ABSTRACT Acidophilic micro-organisms are those (mostly prokaryotes) that grow optimally at pH o3 (extreme acidophiles) or at pH 3–5 (moderate acidophiles). Although once considered to comprise relatively few species of bacteria and archaea, the biodiversity of extreme acidophiles is now recognized as being extensive, both in terms of their physiologies and phylogenetic affiliations. Chemolithotrophy (the ability to use inorganic chemicals as electron donors) is widespread among extreme acidophiles, as ferrous iron and sulfur represent two major available energy sources in many natural and man-made extremely acidic environments. Dissimilatory reduction of iron and sulfur (as a consequence of their use as electron acceptors in oxygen-limited and anoxic environments) are also a major biogeochemical processes in low-pH environments. Acidophiles display considerable diversity in how they assimilate carbon; some are obligate autotrophs, others obligate heterotrophs, while a large number use either organic or inorganic carbon, depending on the availability of the former. This review describes the intricate relationships between carbon, iron and sulfur transformations by acidophilic micro-organisms, and how these are significant in both industrial and environmental contexts.

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Acidophilic micro-organisms: overview . . . . . . . . . . . . . . . . . . . . . . . 1.1. Definition, Habitats and General Physiological Characteristics . . . 1.2. Phylogenetic Diversity of Acidophilic Micro-Organisms . . . . . . . . 1.3. Physiological Diversity of Acidophiles . . . . . . . . . . . . . . . . . . . .

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2.

Carbon metabolism in acidophilic micro-organisms . . . . . . . . . . . . . . 2.1. Carbon Metabolism in Obligately Autotrophic Acidophiles . . . . . . 2.2. Carbon Metabolism in Obligately Heterotrophic Acidophiles . . . . 2.3. Carbon Metabolism in Facultatively Autotrophic Acidophiles . . . . 3. Iron metabolism by acidophilic micro-organisms . . . . . . . . . . . . . . . . 3.1. Dissimilatory Oxidation of Iron . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Dissimilatory Reduction of Iron . . . . . . . . . . . . . . . . . . . . . . . . . 4. Sulfur metabolism by acidophilic micro-organisms . . . . . . . . . . . . . . . 4.1. Dissimilatory Oxidation of Elemental Sulfur and Reduced Inorganic Sulfur Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Dissimilatory Reduction of Sulfur and Sulfate by Acidophilic Micro-Organisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Applied and ecological aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1. Acidophiles and Mineral Processing . . . . . . . . . . . . . . . . . . . . . 5.2. Acidophiles in Natural and Anthropogenic Environments . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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1. ACIDOPHILIC MICRO-ORGANISMS: OVERVIEW 1.1. Definition, Habitats and General Physiological Characteristics While there is no formal definition of what constitutes an acidophilic microorganism, there is a general consensus that optimal growth of an acidophile occurs at a pH that is significantly less than 7. Sub-division between ‘‘moderate’’ and ‘‘extreme’’ acidophiles, with the former having pH growth optima of 3–5 and the latter of o3, has been proposed (Johnson, 2007). Besides these obligate acidophiles, low-pH environments may contain metabolically active ‘‘acid tolerant’’ micro-organisms that have pH optima for growth close to neutral. This review focuses, in the main, on the physiologies of extreme acidophiles, though it also includes reference to some moderate acidophiles, particularly in the context of redox transformations of iron and sulfur. There are a number of recent reviews on the general subject of acidophilic microbiology. Some of these have described the biodiversity of these extremophiles (e.g. Johnson, 2007), while others have a more applied focus (e.g. Schippers, 2007). Acidic environments may occur naturally, for example in volcanic and geothermal areas, or arise due to human activities, the most important of which is mining of metals and coal. In both scenarios, some specialized

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prokaryotes play key roles in determining the geochemistry of these environments. While a number of biological processes, such as nitrification and fermentation, generate acidity, the dissimilatory oxidation of reduced sulfur, ultimately generating sulfuric acid, has the greatest impact on the global biosphere. Sulfur is the 16th most abundant element in the lithosphere (mean abundance 260 mg/g) and all life-forms have a nutritional requirement for this element. Some specialized prokaryotes, as described later in this review, can obtain some or all of their energy requirements from the oxidation of elemental (zero-valent) sulfur and/or various reduced inorganic sulfur compounds (RISCs, where sulfur can be present in a variety of oxidation states). While many of these are neutrophiles, many acidophilic prokaryotes are sulfur-oxidizers. Iron sulfides, chiefly pyrite (FeS2) but also others, such as pyrrhotite (FeS1x), constitute an important reservoir of reduced sulfur in the lithosphere. In these, and related sulfide minerals, iron is present in a reduced form (ferrous iron) and its oxidation to oxidized (ferric) iron is an exergonic reaction that some other acidophilic (and sometimes the same) prokaryotes are able to exploit. It is therefore the case that microbially mediated redox transformations of both sulfur and iron are both particularly important in acidic environments, and microorganisms involved in these processes, and how they mediate these transformations, have been the focus of much of the research carried out in this area of extremophilic microbiology. One curious fact is that chemoautotrophy was recognized as an important metabolic life-style among acidophiles long before phototrophy or heterotrophy. Many organic carbon-assimilating acidophiles are now known, including some that can switch to fixing carbon dioxide when supplies of organic carbon are depleted. These key aspects of acidophile physiologies – carbon metabolism and sulfur and iron transformations – are the three topics addressed in this review. Two physiological characteristics of acidophilic micro-organisms are vital in allowing them to survive in environments that are lethal to most lifeforms. The first is a remarkable ability to prevent ingress of protons (H3Oþ) even when presented with a high inward gradient; the external concentration may be 104–106 times greater than in the cell cytoplasm. These proton gradients are maintained even in resting cells. Acidophiles, like neutrophiles, make use of pH gradients to generate ATP using membrane-bound ATPases, though controlled influx of protons needs to be balanced by accessing electrons (derived, e.g., from the oxidation of ferrous iron to ferric). A downside of living in acidic liquors and possessing large transmembrane pH gradients results, however, in acidophiles being acutely sensitive to many small molecular weight aliphatic acids (such as acetic acid)

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that exist mainly as undissociated, lipophilic molecules at pH values typical of those which extreme acidophiles grow. A second major distinguishing feature between acidophilic and neutrophilic prokaryotes is that the former generate positive membrane potentials (Do), achieved by active influx of cations such as Kþ, whereas the latter possess negative potentials (Albers et al., 2001). Possession of positive membrane potentials confers a degree of protection against many positively charged ions (such as cationic transition metals, which tend to be present at elevated concentrations in many acidic environments), but conversely makes acidophiles more sensitive to many common anions, such as nitrate (Norris and Ingledew, 1992).

1.2. Phylogenetic Diversity of Acidophilic Micro-Organisms Complex, multicellular life-forms are generally absent from extremely lowpH environments, with the notable exception of some macrophytes (e.g. Juncus bulbosus and Eriophorum angustifilium) that have been observed to colonize acidic lakes and wetlands (Fyson, 2000). As with other extreme environments, biodiversity tends to decrease with increasing acidity, though other factors, such as concentrations of soluble (heavy) metals, which also tend to be greater in acidic liquors, may have an overriding influence. Eukaryotes, such as rotifers, protozoa and yeasts, are often found in extremely low-pH (o3) mesophilic environments, though these organisms are rarely observed in thermal acidic waters.

1.3. Physiological Diversity of Acidophiles Acidophiles may be categorized in a number of ways, for example in their pH optima and minima for growth (moderate or extreme acidophiles), and on the basis of their response to temperature. Three groups of acidophiles have often been recognized in this way: mesophilic acidophiles (temperature optima for growth of less than 40 1C), moderate thermo-acidophiles (temperature optima of 40–60 1C), and extreme thermo-acidophiles (temperature optima of W60 1C). Conveniently, most mesophilic acidophiles are Gram-negative bacteria, most moderate thermo-acidophiles are Grampositive bacteria, and extreme thermo-acidophiles are (with the exception of Hydrogenobaculum spp.) exclusively archaea. Figure 1 shows the distribution of acidophilic Gram-negative and Gram-positive bacteria, and of acidophilic archaea, on the basis of pH and temperature growth optima. While the general triphasic distribution just described is apparent, some

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Figure 1 Scatter plot of temperature optima versus pH optima for growth of acidophilic prokaryotes. Key: () archaea; (D) Gram-positive bacteria; (8) Gramnegative bacteria.

recently described acidophiles (such as mesophilic Ferroplasma spp. and some acidophilic Firmicutes) appear to be outside the general pattern. The data plotted in Fig. 1 show linear correlation between increasing temperature and pH optima for both the Gram-positive bacteria (r ¼ 0.75) and the archaea (r ¼ 0.67), but not for the Gram-negative bacteria. The maximum temperature at which growth of an extreme acidophile has been reported is 90 1C (the archaeon Acidianus infernus) which is about 30 1C lower than that observed by the most thermophilic Pyrolobus-like neutrophilic archaeon. Another way to distinguish acidophiles is on the basis of their nutritional characteristics (energy and carbon sources). Four groups may be recognized: (i) chemolithotrophic acidophiles–autotrophic prokaryotes that use an inorganic energy source; (ii) phototrophic acidophiles (micro-algae such as Euglena mutabilis) that use solar energy; (iii) heterotrophic acidophiles that use organic carbon as both carbon and energy source; (iv) mixotrophic acidophiles that use inorganic electron donors and assimilate organic carbon. Again this is, in reality, a rather simplistic approach, as many acidophiles do not display ‘‘black and white’’ metabolic life-styles. For example, a number of heterotrophic acidophiles can oxidize RISCs and appear to obtain energy from these reactions, though these bacteria (most

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Acidiphilium spp.) have no energetic or nutritional requirement for reduced sulfur compounds. Table 1 lists currently recognized species of acidophilic prokaryotes (including some species that have been described but not yet officially named) on the basis of their major physiological characteristics.

2. CARBON METABOLISM IN ACIDOPHILIC MICRO-ORGANISMS 2.1. Carbon Metabolism in Obligately Autotrophic Acidophiles As described above, a general classification of acidophilic prokaryotes may be based on their carbon metabolism. Obligately autotrophic acidophiles are those that obtain cellular carbon from an inorganic carbon source, carbon dioxide, in contrast to obligate heterotrophs that obtain carbon from organic sources and facultative autotrophs that can use both inorganic and organic carbon. In most acidic environments in the absence of light, autotrophic iron- and sulfur-oxidizing micro-organisms are the primary producers, fixing inorganic carbon and being the providers of organic materials used as growth substrates by heterotrophic acidophiles. At the low pH at which acidophiles grow, carbon dioxide is poorly soluble since it exists as gaseous CO2 rather than water-soluble bicarbonate (HCO 3 ), and often growth of autotrophic acidophiles is enhanced by supplementing air (used to supply oxygen) with carbon dioxide, especially at higher temperatures where the solubility of CO2 is further decreased. There are four currently accepted pathways of carbon dioxide fixation by prokaryotes, including the Calvin–Benson–Bassham cycle (‘‘CBB’’; Calvin, 1962), the reductive tricarboxylic acid cycle (Antranikian et al., 1982), the acetyl-CoA pathway (Pezacka and Wood, 1986) and the 3-hydroxypropionate cycle (Strauss and Fuchs, 1993). 2.1.1. Fixation of CO2 by Autotrophic Acidophiles The majority of acidophilic bacteria use the CBB cycle to obtain their cellular carbon. The key enzyme in this pathway is ribulose bisphosphate carboxylase/oxygenase (RUBISCO), which has been detected in both At. thiooxidans (Suzuki and Werkman, 1958) and At. ferrooxidans (Gale and Beck, 1967). While RUBISCO can be found in the cytoplasm of bacteria, it is often found packaged in organelles called carboxysomes (Shively et al., 1998). Carboxysomes, polyhedral inclusion bodies in the

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Table 1 Species of known acidophilic prokaryotic micro-organisms, on the basis of their major physiological characteristics Obligate autotrophs Iron-oxidizers Leptospirillum spp. – L. ferrooxidans – L. ferriphilum – ‘‘L. ferrodiazotrophum’’

Facultative autotrophs

Obligate heterotrophs

Acidimicrobium ferrooxidansa

Ferrimicrobium acidiphiluma

Ferrithrix thermotoleransa

‘‘Thiobacillus ferrooxidans’’ m-1

Ferroplasma spp.a – Fp. acidiphilumb – ‘‘Fp. acidarmanus’’

Sulfur-oxidizers Acidithiobacillus thiooxidans Acidithiobacillus caldusc Hydrogenobaculum acidophilum

Acidiphilium acidophiluma

Acidiphilium spp.a

Alicyclobacillus disulfidooxidans Metallosphaera spp.

Acidicaldus organivoransa

– M. sedula – M. prunae – M. hakonensis

Sulfurococcus spp. – Sc. yellowstonensis – Sc. mirabilis

Iron and sulfur-oxidizers Acidithiobacillus ferrooxidansa

Sulfobacillus spp.

Sulfolobus metallicus

Alicyclobacillus toleransa Thiomonas spp.

– – – – –

Sb. Sb. Sb. Sb. Sb.

Sulfolobus tokodaii

thermosulfidooxidans acidophilus thermotolerans sibiricus benefaciens

– Tm. cuprina – Tm. arsenivorans

Iron-reducers Acidocella spp. – Ac. facilis – Ac. aminolytica – ‘‘Ac. aromatica’’

Acidobacterium capsulatum (Continued )

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Table 1 (Continued ) Obligate autotrophs

Facultative autotrophs

Obligate heterotrophs

Sulfur-reducers Acidianus spp.

Acidianus brierleyid

Thermoplasma spp.

– Ad. infernus – Ad. ambivalens

Stygiolobus azoricus

– Tm. acidophilum – Tm. volcanium

Sulfurisphaera ohwakuensis

Others (no recorded iron/sulfur transformations) Acidisphaera rubrifaciens various Alicyclobacillus spp. (see Table 2) Sulfolobus spp. – S. acidocaldarius – S. solfataricus

Picrophilus spp. – P. oshimae – P. torridus a

Also shown to reduce ferric iron. Also reported to grow autotrophically. c Can incorporate yeast extract. d Inferred. b

cytoplasm of bacteria, are generally produced during CO2-limiting conditions, and are generally considered to enhance CO2-fixation by increasing the concentration of HCO 3 in the organelle where is it converted to CO2 by carbonic anhydrase in the immediate vicinity of RUBISCO (So et al., 2004). This high concentration of the RUBISCO substrate enhances the carboxylation reaction over the oxygenation reaction. Both of these acidithiobacilli have been observed to produce these organelles when grown under carbon-limiting conditions (Shively et al., 1998). Two different forms of RUBISCO are found in At. ferrooxidans, namely, form I (CbbL) and form II (CbbM). The former has a higher ability to catalyze carboxylation than oxidation, while CbbM has a relatively lower capacity. Gene cloning has shown that At. ferrooxidans contains duplicate, and identical, copies of genes coding for CbbL (Kusano et al., 1991), and hybridization with a fragment of the cbbM gene from Rhodospirillum rubrum confirmed that this gene is also in At. ferrooxidans (Shively et al., 1993). The sequence of the genome from the type strain (ATCC 23270) of At. ferrooxidans confirms the hybridization results. Recent genome sequence comparisons show that the three currently recognized Acidithiobacillus spp.

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(At. ferrooxidans, At. thiooxidans and At. caldus) all contain the necessary genetic information for fixation of CO2 by the CBB cycle (Valde´s et al., 2007). Other acidophilic autotrophic bacteria that have been shown to fix CO2 through the CBB cycle include Leptospirillum ferriphilum and Sulfobacillus acidophilus. Community metagenomics, sequencing of genomes extracted from a microbial community, of samples from acid mine drainage at Iron Mountain, California, have allowed the reconstruction of the genome of Leptospirillum ferriphilum and ‘‘Leptospirillum ferrodiazotrophum’’ (Tyson et al., 2004). Both of these acidophiles contain genes for RUBISCO and other CBB enzymes. Recently, the facultative autotroph Sulfobacillus thermosulfidooxidans has been shown to have RUBISCO activity when grown autotrophically (Caldwell et al., 2007). Those authors also cloned the genes coding for carbon dioxide fixation from Sulfobacillus acidophilus and found these bacteria also use form I of RUBISCO. Interestingly, it was found that RUBISCO activity of Sb. thermosulfidooxidans was similar to that of At. ferrooxidans, though the former could not grow as efficiently under autotrophic conditions as the latter. This was attributed, in part, to the lack of carboxysomes in Sb. thermosulfidooxidans, the genes for which were not detected in the genomic region near to the cbbL gene in Sb. acidophilus (Caldwell et al., 2007). Little is known about carbon dioxide fixation in another facultative autotroph Acidimicrobium ferrooxidans, though the gene coding for the large subunit of RUBISCO has been cloned from this acidophile and has been partially sequenced (P.R. Norris, Warwick University, U.K; personal communication).

2.1.2. Fixation of CO2 by Acidophilic Prokaryotes, Using Other Pathways In contrast to acidophilic bacteria, most thermophilic acidophilic archaea described are able to grow as heterotrophs or autotrophs. During growth with CO2 as carbon source, the thermo-acidophilic archaea use the 3-hydroxypropionate cycle for fixation of carbon dioxide. Evidence for this was first provided when the key enzymes acetyl-CoA carboxylase and propionyl-CoA carboxylase were detected in Acidianus brierleyi (Ishii et al., 1997). These activities were also found in Acidianus infernus, Metallosphaera sedula and Sulfolobus metallicus (Menendez et al., 1999). Subsequent work showed that a single enzyme catalyzed both of these activities in M. sedula, in a modified form of this CO2-fixing mechanism (Hu¨gler et al., 2003). These authors also showed that homologous genes coding for this enzyme can be

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found in other acidophilic archaea, including Sulfolobus solfataricus and Sulfolobus todakaii, but not in Thermoplasma acidophilum. It has been reported that the genome of Ferroplasma species in Iron Mountain acid mine drainage contain genes for formate hydrogenlyase, formate dehydrogenase and carbon monoxide dehydrogenase (Tyson et al., 2004), and it has been suggested that these enzymes may form part of the acetyl-CoA pathway for CO2 fixation (Golyshina and Timmis, 2005). This, however, is controversial as, aside from the original report describing Ferroplasma acidiphilum (Golyshina et al., 2000), no evidence has been provided for the fixation of CO2 or autotrophic growth of Ferroplasma isolates (Section 2.2.4). These enzymes are probably more likely to be involved in generation of energy from formate during organotrophic growth of Ferroplasma with formate as energy substrate. Although no direct evidence has been provided for the fixation of carbon dioxide in the thermo-acidophilic bacterium Hydrogenobaculum acidophilum, it probably uses the reductive TCA cycle for carbon assimilation. This is a reasonable assumption since representatives of the three different lineages of the phylum Aquificae use the reductive TCA cycle for CO2 assimilation in these bacteria (Hu¨gler et al., 2007). 2.1.3. Regulation of CO2-Fixing Enzymes Many acidophiles (facultative autotrophs; section 2.3) are capable of obtaining carbon in either inorganic or organic forms. From an energetic point of view, it is clearly beneficial to obtain cellular carbon from an organic source. What is unclear, however, is how the assimilation of carbon from organic or inorganic sources is regulated. In obligate autotrophs, little regulation of CO2-fixing enzymes would be expected, as this is the only source of carbon for cellular growth. It is clear that during growth in the absence of CO2-enhanced air, RUBISCO activities in At. ferrooxidans and Sulfobacillus thermosulfidooxidans are increased (Caldwell et al., 2007). Regulation of RUBISCO genes also appears to be substrate-dependent in At. ferrooxidans, where it was shown that during the switch from growth of At. ferrooxidans on iron to sulfur the expression of the CBB cycle genes was increased (Appia-Ayme et al., 2006). This probably reflects the need for more cellular carbon during growth on the more energetic substrate sulfur compared with iron. Apart from growth with iron, reduced inorganic sulfur and hydrogen, At. ferrooxidans has also been shown to grow with formic acid as electron donor (Pronk et al., 1991). The use of this organic acid might imply that this bacterium is heterotrophic, though this does not appear to be the case.

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At. ferrooxidans uses a formate dehydrogenase to oxidize formate to CO2, while reducing NAD to NAD(H). The CO2 was shown to be assimilated by At. ferrooxidans using RUBISCO, the activity of which was identical to rates of carbon assimilation during growth with formate (Pronk et al., 1991).

2.2. Carbon Metabolism in Obligately Heterotrophic Acidophiles It is something of an anomaly that obligately heterotrophic acidophilic prokaryotes were first isolated and characterized some 60 years after the first report of an acidophilic chemolitho-autotrophic acidophile (At. thiooxidans). The status of the earliest bacterial isolate (Flavobacterium acidurans; Millar, 1973) as a genuine acidophile is questionable, and there are no reports of similar bacteria being isolated from acidic environments. However, following the pioneering work of Arthur Harrison at the University of Missouri, a growing number of extremely and moderately acidophilic heterotrophic species have been characterized. However, while there are many published data on the types of organic substrates used by these prokaryotes, there has been relatively little research into the biochemistry of organic carbon metabolism in acidophilic micro-organisms. 2.2.1. Acidiphilium spp. Acidiphilium spp. are extreme acidophiles that are widely distributed in metal-rich, acidic environments. Species currently recognized fall into two phylogenetic groups on the basis of their 16S rRNA gene sequences. The first includes the type species Acidiphilium cryptum, as well as A. organovorum and A. multivorum, and the second includes A. angustum and A. rubrum (though these are probably the same species, since they have W99% 16S rRNA gene sequence homology and share 100% DNA:DNA homology; Kishimoto et al., 1995), and A. acidophilum. All of these are obligate heterotrophs, with the exception of A. acidophilum, which is considered in Section 2.3.1. A. cryptum was so called as it was found to grow as a contaminant of chemo-autotrophic acidophiles where it apparently grows on the small amounts of organic carbon originating from the iron/sulfur-oxidizers. It was originally considered to be an obligate oligotroph (early reports claimed that growth was inhibited by 0.1%, or 5.5 mM, glucose). However, all bacteria in the A. cryptum cluster can grow to high cell densities in media containing

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W10 mM glucose, though addition of a growth supplement, such as yeast extract, may be necessary to achieve this. The A. cryptum/A. organovorum/ A. multivorum group can grow on a wide range of monosaccharides, as well as dicarboxylic acids (such as malic acid) and tricarboxylic acids (such as citric acid). In contrast, monocarboxylic aliphatic acids, such as acetic and pyruvic acids, tend to inhibit growth of these (and most other) acidophiles when present in micro-molar concentrations, which is probably due to the fact that they occur as lipophilic, non-dissociated molecules in extremely acidic liquors. A. multivorum was shown, in contrast to both A. cryptum and A. organovorum, to grow on small molecular weight alcohols (methanol, ethanol and propanol; Wakao et al., 1994). Interestingly, A. rubrum and A. angustum were reported not to grow (or show poor growth) on monosaccharides, though they can utilize some organic acids (such as citric acid) and glycerol (Wichlacz et al., 1986). Acidiphilium spp. appear not to utilize polymeric organic substrates such as starch or disaccharides such as maltose, lactose or cellobiose (Lobos et al., 1986; Wakao et al., 1994). Although aerobic growth on substrates such as glucose has been reported to result in acidification of the growth media, analysis of cultures has failed to detect organic acids as metabolic by-products (Johnson et al., unpublished data), and acidification of growth media appears to be due to uptake of ammonium by growing cultures. Acidiphilium spp. were originally described as obligate aerobes and, although no strain has been shown to ferment organic substrates, all known species can grow by dissimilatory reduction of ferric iron under conditions of oxygen limitation (Section 3.2). Pathways of glucose catabolism in the type strain of A. cryptum and five other acidophilic heterotrophs were studied by Shuttleworth et al. (1985), who concluded that both the pentose phosphate and Entner–Doudoroff pathways were functional in the isolates, but that the Embden–Meyerhof– Parnas pathway was apparently absent. 2.2.2. Other Alpha-Proteobacteria Two bacteria, originally named Acidiphilium facilis and Acidiphilium aminolytica, were transferred to the new genus Acidocella on the basis of 16S rRNA gene sequence analysis (Kishimoto et al., 1995). A third species (‘‘Ac. aromatica’’) has been described, though not officially named. Acidocella species tend to grow more rapidly than Acidiphilium spp. but are not so acidophilic. Both Ac. facilis and Ac. aminolytica can grow on a wide range of monosaccharides, and also on amino acids such as glutamate. In contrast, ‘‘Ac. aromatica’’ cannot grow on substrates, such as glucose, glycerol and yeast extract, that are commonly used to cultivate acidophilic

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heterotrophs. It does, however, grow of fructose and on a wide range of aromatic compounds, including benzoic acid, phenol and naphthalene (Hallberg et al., 1999). In addition, strains of ‘‘Ac. aromatica’’ have been shown to grow on ethanol, propanol and organic acids, including acetic acid (at up to 5 mM) and hexadecanoic acid (Gemmell and Knowles, 2000). Other recognized genera of acidophilic heterotrophic alpha-proteobacteria include Acidomonas, Acidisphaera and Acidicaldus. Acidomonas methanolicus (a moderate acidophile) was so-named because it grew on methanol (as well as other substrates, including pectin) though this trait had previously been reported for A. multivorum and later shown to be widespread amongst all Acidiphilium spp. (Hallberg and Johnson, 2001). Acidisphaera rubrifaciens is, in contrast, a moderate acidophile that grows well on organic acids including pyruvic acid (though growth was tested at pH 4.5, which is above the pKa of this organic acid (2.5) so that the acid would be predominantly present in its relatively non-toxic dissociated state) and only sporadically on monosaccharides (Hiraishi et al., 2000). As. rubrifaciens is the most closely related cultivated bacterium to the recently described species Acidicaldus organivorans, though the latter is a moderately thermophilic (growing at up to 65 1C) extreme acidophile. Acd. organivorans is also interesting in terms of its nutrition in that, like ‘‘Ac. aromatica’’ it can grow on phenol as sole carbon source, as well as the commonplace substrates such as various monosaccharides and glycerol (Johnson et al., 2006a). 2.2.3. Gram-Positive Acidophiles Obligately heterotrophic Gram-positive acidophilic bacteria can be divided into two groups – Firmicutes (endospore-forming bacteria that contain relatively low ratios of guanine and cytosine in their chromosomal DNA) and Actinobacteria. Thus far, all Firmicutes in this category belong to the genus Alicyclobacillus (which also contains some facultatively autotrophic species, as described below) and are all moderately thermophilic or thermotolerant acidophiles. Alicyclobacillus spp. were first isolated (and classified as Bacillus spp.) from thermal, acidic soils and later also found to be widespread contaminants of pasteurized fruit juices (Darland and Brock, 1971; Goto et al., 2007). Currently, 17 species of Alicyclobacillus are recognized (Table 2), two of which (Alb. tolerans and Alb. disulfidooxidans) are able to utilize inorganic electron donors. These Firmicutes vary in the organic substrates that they utilize, though monosaccharides and disaccharides are commonly used (Table 2). Some species are also able to hydrolyze polysaccharides, such as starch and glycogen. Production of acid, resulting

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Table 2 Physiological characteristics of known Alicyclobacillus spp Alb. acidocaldariusa

Original site of isolation o-fatty acids Growth at pH o3 Inorganic electron donor(s) Growth on: glycerol Glucose mannitol Sucrose Lactose Starch

Alb. acidoterrestrisb

Alb. cycloheptanicusc

Alb. disulfidooxidansd

Alb. Alb. hesperidume herbariusf

Alb. Alb. acidiphilusg sendaiensish

Acidic thermal soil þ þ þ þ

þ 

Waste-water sludge þ þ

Solfataric soil þ þ

Herbal tea þ 

Acidic beverage þ þ

þ þ







þ









þ þ þ þ þ þ

þ þ  þ þ 

 þ þ   

þ þ  þ NR þ

þ þ þ þ þ 

þ þ þ þ þ 

 þ  þ þ 

þ þ NR þ þ NR

NR, not reported. a Darland and Brock (1971). b Deinhard et al. (1987a). c Deinhard et al. (1987b). d Dufresne et al. (1996). e Albuquerque et al. (2000). f Goto et al. (2002). g Matsubara et al. (2002). h Tsuruoka et al. (2003). i Goto et al. (2003). j Simbahan et al. (2004). k Karavaiko et al. (2005). l Goto et al. (2007).

Soil

CARBON, IRON AND SULFUR METABOLISM

Alb. pomorumi

215

Alb. vulcanalisj

Alb. toleransk

Alb. contaminansl

Alb. fastidiosusl

Alb. kakegawensisl

Alb. macrosporangiidusl

þ þ 

Geothermal pool þ þ þ

Leadzinc ore   

Fruit juice

  

þ () 

Apple juice þ  

Agricultural soil   

þ þ þ þ  

NR þ þ þ  NR

(þ) þ þ þ NR þ

þ þ þ þ (þ) 

 þ þ   

 þ þ þ þ 

þ þ 

Fruit juice

þ

þ 

Alb. saccharil

Alb. shizuokensisl

þ () 

Agricultural soil þ  

þ þ þ þ þ þ

 þ þ þ  

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D. BARRIE JOHNSON AND KEVIN B. HALLBERG

from the catabolism of organic substrates, has been widely reported for Alicyclobacillus spp., though there are no reports of anaerobic growth of these bacteria via fermentation (Chang and Kang, 2004). Recently, two novel genera of obligately heterotrophic Actinobacteria, Ferrimicrobium and Ferrithrix have been described (Johnson et al., 2008). These are ‘‘mixotrophs’’ in that they can obtain energy from oxidizing an inorganic electron donor (ferrous iron) but, in contrast to an earlier isolate, Acidimicrobium ferrooxidans, to which they are closely related, neither Ferrimicrobium nor Ferrithrix can assimilate inorganic carbon. Both bacteria can utilize yeast extract as a carbon (and energy) source, though their abilities to use single, defined organic compounds appears to be limited. Neither Ferrimicrobium acidiphilum nor Ferrithrix thermotolerans appear to utilize monosaccharides; growth of the former is enhanced by citric acid, glutamic acid and glycerol, but Fx. thermotolerans appears to use only a limited range of small molecular weight alcohols (ethanol and glycerol). 2.2.4. Archaea Three genera of extremely acidophilic euryarchaeotes are currently recognized: Thermoplasma, Picrophilus and Ferroplasma. There are some similarities between these archaea, such as the absence of cell walls in Thermoplasma and Ferroplasma spp. and the requirement of a complex organic carbon source (such as yeast extract) to support growth. Thermoplasma acidophilum and Tp. volcanium are obligately heterotrophic, facultative anaerobes (Segerer et al., 1988). Growth occurs in yeast-extractcontaining media, and is enhanced by the inclusion of some defined organic substrates such as glucose and fructose, though these archaea are reported not to grow in yeast-extract-free media. Picrophilus oshimae and P. torridus contrast with Thermoplasma spp. in being obligate aerobes, though their requirement for yeast extract and enhanced growth yields when media are supplemented with sugars (including the disaccharides, lactose and sucrose) mimics the behavior of Thermoplasma spp. (Schleper et al., 1995). Supplementing sugar-containing growth media with vitamins or Casamino Acids does not result in growth of Picrophilus spp., indicating a potentially complex growth factor requirement in these archaea. The situation regarding carbon metabolism in the iron-oxidizing acidophilic archaeon Ferroplasma is somewhat more contentious. Ferroplasma acidiphilum was described initially as an obligate chemoautotroph, though it was noted that addition of yeast extract was necessary for the growth of the original (and type) strain (Golyshina et al., 2000). Dopson et al. (2004) failed to detect uptake of 14CO2 by three strains of Fp. acidiphilum, or by the more acid-tolerant species

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‘‘Ferroplasma acidarmanus.’’ Marginal, but statistically significant, enhanced growth yields were obtained when some monosaccharides were added to growth media, and ‘‘Fp. acidarmanus’’ appeared to be more heterotrophically inclined than the strains of Fp. acidiphilum that were examined. Several acidophilic crenarchaeotes appear to be obligately heterotrophic. These include the first-documented acidophilic archaeon, Sulfolobus acidocaldarius (Brock et al., 1972), and Sulfolobus solfataricus (Zillig et al., 1980), both of which can grow on complex organic substrates such as yeast extract and on a range of mono- and disaccharides. S. solfataricus has also been reported to grow on phenol (Izzo et al., 2005). Sulfurisphaera ohwakuensis is also a heterotroph and grows on proteinaceous and complex substrates such as yeast extract or tryptone, but not on amino acids or monosaccharides (Kurosawa et al., 1998). The situation with Sulfolobus tokodaii is somewhat less clear. Although described as a facultative chemoheterotroph by Suzuki et al. (2002), the type strain required several months to adapt to sulfur-containing medium (which also contained organic carbon) and only about 20% of cell carbon appeared to originate from CO2 under such conditions. This archaeon again grows successfully on complex organic carbon, such as yeast extract, and there is some evidence that it can oxidize ferrous iron as well as sulfur. The situation with S. yangmingensis is also rather unclear; there is only tentative evidence that this prokaryote can grow as a chemolithotroph as well as by metabolizing organic substrates (Jan et al., 1999). A unique phenotype (for acidophilic archaea) is exhibited by Acidilobus aceticus, which grows anaerobically on starch, forming acetate as the main metabolic product (Prokofeva et al., 2000).

2.3. Carbon Metabolism in Facultatively Autotrophic Acidophiles There are a number of characterized acidophilic bacteria and archaea that can fix carbon dioxide, though revert to using organic carbon when this is available. Amongst acidophilic bacteria, this trait appears to be more widespread in Gram-positives than in Gram-negatives. 2.3.1. Acidiphilium acidophilum Prior to the advent of gene sequence analysis, A. acidophilum was named Thiobacillus acidophilus on account of the fact that it was an acidophile that could grow autotrophically and catalyze the dissimilatory oxidation of sulfur though, in contrast to other bacteria known at the time, A. acidophilum was shown to grow on organic substrates (Guay and Silver, 1975). Small

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molecular weight organic compounds, including a variety of monosaccharides (such as glucose and fructose), aliphatic acids (citric and malic) and amino acids (aspartic and glutamic acids) can support the growth of this acidophile. Mason and Kelly (1988) showed that, when A. acidophilum was grown on batch or continuous culture in the presence of both glucose and tetrathionate, it obtained energy from the simultaneous oxidation of both compounds and it obtained carbon from both glucose and carbon dioxide. There were indications that, when grown mixotrophically, A. acidophilum assimilated glucose preferentially, at the expense of tetrathionate, allowing an energy saving that resulted in enhanced growth yields. 2.3.2. Gram-Positive Acidophiles Sulfobacillus spp. are acidophilic iron- and sulfur-oxidizing Firmicutes, and most known species are moderate thermophiles. The first species to be described, Sb . thermosulfidooxidans, displayed an absolute requirement for an inorganic electron donor (such as ferrous iron) to support growth on defined organic substrates, such as glucose (Wood and Kelly, 1983). Similar results were reported for Sb. acidophilus (strain ALV; Wood and Kelly, 1984). Even when glucose was present, about 20% of cell carbon was estimated to have been obtained by fixation of CO2. Later, however, Norris et al. (1996) reported that the type strain of Sb. acidophilus (NAL, isolated from the same site as ALV) could be serially subcultured in a medium containing yeast extract and no additional ferrous iron or sulfur, the inference being that this strain is capable of heterotrophic, as well as autotrophic or mixotrophic, growth. Wood and Kelly (1984) showed that glucose was oxidized by the oxidative pentose phosphate cycle by Sb. acidophilus, with further oxidation of glyceraldehyde 3-phosphate via the tricarboxylic acid cycle. In contrast, Zakharchuk et al. (1994) concluded from studies of enzymes involved in glucose metabolism that, in Sb. thermosulfidooxidans, sugars were metabolized via the Embden–Meyerhof–Parnas and/or the Entner–Doudoroff pathway, and that the tricarboxylic acid cycle and the glyoxylate bypass were inoperative. More recently described Sulfobacillus spp. (e.g. Sb. sibiricus and Sb. thermotolerans) display similar mixotrophic metabolisms to Sb. thermosulfidooxidans. As noted above, some Alicyclobacillus spp. have also been shown to be facultative autotrophs, though these are more ‘‘heterotrophically inclined’’ than Sulfobacillus spp. and can readily be grown to high cell densities (W109/ml) in media containing suitable organic substrates. In one such strain (isolate GSM), metabolism of fructose and oxidation of ferrous were found to be uncoupled (Yahya et al., 2008). Cultures pre-adapted to

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heterotrophic growth were found to metabolize fructose immediately but there was a considerable lag period before iron oxidation was evident, while the reverse was the case when cells were harvested during growth phases where iron was being actively oxidized. Acidimicrobium ferrooxidans was originally, erroneously, considered to be a heterotrophic iron-oxidizer, like the related species Fm. acidiphilum and Fx. thermotolerans. Clark and Norris (1996) confirmed that it is able to fix CO2 though, like mixotrophic Alicyclobacillus isolates, it can be readily cultivated as a heterotroph. Yeast extract can serve as a suitable carbon and energy source for Am. ferrooxidans, but there is little published data on what defined carbon compounds are utilized by this acidophile and the pathways by which they are metabolized. 2.3.3. Archaea Many thermo-acidophilic archaea (all members of the Sulfolobaceae) are facultative autotrophs. These include Sulfolobus spp. (S. shibatae and ‘‘S. tengchongensis’’), Metallosphaera spp. (M. sedula, M. prunae and M. hakonensis), Sulfurococcus spp. (S. mirabilis and S. yellowstonensis) and Acidianus brierleyi (Garrity et al., 2001). With most of these prokaryotes, complex organic substrates are used to support heterotrophic growth, though S. shibatae can also metabolize mono- and disaccharides, starch and tryptone (Grogan et al., 1991), while S. tengchongensis grows on some sugars and glutamic acid (Xiang et al., 2003).

3. IRON METABOLISM BY ACIDOPHILIC MICRO-ORGANISMS 3.1. Dissimilatory Oxidation of Iron Due to the widespread occurrence of pyrite and other iron sulfides, iron is often present at elevated concentrations in the low pH of typical acidic environments, where most (cationic) metals are highly soluble. In addition to being highly soluble, ferrous iron is only very slowly oxidized by molecular oxygen at pHo4 (Stumm and Morgan, 1981), so iron-oxidizing acidophiles do not have to compete with chemical oxidation of their growth substrate. Because the free energy associated with the oxidation of iron is rather small (only one mole of electrons is released per mole Fe2þ oxidized) and the Fe2þ/Fe3þ couple redox potential is close to that of oxygen/water couple at low pH ðE0o þ 770 mV and þ 820 mV; respectivelyÞ, growth yields on ferrous iron are inevitably small. Nevertheless, the abundance of ferrous

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D. BARRIE JOHNSON AND KEVIN B. HALLBERG

iron means that it is an important electron donor in many acidic environments. Another important fact is that, due to the small difference in redox couples, only molecular oxygen can serve as electron acceptor for acidophilic micro-organisms that respire on ferrous iron. 3.1.1. Oxidation of Iron by Autotrophic Acidophiles Although a variety of acidophiles are known to catalyze the oxidation of ferrous iron to ferric (Table 1), only few obligately autotrophic iron-oxidizing acidophiles are known to do so. These are all Gram-negative bacteria, and include the specialist iron-oxidizers such as Leptospirillum ferrooxidans, L. ferriphilum and the newly isolated ‘‘L. ferrodiazotrophum’’. These acidophiles are considered to be specialists in that to date they are only known to grow by oxidation of iron. Another iron-oxidizing autotrophic acidophile At. ferrooxidans is the most widely studied of all acidophiles. Also an autotrophic iron-oxidizer, this bacterium is more versatile than the leptospirilli in that it can also oxidize RISCs, hydrogen and formate, and it can also grow anaerobically using ferric iron as an electron acceptor. Although these acidophiles can, in theory, be found co-habiting in acidic environments, significant physiological differences between them allows for the dominance of one over the other. At. ferrooxidans has higher specific iron oxidation activity than L. ferrooxidans at low redox potentials (e.g. high Fe2þ/Fe3þ ratios), but cannot oxidize iron above a redox potential of þ840 mV (Rawlings et al., 1999). Additionally, bacteria of the genus Leptospirillum are known to have higher affinities for Fe2þ than At. ferrooxidans (Km ¼ 0.25 mM and 1.34 mM, respectively) and the former are less inhibited by Fe3þ than the latter (Norris et al., 1988). For these reasons, bacteria of the genus Leptospirillum tend to dominate systems where Fe3þ concentrations are high, for example well-oxygenated acidic solutions, while At. ferrooxidans dominates acidic solutions where Fe2þ predominates (Schrenk et al., 1998; Walton and Johnson, 1992). Similar comparative data for a variety of acidophilic iron-oxidizing prokaryotes have been summarized, showing that these micro-organisms have a wide affinity for ferrous iron and varying sensitivities to ferric iron (Norris, 2007). 3.1.2. Oxidation of Iron by Heterotrophic Acidophiles In contrast to the few autotrophic iron-oxidizing acidophiles, many heterotrophic acidophiles are also capable of oxidizing iron (Table 1). Some of these are not obligately heterotrophic, but can assimilate cellular carbon

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from either CO2 or organic sources (Section 2.3). Obligately heterotrophic iron-oxidizing acidophiles are capable of growth and oxidation of iron only when supplied with an organic source of carbon. Generally, oxidation of iron by heterotrophic iron oxidizers is coupled to growth and organic carbon assimilation (Clark and Norris, 1996; Dopson et al., 2005; Johnson et al., 2008). Growth yields of these acidophilic heterotrophs are greater in the presence of iron, indicating that the micro-organisms are obtaining energy from the oxidation. Care must be taken, though, when determining this as the energy yield from iron is much lower than that obtained from organic carbon sources, and so media must be used that contains a minimal amount of organic carbon (e.g. Johnson et al., 2008). Alternatively, chemostat cultures have been used to determine growth yields of Ferroplasma spp. in the presence or absence of iron, where increased biomass was found in all cultures supplemented with ferrous iron (Dopson et al., 2004). 3.1.3. Energy Acquisition during Iron Oxidation During the oxidation of ferrous iron to ferric, electrons are used to reduce oxygen and protons to water. The consumption of protons allows for the transfer of protons into the cytoplasm through the ATPase as ATP is synthesized. Vesicles prepared from iron-grown At. ferrooxidans contain such an ATPase, which could synthesize ATP from ADP and Pi in response to an artificially induced proton gradient (Apel et al., 1980). It was proposed through the use of this proton-translocating ATPase that At. ferrooxidans gains energy from the oxidation of ferrous iron. Not all electrons, however, are used for the reduction of oxygen in autotrophic iron-oxidizers. Carbon dioxide fixation and other cellular metabolic functions require NAD(P)H. This is produced in the cell by reduction of NAD(P) by electrons obtained during growth. In the case of ferrous iron, where the redox potential is more positive than that of the NAD(P)/NAD(P)H couple, electrons must be transported ‘‘uphill’’ by the electron transport chain to NAD(P). It has been hypothesized that this occurs by reverse electron transport through the cytochrome bc1 complex, the quinone pool and the NAD(P)H dehydrogenase, and is energized by the proton-motive force generated by the hydrolysis of ATP (Ingledew, 1982). For this reason, growth yields of autotrophic bacteria that use ferrous iron as sole energy source are invariably small. The characterization of the electron transport chain of iron-grown At. ferrooxidans provided evidence for the existence of a bc1 complex (Elbehti et al., 1999). Further studies indicated that electrons could be transported from an exogenously added ferrocytochrome to NADþ in

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spheroplasts of iron-grown At. ferrooxidans (Elbehti et al., 2000). Through the use of metabolic inhibitors, it was found that this electron transport occurred via the bc1 complex, and the electron transport was accelerated by addition of ATP. It was hypothesized that the hydrolysis of ATP by the ATP synthase provided the proton motive force necessary for the uphill electron transport, and that the direction of electron flow (e.g. to oxygen or uphill) was regulated by the ratio of ATP/ADP in the cell (Elbehti et al., 2000). While no similar direct evidence for the uphill electron transport has been provided in other iron-oxidizing autotrophs, genes coding for similar uphill electron transport chain components have been detected in the reconstructed genomes of Leptospirillum species found in Iron Mountain (Tyson et al., 2004) and were found to be expressed in situ (Ram et al., 2005). Aside from the enhanced growth yields of heterotrophic acidophiles in the presence of iron oxidation, nothing further is known about how these microorganisms obtain energy from the oxidation of iron. 3.1.4. Enzymology of Iron Oxidation The enzymology of iron oxidation is best understood in At. ferrooxidans. For many years, it has been known that the blue copper protein rusticyanin is involved in transport of electrons from ferrous iron to oxygen (Cobley and Cox, 1983). This is an acid-stabile protein that is found in the periplasm of At. ferrooxidans, constituting up to 5% of the total protein of the cell. Initially, it was thought that rusticyanin was the electron acceptor from ferrous iron because it is reduced when cells are exposed to ferrous iron, and because ferrous iron can reduce purified rusticyanin. However, biochemical evidence showed that the rate at which electrons are transferred from ferrous iron to rusticyanin is too slow to account for oxidation rates of intact cells (Blake and Shute, 1987). Further work identified an acid-stable cytochrome c that was able catalyze the ferrous iron-dependent reduction of rusticyanin, suggesting that this molecule is the actual electron acceptor from ferrous iron (Blake and Shute, 1994). Cloning and sequencing of the DNA, around the region of the At. ferrooxidans chromosome where the rusticyanin gene can be found, revealed an operon coding for other electron transport proteins (Appia-Ayme et al., 1999). These include a cytochrome c and a cytochrome c4, as well as proteins that form a cytochrome oxidase. It was postulated that either of these could be the actual electron acceptor from ferrous iron. Purification of the cytochrome c4 and biochemical characterization of this protein showed that it could interact with rusticyanin, hinting that it is this cytochrome that accepts electrons from ferrous iron and transfers them to

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rusticyanin (Giudici-Orticoni et al., 2000). Further work, however, has shown that cytochrome c (called Cyc2) is actually a membrane-bound protein that is most likely to be the oxidant of ferrous iron (Yarzabal et al., 2002). These data, coupled with expression studies, where At. ferrooxidans is shifted from growth with sulfur as electron donor to ferrous iron, have led to the creation of a model pathway for electron transport from ferrous iron to oxygen (Quatrini et al., 2006). This model also incorporates the path of electrons from ferrous iron to NAD(P) (Fig. 2). Interestingly, it has been proposed that this electron transport chain (or at least a portion of it) operates during anaerobic growth of At. ferrooxidans with either sulfur or formate as the electron donor (Pronk et al., 1992), since bacteria grown under either condition exhibit nearly identical levels of iron oxidation activity as do iron-grown cells. While the enzymology of iron oxidation is best understood in the acidophile At. ferrooxidans, information on iron-oxidation enzymes in other acidophiles is relatively scarce. It has been known for some time that other iron-oxidizing acidophiles, including some strains of At. ferrooxidans, L. ferrooxidans, ‘‘ Thiobacillus ferrooxidans’’ m-1 and the archaea Sulfolobus metallicus and M. sedula, all contain redox-active proteins that differ from each other in spectral Fe2+ OM

Fe3+ Cyc 2

Rus Periplasm

IM

Cyc c4

Cyc c4

cytochrome oxidase

bc1 complex

2 H+ + ½ O2

H2 O

UQ

NADH dehydrogenase NAD(P) + H+

NAD(P)H

Cytoplasm

Figure 2 Model of electron transport (broken arrows) in Acidithiobacillus ferrooxidans showing the exergonic path to reduction of oxygen and the ‘‘uphill’’ path to the reduction of NAD(P) via ubiquinone (UQ). Key: OM, outer membrane; IM, inner membrane; Rus, rusticyanin; Cyc, cyctochrome c.

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characteristics (Barr et al., 1990; Blake et al., 1992, 1993). With the availability of genome sequence data and some experimental evidence, different pathways of electron transport from iron to oxygen have been proposed. Up-regulation of proteins in ‘‘Ferroplasma acidarmanus’’ strain Fer1 in yeast extract- and ferrous-iron-containing medium, coupled with metabolic inhibitors and spectroscopic data, have led to the development of a model of electron flow in this acidophilic archaeon (Dopson et al., 2005). In this model, electrons are passed from ferrous iron to a blue copper protein (sulfocyanin), which transfers the electrons to a cbb3-type terminal oxidase. The shorter electron transport chain in this archaeon compared with At. ferrooxidans could be related to the lack of a periplasm in the archaeon, allowing the cell membrane to contact iron directly. Differential expression of genes during growth of the thermophilic archaeon Sulfolobus metallicus on iron or sulfur has led to the identification of several proteins that are potentially involved in iron oxidation (Bathe and Norris, 2007). These genes, called fox genes, code for electron transport proteins including a cytochrome b and two subunits of a cytochrome c oxidase. Homologs of these genes can be found on the chromosomes of Sulfolobus tokodaii, Acidianus ambivalens and Metallosphaera sedula. Expression of these genes was high when S. metallicus was grown in a liquid medium with both ferrous iron and pyrite as electron donors, as opposed to sulfur as electron source. It appears, therefore, that thermoacidophilic archaea use a different set of proteins for the transport of electrons than the mesophilic archaeon, Ferroplasma.

3.2. Dissimilatory Reduction of Iron 3.2.1. Autotrophic Acidophiles The first report of the dissimilatory reduction of iron by acidophilic bacteria was by Brock and Gustafson (1976) who showed that some Acidithiobacillus spp. and the archaeon Sulfolobus acidocaldarius could reduce ferric iron to ferrous in anaerobic cultures. It was not ascertained at the time whether these prokaryotes were able to use ferric iron respiration as an energytransducing process. At. thiooxidans was also noted to reduce ferric iron to ferrous under aerobic conditions, suggesting that ferric iron and oxygen could be co-respired. Sand (1989) showed that At. ferrooxidans accumulated significant concentrations of ferrous iron when grown aerobically in low-pH (o1.3) liquid media containing both elemental sulfur and ferric iron. At higher pH values, iron oxidation (which was inhibited at very low pH)

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masked any accumulation of ferrous iron. Pronk and co-workers were the first to confirm that At. ferrooxidans could indeed grow by ferric iron respiration under strictly anaerobic conditions, using elemental sulfur as electron donor (Pronk et al., 1992). Ohmura et al. (2002) later showed that hydrogen, which is used as electron donor for oxygen-coupled respiration, could also be coupled to ferric iron reduction to support growth of this acidophile under anoxic conditions. The fact that At. ferrooxidans, the most well-studied of all acidophiles, is a facultative anaerobe is frequently overlooked, but does help explain its frequently reported dominance in anoxic mine waters (e.g. Coupland and Johnson, 2004; Rowe et al., 2007). The situation with other autotrophic acidophilic bacteria appears to be less clear, however. Hallberg et al. (2001) compared growth of At. ferrooxidans, At. thiooxidans and the moderate thermophile At. caldus in media containing sulfur or tetrathionate as electron donor and ferric iron, incubated under aerobic and anaerobic conditions. While growth of all three acidophiles occurred in aerobically incubated cultures, only At. ferrooxidans grew in anaerobic cultures. Ferric iron reduction was observed only in anaerobic cultures of At. ferrooxidans and cell suspensions of At. thiooxidans maintained under aerobic conditions. It appears therefore that, of currently classified Acidithiobacillus spp., only At. ferrooxidans can grow via ferric iron respiration. 3.2.2. Heterotrophic Acidophiles Johnson and McGinness (1991) screened 50 mesophilic and five moderately thermophilic strains of acidophilic heterotrophic bacteria for their abilities to reduce ferric iron in liquid and on solid media, incubated under aerobic conditions. About 40% of mesophilic isolates displayed variable capacities to reduce iron under such conditions, though none of the moderate thermophiles appeared to have this ability. Mixed cultures of iron-oxidizers (At. ferrooxidans or L. ferrooxidans) and a heterotrophic isolate that was particularly adept at reducing iron in aerobic media (Acidiphilium SJH) produced sequences of iron cycling in liquid media containing ferrous iron and glucose, presumably due to fluctuations in dissolved oxygen concentrations in these cultures. It was noted that both growth and ferric iron reduction by the mesophilic heterotrophs were very limited in cultures incubated under strictly anoxic conditions. However, Ku¨sel et al. (1999) isolated a strain (JF-5) of Acidiphilium cryptum from pit lake sediment that could grow by ferric iron respiration under anoxic conditions using a range of sugars, some other small molecular weight organic compounds, or hydrogen, as electron donors. Co-respiration of ferric iron and oxygen was

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later demonstrated in A. cryptum JF-5 (Ku¨sel et al., 2002). Further research with Acidiphilium SJH showed that it was able to accelerate the reductive dissolution of a number of ferric iron minerals (akageneite, goethite, jarosite, natrojarosite and amorphous ferric hydroxide) though hematite was not reduced (Bridge and Johnson, 2000). The relative rates of dissolution varied with the structural stabilities of the ferric iron minerals tested (Table 3 and Fig. 3). Contact between bacterial cells and the solid mineral was not necessary to bring about reductive dissolution of the latter, and addition of the iron-chelators EDTA or DPTA enhanced the rates at which goethite and amorphous ferric hydroxide were solubilized. It was found that reductive dissolution of ferric iron minerals by Acidiphilium SJH was more rapid at pH 2 than at pH 3, even though the pH growth optimum for this acidophile is close to pH 3. It was concluded that Acidiphilium SJH mediated the reductive dissolution of ferric-iron-containing minerals by an indirect mechanism, wherein reduction of ferric iron causes a shift in equilibrium between the solid (mineral) phase and soluble ferric iron, inducing further chemical dissolution of the mineral (Equation (1)). The ultimate rate of dissolution depended on the latter, which varied with the stability of the mineral and pH of the medium. FeðIIIÞ-mineral 2 Fe3þ soluble ðþAcidiphilium SJHÞ ! Fe2þ

(1)

One of the heterotrophic acidophiles that tested negative for iron reduction in the report of Johnson and McGinness (1991) was Acidocella facilis (classified at that time as Acidiphilium facilis). Soluble ferric iron (requiring culture pH of r2.5) was used in these early screening experiments, and reduction of soluble ferric iron is an acid-generating process (e.g. Equation (2), Table 3 Specific rates of reduction of soluble ferric iron and ferric iron-containing minerals by Acidiphilium SJH (data, showing mean values and standard deviations, from Bridge and Johnson (2000)) Iron Form

Specific Rate of Iron Reduction (mg Fe3þ reduced /min /mg protein)

Soluble Fe3þ Amorphous Fe(OH)3 Magnetite Goethite Natrojarosite Jarosite Akaganeite Hematite

1.4670.46 0.7670.10 0.3370.04 0.1770.05 0.1770.01 0.0670.02 0.0670.05 0.0170.01

n ¼ 4, except for natrojarosite, where n ¼ 2.

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Figure 3 Reduction of () soluble ferric iron, (o) amorphous ferric hydroxide, (7) magnetite, (r) goethite, (’) natrojarosite, (&) jarosite, (~) akaganeite and (B) hematite by Acidiphilium SJH. Reproduced from Bridge and Johnson (2000) with permission from the publishers.

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which shows ferric iron reduction coupled to oxidation of glucose): 24Fe3þ þ C6 H12 O6 þ 6H2 O ! 24Fe2þ þ 6CO2 þ 24Hþ

(2)

Both of these factors would have mitigated against Ac. facilis, as Acidocella spp. are, in general, less acid-tolerant than Acidiphilium spp. (Hallberg and Johnson, 2001), and the negative result obtained for Ac. facilis and some other bacteria screened could therefore have been due to the pH of the media being, or becoming, too low to support their growth. Recently, several classified Acidocella spp. and Acidocella-like isolates have been screened for iron reduction using synthetic schwertmannite as the source of ferric iron (Coupland and Johnson, 2008). Schwertmannite is a poorly crystalline ferric iron, and is frequently the dominant ferric iron mineral that forms in acid mine drainage-impacted environments (Bigham et al., 1996) from where Acidocella and other acidophilic heterotrophs have frequently been isolated (e.g. Johnson et al., 2001). In contrast to soluble iron, the reductive dissolution of schwertmannite is an alkali-generating reaction (Equation (3)): 3Fe8 O8 ðOHÞ6 ðSO4 Þ þ C6 H12 O6 þ 6H2 O !  24Fe2þ þ 6CO2 þ 3SO2 4 þ 42OH

ð3Þ

Ac. facilis, a strain of ‘‘Ac. aromatica’’ and two Acidocella-like environmental isolates were all found to be able to accelerate the reductive dissolution of schwertmannite under oxygen-limiting conditions, as were Acidobacterium capsulatum and related environmental isolates, and some Frateuria-like acidophiles (Coupland and Johnson, 2008). In contrast, Acidisphaera rubrifaciens and three Acidisphaera-like isolates showed no evidence of iron reduction under the experimental conditions used. The ability to catalyze the dissimilatory reduction of ferric iron appears, therefore, to be widespread among acidophilic heterotrophs, though it remains to be demonstrated whether this can facilitate growth of these bacteria in anaerobic or micro-aerobic environments. Most moderately thermo-acidophilic heterotrophic bacteria that have been described are Gram-positive Alicyclobacillus spp. (Section 2.2.3). The five moderately thermo-acidophilic heterotrophs (all of which were isolated from Yellowstone National Park) that were found by Johnson and McGinness (1991) not to reduce ferric iron were all later confirmed (from 16S rRNA gene sequence analysis) to be Alicyclobacillus spp. A related Yellowstone isolate (Y004) was later reported to display a limited capacity for iron reduction (Johnson et al., 2003). In the same study, Gram-negative thermo-acidophilic heterotrophic bacteria were isolated that were later named as strains of the novel species, Acidicaldus organivorans

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(Johnson et al., 2006a). Acd. organivorans is an adept iron-reducer and, in contrast to most other heterotrophic acidophiles, can grow under strictly anaerobic conditions using ferric iron as sole electron acceptor. 3.2.3. Mixotrophic Acidophiles The ability to reduce ferric iron is widespread among acidophilic bacteria that can use inorganic electron donors (ferrous iron or reduced sulfur) and assimilate either organic or inorganic carbon. Acidophilic iron- and sulfuroxidizing Firmicutes include all Sulfobacillus spp. and some Alicyclobacillus spp. Bridge and Johnson (1998) showed that both Sulfobacillus thermosulfidooxidans and Sulfobacillus acidophilus could grow anaerobically by ferric iron respiration, and accelerate the reductive dissolution of some ferric iron minerals. Ferric iron reduction was optimum when glycerol was used as the electron donor, but was also found to occur in inorganic culture media containing tetrathionate. Other Sulfobacillus spp. (e.g. ‘‘Sb. montserratensis’’) have also been shown to reduce iron (Yahya et al., 1999). Mixotrophic Alicyclobacillus spp. can also use either iron or reduced sulfur as electron donors, though these have more ‘‘heterotrophically inclined’’ physiologies than Sulfobacillus spp. Alicyclobacillus tolerans was reported to reduce iron, though whether this was used to support growth in oxygen-limited situations was not ascertained (Karavaiko et al., 2005). However, growth and ferric iron reduction were shown to be tightly correlated when a closely related strain (Alicyclobacillus GSM) was grown in anaerobic cultures (Yahya et al., 2008). Three species of iron-oxidizing acidophilic Actinobacteria have been described (Acidimicrobium ferrooxidans, Ferrimicrobium acidiphilum and Ferrithrix thermotolerans; Sections 2.2.3 and 2.3.2), all of which can use ferric iron reduction as an energy-transducing reaction to support growth in the absence of oxygen (Bridge and Johnson, 1998; Johnson et al., 2008). A strain of the iron-oxidizing archaeon Ferroplasma acidiphilum, isolated from a pilot-scale commercial mineral bioleaching system, was reported to reduce ferric iron to ferrous using yeast extract as substrate (Okibe et al., 2003). Dopson et al. (2004) later showed that Ferroplasma spp. are facultative anaerobes and can grow in the absence of oxygen by ferric iron respiration. 3.2.4. Enzymology of Ferric Iron Reduction in Acidophilic Prokaryotes Very little is known about the enzymology of ferric iron reduction in acidophilic bacteria. The fact that all iron-oxidizing acidophilic bacteria (with the exception of Leptospirillum spp. that appear to use no electron

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donors other than ferrous iron) and the iron-oxidizing mesophilic/ thermotolerant archaeon Ferroplasma can reduce ferric iron may suggest a reverse role of a Fe2þ/Fe3þ cycling system, as suggested by Sugio and co-workers (Sugio et al., 1987, 1989). This cannot be the case, however, in heterotrophic bacteria such as Acidiphilium spp. and Acd. organivorans that do not oxidize ferrous iron. Little is known about the enzymology of ferric iron reduction in these acidophiles. A comparative study of iron reduction by Acidiphilium SJH (which is closely related to the type species, A. cryptum) and A. acidophilum (which falls into a different group of Acidiphilium spp., on the basis of 16S rRNA gene sequences, and which is a facultative autotroph) showed distinct differences between the two bacteria (Johnson and Bridge, 2002). In Acidiphilium SJH, the capacity for ferric iron reduction appeared to be constitutive; specific rates of iron reduction were similar in cultures grown under different oxygen status and in the presence or absence of ferric iron. In contrast, ferric iron reduction by A. acidophilum was very dependent on dissolved oxygen concentrations in the growth medium (though not by the ionic form of iron present); the capacity for iron reduction was induced under micro-aerobic conditions and repressed when grown in well-aerated cultures. Whole cell protein profiles of Acidiphilium SJH grown under different oxygen status were very similar, while those of A. acidophilum showed additional protein bands in micro-aerobic compared to aerobic cultures. Whether these additional proteins had a role in mediating ferric iron reduction was not ascertained. Interestingly, Ku¨sel et al. (2002) concluded that the enzymes responsible for ferric iron reduction in A. cryptum JF-5 (which is more closely related to Acidiphilium SJH than to A. acidophilum) are not constitutive. Clearly, further work is required to elucidate the biochemistry of ferric iron reduction in acidophilic prokaryotes.

4. SULFUR METABOLISM BY ACIDOPHILIC MICRO-ORGANISMS 4.1. Dissimilatory Oxidation of Elemental Sulfur and Reduced Inorganic Sulfur Compounds Naturally acidic environments are formed in areas of high sulfur concentration, and are usually associated with areas of geothermal activity. Those sites that have low neutralization potential (e.g. lack carbonatecontaining minerals) become acidic due to the oxidation of sulfide by

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acidophilic micro-organisms, producing sulfate and protons (e.g. sulfuric acid). In addition, geothermal sites may also contain sulfur, which is formed by the condensation of hydrogen sulfide with sulfur dioxide. This sulfur can also be oxidized by acidophilic micro-organisms to sulfuric acid. Sulfur and RISCs can also accumulate in areas where pyrite and other sulfide minerals are oxidized by ferric iron (Schippers and Sand, 1999). Generally, acidophilic sulfur-oxidizing micro-organisms are capable of oxidizing these RISCs (e.g. polythionates such as thiosulfate and tetrathionate) as well as oxidizing sulfur and sulfide. 4.1.1. Oxidation of Sulfur and RISCs The first acidophile to be described was a sulfur-oxidizing bacterium isolated from soil, and was named Thiobacillus thiooxidans (Waksman and Joffe, 1922). In addition to being able to oxidize sulfur to sulfuric acid, At. thiooxidans was also shown to oxidize thiosulfate. An iron-oxidizing acidophile that was also able to oxidize thiosulfate was isolated some years later from acid mine drainage in Pennsylvania, U.S.A. (Colmer et al., 1950), and because of the former trait, was subsequently named Thiobacillus ferrooxidans (Temple and Colmer, 1951). Recent phylogenetic analysis based on 16S rRNA gene sequencing has shown that these bacteria are a distinct group of bacteria from the type species of Thiobacillus, namely Thiobacillus thioparus, and thus they were placed into a new genus called Acidithiobacillus to reflect their acidophilic nature (Kelly and Wood, 2000). Bacteria of this genus, including the moderately thermophilic At. caldus (Hallberg and Lindstro¨m, 1994), are capable of oxidizing a wide range of RISCs in addition to sulfur, including sulfide, thiosulfate, tetrathionate, trithionate, and sulfite. When each of the three acidithiobacilli were cocultured in sulfur-containing medium at 30 1C, cell numbers of At. caldus remained constant during successive sub-cultures, while the numbers of At. thiooxidans and At. ferrooxidans were found to decrease rapidly (Hallberg et al., 2001). The ability of At. caldus to out-compete the other strains under these conditions is interesting, as this moderate thermophile is often thought to be exclusive to higher temperature environments because it has a higher optimum growth temperature than the other two Acidithiobacillus spp.. Given the widespread distribution of sulfur and RISCs in acidic environments, it is not surprising that many acidophiles besides those of the genus Acidithiobacillus can also oxidize these substrates (Table 1). Many of these are capable of autotrophic growth on sulfur and RISCs, and these usually are also able to grow with ferrous iron as electron donor. Interestingly, Alicyclobacillus disulfidooxidans, which is capable of autotrophic

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growth on sulfur, was also shown to grow using organic sulfur-containing compounds (Dufresne et al., 1996). No data were presented, however, to show that this bacterium could oxidize the organic sulfur. Typically, these acidophiles will oxidize iron in preference to sulfur and RISCs when provided both as alternative substrates, even though sulfur is a more energetic electron donor (see below). While this may seem paradoxical, it may reflect the relative reactivity of these substrates (iron is much more chemically reactive with oxygen than is sulfur). Some heterotrophic acidophiles are also capable of oxidizing sulfur and RISCs. One heterotroph, Acidiphilium acidophilum (formerly Thiobacillus acidophilus), is also capable of autotrophic growth with these substrates using RUBISCO to assimilate carbon from CO2. Other Acidiphilum species are also capable of oxidizing sulfur, though they require organic carbon to do so (Hallberg et al., 2001), most likely reflecting the lack of RUBISCO in these species as opposed to A. acidophilum. During growth in media containing both glycerol and sulfur, sulfate and protons are produced concurrently with the decrease in glycerol concentration, indicating that these bacteria oxidize sulfur and glycerol simultaneously (Hallberg, unpublished data). A moderately thermophilic heterotrophic acidophile, Acidicaldus organivorans, is also able to oxidize tetrathionate (Johnson et al., 2003), though in the description of the species it was only indicated that it could oxidize sulfur (Johnson et al., 2006a). Like the majority of Acidiphilum species, Acd. organivorans was only able to oxidize sulfur and tetrathionate in the presence of organic carbon, which correlated with the observation that RUBISCO genes could not be amplified from this acidophile. In addition to aerobic growth by oxidation of sulfur and RISCs, some acidophiles are also able to couple the oxidation of sulfur to reduction of iron. This was first observed with At. thiooxidans, At. ferrooxidans and an isolate of Sulfolobus (Brock and Gustafson, 1976). The reduction of iron was observed in aerobic cultures of At. thiooxidans and microaerophilic cultures of Sulfolobus, while reduction was only observed in cultures of At. ferrooxidans when oxygen was excluded. There appears to be a link to the ability to oxidize and reduce iron as the sulfur-oxidizers At. thiooxidans and At. caldus were shown to be unable to reduce iron and grow under anaerobic conditions (Hallberg et al., 2001). 4.1.2. Energetics of Sulfur Oxidation Elemental sulfur and RISCs are more energetically favorable substrates for acidophilic prokaryotes than is ferrous iron. There are two reasons for this: first, more electrons are available per mole of sulfur and RISC than from

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ferrous iron (e.g. one from the oxidation of Fe2þ to Fe3þ, as opposed to six from the oxidation of S0 to SO2 4 ) and second, since electrons obtained from the oxidation of sulfur and RISCs enter the electron transport chain at a higher redox level than those derived from ferrous iron, more ATP is produced per mole of substrate due to increased proton translocation (Kelly, 1999). The free energy available from the oxidation of thiosulfate to sulfuric acid at pH 2 is 762.47 kJ/mol, while that of oxidation of Fe2þ to Fe3þ at the same pH is 138.89 kJ/mol. The coupling of the oxidation of sulfur and RISCs to the electron transport chain at a higher redox level also has important energetic implications in relation to fixation of carbon dioxide. Since electrons from these substrates enter the electron transport chain at cytochrome b (see below), they are readily available for the reduction of NAD(P) to NAD(P)H, thus eliminating the need for the uphill electron transport cascade required during iron oxidation (Section 2.1). For these reasons, growth yields of acidophiles on sulfur (and RISCs) is much higher than when using iron as growth substrate, for example 0.92 g dry weight cells of At. ferrooxidans/mol electrons from tetrathionate vs 0.23/mol electrons from iron (Hazeu et al., 1987). This is why sulfur-oxidizing acidophiles can often be found in greater numbers than iron oxidizers in bioreactors where sulfide mineral oxidation is occurring (e.g. Okibe et al., 2003). 4.1.3. Biochemistry of Sulfur Oxidation Since the variety of RISCs is great, more elaborate mechanisms are needed for the oxidation of these compared with the oxidation of iron. Early studies that examined the oxidation of RISCs and sulfur made use of whole cells. Such work with At. ferrooxidans (as reviewed in Pronk et al., 1990) and A. acidophilum (Meulenberg et al., 1992) led to a model of RISC and sulfur oxidation in these acidophiles (Fig. 4). This model was shown to also operate in the acidophiles At. thiooxidans (Chan and Suzuki, 1994) and At. caldus (Hallberg et al., 1996). There is evidence that this path of RISC and sulfur oxidation is common among acidophiles, since a Sulfolobus isolate was also shown to oxidize thiosulfate to tetrathionate, followed by oxidation of tetrathionate with thiosulfate as an intermediate (Nixon and Norris, 1992). However, while it is apparent that the acidophilic microorganisms share many common features in the oxidation of RISCs and sulfur, these are different from those used by neutrophilic sulfur-oxidizing bacteria (Friedrich et al., 2005). Sulfide is oxidized by At. ferrooxidans by a sulfide-quinone reductase, a membrane-bound protein that transfers electrons to ubiquinone (Quatrini

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Figure 4 Sequential oxidation of sulfur and reduced inorganic sulfur compounds to sulfuric acid by acidophilic prokaryotes.

et al., 2006). The product of this enzyme reaction is sulfur. Sulfur is then oxidized by the periplasmic sulfur dioxygenase to sulfite (Rohwerder and Sand, 2003). This enzyme activity was also detected in the acidophilic bacteria At. thiooxidans, A. acidophilum and A. cryptum. The thermophilic acidophilic archaea Acidianus brierleyi and Acidianus ambivalens use a sulfur dioxygenase that differs from that used by the bacteria in that it does not require reduced glutathione (Emmel et al., 1986; Urich et al., 2004). Finally, sulfite is oxidized to sulfate by the enzyme sulfite:acceptor oxidoreductase. A similar coupling of enzymes is used for the oxidation of the polythionates, such as trithionate, tetrathionate and thiosulfate. A trithionate dehydrogenase that produces thiosulfate and sulfate was purified from A. acidophilum. This enzyme was assumed to be periplasmic due to the low pH required for its activity (Meulenberg et al., 1992). The thiosulfate thus produced is oxidized by another periplasmic enzyme, thiosulfate dehydrogenase, which produces tetrathionate (Meulenberg et al., 1993). A similar enzyme was also purified from At. thiooxidans (Chan and Suzuki, 1994), and also from the thermo-acidophilic archaeon Acidianus ambivalens, though in the latter case the enzyme could use quinones as electron acceptors (Mu¨ller et al., 2004). The final enzyme in this series is tetrathionate dehydrogenase, which produces sulfur, sulfate and thiosulfate as products. A comparison of this enzyme from several acidophiles reveals that it is similar in all acidophiles, though recent evidence suggests that it is membrane-bound rather than periplasmic as previously assumed (Kanao et al., 2007). The gene for this enzyme in At. caldus has been sequenced (Rzhepishevska et al., 2007), and bioinformatic analysis by these authors showed that a homolog of the tetrathionate dehydrogenase could be found in other acidophiles, including At. ferrooxidans as well as the archaea Sulfolobus tokodaii and Metallosphaera sedula.

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In contrast to the enzymes and electron transport pathways of iron oxidation in acidophiles, it appears that there is more congruence among the enzymes of sulfur and RISC oxidation in acidophilic micro-organisms. This would seem to suggest that sulfur and RISC oxidation is perhaps a trait that has been shared among acidophiles for longer than iron oxidation.

4.2. Dissimilatory Reduction of Sulfur and Sulfate by Acidophilic Micro-Organisms Most extremely acidic natural and man-made environments tend to contain elevated concentrations of sulfur in various oxidation states, though the þ6 state (sulfate) is often the dominant form present. Against this backdrop, it is curious that few prokaryotes are known to catalyze the dissimilatory reduction of oxidized forms of sulfur at low pH. Anaerobic growth by reduction of elemental sulfur has been described in four genera of acidophilic archaea: Acidianus, Stygiolobus, Sulfurisphaera (all thermoacidophilic crenarchaeotes) and Thermoplasma (a moderately thermoacidophilic euryarchaeote). Acidianus spp. and Sulfurisphaera ohwakuensis are all facultative anaerobes that couple the oxidation of hydrogen to the reduction of sulfur in anoxic environments. Stygiolobus azoricus is, in contrast, an obligately anaerobic thermo-acidophile that grows by coupling the oxidation of hydrogen to the reduction of sulfur. Both classified species of Thermoplasma (Tp. acidophilum and Tp. volcanium) are facultative anaerobes and obligately heterotrophic oxidation of organic carbon is coupled to the reduction of elemental sulfur for growth in anaerobic environments (Segerer et al., 1988). No acidophilic archaea are known to catalyze the dissimilatory reduction of sulfate. Sulfidogenesis in cool temperature, low-pH anaerobic sediments and microbial mats has been described on a number of occasions (e.g. Tuttle et al., 1969) and there is evidence that sulfate, rather than elemental sulfur, is the source of sulfide in these situations. However, isolation and characterization of acidophilic or acid-tolerant sulfate reducing bacteria (SRB) has mostly proved elusive, with the majority of isolates from acidic sites behaving similarly to known SRB in being highly sensitive to even mild acidity (no growth in synthetic media poised at pH o5). This has led some authors to conclude that sulfidogenesis in these situations is mediated by neutrophilic SRB that inhabit and maintain (by way of their alkali-generating metabolisms) micro-sites of significantly higher pH than that of the bulk environment.

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There are no thermodynamic reasons why sulfate cannot be used as a terminal electron acceptor at low pH. The redox potential (Eo) of the  SO2 4 /HS couple at pH 7 is well known to be very low (0.217 V) making it a less favorable electron sink than possible alternatives such as nitrate. However, this value is affected by pH (due to the fact that protons are þ  involved in half-cell reaction: SO2 4 þ 10H þ 8e 2H2 S þ 4H2 O) and it becomes more positive (and therefore thermodynamically more attractive) as pH decreases (e.g. at pH 3.0, the theoretical value of the sulfate/sulfide couple is þ75 mV). It is necessary, therefore, to identify what other factors could be responsible for the apparent absence of genuinely acidophilic SRB. There are at least two possibilities – the toxicity of organic acids and of hydrogen sulfide. At the pH at which many acidophiles grow, small molecular weight organic acids (such as acetic) tend to be present as uncharged/undissociated molecules, due to their pKa values being greater than those of the liquid medium. Uncharged small molecular weight aliphatic acids are lipophilic and migrate through membranes into the cell cytoplasm that, even in the majority of acidophiles, has a pH close to neutral, causing the inflowing acids to dissociate. This results in dis-equilibrium, inducing continued influx of protons and potential severe acidification of cell cytoplasm and cell death. As a result, small molecular weight aliphatic acids tend to be highly toxic to many acidophilic bacteria when present in very small (micro-molar) concentrations. Media formulations designed to enrich for and cultivate SRB often contain organic acids (such as lactic acid) as carbon/energy sources, and these are, for this reason, inappropriate for acidophilic SRB. Even when a non-acid carbon source (such as glycerol) is provided as an alternative, problems due to organic acid toxicity may arise due to the fact that many SRB are ‘‘incomplete substrate oxidizers,’’ and generate and accumulate stoichiometric concentrations of (usually) acetic acid in their growth milieu resulting, again, in inhibition of growth. Hydrogen sulfide is also a toxic byproduct generated by SRB, and can inhibit growth of these bacteria when present at as little as 3 mM (Okabe et al., 1995). Hydrogen sulfide is itself a weak acid, with two pKa values (6.9 and 14; Equation (4)): H2 S 2 HS þ Hþ 2 S2 þ 2Hþ

(4)

Of these, the most toxic form is H2S (Moosa and Harrison, 2006) and, at pH 7, H2S and HS each account for about 50% of sulfide species present, and S2 less than 1%. At pH 4, however, W95% of the sulfide generated by SRB is present as H2S. Ferrous iron is included in many media formulations for cultivating SRB specifically to act as a sink for biogenic sulfide. However, FeS has a relatively large solubility product and does not form in

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acidic liquors (less than BpH 5.5) and is therefore ineffective at removing sulfide where SRB are cultivated in media of low pH. Sulfides of other metals, such as copper and zinc, have smaller solubility products than FeS and could, in theory, substitute for iron, though at least some SRB are highly sensitive to both of these metals (Hao, 2000). There has been at least one demonstration, however, of dissimilatory sulfate reduction occurring under carefully controlled, acidic conditions, by acidophilic/acid-tolerant bacteria (Johnson et al., 2006b; Kimura et al., 2006). In this case, the sulfidogenic population was a bacterial consortium, comprising two species: an apparently novel species of Desulfosporosinus (isolate M1) and a heterotrophic acidophile (strain PFBC) that is closely related to ‘‘Acidocella aromatica.’’ The growth medium used contained glycerol as carbon/energy source and zinc as a sink for hydrogen sulfide (both species were shown to tolerate at least 10 mM zinc). Desulfosporosinus M1 was found, like its close relatives, to be an incomplete substrate oxidizer and generated equimolar concentrations of acetic acid when oxidizing glycerol. In co-culture with Acidocella PFBC, however, little or no acetic acid was found to accumulate in the culture medium, and the stoichiometry of glycerol oxidized to zinc precipitated in cultures grown at pH 4.0 was close to 1:1, which is the theoretical value obtained when glycerol is completely oxidized to CO2 (Equation (5)): þ 4C3 H8 O3 þ 7SO2 4 þ 14H ! 12CO2 þ 7H2 S þ 16H2 O

(5)

Curiously, Acidocella spp. had previously been described as obligate aerobes, and attempts to grow pure cultures of strain PFBC in anoxic media either by fermentation or by anaerobic respiration (using soluble ferric iron, nitrate or sulfur as potential electron acceptor) were all unsuccessful (though later it was shown that Acidocella PFBC can accelerate the reductive dissolution of the ferric iron mineral schwertmannite under oxygen-limiting conditions; Section 3.2.2). Analysis of the consortium during active sulfidogenesis using fluorescent in situ hybridization (FISH) showed that both Desulfosporosinus M1 and Acidocella PFBC were growing concurrently. Using these data, a scheme was proposed in which the two bacteria assumed a syntrophic relationship, involving the interspecies transfer of hydrogen (Fig. 5). In this scheme, Desulfosporosinus M1 first partially oxidized glycerol to acetic acid (using sulfate as electron acceptor), which was then converted to hydrogen and carbon dioxide by Acidocella PFBC. The hydrogen produced was used as a supplementary electron donor by Desulfosporosinus M1, again coupled to sulfate reduction. The thermodynamics of these reactions are all favorable, with the exception of the acetoclastic reaction, which has a theoretical positive DG value. However, as demonstrated with other syntrophic associations of

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Glycerol

Desulfosporosinus M1

4C3H8O3 + 3SO42- + 6H+ 4CH3COOH + 4CO2 + 3H2S + 8H2O

Acetic acid

4CH3COOH + 8H2O 16H2+ 8CO2

Acidocella PFBC

Hydrogen

16H2 + 4SO42- + 8H+ 16H2O + 4H2S Water

ZnS

Figure 5 Proposed syntrophic relationship between Desulfosporosinus M1 and Acidocella PFBC grown anaerobically in an acidic (pH 3.5–4.0) liquid medium containing zinc sulfate and glycerol as substrate. Modified from Johnson et al. (2006a) with permission from the publishers.

anaerobic micro-organisms (e.g. Schink, 1997), the DG value becomes negative and the reaction thermodynamically viable, where there is a highly effective sink for hydrogen (i.e. Desulfosporosinus M1 in this particular acidophilic sulfidogenic consortium) that results in the maintenance of a very low partial pressure of the hydrogen. It was shown that a mixed culture of these bacteria (though not a pure culture of Desulfosporosinus M1) could grow using acetic acid as sole carbon/energy source, and that Desulfosporosinus M1 could use hydrogen as an electron donor, both observations supporting the proposed hypothesis. The lowest pH at which the Desulfosporosinus M1/Acidocella PFBC could operate as an effective sulfidogenic consortium was found to be B3.6.

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The reason for this was possibly due to the fact that ZnS is quite soluble at this pH (and below) so that zinc no longer acted as an effective sink for sulfide. Desulfosporosinus M1 was shown to be highly sensitive to some other metals that formed less soluble sulfides (e.g. 1 mM Cu2þ inhibits its growth) and no suitable alternative metal was identified that could serve as an effective sink, at least when the consortium was grown in batch culture. However, using the same techniques as those used to isolate Desulfosporosinus M1, Rowe et al. (2007) obtained phylogenetically distinct strains of Gram-positive sulfidogens that were seemingly generating copper sulfide in the anaerobic zone of a streamer mat growing in a highly acidic (pH 2.5) mine drainage stream in southern Spain. It will be interesting to ascertain whether, by using these or other novel strains of acidophilic SRB, it will be possible in future to obtain sulfidogenesis at extremely low pH in vitro by consortia or pure cultures of bacteria.

5. APPLIED AND ECOLOGICAL ASPECTS Acidophilic micro-organisms are the key drivers of a rapidly expanding area of biotechnology, generically referred to as ‘‘biomining.’’ In addition, their role in generating acidic, metal-rich effluents that drain abandoned mine spoils and mineral tailings have been documented for over 50 years. The extremophiles exploit natural as well as man-made acidic environments, such as solfatara fields in geothermal areas, and sites where there are surface exposures of sulfide-rich rocks (gossans). In nearly all such situations, carbon, iron and sulfur transformations catalyzed by acidophilic micro-organisms are critical in determining the geochemistry of the local environment. There are a large number of review articles on applied and ecological aspects of acidophilic microbiology, including Rawlings (2002) and Olsen et al. (2003), both of which focus on aspects of biomining, and Johnson (2006) which has a more environmental bias.

5.1. Acidophiles and Mineral Processing ‘‘Biomining’’ can be defined as biotechnology that uses the ability of certain prokaryotic micro-organisms to accelerate the oxidative dissolution of sulfide minerals, either to solubilize target metal(s), such as copper and nickel, or to dissolve minerals in order to expose the target (noble) metal,

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such as gold. The advantages of bioprocessing of ores and concentrates over more conventional approaches such as smelting include the potential for processing low-grade deposits, re-processing waste materials from mining carried out in previous times where metal extraction tended to be less efficient, potentially lower energy inputs and other environmental benefits (zero production of noxious gases, etc.). Acidophiles were exploited for the extraction and recovery of metals from ores and wastes long before their roles were first recognized (in the mid-20th century) at both the Rio Tinto mine (southern Spain) and the Parys mine (Anglesey, north Wales). During the 18th and 19th centuries, the practice of allowing underground shafts and adits at these mines to flood, and then periodically releasing and capturing the metal-rich waters to recover copper from the leached subterranean rocks by adding scrap iron (cementation) was adapted. This was, essentially, an unwitting application of ‘‘in situ’’ leaching, which later became popular for extracting uranium from worked-out mines in Canada in the 1970s, and remains as an engineering option in commercial biomining. However, most current biomining operations use one of two other engineering alternatives – stirred tanks and irrigated heaps and dumps. Since the 1980s, aerated stirred tanks have been used to process sulfidic ore concentrates. These tanks, which may be extremely large (W1,000 m3), allow precise control of temperature, pH, etc., though, in common with all biomining options, they operate as non-sterile ‘‘open’’ systems with little or no control of bioleaching microbial populations. Most stirred tank bioreactors used for mineral processing have tended to operate at between 40 1C and 50 1C (i.e. where moderate thermophiles and thermotolerant acidophiles would tend to be of greatest significance) and have been used to process refractory gold concentrates. Two notable exceptions are: (i) Kasese, Uganda, where a cobalt-rich pyrite concentrate is being bioleached in stirred tanks (Morin and d’Hugues, 2007); (ii) a thermophilic stirred tank, operating at about 80 1C, that was used to extract copper from chalcopyrite, a mineral that is notoriously difficult to bioleach at lower temperatures (Batty and Rorke, 2006). Most heap and dump leaching operations, in contrast, have been used to leach copper from secondary sulfide minerals (e.g. chalcocite; Cu2S), though full-scale heap leaching of a polymetallic black schist ore (containing nickel, and some other base metals) in Talvivaara, Finland, has recently been initiated (Riekkola-Vanhanen, 2007). Somewhat surprisingly, there have been relatively few accounts of the compositions and dynamics of microbial populations in commercial biomining operations. These have tended to show, as would be expected, that microbial diversity is far greater in heaps and dumps, which are highly

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heterogeneous and uncontrolled environments, than in stirred tanks where conditions are far more homogeneous. Both operate essentially as ‘‘inorganic’’ systems in that, while inorganic nutrients (ammonium and phosphate) are added to stimulate microbial activity, organic carbon is not. This, together with the primary energy sources available being the sulfide minerals themselves, means that the dominant prokaryotes present are invariably chemo-autotrophic iron and sulfur oxidizers. However, organic carbon derived from living (as exudates) and dead (as lysates) primaryproducers can accumulate in leachate liquors, and can support the growth of mixotrophic and heterotrophic acidophiles. It is possible, therefore, to divide micro-organisms in biomining operations into three groups: (i) ‘‘primary acidophiles,’’ iron-oxidizing prokaryotes that generate ferric iron and are responsible for initiating mineral dissolution; (ii) ‘‘secondary acidophiles,’’ sulfur-oxidizing acidophiles that generate sulfuric acid from reduced sulfur produced during mineral dissolution and help maintain pH conditions that are conducive for the bio-oxidation of sulfide minerals; (iii) ‘‘tertiary acidophiles,’’ heterotrophic and/or mixotrophic microorganisms that degrade soluble organic carbon wastes originating from the autotrophs, thereby detoxifying the environment for some of the more organic-sensitive primary and secondary prokaryotes. As noted previously, some acidophiles may assume more than one role in this consortium hierarchy, for example At. ferrooxidans is capable of oxidizing both iron and sulfur, and both Ferroplasma and Ferrimicrobium are heterotrophic acidophiles that can oxidize ferrous iron. The importance of microbial consortia and interactions for optimizing sulfide mineral dissolution has been demonstrated in laboratory studies (e.g. Okibe and Johnson, 2004). Interestingly, microbial analysis of commercial-scale bioprocessing operations have shown, in all cases so far reported, that bacteria and/or archaea that fulfil these primary, secondary and tertiary roles are present (Table 4). Figure 6 is a schematic representation of microbial interactions between primary, secondary and tertiary acidophiles that occur during commercial bioprocessing of minerals. In a different approach to bioprocessing minerals (a two-stage ‘‘indirect’’ design) mineral oxidation (a chemical process) and regeneration of the oxidant ferric iron (a biological process) are physically separated. Perceived advantages of such an approach are (i) conditions (such as temperature) for each of these steps can be optimized independently; (ii) sulfur oxidation can be more readily controlled to facilitate the production of elemental sulfur rather than sulfate. A critical component of an indirect two-stage system is the efficiency of the ferric iron-regenerating bioreactor. One of the most efficient of these operated at 37 1C, regenerating ferric iron at the rate of

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Table 4 Acidophilic prokaryotes identified in stirred tank mineral bioleaching and biooxidation operations (from Rawlings and Johnson (2007) reproduced with permission of the publishers) Mineral concentrate

T ( 1C)

Prokaryotes identified

Reference

Zinc/lead pyrite

35–40

Leptospirillum ferrooxidansa Acidithiobacillus thiooxidansb Acidiphilium cryptumc Acidithiobacillus ferrooxidansc

Goebel and Stackebrandt (1994)

Pyrite/arsenpyrite (gold) Bioxs culture

40

L. ferrooxidansa At. thiooxidansb At. ferrooxidans

Dew et al. (1997)

Cobaltiferous pyrite

35

L. ferrooxidans At. thiooxidans Sulfobacillus thermosulfidooxidans

Battaglia-Brunet et al. (2002)

Polymetallic (copper, zinc and iron sulfides)

45

Leptospirillum ferriphilum Acidithiobacillus caldus Sulfobacillus sp. Ferroplasma acidiphilum

Okibe et al. (2003)

Pyrite, arsenical pyrite and chalcopyrite

45

At. caldus Sb. thermosulfidoooxidans ‘Sulfobacillis montserratensis’

Dopson and Lindstro¨m (2004)

Chalcopyrite

78

(Sulfolobus shibitaed,e) (Sulfurisphaera ohwakuensisd,e) Stygiolobus azoricusd Metallosphaera sp.d Acidianus infernusd

Mikkelsen et al. (2006)

a L. ferrooxidans was almost certainly L. ferriphilum as identification methods at the time did not permit the two species to be distinguished from each other. b At. thiooxidans was almost certainly At. caldus for the same reason as footnote a. c These two species were found in batch tanks but not in continuous flow tanks. d Nearest affiliated cultivated archaea to recovered clones. e Clones probably represent new species within the order Sulfolobales.

8.2 g/L/ h with a hydraulic retention time of 36 minutes (Kinnunen and Puhakka, 2004). Analysis of the bioreactor population revealed that it comprised only iron-oxidizing acidophiles, though one (L. ferriphilum) was an autotroph while the other (a Ferroplasma-like archaeon) was heterotrophic.

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(Ia)

CO2

(Ib)

(III) 2Fe2+

Fe2+

FeS2 FeS 2

DOC

Fe3+

Fe2+

Fe3+

Fe3+ S2O32-

Polythionates, S0

DOC

(II) DOC

Figure 6 Schematic representation of the roles of and interactions between different physiological groups of acidophilic prokaryotes during the oxidative dissolution of a representative sulfide mineral (pyrite; FeS2). Iron-oxidizing prokaryotes, either attached to the mineral surface (Group Ia) or free-swimming (Group Ib) have the primary role in mineral dissolution in that they generate ferric iron, the chemical oxidant that attacks and degrades the mineral. Sulfur-oxidizing prokaryotes (Group II) have a secondary role whereby they oxidize reduced sulfur produced during pyrite dissolution, producing sulfuric acid which maintains the required acidic conditions. Both the primary and secondary micro-organisms are predominantly autotrophs and leak dissolved organic compounds (DOC) into their environment. This is utilized by tertiary (Group III) prokaryotes (heterotrophic and facultatively autotrophic acidophiles); mineralization of DOC helps maintain a suitable environment for the autotrophic members of the consortium.

5.2. Acidophiles in Natural and Anthropogenic Environments Acidophilic micro-organisms, including novel species and genera, have been isolated from various environments, including acidic springs and soils in Yellowstone National Park and other geothermal areas (e.g. Johnson et al., 2003); gossans in the high Arctic (Langdahl and Ingvorsen, 1997) and at derelict and abandoned mine sites (e.g. Bond et al., 2000; Johnson, 2006). The Richmond Iron Mountain mine in California has been one site that has been extensively studied, as it is one of the most extreme environments yet discovered and it contains underground pools where water pH values are negative (Nordstrom et al., 2000). Despite this, a large amount of microbial

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biodiversity has been found within Iron Mountain, including new species of iron-oxidizing prokaryotes (‘‘Ferroplasma acidarmanus’’ and ‘‘Leptospirillum ferrodiazotrophum’’; Edwards et al., 2000; Tyson et al., 2005). In contrast to biomining operations, where forced aeration of heaps and stirred tanks occurs, resulting in predominantly aerobic conditions that are conducive to the activities of mineral-oxidizing prokaryotes, geothermal and post-mining environments will often include zones where oxygen is abundant, limiting or absent, facilitating colonization by a more diverse range of acidophiles. Phototrophic acidophiles (micro-algae) may contribute to, or dominate, primary production in such situations, supplying additional fixed, dissolved organic carbon that can support extensive growths of heterotrophic acidophiles (e.g. Rowe et al., 2007). Under such conditions, oxidation and reduction of iron and sulfur can occur in closely juxtaposed micro-environments, and cycling of these macro-elements may be observed in these environments. Natural and anthropogenic acidic environments are also suitable locations for bioprospecting for novel acidophilic micro-organisms that can be used, potentially, for improving biomining operations or for developing new bioremediation systems. For example, there is considerable interest in halo-acidophilic acidophiles that could be used for processing minerals in parts of the world where the available water supply is either brackish or saline (most of the iron-oxidizing bacteria commonly found in stirred tanks and heaps are sensitive to relatively small concentrations of chloride). There is also continued interest in obtaining acid-tolerant and acidophilic anaerobic acidophiles (such as SRB) that could be used for bioremediation of acidic, metal-rich waste-waters. Advances in the past decade in understanding the physiologies of acidophilic micro-organisms, and in microbiological and biomolecular techniques used to isolate and analyze these extremophiles, have given new opportunities to obtain and exploit pure cultures and consortia of acidophilic micro-organisms for both current and new biotechnologies in the new millennium.

ACKNOWLEDGMENTS We acknowledge the financial support provided by the European Commission under the Sixth Framework Programme for Research and Development in the frame of the research projects BioMinE (European project contract NMP1-CT-500329-1) and Bioshale (European project contract NMP2-CT-2004 505710). DBJ is grateful to the Royal Society (U.K.) for the provision of an Industrial Fellowship.

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Valde´s, J., Pedroso, I., Quatrini, R., Hallberg, K.B., Valenzuela, P.D.T. and Holmes, D.S. (2007) Insights into the metabolism and ecophysiology of three acidithiobacilli by comparative genome analysis. Adv. Mater. Res. 20–21, 439–442. Wakao, N., Nagasawa, N., Matsuura, T., Matsukura, H., Matsumoto, T., Hiraishi, A., Sakurai, Y. and Shiota, H. (1994) Acidiphilium multivorum sp. nov., an acidophilic chemoorganotrophic bacterium from pyritic acid-mine drainage. J. Gen. Appl. Microbiol. 40, 143–159. Waksman, S.A. and Joffe, J.S. (1922) Microorganisms concerned in the oxidation of sulfur in the soil. II. Thiobacillus thiooxidans, a new sulfur-oxidizing organism isolated from the soil. J. Bacteriol. 7, 239–256. Walton, K.C. and Johnson, D.B. (1992) Microbiological and chemical characteristics of an acidic stream draining a disused copper mine. Environ. Pollut. 76, 169–175. Wichlacz, P.L., Unz, R.F. and Langworthy, T.A. (1986) Acidiphilium angustum sp. nov., Acidiphilium facilis sp. nov., and Acidiphilium rubrum sp. nov.-acidophilic heterotrophic bacteria isolated from acidic coal mine drainage. Int. J. Syst. Bacteriol. 36, 197–201. Wood, A.P. and Kelly, D.P. (1983) Autotrophic and mixotrophic growth of three thermoacidophilic iron-oxidizing bacteria. FEMS Microbiol. Lett. 20, 107–112. Wood, A.P. and Kelly, D.P. (1984) Growth and sugar metabolism of a thermoacidophilic iron-oxidizing mixotrophic bacterium. J. Gen. Microbiol. 130, 1337–1349. Xiang, X., Dong, X. and Huang, L. (2003) Sulfolobus tengchongensis sp. nov., a novel thermoacidophilic archaeon isolated from a hot spring in Tengchong, China. Extremophiles 7, 493–498. Yahya, A., Roberto, F.F. and Johnson, D.B. (1999) Novel mineral-oxidising bacteria from Montserrat (W.I.): Physiological and phylogenetic characteristics. In: Biohydrometallurgy and the Environment Toward the Mining of the 21st Century (R. Amils and A. Ballester, eds), vol. 9A, pp. 729–740. Elsevier, Amsterdam. Yahya, A., Hallberg, K.B. and Johnson, D.B. (2008) Iron and carbon metabolism by a mineral-oxidizing Alicyclobacillus-like bacterium. Arch. Microbiol. 189, 305–312. Yarzabal, A., Brasseur, G., Ratouchniak, J., Lund, K., Lemesle-Meunier, D., DeMoss, J.A. and Bonnefoy, V. (2002) The high-molecular-weight cytochrome c Cyc2 of Acidithiobacillus ferrooxidans is an outer membrane protein. J. Bacteriol. 184, 313–317. Zakharchuk, L.M., Tsaplina, I.A., Krasil’nikova, E.N., Bogdanova, T.I. and Karavaiko, G.I. (1994) Carbon metabolism in Sulfobacillus thermosulfidooxidans. Mikrobiologiya 63, 573–580. Zillig, W., Stetter, K.O., Wunderl, S., Schulz, W., Priess, H. and Scholz, L. (1980) The Sulfolobus- ‘‘Caldariella’’ group: Taxonomy on the basis of the structure of DNA-dependent RNA polymerases. Arch. Microbiol. 125, 259–269.

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Chemostat-Based Micro-Array Analysis in Baker’s Yeast Pascale Daran-Lapujade, Jean-Marc Daran, Antonius J.A. van Maris, Johannes H. de Winde and Jack T. Pronk Department of Biotechnology, Delft University of Technology and Kluyver Centre for Genomics of Industrial Fermentation, Delft, The Netherlands

ABSTRACT Chemostat cultivation of micro-organisms offers unique opportunities for experimental manipulation of individual environmental parameters at a fixed, controllable specific growth rate. Chemostat cultivation was originally developed as a tool to study quantitative aspects of microbial growth and metabolism. Renewed interest in this cultivation method is stimulated by the availability of high-information-density techniques for systemic analysis of microbial cultures, which require high reproducibility and careful experimental design. Genome-wide analysis of transcript levels with DNA micro-arrays is currently the most commonly applied of these highinformation-density analysis tools for microbial gene expression. Based on published studies on the yeast Saccharomyces cerevisiae, a critical overview is presented of the possibilities and pitfalls associated with the combination of chemostat cultivation and transcriptome analysis with DNA micro-arrays. After a brief introduction to chemostat cultivation and micro-array analysis, key aspects of experimental design of chemostat-based micro-array experiments are discussed. The main focus of this review is on key biological concepts that can be accessed by chemostat-based micro-array analysis. These include effects of specific growth rate on transcriptional regulation, context-dependency of transcriptional responses, correlations between transcript profiles and contribution of the corresponding proteins to cellular function and fitness, and the analysis ADVANCES IN MICROBIAL PHYSIOLOGY, VOL. 54 ISBN 978-0-12-374323-7 DOI: 10.1016/S0065-2911(08)00004-0

Copyright r 2009 by Elsevier Ltd. All rights reserved

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and application of evolutionary adaptation during prolonged chemostat cultivation. It is concluded that, notwithstanding the incompatibility of chemostat cultivation with high-throughput analysis, integration of chemostat cultivation with micro-array analysis and other high-informationdensity analytical approaches (e.g. proteomics and metabolomics techniques) offers unique advantages in terms of reproducibility and experimental design in comparison with standard batch cultivation systems. Therefore, chemostat cultivation and derived methods for controlled cultivation of micro-organisms are anticipated to become increasingly important in microbial physiology and systems biology.

Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Experimental tools . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Chemostat Cultivation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Micro-Array Technology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Reproducibility of Chemostat-Based Micro-Array Analysis . . . . . . 3. Specific growth rate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Specific Growth Rate and Transcriptional Regulation . . . . . . . . . 3.2. Specific Growth Rate and Environmental Stress Responses . . . . 4. Context dependency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Functional analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1. Assignment of Gene Function . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2. Analysis of Transcriptional Regulation Networks . . . . . . . . . . . . 5.3. Correlation of mRNA Levels and Metabolic Fluxes . . . . . . . . . . . 5.4. Correlation of Transcript Levels and Fitness of Null Mutants . . . . 6. Evolutionary and reverse engineering . . . . . . . . . . . . . . . . . . . . . . . . 7. Outlook . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1. Beyond the Transcriptome: Multi-Level Approaches . . . . . . . . . . 7.2. Beyond Steady-State Analysis: Perturbation Experiments . . . . . . 8. Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. 2.

ABBREVIATIONS a-IPM 5-HMF ChIP ESR GIGO

a-isopropylmalate 5-hydroxy methyl furfural chromatin immuno-precipitation environmental stress response garbage in–garbage out

258 259 261 261 271 274 276 276 278 279 282 282 284 286 288 290 295 295 298 298 299 299

MICRO-ARRAY ANALYSIS IN BAKER’S YEAST

PCR RiBi RP RT-PCR SAGE SGD TCA

259

polymerase chain reaction ribosome biogenesis ribosomal protein reverse transcriptase polymerase chain reaction serial analysis of gene expression Saccharomyces genome database tricarboxylic acid

1. INTRODUCTION Microbial physiology started its quest to discover, describe and understand pathways and processes in micro-organisms at the end of the 19th century. After a vibrant period, characterized by discovery of biochemical pathways and metabolic processes, a shift towards quantitative aspects of microbial growth and product formation was strongly inspired by industrial fermentation as well as by microbial ecology. In both these disciplines, quantitative relationships between environmental conditions and microbial performance are of paramount importance. Consequently, new theoretical and experimental approaches were developed and implemented to quantitatively analyze how metabolic fluxes, yields and other key parameters change in response to growth conditions (Kurowski and Pirt, 1975; Monod, 1949; Novick and Szilard, 1950a; Stouthamer, 1973). At the onset of the 21st century, the complete sequencing of many microbial genomes has stimulated the development of novel methods that enable a systemic analysis of gene expression and cellular performance. These ‘‘-omics’’ techniques, some of which are still in their infancy and undergoing rapid development, should ultimately enable a quantitative analysis of all messenger RNAs, all proteins and all intracellular lowmolecular-weight metabolites in microbial cultures. This will provide microbial physiology with unprecedented tools to accomplish its ultimate goal: an understanding of microbial cells as integral systems that respond to changes in their natural and industrial environments by intricate, multi-level regulation mechanisms (Nielsen and Olsson, 2002). Commonly used model organisms such as Escherichia coli and Saccharomyces cerevisiae contain thousands of genes, mRNAs and proteins and hundreds of low-molecularweight metabolites. Given the number of possible functional interactions between these components, bioinformatics and mathematical modeling are indispensable for data interpretation and organization. Since mathematical models rigorously adhere to the garbage in–garbage out theorem

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(GIGO; Vodovotz et al., 2007), they should be based on and be validated with accurate, reproducible data. More importantly, when highinformation-density approaches are used to analyze microbial responses to various environmental conditions or to genetic interventions, the experimental setup should ideally allow for a systematic dissection of the responses to various and varying stimuli. These requirements have rekindled broad interest in the fundamental concepts and experimental tools of quantitative physiology and, in particular, in chemostat cultivation (Castrillo and Oliver, 2004; Hoskisson and Hobbs, 2005; Nielsen and Olsson, 2002). The hitherto most extensively developed and, hence, utilized tool for genome-wide expression studies is DNA-micro-array analysis (DeRisi et al., 1997), which allows for an integral and quantitative analysis of all mRNAs in microbial cultures. Since chemostat cultivation offers unique possibilities for control and manipulation of cultivation conditions, the combination of DNA-micro-array analysis with chemostat cultivation has gained rapidly increasing interest (Fig. 1). With this review, we aim to provide an overview of

Citations per year

200

150

100

50

7 00 *2

06 20

05 20

04 20

03 20

02 20

01 20

00 20

19

99

0

AD

Figure 1 Citation rates of publications on chemostat-based yeast transcriptome studies. The data shown in the graph are derived from a manually checked search using ISI Web of Knowledge (The Thomson Corporation). In addition to the terms ‘‘yeast’’ and/or ‘‘cerevisiae,’’ the cited papers contain the terms ‘‘chemostat,’’ ‘‘continuous culture’’ and/or ‘‘continuous cultivation.’’ Furthermore, at least one of a plethora of terms used to describe genome-wide transcriptional analysis was present (‘‘transcriptome,’’ ‘‘transcriptomics,’’ ‘‘genome-wide,’’ ‘‘microarray,’’ ‘‘DNA-array,’’ ‘‘transcription analysis,’’ ‘‘transcript profiles,’’ ‘‘global gene expression’’ or ‘‘transcriptional response’’).  search performed until august 2007.

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the possibilities of chemostat-based micro-array analysis in the yeast S. cerevisiae. A list of the publications covering these analyses and the nature of the analyses is supplied in Table 1. Emphasis is largely on aspects of experimental design and underlying physiological concepts rather than on the biological mechanisms that have hitherto been investigated via this approach. While the review is restricted to S. cerevisiae, the discussed concepts and experimental approaches are equally applicable to other micro-organisms.

2. EXPERIMENTAL TOOLS 2.1. Chemostat Cultivation ‘‘We have developed a device for keeping a bacterial population growing at a reduced rate over an indefinite period of time’’ At a time when the structure of DNA (Watson and Crick, 1953) was still unknown and micro-arrays were not even a distant dream, Novick and Szilard (1950a) used these words to announce their invention of the chemostat. Like many important findings, the chemostat was simultaneously invented in two locations. Monod (1950), the other inventor, introduced the term ‘‘culture continue’’. In quantitative physiology, ‘‘chemostat’’ and ‘‘continuous culture’’ are now often used as synonyms. However, since the latter term has an inherently broader meaning (including, e.g., plug-flow bio-reactors and cultivation systems with cell retention), we will use the term ‘‘chemostat’’ throughout this review. 2.1.1. Basic Principles Batch cultivation of micro-organisms on agar plates, in shake flasks or in bioreactors is, at least initially, characterized by growth at the maximum specific growth rate possible under the experimental conditions (mmax, h1). However, the conditions in batch cultures inevitably change over time. Concentrations of products and nutrients, including, in aerobic cultures, the dissolved-oxygen concentration, all change as growth proceeds. Depending on the cultivation system, limited rates of diffusion (e.g. in colonies on agar plates) and/or gas transfer may cause spatial and/or temporal concentration gradients. Unless such heterogeneity and dynamics are themselves the subject of study, they represent an undesirable complication in quantitative studies. As the term ‘‘chemostat’’ and the above quote of Novick and Szilard already indicate, chemostats enable studies on microbial growth under

262

Table 1 Chemostat-based microarray studies in Saccharomyces cerevisiae. Range tested

Specific growth rate

Temperature

Affymetrix YGS98 0.02–0.05–0.1–0.25– 0.33 h1 0.03, 0.1, 0.2 h1 in C, N, Affymetrix YGS98 S and P limitations 12 and 30 1C Affymetrix YGS98

Oxygen availability

Aerobic and anaerobic

Nutrient limitation

Macronutrient limitation (C, N, O, P, S) – – – Glucose, ammonium Glucose Zinc Potassium Ammonium, leucine Ammonium, leucine, alanine, glutamine Glucose, maltose, acetate, ethanol Glucose, oleate

Array type

Affymetrix Affymetrix Affymetrix Affymetrix

YGS98 YGS98 YGS98 YGS98

Reference

Accession number

Regenberg et al. (2006)

E-MEXP593d

Castrillo et al. (2007)

E-MEXP-115d

Tai et al. (2007a); Tai et al. GSE6190e (2007b) ter Linde et al. (1999) – f Piper et al. (2002) Tai et al. (2005) GSE1723e Boer et al. (2003) GSE1723e GSE1723e E-MEXP-115d

Affymetrix YGS98 Affymetrix YGS98 Custom made filtersa Custom made filtersb Stanford slide arrayc Affymetrix YGS98

Tai et al. (2005) Castrillo et al. (2007) Wu et al. (2004) Hayes et al. (2002) Brauer et al. (2005) De Nicola et al. (2007)

V2 agilent microarray Stanford slide arrayc Affymetrix YGS98

Hess et al. (2006) Saldanha et al. (2004) Usaite et al. (2006)

i

Affymetrix YGS98

Daran-Lapujade et al. (2004) Koerkamp et al. (2002)

GSE8895f

Custom made filtersc

g

– h

GSE8035/GSE8088/ GSE8089e

j

E-MEXP-724d

k

PASCALE DARAN-LAPUJADE ET AL.

Control option

Ammonium, leucine, Affymetrix YGS98 phenylalanine, proline, asparagine, methionine Organic acid stress Acetate, benzoate, Affymetrix YGS98 propionate, sorbate Carbon dioxide stress 0–100% CO2 in sparging Affymetrix YGS98 gas Oxidative stress Custom made filtersc Chemical stress 0 or 0.5% of 5Affymetrix YGS98 hydroxymethyl furfural Glucose pulse Glucose-lim steady Affymetrix YGS98 stateþ10 mM glucose pulse 0 to 330 seconds Galactose-lim steady Stanford slide arrayc stateþ0.2 and 2.0 g/l glucose pulse Engineered strains hap4D Affymetrix YGS98 leu3D Affymetrix YGS98 rox1D Affymetrix YGS98

Boer et al. (2007)

GSE6405e

Abbott et al. (2007)

GSE5926e

Aguilera et al. (2005a)

GSE8900e

Koerkamp et al. (2002) Petersson et al. (2006)

k

–

Kresnowati et al. (2006)

GSE3821e

Ronen and Botstein, (2006)

l

m

hap1D

Affymetrix YGS98

sfp1D bmh2ts

Affymetrix YGS98 V2 agilent microarray

Raghevendran et al. (2006) Boer et al. (2005) ter Linde and Steensma, (2002) ter Linde and Steensma, (2002) Cipollina et al. (2008) Bruckmann et al. (2004)

gdh1D P. stipitis XYL1, P. stipitis XYL2, overexpression XSK1

Affymetrix YGS98 YG Filter (ResGen)

Bro et al. (2004) Salusjarvi et al. (2006)

MICRO-ARRAY ANALYSIS IN BAKER’S YEAST

Nitrogen sources

GSE2076e – – GSE5238e GSM13009 to GSM13012e n

–

263

(Continued )

Table 1 (Continued ) Range tested

a

Reference

Accession number

Affymetrix YGS98

Ferea et al. (1999)

GSE29e

Affymetrix YGS98

Jansen et al. (2005)

GSE8898e

Affymetrix YGS98

Jansen et al. (2004)

GSE8897e

Affymetrix YGS98

van Maris et al. (2004a)

–

Affymetrix YGS98

Sonderegger et al. (2003)

–

Affymetrix YGS98

Wahlbom et al. (2003)

–

Custom made yeast gene-filter array supplied by J. Hoheisel (Deutsches Krebsforschungszentrum, Heidelberg) (Hauser et al., 1998). Custom made yeast gene-filter array. c Custom made array described at http://cmgm.stanford.edu/pbrown/mguide/described in (DeRisi et al., 1997). d Array express database, http://www.ebi.ac.uk/arrayexpress/ e Genome expression omnibus database, http://www.ncbi.nlm.nih.gov/geo/ f The respective dataset can be retrieved at http://www.cbs.dtu.dk/yeast/ g The respective dataset can be retrieved at http://www.cs.man.ac.uk/cogeme/data h The respective dataset can be retrieved at http://genomics-pubs.princeton.edu/DiauxicRemodeling/home.shtml i he respective dataset can be retrieved at http://puma.princeton.edu/cgi-bin/publication/viewPublication.pl?pub_no ¼ 508 j The respective dataset can be retrieved at http://microarray-pubs.stanford.edu/yeast_bc/home.shtml k The respective data can be retrieved at http://www.molbiolcell.org/cgi/content/full/E02-02-0075/DC1 l The respective data can be retrieved at http://smd.stanford.edu/cgi-bin/publication/viewPublication.pl?pub_no ¼ 502 m The respective data can be retrieved at http://www.jbc.org/cgi/data/M512972200/DC1/1 n The respective dataset can be retrieved at http://www.cpb.dtu.dk/data/gdh1.htm b

PASCALE DARAN-LAPUJADE ET AL.

Selection experiments Prolonged glucoselimited chemostat cultures Prolonged glucoselimited chemostat cultures Prolonged maltoselimited chemostat cultures Directed evolution of pyruvate decarboxylasenegative S. cerevisiae Anaerobic growth of S. cerevisiae on xylose S. cerevisiae mutant with improved ability to utilize xylose

Array type

264

Control option

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constant physico-chemical conditions. Furthermore, chemostat cultures enable studies on the growth of micro-organisms at specific growth rates below their mmax. Thus, even when microbial strains or growth conditions are compared that have or cause a different mmax, chemostat cultivation enables a comparison of their physiology at an identical specific growth rate chosen and set by the researcher. A chemostat (Fig. 2) can be defined by five key elements: (i) an ideally mixed fermentation vessel (bio-reactor), (ii) a continuous inflow of fresh growth medium, (iii) a medium composition in which a single nutrient of choice limits biomass formation and in which all other nutrients are present in excess, (iv) a continuous outflow of culture broth with an identical chemical composition and biomass concentration to that in the bio-reactor and (v) a constant ratio between effluent flow rate and culture volume in the bio-reactor. The unique option of chemostats to ‘‘dial in’’ a specific growth rate can best be understood by combining these five defining elements with a mass balance of the biomass: dðVcx Þ=dt ¼ jin cx;in  fout cx;out þ mcx;bioreactor V

Figure 2 Experimental setup for chemostat cultivation. Panel A, early design by Monod (from Monod, 1950) where N indicates the medium reservoir; B the bioreactor in which cells are grown; P the effluent reservoir; E the inoculation flask and M the stirring engine. Panel B, photograph of a current chemostat setup where 1 indicates the medium reservoir, 2 the bio-reactor, 3 the effluent reservoir, 4 the inoculation flask and 5 the stirring engine. Adapted from Monod (1950) with permission from Institut Pasteur.

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In chemostat cultivation, the constant ratio between effluent flow (fout) and the culture volume in the bio-reactor (V) is commonly referred to as the dilution rate D ¼ (jout/V), h1. Combined with the requirement that cx,out equals cx,bioreactor, the assumption of constant volume (V) and sterile fresh medium (cx,in ¼ 0), this becomes dðcx Þ=dt ¼ Dcx þ mcx ¼ ðm  DÞcx When the specific growth rate (m) exceeds the dilution rate (D), the biomass concentration (cx) in the bio-reactor will increase (d(cx)/dtW0). Whenever the specific growth rate is lower than the dilution rate, the biomass concentration will decrease (d(cx)/dto0). The latter situation occurs when the biomass concentration is higher than can be sustained by the inflow of fresh medium. This is, for example, the case after starting up the culture as a batch culture on a high substrate concentration. In both scenarios, the system will asymptotically approach a steady-state situation (d(cx)/dt ¼ 0; discussed in Section 2.1.2) in which the specific growth rate equals the dilution rate (m ¼ D). In a steady-state chemostat culture, the residual concentration of the growth-limiting nutrient in the extracellular medium will settle on a low (but non-zero) value that is unique for the combination of micro-organism, environmental conditions (temperature, pH, etc.) and specific growth rate. The relationship between specific growth rate and the residual concentration of the growth-limiting nutrient can often be approximated by an empirical equation proposed by Monod (1949) m ¼ mmaxcs/(cs þ Ks). In the Monod equation, Ks is the concentration of the growth-limiting nutrient at which the specific growth rate equals (1/2)mmax. The highest dilution rate in a chemostat at which a steady state can be achieved is determined by mmax, the concentration of the limiting nutrient in the medium reservoir and Ks. In most cases, the concentration of the growthlimiting nutrient in the medium reservoir exceeds Ks by at least an order of magnitude, so that the highest possible dilution rate is close to mmax. When the dilution rate exceeds this maximum value, the biomass concentration in the culture will progressively decrease, a phenomenon known as ‘‘wash-out’’. Analysis of wash-out kinetics can provide useful information on near-mmax specific growth rates (see, e.g., Flikweert et al., 1996). However, unless mutants appear with improved growth kinetics, wash-out will ultimately lead to a biologically uninspiring zero-biomass steady state. 2.1.2. Experimental Design To fully exploit the control options offered by chemostat cultivation, careful experimental design is of crucial importance. This is not a trivial statement,

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since the design of chemostat cultures encompasses at least five important decisions: (i) choice of microbial strain(s), (ii) choice of the growth-limiting nutrient and medium composition, (iii) choice of dilution rate (and thereby the specific growth rate), (iv) choice of setpoints for physical and chemical culture parameters (temperature, pH, dissolved oxygen concentration) and (v) choice of bio-reactor hardware and software for process control. The last decision does not fall within the scope of this review and, hence, is not discussed here. When chemostat cultures are used to investigate the effect of genetic modifications (e.g. gene deletion or overexpression), it is of course essential that the tested strains are congenic (e.g. share the same genetic background). Especially in the case of S. cerevisiae, several additional factors need to be taken into account. The rapid developments in yeast molecular genetics have contributed to the popularity of yeast strains and strain families that carry auxotrophic markers as model strains for laboratory research. While the presence of auxotrophic markers (i.e. requirements for uracil or specific amino acids) facilitates genetic research because of swift genetic accessibility, they complicate quantitative physiology studies (Cakar et al., 1999; Pronk, 2002). Therefore, if at all possible, yeast strains used in chemostat experiments should be prototrophic (with the exception of the vitamin auxotrophies that occur in all S. cerevisiae strains; Barnett et al., 1983). When use of auxotrophic strains is inevitable, meaningful comparison of different strains (e.g. reference strains and mutants) is only possible when they have exactly matching sets of auxotrophic markers (Cakar et al., 1999; Pronk, 2002). In experiments with auxotrophic strains, the low concentrations of uracil and amino acids that are used in common batch-cultivation media (Sherman, 1991) will often result in growth limitation by these growth factors rather than in the intended growth limitation (Pronk, 2002). The choice of a growth-limiting nutrient is a crucial determining factor in the design of chemostat experiments. Irrespective of which nutrient limitation is studied, it is crucial to confirm that the nutrient of choice really is the growth-limiting component in the medium. This can be established by reducing the concentration of the envisaged limiting nutrient (e.g. by 50%) while keeping the concentrations of all other nutrients in the medium reservoir the same. True growth limitation by a single nutrient is confirmed when, in the resulting steady-state culture, the following three parameters are identical to those in the original culture (i) the biomass yield on the chosen nutrient, (ii) the residual concentration of the growth-limiting nutrient and (iii) the (macro-)molecular composition of the biomass. A low residual concentration of a nutrient per se is not a reliable indicator for nutrient limitation as it may, alternatively, reflect accumulation of a storage

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material (resulting in a different biomass composition) or overflow metabolism. For example, under aerobic, nitrogen-limited conditions, chemostat cultures of S. cerevisiae convert a mild excess of glucose into ethanol (Larsson et al., 1993). In comparing various nutrient-limitation regimes, concentrations of the non-limiting nutrients should ideally be the same in all scenarios. For example, when ammonium- and sulfate-limited cultures are compared, flexibility in medium design may require the use of ammonium chloride and potassium sulfate rather than ammonium sulphate (see, e.g., Boer et al., 2003). The combination of independent nitrogen and sulfur sources allows the researcher to keep the concentrations of counter ions (chloride and potassium) the same under the two nutrient-limitation regimes. The specific growth rate (dilution rate) constitutes a very important parameter in the design of chemostat experiments. In aerobic, sugar-limited chemostat cultures of S. cerevisiae, the specific growth rate determines the mode of sugar catabolism. A fully respiratory metabolism is only observed at relatively low dilution rates in sugar-limited cultures (Barford and Hall, 1979; Rieger et al., 1983; van Dijken et al., 1993). Under all other nutrient limitations, aerobic sugar metabolism is respiro-fermentative. Also in aerobic glucose-limited chemostats grown at high dilution rates, S. cerevisiae displays a mixed respiro-fermentative glucose metabolism. The threshold dilution rate above which respiro-fermentative metabolism occurs in aerobic, glucose-limited cultures is strain-dependent and may vary between 0.20–0.38 h1 (Postma et al., 1989b; van Dijken et al., 2000). In addition to the mode of sugar catabolism, other cellular processes are affected by specific growth rate, including the expression of glucose transporters (Diderich et al., 1999), residual substrate concentrations (Mashego et al., 2003; Monod, 1949), protein and RNA content (Lange and Heijnen, 2001) and relative impact of free-energy use for cellular maintenance processes (Pirt, 1982). The lower limit for the dilution rate that can be applied in chemostat cultures is primarily based on a practical constraint: the time required to achieve a steady state. For example, at a dilution rate of 0.01 h1, 10 bioreactor volume changes require some six weeks of cultivation. As discussed above, the upper limit for the dilution rate is primarily determined by the maximum specific growth rate (mmax). ‘‘Optimal’’ conditions for growth of S. cerevisiae are usually defined based on mmax in batch cultures, resulting in a temperature of ca. 30 1C and a pH of 4.5–6.0. In addition to scientific interest, industrial relevance provides incentives to study yeast growth outside these ‘‘standard’’ conditions. For example, beer fermentation often involves temperatures of 10–12 1C.

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Conversely, simultaneous saccharification and fermentation (SSF) processes for fuel-ethanol production should ideally occur at temperatures considerably above 30 1C to optimize activity of saccharification enzymes. Similary, industrial applications may require pH values that are considerably lower than 5 (van Maris et al., 2004b) and oxygen-supply regimes in industry vary from virtually anaerobic (e.g. bioethanol production, beer fermentation) to fully aerobic (e.g. production of bakers’ yeast and pharmaceutical proteins). Chemostat cultivation offers the possibility to study the impact of these important process parameters, while keeping other parameters (including the specific growth rate) constant. When yeast strain(s), medium composition, dilution rate and growth conditions have been chosen, the pumps of the chemostat culture can be started. But what criteria should be used to determine whether or not a chemostat has reached ‘‘steady-state’’? Although criteria for steady state employed by different researchers may vary, they all take into account that steady-state values are approached asymptotically. Cultures are commonly assumed to be in steady state when, after a minimum of five bio-reactor volume changes since the last change in growth conditions, the biomass concentration, the concentration of the growth-limiting nutrient and key biomass-specific production and consumption rates change by less than 2% during at least one additional volume change. 2.1.3. Pitfalls While the basic experimental design of chemostat cultures outlined in the preceding paragraph seems simple, several pitfalls may compromise a welldesigned chemostat experiment. Several frequently occurring experimental problems that may preclude successful steady-state chemostat cultivation are discussed below. Ideal mixing, one of the first criteria for operating a chemostat, is relatively easy to achieve by a combination of reactor design and power input for mixing. However, many problems with chemostat cultivation can be traced back to either unintended biomass retention or selective removal of biomass. Especially in chemostats equipped with a simple overflow device, non-representative removal of biomass can be caused by sedimentation or flotation, leading to differences between the dilution rate and the specific growth rate (Noorman et al., 1991). However, even when reactors are equipped with a mass- or volume-controlled effluent system that removes culture broth from the center of the culture, biomass concentrations in samples taken from the culture should be routinely compared to those in samples taken from the effluent line. Although it is not a common

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problem in chemostat cultures of S. cerevisiae, wall growth is another phenomenon that may cause undesirable cell retention and, additionally, culture heterogeneity. Other pitfalls related to chemostat cultivation originate from supply of the growth-limiting nutrient. As discussed above, it is crucial to establish that the nutrient of choice is indeed limiting growth. Chemostat cultures of S. cerevisiae used for micro-array analysis are usually operated at biomass concentrations of several grams dry weight per liter. In such cultures the amount of the growth-limiting substrate that is consumed per minute can be several orders of magnitude higher than the amount of substrate present in the bio-reactor as residual substrate. This rapid turn-over of the growthlimiting nutrient implies that immediate quenching of metabolism is necessary for accurate measurements of residual substrate concentrations (Mashego et al., 2003; Weusthuis et al., 1994). In addition, the theory of chemostat cultivation assumes a continuous supply of fresh medium. However, in practice, the supply of medium is mostly dropwise. Especially at low dilution rates and in small bio-reactors, a low frequency of falling drops can result in a succession of ‘‘feast’’ and ‘‘famine’’ situations instead of a steady state, which may result in a different culture physiology (van Kleeff et al., 1996) and should therefore be avoided. The specific growth rate of a chemostat equals the outflow of biomass divided by the total biomass in the reactor. In an ideally mixed chemostat, with identical biomass concentrations in culture and effluent line, this equals the ratio of the effluent flow over the working volume of the bio-reactor. This value may differ significantly from the ratio of the inflow of fresh medium and the working volume of the reactor, which is often used to estimate dilution rate and, thereby, the specific growth rate. Two important factors that can contribute to such differences are addition of pH titrant and water evaporation (the latter can be minimized by cooling the off-gas with a condenser). Especially when culture volume is controlled by a level-sensor system, foaming may influence the dilution rate by causing fluctuations in the reactor volume. Antifoaming agents are therefore essential ingredients in media for cultivation of yeasts in chemostat cultures. S. cerevisiae has the unique ability to grow in the complete absence of oxygen as long as fatty acids and ergosterol are provided in the growth media (Andreasen and Stier, 1953, 1954; Visser et al., 1990). This ability is crucial for several industrial applications of S. cerevisiae, which have inspired a considerable number of anaerobic chemostat studies. In the design of such anaerobic chemostat experiments, specific care should be taken to radically minimize entry of oxygen (Visser et al., 1990). For example, silicon tubing is highly permeable for oxygen and must be replaced

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by other materials such as norprene (Visser et al., 1990). Moreover, technical-grade nitrogen, which is often used to sparge such anaerobic cultures, may contain sufficient oxygen to meet biosynthetic oxygen requirements. A specific problem associated with chemostat cultivation of S. cerevisiae is that many strains have a strong tendency to spontaneously synchronize their cell cycle in aerobic, sugar-limited chemostat cultures (von Meyenburg, 1969). This synchronization is evident from periodic oscillations of oxygen consumption and carbon-dioxide production rates, with a period close to the doubling time. Since, obviously, oscillatory behavior precludes steady-state analysis (Klevecz and Murray, 2001; Li and Klevecz, 2006) spontaneous synchronization of cultures should be avoided – unless synchronization itself is the subject of investigation (Rouwenhorst et al., 1991; Wittmann et al., 2005). Some yeast strains, such as haploid strains of the CEN.PK family (unpublished results from our laboratory) are less prone to synchronization than many other laboratory and industrial strains. Another important potential pitfall of chemostat cultivation related to micro-organisms was already described in the second chemostat publication by Novick and Szilard (1950b): ‘‘It may be said that our strain, if grown in the Chemostat at low tryptophan concentration for a long period of time, undergoes a number of mutational steps, each leading to a strain more ‘fit’ than the previous oney.’’ Evolution may represent a source of undesirable – and even unacceptable – biological variation in experiments aimed at quantitative physiological analysis of different strains, mutants or conditions, especially when such analysis involves sensitive high-information-density tools such as micro-arrays. Therefore, in chemostat-based micro-array experiments, it is strongly recommended to start fresh runs for each condition tested and to limit the length of chemostat runs to fewer than 20 generation times (ca. 14 volume changes). As will be discussed in Section 6, evolution in chemostat cultures not only presents experimental complications, but also opens up unique possibilities for studying evolution and for directed selection of strains with industrially relevant properties.

2.2. Micro-Array Technology Several methodologies are available for measuring mRNA (transcript) levels in microbial cultures, including PCR-based analysis (RT-PCR), sequencingbased serial analysis of gene expression (SAGE; Velculescu et al., 1995) and hybridization-based approaches. The latter include classical Northern analysis (Kroczek, 1993), macro-arrays on filters (Gress et al., 1992) and

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micro-arrays. The unique feature of micro-arrays is that the abundance of thousands of transcripts can be quantitatively analyzed on a single array. This high information density enables genome-wide investigations into genetic interactions, functional analysis and dissection of regulatory networks. The first genome-wide transcript analyses on S. cerevisiae were published only a year after the release of the genome sequence (DeRisi et al., 1997; Lashkari et al., 1997). Micro-arrays have since rapidly become established as a technology standard for genome-wide transcriptome analysis in organisms whose genome has been completely sequenced. Several micro-array techniques are now in use, ranging from genomic libraries or full-length cDNAs spotted on glass slides to oligonucleotide micro-arrays. Because of their higher combined specificity and sensitivity, oligonucleotide micro-arrays have become the standard in quantitative transcriptomics. The two most commonly used approaches for oligonucleotide micro-array analysis can be distinguished based on the number of dyes (fluorescent probes) used for detection. Some technological aspects of oligonucleotide micro-array analysis will be briefly discussed below (Fig. 3). For more detailed information, the reader is referred to Dalma-Weiszhausz et al. (2006) and Wolber et al. (2006). In experiments with double-dye arrays, mRNA from two different biological samples is reverse transcribed into cDNA and fluorescently labeled with either Cy3 (green) or Cy5 (red) dye. The two differently labeled cDNA pools are then mixed and hybridized to oligonucleotide fragments of known sequence that have been spotted on a glass slide. While firstgeneration spotted arrays used full-length cDNAs, the probe-length on glass-slide arrays is now commonly shortened to 60-mer oligonucleotides (‘‘Yeast oligo-micro-array,’’ AgilentTM Technology). Because of limited information density on glass-slide arrays, the average number of oligonucleotides by which each gene is represented on the array is usually below 2. The double-dye approach allows measurement of relative abundance of sequences in the two cDNA samples, which can be expressed as gene intensity ratios. Cy3/Cy5 micro-arrays enable direct comparison of only two conditions in each experimental context. This constraint complicates experimental design and increases the cost of quantitative comparisons that involve three or more conditions (Bowtell, 1999). Single-dye DNA-micro-array technology is epitomized by Affymetrix arrays. High-density Affymetrix Genechipss are produced by photolithographic synthesis on a silica substrate (Dalma-Weiszhausz et al., 2006). Transcript levels of yeast genes are analyzed comparing hybridization of cRNA to an array-borne probe set comprising 20–40 gene-specific, 25-mer oligonucleotides for each yeast gene. Half of these probes are

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Figure 3 Diagram of typical single-dye (A) and double-dye (B) micro-array experiments. 1, total RNA extraction from biomass samples. 2, cDNA synthesis. 3, in vitro transcription (cRNA synthesis). 4, metal-induced cRNA fragmentation. 5, hybridization of the prepared targets. 6, staining with streptavidin-phycoerythrin. 7, washing. 8, scanning. 9, mixing Cy3, Cy5-labeled cDNA samples. (See plate 1 in the color plate section.)

‘‘perfect match’’ probes that correspond exactly to the gene sequence. The remaining ‘‘mismatch’’ probes carry a substitution of the central nucleotide (13th) of the oligomer, thus strongly reducing specific binding of cRNA for the corresponding gene and providing an internal control for non-specific binding. In contrast to the approach with double-dye arrays, a single sample is hybridized on each array. The typical experimental output for each gene consists of a number that represents hybridization intensity. Currently, two S. cerevisiae Genechipss are commercially available. The Yeast genome S98 (YGS98) array is based on the December 1998 version of the Saccharomyces genome database (SGD), while the Yeast Genome 2.0 (YG2) micro-array is based on a May 2004 annotation. The main difference between these micro-arrays is that 500 fewer ORFS are represented on the YG2 micro-arrays, following the removal of these ORFs from the yeast gene directory based on a comparative genome

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analysis of several Saccharomyces genomes (Cliften et al., 2003; Kellis et al., 2003). Recent manufacturing developments have enabled an increase in the information density to over 3.106 probes per 1 cm2 micro-array. This has enabled development of ‘‘tiling’’ arrays on which the entire genome is represented by overlapping (tiling) probes (Gresham et al., 2006). For example, on Affymetrix GeneChips Tiling 1.0R arrays for S. cerevisiae, 25-mer oligonucleotide probes are used with a sequence overlap of 20 base pairs (bp), thus resulting in coverage of the entire genome with a 5 bp resolution. In addition to transcriptome analysis, applications of tiling arrays include discovery of hitherto unknown transcripts (David et al., 2006; Juneau et al., 2007; Zhang et al., 2007), mapping of single-nucleotide polymorphisms (Gresham et al., 2006) and mapping of sites for protein/ DNA interaction in chromatin immunoprecipitation (ChIP) experiments (Negre et al., 2006). Commercial single-dye micro-arrays are expensive in terms of design costs and costs for consumables as compared to dual-dye glass-slide arrays. Despite their accuracy and flexibility, this may limit their spread in academic yeast research. Unfortunately, the different properties of the applied fluorescent dyes, the hybridization specificity and sensitivity make it practically impossible to compare transcript data obtained with different array platforms (Bammler et al., 2005; Ja¨rvinen et al., 2004; Tan et al., 2003). An illustrative example is provided by transcriptome studies on sulfur-limited growth of S. cerevisiae. Transcriptome data on sulfur-limited S. cerevisiae chemostat cultures were reported in three independent studies by Boer et al. (2003), Castrillo et al. (2007) and Wu et al. (2004). Based on at least triplicate YGS98 (single-colour) array data, Boer et al. (2003) and Castrillo et al. (2007) concluded that over 85% of the genome was transcribed in sulfur-limited cultures. Using custom-made cDNA macroarrays, Wu et al. (2004) concluded that, under the same nutrient-limitation regime, less than 30% of the genome was transcribed.

2.3. Reproducibility of Chemostat-Based Micro-Array Analysis Already in the first publication on chemostat-based transcriptome analysis (ter Linde et al., 1999), the reproducible, controlled conditions in chemostat cultures were cited as an important advantage as compared to microarray studies in batch cultures. This assumption was systematically tested in an inter-laboratory study on the reproducibility of chemostat-based

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micro-array analysis (Piper et al., 2002). In this study, two independent research groups used the same S. cerevisiae strain, chemostat equipment, synthetic media and micro-array platform (Affymetrix S98 GeneChip) to perform a transcriptome analysis on aerobic and anaerobic, glucose-limited chemostat cultures. The cumulative experimental variation introduced by the in vitro handling steps from RNA extraction to micro-array analysis was found to be small and unbiased, implying it can be eliminated by replication of experiments. These findings were in agreement with the conclusion of Baugh et al. (2001) that a single round of amplification and labeling by in vitro transcription merely caused low, unbiased variation. In our experience, introduction of routine, automated checks for sample quality (i.e. integrity, size distribution and amount of RNA, cDNA and cRNA) at all stages of nucleic-acid processing helps to minimize variation among replicate analyses and, importantly, reduces costs due to failed micro-array hybridizations. In the study by Piper et al. (2002), each research group analyzed independent triplicate cultures for aerobic and anaerobic, glucose-limited chemostat cultures. A comparison of the experimental variation of the chemostat-based experiments with literature data sets on shake-flask culture confirmed the expectation that chemostat cultivation resulted in much higher reproducibility (Fig. 4). Using appropriate statistical analysis, the change calls from anaerobic versus anaerobic comparisons yielded over 95% agreement between the two laboratories. The general conclusion was that analysis of samples from independent triplicate cultures enabled statistically sound difference calls for genes that exhibited a fold difference of two and higher. Provided that strains, cultivation conditions and (singledye) micro-array protocols are rigorously standardized, this result indicates that chemostat-based micro-array databases can be constructed based on experiments from different laboratories (Bammler et al., 2005; Piper et al., 2002). At a fixed dilution rate, the total amount of mRNA within the cell can be assumed to remain approximately constant. In this case, relatively simple global fluorescence intensity scaling of the micro-arrays is acceptable as a normalization procedure. However, when cultures with different (or varying) specific growth rates are being compared, the total mRNA pool is unlikely to remain constant in both size and content (Regenberg et al., 2006; van de Peppel et al., 2003). To ensure the retrieval of biologically relevant information from such chemostats, the systematic addition of external mRNA-spiking controls is essential and should preferentially become a standard procedure in chemostat-based transcriptome data analysis.

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Figure 4 Comparison of the variation of micro-array-derived transcript levels in independent replicates of chemostat cultures and shake-flask cultures. Independent triplicate aerobic glucose-limited chemostat cultures were run in two laboratories (A, blue line and B, red line; Piper et al., 2002) and compared with shake-flask data (green line; Holstege et al., 1998) (the data can be retrieved from http:// www.wi.mit.edu/young/expression.html). The coefficient of variation (standard deviation divided by the mean) was calculated for each transcript and plotted as a function of increasing average transcript abundance. A trend line (of the coefficient of variation) was generated using the average from a moving window of 50 transcripts and overlaid on each scatter plot. The gene numbers (from 1 to 6383) on the x-axe were generated by ranking the genes by increasing average transcript level. (See plate 2 in the color plate section.)

3. SPECIFIC GROWTH RATE 3.1. Specific Growth Rate and Transcriptional Regulation The vast majority of published yeast micro-array studies on S. cerevisiae have been performed with shake-flask cultures. When shake flasks or other batch-cultivation methods are used to study the effects of genetic mutations or changes in cultivation conditions, these changes are often accompanied by changes in the specific growth rate. As will be discussed below, such inherent changes of the specific growth rate may hinder the interpretation of transcriptome data from batch cultures. Already before the advent of genome-wide analyses, it was established that transcription of certain S. cerevisiae genes is strongly dependent on the specific growth rate. A well-documented example concerns the expression of ribosomal protein genes, which shows a strong positive correlation

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with specific growth rate (Kraakman et al., 1993; Mager and Planta, 1991; Planta and Mager, 1998). This, however, does not imply that decreased expression of ribosomal-protein-encoding genes is always indicative for a decreased specific growth rate. Indeed, factors other than specific growth rate have been shown to influence expression of these genes in wild-type S. cerevisiae strains (Saldanha et al., 2004). The tight correlation between specific growth rate and transcription of genes involved in protein synthesis dramatically complicates the interpretation of batch-culture transcriptome data obtained with mutants affected in protein synthesis or its regulation. This complexity is exemplified by studies on transcriptional regulation of genes involved in ribosomalprotein biosynthesis (RP genes) and ribosome biogenesis (RiBi genes) in S. cerevisiae. Based on batch-culture transcriptome analysis of wild-type S. cerevisiae and sfp1 strains, Jorgensen et al. (2004) proposed that the transcriptional regulator Sfp1p activates both RP and RiBi genes. However, sfp1 mutants exhibit a much lower mmax than wild-type S. cerevisiae strains and their study did not reckon with the possibility that the difference in transcript profiles for the RP and RiBi genes in wild-type and mutant strains was caused by a difference in specific growth rate, rather than by a direct role of Sfp1 in their transcriptional regulation. Indeed, careful comparison of transcriptome data of a wild-type S. cerevisiae strain and its sfp1 derivative in glucose-limited chemostat cultures, in which the specific growth rate for the two strains was kept at an identical value (0.025 h1), confirmed involvement of Sfp1p in transcriptional regulation of RiBi genes, but not of the RP genes (Cipollina et al., 2008). The results of this chemostat study are supported by other expression data sets and by the promoter structure of the RP genes (Rudra et al., 2005; Saldanha et al., 2004). This Sfp1p example illustrates how chemostat cultivation enables the elimination of specific growth rate as a variable in comparative analysis of strains or growth conditions. The unique ability to precisely control specific growth rate at any desired value between 0 and a value close to mmax makes the chemostat an excellent tool to quantify the effects of specific growth rate on genome-wide transcriptional regulation. This possibility has been explored in two recent studies. Regenberg et al. (2007) studied yeast transcriptional responses to a wide range of specific growth rates ranging from 0.02 h1 to 0.33 h1 (corresponding to generation times ranging from 2 h to 35 h) in aerobic, glucose-limited chemostat cultures. Castrillo et al. (2007) investigated a more narrow range of specific growth rates (0.07 h1, 0.1 h1 and 0.2 h1), but included several nutrient-limitation regimes (aerobic cultures in which growth was limited by glucose, ammonium, phosphate or sulfate). As will be discussed in Section 4, this approach allowed a ‘‘filtering out’’ of

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responses that are specifically linked to the identity or concentration of the growth-limiting nutrient. Both studies demonstrated that the impact of specific growth rate on transcriptional regulation extends well beyond ribosomal-protein genes and involves clear effects on the synthesis of macromolecular components (proteins, nucleic acids, lipids). Another important class of genes whose transcription is correlated with specific growth rate is discussed in the next section.

3.2. Specific Growth Rate and Environmental Stress Responses Transcriptome studies in batch cultures of S. cerevisiae that were exposed to a large variety of chemical and physical stresses resulted in the identification of a large set of genes that exhibited a common up-regulation in response to a multitude of stress factors. Based on these observations, these genes were called environmental stress response (ESR) genes (Causton et al., 2001; Gasch et al., 2000; Gasch and Werner-Washburne, 2002). In chemostat cultures grown at various specific growth rates, transcript levels of the majority of ESR genes exhibited a strong negative correlation with specific growth rate (Castrillo et al., 2007; Regenberg et al., 2006). Regenberg et al. (2007) reported that as many as 89% of the ESR genes reported by Gasch et al. (2000) were transcriptionally up-regulated at low specific growth rates in chemostat cultures. Interpretation of these results appears straightforward: since environmental stress will generally result in a reduction of mmax, it may be expected that specific-growth-rate-responsive transcripts will be identified in transcriptome studies on environmental stress in batch cultures. In contrast, at a fixed specific growth rate in chemostat cultures, such transcript would not necessarily be expected to show a transcriptional response when the same stresses are applied. This hypothesis was recently tested in studies on low-temperature responses of S. cerevisiae. In addition to being of academic interest, yeast physiology at low temperature is relevant for several industrial applications (e.g. brewing and wine making). Genome-wide transcriptional responses of S. cerevisiae to low temperature have been extensively studied in batch cultures (Murata et al., 2006; Sahara et al., 2002; Schade et al., 2004). These experiments mostly focused on the transcriptional responses that occur upon cold shock, that is, a sudden decrease of the cultivation temperature. Since cold shock results in a rapid decrease of the specific growth rate, a substantial fraction of the response originally attributed to cold shock may in fact be caused by the concomitant decrease in specific growth rate. Indeed, in an analysis of

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temperature-dependent transcriptional regulation (Tai et al., 2007b) at least 15% of the genes responding to low temperature in batch-culture data sets (Murata et al., 2006; Sahara et al., 2002; Schade et al., 2004) were found to exhibit specific-growth-rate-dependent transcript levels in chemostat studies performed at 30 1C (Regenberg et al., 2006). Conversely, when S. cerevisiae was grown at 30 1C and 12 1C in steady-state, glucose-limited chemostats at a fixed specific growth rate of 0.03 h1 (Tai et al., 2007b), less than 1% of the low temperature-responsive genes overlapped with the previously identified sets of growth-rate responsive transcripts (Regenberg et al., 2006). Obviously, elimination of growth-rate-dependent effects is not just relevant in studies on the transcriptional effects of environmental parameters, but also in the characterization of mutants that have a decreased specific growth rate relative to the wild type. For the analysis of chemical stress factors, chemostats also enable variation of the degree of stress that yeast cultures experience at a fixed specific growth rate. This option was explored by Abbott et al. (2007) in a study of the transcriptional responses to various organic acids (acetate, benzoate, propionate and sorbate) in anaerobic, glucose-limited chemostats. The different organic acids were added to the reservoir medium at concentrations that resulted in the same relative decrease of the biomass yield on glucose for each organic acid. Through transcriptome analysis, performed at the same specific growth rate and at the same degree of uncoupling of growth and glucose consumption for each organic acid, a clear distinction was observed between (rare) generic effects of organic acids and (abundant) transcriptional responses to these four organic acids (Abbott et al., 2007).

4. CONTEXT DEPENDENCY In its most basic and most commonly applied form, transcriptome analysis involves single, pairwise comparisons of two experimental conditions or strains, in which one of the situations serves as the reference. After application of appropriate statistical analysis, this approach results in sets of genes whose transcript levels are higher or lower than in the reference condition (often loosely referred to as ‘‘up-regulated’’ or ‘‘down-regulated,’’ respectively). This approach does not take into account the fact that many genes are controlled by multiple regulatory systems acting in a hierarchical manner. As a consequence, transcriptional responses to any environmental or genetic change may be strongly dependent on additional environmental parameters (i.e. on the experimental context).

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Controlled variation of individual cultivation parameters in chemostat cultures enables a quantitative assessment of the importance of context dependency. This possibility was explored with a study on combinatorial effects of oxygen availability and nutrient-limitation regimes by Tai et al. (2005), which was inspired by earlier chemostat-based transcriptome studies. Previously, ter Linde et al. (1999) and Piper et al. (2002) compared the transcriptomes of aerobic and anaerobic chemostat cultures of S. cerevisiae grown on glucose as the limiting nutrient, while Boer et al. (2003) analyzed transcriptomes of the same S. cerevisiae strain in aerobic chemostat cultures in which growth was limited by glucose, ammonium, phosphate or sulfate. To investigate the extent to which responses to different macro-nutrient limitations were affected by oxygen availability and vice versa, Tai et al. (2005) performed an additional transcriptome analysis on anaerobic chemostat cultures grown under ammonium, phosphate and sulfate limitation. Analysis of the resulting data set, based on 24 independent chemostat cultures comprising eight different growth conditions (four nutrient-limitation regimes, each analyzed in triplicate aerobic and anaerobic chemostat cultures), revealed that responses to nutrient limitation and oxygen availability were strongly interdependent (Fig. 5). Via combinatorial design of chemostat-based micro-array studies as shown in Fig. 5 (Tai et al., 2005), robust ‘‘core’’ sets of genes can be identified that show a consistent response to a specific environmental stimulus (or genetic intervention) in multiple experimental contexts. These gene sets are then expected to be enriched for genes (pre)dominantly regulated by transcriptional regulation pathways that are directly related to a single environmental stimulus. Indeed, gene sets exhibiting a consistent response to oxygen availability under all four nutrient-limitation regimes showed a much stronger over-representation of promoter-binding sites for anaerobiosis-related transcription factors than gene sets whose response to oxygen availability depended on the macro-nutrient-limitation regime. Conversely and consistently, gene sets that, irrespective of oxygen availability, showed a consistent response to each of the four nutrientlimitation regimes showed a strong over-representation of specific nutrientdependent transcription factor-binding sites in their promoter regions (Tai et al., 2005). Identification of genes that show a robust, contextindependent response to a single environmental parameter is helpful for transcript-based process diagnostics, since the identified transcripts are ideally suited to monitor a single process parameter in complex cultivation systems. The question of whether a robust ‘‘context-independent’’ (obviously, it is impossible to experimentally test all possible environmental contexts) up-regulation of a gene, in response to a defined set of culture

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Figure 5 Context dependency of transcriptional responses in S. cerevisiae: (A) Experimental design for two-dimensional transcriptome analysis. Each corner of the cube represents a unique chemostat cultivation regime. The upper horizontal surface represents four aerobic macro-nutrient-limitation regimes (carbon, nitrogen, phosphorus and sulfur). The lower horizontal surface represents the same macronutrient-limitation regimes analyzed under anaerobic conditions. The arrows indicate the pair-wise comparisons included in the two-dimensional transcriptome analysis. (B) Venn diagram of signature anaerobic genes. Red and green represent up-regulation and down-regulation, respectively, under anaerobic conditions. Each of the four circles corresponds to a cluster of genes that showed a transcriptional response to oxygen availability under one of the four macro-nutrient-limitation regimes. The overlap of the four clusters represents genes that showed a consistent response to oxygen availability irrespective of the nutrient limitation regime. Lim Ae, limited aerobic; Lim Anae, limited anaerobic. (C) Signature genes that consistently respond to anaerobic conditions. Figure adapted from Tai et al., 2005 with permission from Elsevier. (See plate 3 in the color plate section.)

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conditions, has a predictive value for its contribution to cellular fitness under those conditions will be addressed in Section 5.4. Context dependency has a major impact on the experimental design of chemostat-based transcriptome studies. Even in tightly controlled chemostat cultures, it is impossible to change a single cultivation parameter without any impact on other parameters. This is exemplified by the chemostat studies of Tai et al. (2007b) on growth of S. cerevisiae at 12 1C and 30 1C (see previous paragraph). Growth at both temperatures was studied at the same specific growth rate of 0.03 h1. This specific growth rate corresponds to ca. 75% of mmax at 12 1C, but to only 10% of mmax at 30 1C. This difference in the ‘‘relative specific growth rate’’ between these two cultivation temperatures may affect the residual concentration of the growth-limiting nutrient in the chemostat cultures. Indeed, consistent with Monod-type growth kinetics (Monod, 1949; see Section 2.1.1), the residual glucose concentration in cultures grown at 12 1C was clearly higher than at 30 1C. Consequently, gene sets whose transcript levels were temperature-dependent in glucose-limited cultures were ‘‘contaminated’’ with genes whose transcription is regulated by glucose availability (e.g. via glucose catabolite repression; Gancedo, 1998). By analyzing nitrogen-limited (i.e. glucose-excess) cultures at the same temperatures, it was possible to filter out such specific effects of the growth-limiting nutrients. A similar approach was followed in a recent analysis of the effects of specific growth rate on multi-level gene expression in chemostats of S. cerevisiae (Castrillo et al., 2007). The strong context dependency of transcriptional responses is likely to be highly relevant in nature as well as in industrial environments. For example, a large difference in transcriptional responses of aerobic and anaerobic chemostat cultures of S. cerevisiae to zinc limitation was partly attributed to the role of this element in mitochondrial activity and protection against reactive oxygen species (de Nicola et al., 2007). Furthermore, combinatorial analysis of transcriptional responses to various environmental stimuli presents an interesting and promising tool for in vivo analysis of the structure and hierarchy of transcriptional regulation networks (Knijnenburg et al., 2007).

5. FUNCTIONAL ANALYSIS 5.1. Assignment of Gene Function More than a decade after the completion of the genome sequence of S. cerevisiae (Goffeau et al., 1996), the biological function of at least 21% of

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its genes remains unknown (Pen˜a-Castillo and Hughes, 2007). This percentage probably grossly overestimates our current knowledge, as the number of yeast genes encoding proteins with multiple, seemingly unrelated functions, continues to grow. Correlations between environmental conditions and gene expression can be investigated through micro-array analysis. This may indicate a physiological role for gene products and, thus, guide functional analysis studies. Since transcript profiling only leads to the hypotheses and not to evidence, follow-up research (e.g. via gene deletion studies and biochemical characterization of the encoded proteins) will always be essential to conclusively establish protein function. A number of recent studies have successfully applied the experimental design options offered by chemostat-based micro-array analysis to formulate hypotheses on gene function. The Ehrlich pathway for amino acid catabolism plays an important role in the production of flavor precursors in beer and wine fermentation. Following transamination of aromatic, branched-chain or sulfur-containing amino acids, the resulting 2-oxo acid is decarboxylated, yielding an aldehyde. Depending on cultivation conditions, this aldehyde will either be reduced to an alcohol (‘‘fusel alcohol’’) or oxidized to an acid (‘‘fusel acid’’). Although this short catabolic pathway has been known for a full century (Ehrlich, 1907), its molecular biology remains to be fully elucidated. Boer et al. (2007) studied growth of S. cerevisiae on six different nitrogen sources (ammonia, asparagine, proline, phenylalanine, leucine and methionine) in aerobic, glucose-limited chemostat cultures. Cultures grown on phenylalanine, leucine and methionine produced substantial amounts of the ‘‘fusel acids’’ phenylacetate, 2-methyl propanoate and 3-methyl-butanoate, whereas the cultures grown on ammonia, proline or asparagine did not. A set of 33 genes showing higher transcript levels specifically in phenylalanine, leucine and methionine grown cultures included ARO10, which has strong sequence similarity with thiamine-pyrophosphate decarboxylase genes (Dickinson et al., 2003; Vuralhan et al., 2003), and PDR12, encoding a well-characterized ATP-Binding-Cassette (ABC) transporter involved in export of weak-organic acid preservatives (Holyoak et al., 1999). Subsequent functional analysis confirmed that the Aro10p protein is a broad-substrate-specificity 2-oxo acid decarboxylase involved in the Ehrlich pathway (Vuralhan et al., 2005). Moreover, Pdr12p was shown to catalyze export of branched-chain and aromatic ‘‘fusel acids’’ generated by amino acid catabolism. This provided a plausible physiological function for this transporter in the yeast’s natural environments (Hazelwood et al., 2006). 5-hydroxymethyl furfural (5-HMF) is an important inhibitor of yeast metabolism and growth in plant biomass hydrolysates. Identification of

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yeast genes involved in 5-HMF tolerance is, therefore, highly relevant for yeast-based production of ethanol from these feedstocks (Ulbricht et al., 1984). Petersson et al. (2006) combined several S. cerevisiae strains and cultivation conditions in a chemostat-based micro-array analysis to identify candidate genes for yeast 5-HMF reductase. Two strains that differed with respect to their intrinsic 5-HMF tolerance were grown in anaerobic, glucoselimited chemostat cultures in the presence and absence of 5-HMF. A total of 15 oxido-reductase-encoding genes that showed at least twofold higher transcript levels in cultures of the 5-HMF tolerant strain were overexpressed, followed by enzymatic analysis of 5-HMF reductase activity. This work has led to the identification of Adh6p as a highly active 5-HMF reductase (Petersson et al., 2006). Growth of micro-organisms at constant, controlled concentrations of dissolved gases is much more straightforward in chemostat cultures than in other cultivation systems. Hence, chemostat cultivation is the method of choice for identification of genes that respond to changes in the dissolved concentrations of oxygen or carbon dioxide (pCO2). Aguilera et al. studied transcript profiles of glucose-limited chemostat cultures grown at pCO2 values of o1 to 100%. Transcript levels of NCE103 which encodes a carbonic anhydrase, showed strong negative correlation with pCO2 (Aguilera et al., 2005a). The role of carbonic anhydrase in yeast was poorly understood. The transcriptome study led to the hypothesis that, at low ambient carbon dioxide concentrations, Nce103p is essential for providing the bicarbonate needed in three bicarbonate-dependent biosynthetic carboxylation reactions, catalyzed by pyruvate carboxylase, acetyl-CoA carboxylase and carbamoyl phosphate synthase. This hypothesis was subsequently confirmed in a physiological study on nce103 null mutants (Aguilera et al., 2005b). These examples illustrate that chemostat-based micro-array studies can be specifically designed to address gene functional analysis. We anticipate that, as the number of chemostat-based micro-array studies in S. cerevisiae increases, systematic exploration of the resulting data sets will continue to generate interesting leads for further functional analysis.

5.2. Analysis of Transcriptional Regulation Networks Understanding the structure and coordinated behavior of gene expression networks is fundamental to systems biology approaches (see also Section 7). S. cerevisiae has proved to be an excellent model for elucidating eukaryotic transcriptional regulation networks. Chemostat cultures present various

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advantages (see previous sections) in the study of function and activity of transcriptional regulators but, surprisingly, have thus far been grossly under-used for this type of analysis. Only three chemostat-based studies have been published to elucidate transcription factor activities of Leu3p (Boer et al., 2005), Hap4p (Raghevendran et al., 2006), Rox1p and Hap1p (ter Linde and Steensma, 2002). Transcriptional regulation of branched-chain amino-acid metabolism in S. cerevisiae involves two key regulator proteins, Leu3p and Gcn4p. Leu3p is a pathway-specific transcriptional regulator, regulating genes involved in branched-chain amino-acid metabolism and nitrogen assimilation. The activity of Leu3p is modulated by a-IPM, an intermediate of the branched-chain amino acid pathway. Leu3p binds to the DNA irrespective of the presence or absence of a-IPM (Friden and Schimmel, 1987; Kirkpatrick and Schimmel, 1995; Wade and Jaehning, 1996). Genome-wide expression analysis of carbon-limited chemostats yielded a robust set of Leu3p-regulated genes. These included previously characterized Leu3p targets (LEU1, LEU2, LEU4, ILV2, ILV5, BAP2) as well as three new candidate targets (GAT1, OAC1 and BAT1) (Boer et al., 2005). Meanwhile, the transcriptome analysis of the leu3D mutant revealed a clear interplay between Leu3p and Gcn4p regulatory networks under nitrogen limitation. In this condition ILV2 and ILV5, two known Leu3p and Gcn4p targets, were significantly up-regulated in the leu3D mutant. Over a hundred putative Gcn4p target genes showed a similar transcriptional upregulation. This example illustrates the large impact of growth conditions in combination with leu3 deletion and indicates how a two-way transcriptome comparison can lead to the identification of sets of genes that specifically respond to a single genetic or environmental change (Boer et al., 2005). As already mentioned in Section 4, combinatorial analysis of genomewide expression profiles to various environmental stimuli is very important. Knijnenburg et al. (2007) showed that incorporating the actual growth conditions in inferring regulatory relationships provides detailed insight into the functionality of the transcription factors that are triggered by changes in the employed cultivation conditions. Integration and comparison of transcriptome data from eight different chemostat culture conditions with large-scale genome-wide chromatin localization data set (Harbison et al., 2004; Lee et al., 2002) revealed the presence of a complicated growthcondition-dependent transcriptional network (Knijnenburg et al., 2007). Although genome-wide chromatin localization data were not derived from chemostat cultures, several interesting correlations were found between gene expression and transcription factor activity.

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Accurate and robust resolution of a regulatory network should ideally encompass the simultaneous recording of genome-wide transcriptional expression as well as localization data under a single growth condition. Indeed, a recent investigation of the transcriptional responses to fatty acids in yeast shows the efficiency of this approach to define the structure of the transcription regulatory network (Smith et al., 2007). No such integral data sets have hitherto been reported for chemostat cultures.

5.3. Correlation of mRNA Levels and Metabolic Fluxes In the interpretation of transcriptome data, it is often tacitly assumed that a protein makes a positive contribution to cellular function and fitness under conditions where its corresponding mRNA is up-regulated. As illustrated in the previous paragraph, this concept of ‘‘transcriptome-inferred function’’ has generated useful leads for more detailed functional analysis. However, this does not mean that it is generally applicable. In attempts to correlate mRNA profiles with the physiological role of the encoded proteins, a distinction can be made between the in vivo metabolic flux catalyzed by the protein and, more generally, the contribution of the encoded protein to cellular fitness. While the former is only applicable to proteins with a known catalytic function (i.e. enzymes, discussed in this section) the latter can be quantified for any gene or encoded protein, even if its biochemical function is unknown (see Section 5.4). In batch cultures, concentrations of mRNAs, proteins and fluxes continuously change with different time constants. Hence, it is extremely difficult to accurately quantify correlations between transcript levels and in vivo enzyme activity in, for example, shake-flask cultures. In contrast, steady-state chemostat cultivation makes such a direct comparison relatively straightforward. Using chemostat cultures of S. cerevisiae in which growth was limited by different carbon sources (glucose, maltose, acetate or ethanol), DaranLapujade et al. (2004) investigated the correlation between in vivo fluxes through enzymes in central carbon metabolism and the levels of the corresponding transcripts. The extent to which mRNA levels correlated with the in vivo activity of the encoded enzymes strongly depended on the metabolic pathway that was investigated. Transcripts encoding enzymes involved in utilization of 2-carbon substrates (gluconeogenesis and glyoxylate cycle) showed a good correlation with corresponding in vivo fluxes. However, such a correlation was not observed for other important pathways, including the pentose-phosphate pathway, TCA cycle and glycolysis. For these pathways, large differences in the in vivo activities of

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key enzymes between the different growth conditions were not or hardly reflected by changes in the corresponding mRNA levels (Fig. 6). These results indicate that in vivo fluxes in the central carbon metabolism of S. cerevisiae can be controlled to a large extent via post-transcriptional mechanisms (i.e. translational efficiency, post-translational modifications, (in)activation of enzymes by metabolites, etc.). Application of chemostat cultures to quantify the contributions of these post-transcriptional regulation mechanisms is discussed in Section 7.

Figure 6 Evidence for post-transcriptional regulation of the glycolytic fluxes in S. cerevisiae grown in chemostat. S. cerevisiae was grown in aerobic and anaerobic, glucose-limited chemostat cultures (dilution rate: 0.10 h1) with or without addition of benzoic acid. For each condition, in vivo fluxes were estimated using a stoichiometric model and transcript levels were quantified with microarrays. For each reaction step, the ratio in in vivo flux between anaerobic and aerobic cultures or anaerobic cultures without and with benzoic acid cultures is plotted against the ratio in transcript levels (hybridization intensity of signals for transcripts that encode isozymes of glycolytic enzymes have been pooled). Data recalculated from DaranLapujade et al. (2007).

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Studies on cultures whose growth is limited by macro-elements such as nitrogen, sulfur and phosphate provide another clear illustration that transcript levels for enzymes in the S. cerevisiae metabolic network cannot be considered as reliable ‘‘flux indicators.’’ When growth in a chemostat is limited by a particular macro-element, this typically leads to the transcriptional up-regulation of pathways for uptake and metabolism of alternative chemical forms of that element. For example, ammonium-limited growth leads to a concerted transcriptional up-regulation of pathways for uptake and assimilation of alternative, organic nitrogen sources (e.g. the allantoate permease gene DAL5, the proline transporter gene PUT4 and the urea and polyamine permease gene DUR3) – even if these are not available in the culture medium (Boer et al., 2003).

5.4. Correlation of Transcript Levels and Fitness of Null Mutants The relative fitness of yeast mutants can be quantified by monitoring the abundance of tagged deletion mutants in competitive cultivation experiments (Baganz et al., 1998; Birrell et al., 2002; Giaever et al., 2002; Tai et al., 2007c; Winzeler et al., 1999). While competition experiments can be performed in batch cultures, the constant growth conditions in chemostat cultures enable a more accurate quantification of the results. In chemostat competition experiments, mutations that have a negative effect on either the maximum specific growth rate mmax or on the substrate saturation constant Ks will directly affect fitness because they lead to a lower affinity (mmax/Ks) for the growth-limiting nutrient (Button, 1991; Monod, 1942). Competitive chemostat cultivation has been used extensively in microbial ecology to investigate competition of mixed microbial populations for a common growth-limiting nutrient (Harder et al., 1977; Kuenen et al., 1982; Postma et al., 1989a). Its application to quantify the fitness of yeast deletion mutants was pioneered by Baganz et al. (1997, 1998) and slightly modified by Tai et al. (2007c). Monitoring of population dynamics in mixed cultures of yeast mutants is greatly facilitated by the use of deletion cassettes that carry molecular bar codes (Shoemaker et al., 1996), whose abundance can be monitored by quantitative RT-PCR. The high degree of sensitivity obtained with this method is crucial since the relative abundance of the individual mutants should ideally remain low to minimize the impact of cellular interactions between different mutants (e.g. nutritional complementation via cross-feeding; Pronk, 2002). In their pioneering experiments, Baganz et al. (1998) inoculated tagged deletion mutants at the start-up of the chemostat. The cultivation conditions

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during start-up of a chemostat are dynamic and the selective pressure is likely to differ from that under steady-state conditions. To circumvent this problem, Tai et al. (2007c) first grew the reference strain to steady state, followed by injection of a mixture of tagged deletion strains into the chemostat. To investigate the correlation between transcript levels and fitness of the corresponding null mutants, Tai et al. (2007c) focused on a set of 24 S. cerevisiae genes that, irrespective of the nutrient limitation (glucose, ammonium, sulfate or phosphate), showed a strong and consistent transcriptional upregulation in anaerobic chemostat cultures (see Section 4, Tai et al., 2005), but whose function or role under anaerobic conditions was unknown. Only 5 of the corresponding 24 deletion mutants showed a significant (W20%) decrease of fitness during anaerobic competitive chemostat cultivation (Fig. 7). This result indicated that, even for this set of 24 genes

Figure 7 Competitive chemostat cultivation for fitness analysis. A pool of tagged deletion mutants was injected into a steady-state anaerobic chemostat culture of a reference S. cerevisiae strain. Subsequently, abundance of the oligonucleotide tags of each of the mutants was monitored by quantitative PCR and fitness was plotted as the log ratio [DC(t)mutant/DC(t)reference] as a function of time. Graph areas (Roman numerals) indicate the following reductions of fitness relative to the reference strain: (I) o20%, (II) 20–30%, (III) 30–40%, (IV) 40–50%, (V) W50%. The dashed line denotes the theoretical wash-out curve (zero specific growth rate). Error bars indicate mean 7SD of two independent chemostat cultures with triplicate measurements , for each time point. ’, ura3 ; &, ylr413w ; K, izh2 ; , yor012w ; eug1 ; W, plb2 . Figure adapted from Tai et al. (2007c) with permission from Society for General Microbiology.

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that did not exhibit context dependency of their anaerobic response under the experimental conditions (Tai et al., 2005, see Section 4), increased transcript levels could not be interpreted as evidence for a contribution of the encoded proteins to cellular fitness in the immediate experimental context. This result strongly corroborated the conclusions from genome-scale comparisons between transcript profiles and fitness in batch cultures, in which S. cerevisiae was exposed to DNA-damaging agents (Birrell et al., 2002) and to various other stress conditions (Giaever et al., 2002). Apparently, increased transcript levels as such cannot be interpreted as evidence for a high physiological relevance of the encoded protein under the experimental conditions. As stressed by Tai et al. (2007c), this does not imply that transcriptional up-regulation is without biological significance. First, functional redundancy is an inherent problem in the analysis of (single) deletion mutants. Indeed, a quarter of gene deletions in S. cerevisiae that do not confer a clear phenotype are compensated by expression of related genes (Gu et al., 2003). Second, the impact of up-regulation of a gene on cellular fitness may itself be context dependent. For example, as mentioned above, ammonium-limited growth of S. cerevisiae leads to a coordinated up-regulation of transporters and enzymes involved in the assimilation of alternative nitrogen sources, even if these are not available in the growth medium (Boer et al., 2003; Magasanik and Kaiser, 2002; ter Schure et al., 1998). Similar mechanisms may underlie the transcriptional up-regulation under anaerobic conditions of some of the genes included in this study. Finally, transcriptional regulation networks may have evolved to generate coordinated responses to environmental stimuli that tend to coincide in the natural environment. When these stimuli are separated in the laboratory or in industry, not all transcriptional responses will have a direct bearing on each individual stimulus. Ideally, functional redundancy and context dependency could be circumvented by testing strain collections that carry multiple, combinatorial deletions under a range of different experimental conditions. However, such experiments would require high-throughput analysis of cellular fitness under a range of cultivation regimes, which is currently not feasible in a chemostat setup.

6. EVOLUTIONARY AND REVERSE ENGINEERING It has long been recognized that prolonged chemostat cultivation leads to a strong selective pressure on micro-organisms (Fig. 8; Novick and Szilard, 1950b). In particular, nutrient-limited cultivation imposes a positive

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Figure 8 Selection of mutants during prolonged cultivation in chemostat. Rise in b-galactosidase activity in E. coli grown in a lactose-limited chemostat, indicating the selection of spontaneous mutants with an improved affinity for lactose. Adapted from Novick (1961) with permission from Basic Books.

selective pressure on spontaneous or induced mutants that have an improved affinity for the growth-limiting nutrient (Dykhuizen and Hartl, 1983; Harder et al., 1977; Novick and Szilard, 1950b). Affinity is a measure of the ability to grow at relatively high rates at a very low concentration of the growth-limiting nutrient. Monod kinetics (Monod, 1942) imply that a high affinity can be achieved by decreasing the substrate-saturation constant Ks and/or by increasing mmax. Affinity can therefore be defined as mmax/Ks (Button, 1991). Mutants with an increased affinity for the growth-limiting nutrient can achieve the required specific growth rate in chemostat, dictated by the dilution rate, at a lower residual substrate concentration than the original population, which they can therefore completely outcompete. Evolution in chemostat cultures is not only a complication in chemostat studies (see Section 2.1.3), but can also be harnessed to select for mutants with industrially or academically interesting properties. The use of chemostat cultures for such ‘‘evolutionary engineering’’ approaches (Butler et al., 1996) has been discussed extensively, along with other (cultivation) techniques, in an excellent review by Sauer (2001). Examples of chemostat-based evolutionary engineering of S. cerevisiae include the selection of strains with an increased affinity for glucose and maltose (Adams et al., 1985; Jansen et al., 2004, 2005), improved utilization of xylose

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(Kuyper et al., 2005a; Sonderegger and Sauer, 2003), increased contribution of respiration to glucose dissimilation in cultures grown under an unidentified limitation (Ferea et al., 1999), increased stress resistance (Cakar et al., 2005) and independence from a growth factor (van Maris et al., 2004a). The scientific challenge in evolution-based approaches is not only to select mutants with interesting properties, but also to understand the molecular basis for their altered performance. Such an understanding is not only interesting from an academic point of view. Upon identification of the responsible mutations, these can be applied in rational metabolic engineering strategies, for example, to combine different relevant genotypes by transfer of mutations to industrial strains. Genotypic analysis of evolved strains and the subsequent use of key mutations in targeted metabolic engineering strategies is known as reverse metabolic engineering (Bailey et al., 2002; Bro and Nielsen, 2004). In addition to their role as ‘‘evolution machines,’’ chemostat cultures provide a useful tool for genotypic analysis of evolved phenotypes. Several studies have combined chemostat cultivation of evolved S. cerevisiae strains with micro-array analysis to gain insight into the mutations responsible for the acquired phenotypes. Some of these studies are briefly discussed below. Several groups have used a combination of evolutionary engineering and chemostat-based micro-array analysis to improve xylose utilization by metabolically engineered S. cerevisiae strains. Wahlbom et al. (2003) used chemostat cultures to compare strains that were originally evolved in batch cultures for improved utilization of, and specific growth rate on, xylose. Micro-array analysis showed increased transcript levels for XKS1 encoding xylulokinase and for two genes encoding enzymes of the pentose phosphate pathway (TAL1 and TKL1). Indeed, overexpression of these genes has been shown to lead to improved xylose fermentation by S. cerevisiae (Ho et al., 1998; Karhumaa et al., 2005; Kuyper et al., 2005a). Sonderegger and Sauer (2003) gradually decreased the oxygen feed to xylose-limited chemostat cultures of an engineered strain overexpressing Pichia stipitis genes for xylose reductase and xylitol dehydrogenase. While the original strain required oxygen for growth on xylose, the evolved culture obtained after 460 generations of selection was capable of anaerobic growth on xylose. Micro-array analysis of the evolved strain indicated multiple changes in central metabolism, many of which could be related to redox co-factor use by the P. stipitis enzymes. In other studies, anaerobic conversion of xylose into ethanol was achieved by combined metabolic and evolutionary engineering of S. cerevisiae strains expressing a fungal xylose isomerase (Kuyper et al., 2005b). Subsequent chemostat-based

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micro-array analysis revealed drastic changes in the transcript levels of several hexose-transporter genes (van Maris et al., 2007). However, transcriptome analysis did not enable unequivocal conclusions on the molecular basis for anaerobic growth on xylose, neither in the xylosereductase/xylitol dehydrogenase-based strains nor in the xylose-isomerasebased strain. Pyruvate-decarboxylase-negative S. cerevisiae strains provide an interesting platform for metabolic engineering. However, Pdc strains are unable to grow at high glucose concentrations and, moreover, require a 2-carbon compound (ethanol or acetate) for growth in glucose-limited cultures (Flikweert et al., 1999; van Maris et al., 2004a). van Maris et al. (2004a) used evolutionary engineering in chemostats and batch cultures to select Pdc strains that were two-carbon compound-independent and glucose tolerant. Subsequent micro-array analysis of the evolved and parental strains was performed in nitrogen-limited cultures. These conditions were specifically chosen to enable a comparison at a fixed, high residual glucose concentration. Micro-array analysis indicated altered transcription of many targets of the glucose-dependent repressor Mig1p as well as drastically reduced transcription of several hexose transporter genes in the evolved strain (van Maris et al., 2004a). This, however, did not provide a mechanistic explanation for the acquired phenotype. The transcript profiles of S. cerevisiae strains evolved during prolonged sugar-limited growth in chemostat cultures have been the topic of three separate studies. Ferea et al. (1999) described evolution of a laboratory strain in which aerobic alcoholic fermentation already occurred at very low specific growth rates in glucose-limited chemostat cultures. Prolonged aerobic, glucose-limited cultivation resulted in fully respiratory growth at dilution rates that originally resulted in a respiro-fermentative sugar metabolism. The evolved strain showed reduced transcript levels for genes involved in glycolysis and an increased transcript level for genes involved in the tricarboxylic acid cycle and oxidative phosphorylation (Ferea et al., 1999). In another study, starting with a laboratory strain with a much higher critical dilution rate, prolonged cultivation of respiratory, glucose-limited cultures also resulted in a strong and coordinated decrease of the transcript levels of genes encoding glycolytic enzymes (Jansen et al., 2005). Subsequent analysis of intracellular metabolite levels in aerobic, glucose-limited chemostat cultures of the parental and evolved strains indicated that the reversible reactions in glycolysis were close to thermodynamic equilibrium in the two strains. Apparently, the high levels of these enzymes represent a true ‘‘overcapacity’’ in steady-state, glucose-limited cultures. Moreover, glycolytic enzymes represent a significant fraction of the soluble protein in

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wild-type S. cerevisiae (Bakker et al., 2000). The energy cost for glycolytic protein synthesis might therefore explain the selective advantage of mutants with decreased levels of glycolytic enzymes under energy-limited cultivation conditions (Mashego et al., 2005). Jansen et al. (2005) demonstrated that the affinity for glucose increased during evolution in aerobic, glucose-limited chemostat cultures of S. cerevisiae. Improved affinity was reflected by an almost threefold reduction of the residual glucose concentration in the cultures, a decreased Km and an increase of the maximum capacity of glucose transport. However, these changes were not reflected by corresponding changes in the transcript levels of hexose transporter genes (Jansen et al., 2005). Conversely, in a similar chemostat-based evolution study with maltose as the growth-limiting nutrient, an increased affinity for maltose was accompanied by increased transcript levels of genes encoding maltose permease, maltase and the MAL regulator (Jansen et al., 2004). The evolved strains became hypersensitive to sudden exposure to excess maltose, probably due to substrate-accelerated death caused by unrestricted maltose uptake (Jansen et al., 2004; Postma et al., 1989b), thus clearly illustrating how prolonged, steady-state nutrientlimited growth can result in a trade-off between affinity and robustness. The cases discussed in this section show that, although chemostat-based micro-array analysis of evolved strains can provide valuable information, it rarely provides a sufficient knowledge base for reverse engineering. Several aspects may explain why transcriptome analysis by itself is not sufficient. Long-term evolution can result in many mutations, not all of which are relevant for the observed phenotype. For example, Jansen et al. (2005) showed that many genes with altered transcript levels were physically clustered on the chromosomes, suggesting deletion or duplication of relatively large chromosomal regions. The technical complexity and cost of chemostat cultures preclude their use as a true high-throughput cultivation method. Nevertheless, it seems prudent to include replicate evolution runs in the design of evolutionary engineering experiments in order to facilitate the identification of biologically relevant mutations and/or processes. Standard yeast microarrays cannot identify point mutations, which may result in drastic functional changes in the encoded protein. Application of tiling arrays (see Section 2.2.1) does enable the identification of single- nucleotide polymorphisms (Gresham et al., 2006) and will provide an invaluable addition to the toolbox for genotypic analysis of evolved strains. Finally, as illustrated by the study on strains with reduced glycolytic activity (Jansen et al., 2005; Mashego et al., 2005) discussed above, combination of micro-array analysis with posttranscriptional analytical tools can make decisive contributions to the interpretation of evolved phenotypes.

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7. OUTLOOK 7.1. Beyond the Transcriptome: Multi-Level Approaches Although transcriptome analysis provides valuable insights into the responses of living cells to environmental changes and genetic interventions, transcription is but the first step in a long chain of events leading from gene to in vivo function. In addition to transcriptional regulation, regulation of cellular processes can occur at multiple levels, including translation, protein degradation, post-translational modification and modification of in vivo activity of proteins via low molecular weight substrates, products and effectors. One of the major challenges in systems biology is to dissect and quantify different levels of cellular information (‘‘-omes’’) and assess their importance in overall regulation of cellular processes. Especially for this type of research, in which a robust statistical analysis of data is crucial, the high degree of reproducibility of chemostat-based experiments (Piper et al., 2002) is attractive. Another important advantage of chemostat cultivation concerns the dynamics of cellular regulation. Mechanisms such as modification of enzyme activities by low-molecular weight effectors and posttranslational modification (e.g. via (de)phosphorylation events) may occur within seconds and minutes, respectively. Depending on growth conditions, half an hour or longer may pass before a newly induced protein is fully expressed. Under the dynamic conditions existing in batch cultures, these different time constants will inevitably lead to time delays between the responses at different levels of information, which makes an uncluttered comparison of different cellular information levels extremely challenging. While, eventually, the dynamics of multi-level regulation will need to be addressed (see Section 7.2), chemostat cultivation provides an ideal platform for an initial analysis of multi-level regulation under steady-state conditions. In batch studies for which transcriptome and proteome data from the same samples were compared, a rather poor correlation between the levels of mRNAs and the corresponding proteins was observed (Bro et al., 2003; Chen et al., 2002; Greenbaum et al., 2003; Griffin et al., 2002; Gygi et al., 1999; Ideker et al., 2001). Recent multi-level analyses in chemostat cultures showed that this imperfect correlation is not solely due to the dynamics of batch cultures. Also in steady-state chemostat cultures, comparisons between different cultivation regimes revealed consistent differences in the fold changes found for mRNA and protein levels (Castrillo et al., 2007;

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de Groot et al., 2007; Kolkman et al., 2006; Salusjarvi et al., 2006). Importantly, regulation at the protein level was implicated in the regulation of entire pathways and cellular processes, including glycolysis and charging of tRNAs (Castrillo et al., 2007; de Groot et al., 2007). The proteome is the net result of protein synthesis and protein degradation. Pratt et al. (2002) have dissected these two processes by pulsing unlabeled amino acids to isotope-labeled steady-state chemostat cultures of S. cerevisiae. Their study revealed a remarkable heterogeneity in the turnover rates of abundant yeast proteins, thus underscoring the potential relevance of this process in cellular regulation. Because of the multiplicity of posttranslational modification mechanisms (e.g. phosphorylation, glycosylation, etc.) and the analytical challenges involved in their quantification, there are as yet no genome-wide studies to quantify their impact on cellular regulation. Ter Kuile and Westerhoff (2001) have proposed that metabolic regulation can be dissected in hierarchical and metabolic regulation. Hierarchical refers to regulation of processes that affect enzyme or pathway capacity (Vmax), while metabolic regulation refers to the modification of this capacity by low molecular weight metabolites and effectors (ter Kuile and Westerhoff, 2001). Provided that quantitative information is available, this concept enables a straightforward ‘‘bookkeeping’’ of the contribution of different levels of regulation in different experimental contexts (Rossell et al., 2005, 2006). When potential pitfalls related to differential expression of isozymes and differences between in vivo and in vitro activities are taken into account, enzyme assays in cell extracts can provide an estimate of relative Vmax values, while the in vivo flux through individual enzymes can be calculated and/or measured via in vivo labeling and metabolic flux analysis. Genome-scale or core stoichiometric models of the metabolic network of S. cerevisiae (Fo¨rster et al., 2003; Stu¨ckrath et al., 2002) enable a fast estimate of in vivo intracellular fluxes in steady-state chemostat cultures. Two chemostat studies in which calculated in vivo fluxes under different conditions were compared to relative Vmax values calculated from enzyme assays in cell extracts (Daran-Lapujade et al., 2007; Jansen et al., 2005; Tai et al., 2007a) demonstrated that the in vivo glycolytic activity under steady-state conditions was mainly regulated at the metabolic level. Further development of analytical tools that enable analysis of the entire metabolome and, in eukaryotes such as S. cerevisiae, its subcellular compartmentation, represents a major challenge for the unraveling and quantification of different metabolic regulation mechanisms.

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Figure 9 Chemostat based perturbation studies. Transcriptional responses of S. cerevisiae grown in glucose-limited chemostat (D ¼ 0.05 h1) after exposure to a 5.6 mM glucose pulse, shown as a two-dimensional clustering heat-map of the differentially transcribed genes. Each transcript level represents the average of at least two independent replicates. Orange (relatively high expression) and blue (relatively low expression) squares were used to represent the transcription profiles of genes deemed specifically changed. K-means clusters of genes with ascendant profiles (A, B and C) and descendent profiles (D, E). The thick black line represents the average of the median-normalized expression data of the genes comprising the cluster. Adapted from Kresnowati et al. (2006) with permission from Nature Publishing Group. (See plate 4 in the color plate section.)

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7.2. Beyond Steady-State Analysis: Perturbation Experiments Dynamic situations are, inherently, richer in information than steady-state cultivation experiments. It is, therefore, logical that transcriptome analysis is progressing from comparison of (pseudo) steady-state situations to studies of the dynamics of transcriptional responses upon a perturbation of the cultivation conditions. Since, in dynamic experiments, time is introduced as an additional variable, they are more sensitive to experimental variation than steady-state analysis. Hence, data quality and tight control of experimental conditions are of paramount importance. Accordingly, it has therefore been proposed that steady-state chemostat cultures are excellently suited as a reproducible platform for dynamic perturbation studies (Kresnowati et al., 2006; Ronen and Botstein, 2006). A typical experimental design for a chemostat-based dynamic study consists of the application of perturbation (e.g. a glucose pulse) to a steadystate chemostat culture, followed by rapid sampling, quenching of metabolism and analysis of relevant intracellular and extracellular components (Theobald et al., 1997). Until recently, chemostat-based perturbation studies mainly focused on changes of metabolite levels. This has led to the development of techniques that enable analysis of changes in intracellular metabolite concentrations over a very short time scale (1 s to 5 min; Mashego et al., 2003; Theobald et al., 1997). Kresnowati et al. (2006) have recently integrated metabolite analysis and micro-array analysis to study the dynamic responses of glucose-limited chemostat cultures of S. cerevisiae to a glucose pulse. By monitoring the transcriptome within a 5 min time window (Fig. 9), evidence was obtained for wide-spread active degradation of mRNAs. Moreover, the combination of transcriptome and metabolome data provided clear insights into the main cellular adaptations that occur upon a sudden relief from glucose limitation. By expanding the time scale of perturbation studies, it will become relevant to also include proteome data.

8. CONCLUDING REMARKS The reproducibility of chemostat cultivation and its flexible options for experimental design form a powerful combination with high-informationdensity analytical approaches such as DNA-micro-array analysis. However, progress in microbial physiology would be ill served by dogmatism about the choice of a cultivation technique. A major limitation of chemostat

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cultivation is that it remains a fairly complicated way of growing microorganisms, which is at present not compatible with high-throughput analysis. Therefore, systems such as shake flasks and microtiter plates will remain indispensable for high-throughput, exploratory research in microbial physiology. Quantitative analysis in chemostat-based studies will contribute to the interpretation of data generated in such less-controlled cultivation systems, for example via the identification of clear growth-rate-related effects (Castrillo et al., 2007; Regenberg et al., 2006). Paradoxically, one of the strengths of chemostat cultures – the option to study highly reproducible steady-state situations – also represents one of its important limitations. Today, many important questions in microbial physiology should preferably be addressed in steady-state situations, with the elimination of dynamic effects. At the same time, these dynamic aspects are themselves extremely interesting and merit research. As indicated in the preceding paragraph, steady-state chemostat cultivation can be used as a reproducible platform for dynamic stimulus-response studies, thus expanding the applicability of chemostat cultures beyond steady-state analysis. We anticipate that the combination of micro-array analysis with other non-steady-state methods for continuous cultivation such as retentostats (for analysis of microbial physiology at extremely low specific growth rates; van Verseveld et al., 1984) and accelerostats (for investigating dynamic growth rate profiles; Kasemets et al., 2003; Paalme et al., 1997) will yield further interesting results.

ACKNOWLEDGEMENTS We thank Derek Abbott for making Fig. 2 and our colleagues in the Delft Industrial Microbiology Section for stimulating discussions.

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A Predatory Patchwork: Membrane and Surface Structures of Bdellovibrio bacteriovorus Carey Lambert, Laura Hobley, Chien-Yi Chang, Andrew Fenton, Michael Capeness and Liz Sockett Institute of Genetics, School of Biology, University of Nottingham, Queen’s Medical Centre, Nottingham NG7 2UH, UK

ABSTRACT Predatory Bdellovibrio bacteriovorus bacteria are remarkable in that they attach to, penetrate and digest other Gram-negative bacteria, living and replicating within them until all resources are exhausted, when they escape the prey ghost to invade fresh prey. Remarkable remodeling of both predator and prey cell occurs during this process to allow the Bdellovibrio to exploit the intracellular niche they have worked so hard to enter, keeping the prey ‘‘bdelloplast’’ intact until the end of predatory growth. If one views motile non-predatory bacteria in a light microscope, one is immediately struck by how rare it is for bacteria to collide. This highlights how the cell surface of Bdellovibrio must be specialized and adapted to allow productive collisions and further to allow entry into the prey periplasm and subsequent secretion of hydrolytic enzymes to digest it. Bdellovibrio can, however, also be made to grow artificially without prey; thus, they have a large genome containing both predatory genes and genes for saprophytic heterotrophic growth. Thus, the membrane and outer surface layers are a patchwork of proteins encompassing not only those that have a sole purpose in heterotrophic growth but also many more that are specialized or employed to attach to, enter, remodel, kill and ultimately digest prey cells. There is much that is as yet not understood, but molecular genetic and post-genomic approaches to ADVANCES IN MICROBIAL PHYSIOLOGY, VOL. 54 ISBN 978-0-12-374323-7 DOI: 10.1016/S0065-2911(08)00005-2

Copyright r 2009 by Elsevier Ltd. All rights reserved

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microbial physiology have enhanced the pioneering biochemical work of four decades ago in characterizing some of the key events and surface protein requirements for prey attack.

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4. 5. 6. 7. 8.

9.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The role of flagellar motility in predation . . . . . . . . . . . . . . . . . . . . . . 2.1. The Bdellovibrio Flagellum . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Genes for Flagellar-Related Proteins Found in B. bacteriovorus HD100 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. The Role of Each of the Six fliC Genes and their Importance in Motility . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. The Bdellovibrio Chemotaxis System . . . . . . . . . . . . . . . . . . . . 2.5. Genes for Chemotaxis-Related Proteins Found in B. bacteriovorus HD100 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6. The Role of mcp2 in Chemotaxis . . . . . . . . . . . . . . . . . . . . . . . 2.7. The Importance of Motility and Chemotaxis . . . . . . . . . . . . . . . . The role of type IV pili in predation . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. The Pilus and Bdellovibrio . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. The Structure of a Type IV Pilus and the Gene Complement in Bdellovibrio . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Proposed Regulation of Bdellovibrio Pili . . . . . . . . . . . . . . . . . . 3.4. Flp Pili . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5. Expression of Pilus Genes in the Predatory Cycle . . . . . . . . . . . 3.6. The Role of PilA in Predation . . . . . . . . . . . . . . . . . . . . . . . . . . Outer membrane proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Autotransporters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The role of the sec system . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Membrane chemistry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Peptidoglycan chemistry and metabolism . . . . . . . . . . . . . . . . . . . . . 8.1. Cell Wall Precursor Biosynthesis Retained by Bdellovibrio . . . . . 8.2. Cell Wall Hydrolase Activities in Bdellovibrio . . . . . . . . . . . . . . . 8.3. Aminidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.4. DD-Endopeptidases and DD-Carboxypeptidases . . . . . . . . . . . . 8.5. Bdellovibrio Cell Wall Hydrolyases Act upon the Prey during Predation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

314 316 317 318 321 322 323 325 326 326 327 328 331 332 333 334 335 340 344 345 349 349 350 352 352 353 354 355 355

1. INTRODUCTION Bdellovibrio are small, highly motile Gram-negative bacteria which prey upon other Gram-negative bacteria, entering the prey’s periplasm,

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replicating within and using the contents of the prey as the sole nutrient source. In order to carry out this unusual lifestyle, they have evolved specialized surface properties and structures to achieve several steps including contact with the prey, entry into it and transport of various substances into and out of the killed prey. Thus the membranes and surface structures of Bdellovibrio are more than simple semi-permeable barriers, as in other cells, and can be thought of almost as an organelle of prey interaction, with regular membrane functions slotted in between – hence, the choice of title. We review what is known about the proteins of the Bdellovibrio surface, their roles in predation and we try to highlight areas worthy of further work. Bdellovibrio bacteriovorus can exist in two different growth phases, as host-dependent (HD) cells that require prey for growth and division and as host-independent (HI) cells, growing in very rich nutrient media (see Fig. 1). When growing host-dependently, Bdellovibrio exhibit a bi-phasic lifestyle consisting of a free-swimming attack-phase and an intraperiplasmic growth phase, during which they reside within the periplasm of their prey. During the free-swimming attack phase, prey location, attachment and recognition are vital to the successful replication of the Bdellovibrio, as they typically

Figure 1 The lifecycle of Bdellovibrio bacteriovorus growing both hostdependently (HD) and host-independently (HI). During both forms of growth the Bdellovibrio cell elongates and then divides at multiple fission sites.

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have a half-life of about 10 hours (Hespell et al., 1974; Hobley et al., 2006) during starvation in buffered environments. After a brief recognition period, during which the Bdellovibrio identifies its prey by mechanisms as yet unknown, attachment between Bdellovibrio and prey becomes irreversible, and the invasion process begins. Invasion begins by the production of a small pore in the outer membrane of the prey, through which the Bdellovibrio squeezes. Once inside, the pore is resealed and modification of prey peptidoglycan causes the rounding of the prey cell and the formation of the structure known as the bdelloplast. This is followed by a period of intense hydrolytic enzyme production and subsequent secretion into the prey cell’s cytoplasm, and the uptake of the degraded cytoplasmic contents. The Bdellovibrio cell is seen to elongate, and then septates at multiple fission sites, whereupon the multiple progeny cells become flagellate before releasing a final volley of lytic enzymes to burst from the confines of the bdelloplast. When growing host-independently, the growth of the Bdellovibrio appears to mimic that of the HD lifecycle; highly motile, attack-phase-like cells are seen to elongate into long filamentous cells (both straight and spiralled), which eventually septate to give multiple progeny in a manner similar to that employed within the bdelloplast. Many features of this unique lifecycle require the use of specialized membrane proteins and a highly optimized flagellum. A chemotaxis system is required for the location of environmental niches rich in prey and for productive collisions with prey. Type IV pili are known to be vital for prey cell entry, and a variety of transporters and membrane proteins are likely to be needed to both facilitate attachment and entry and to deploy the arsenal of proteases, nucleases and other hydrolases into the prey cytoplasm. In this study, we present our first analyses of the surface protein genes within the B. bacteriovorus HD100 genome, along with a study of gene expression patterns and propose potential roles for them in the Bdellovibrio lifecycle. The reader is referred to a recent review by Barabote et al. (2007) for a further treatment of the inventory of transport proteins in the Bdellovibrio genome which is beyond our scope here.

2. THE ROLE OF FLAGELLAR MOTILITY IN PREDATION Bdellovibrio bacteriovorus are among the fastest self-propelled organisms known to science. B. bacteriovorus HD100 has been recorded swimming at over 160 mm/s, approximately 160 body lengths per second, roughly the

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equivalent of a human running at the speed of sound. The role of motility in predation has been studied by several laboratories over the 45 years since Bdellovibrio were first isolated. The contribution of chemotaxis, rather than simple motility, in predation is more cryptic, but has been shown to be of some importance in prey location in certain experimental setups. Here, we review the evidence for the importance of flagellar motility in the predatory lifestyle.

2.1. The Bdellovibrio Flagellum Bdellovibrio characteristically have a single, polar flagellum which is surrounded by a continuous sheath (Seidler and Starr, 1968; Thomashow and Rittenberg, 1985c) (described further below) similar to that seen for various Vibrio spp. When viewed by electron microscopy, the Bdellovibrio flagellum has a dampened waveform (Thomashow and Rittenberg, 1985a), with larger amplitude waves closest to the cell and a decrease in amplitude towards the distal end of the flagellum, as in Fig. 2.

Figure 2 An electron micrograph of a single cell of B. bacteriovorus 109 JK (KanR wild-type control) showing the sheathed flagellum and the typical dampened waveform. Cell stained with 0.5% Uranyl Acetate and the scale bar represents 1 mm.

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2.2. Genes for Flagellar-Related Proteins Found in B. bacteriovorus HD100 The Bdellovibrio genome sequence revealed that most of the genes required to form a typical, functional Gram-negative bacterial flagellum were located within three operons (see Fig. 3; Rendulic et al., 2004), while, in contrast to the more usual single copy, six copies of the gene encoding the filament structural protein FliC were found at four loci, and three copies of each of the genes for the motor proteins MotA and MotB were found arranged in pairs around the genome. Homologs of most of the known regulators of the flagellar morphogenetic pathway can be found within the B. bacteriovorus HD100 genome. Two exceptions are the genes flhC and flhD which act as master regulators of flagellar synthesis in Escherichia coli and for which genes of significant homology cannot be found within the genome. Thus, highly divergent genes or alternative regulatory mechanisms must exist. The genes coding for all of the main proteins that make up the flagellar structure and export apparatus can be found on two operons in the HD100 genome (flh-fli and flg-fli, see Table 1 and Fig. 3) and are similar to those

Figure 3 Position of genes involved in flagella motility and chemotaxis in the genome of B. bacteriovorus HD100. The genes that are underlined are those for which insertion/deletion mutants have been made and characterized in B. bacteriovorus 109 J.

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Table 1 Genes from B. bacteriovorus HD100 genome with homology to genes for known proteins used to form a bacterial flagellum and motor. Those underlined have been studied in B. bacteriovorus 109J, and null phenotypes characterized. Conserved domains were found using the NCBI blast service, and the E-value is that to the numbered domain Bd number

Annotation

Likely gene homolog

COG number

E-value

Regulators Bd0537 Bd3318 Bd3319 Bd3320

flgM fliA fleN flhF

flgM fliA fleN flhF

COG2747 COG1191 cd02038 COG1419

1e–03 4e–49 1e–36 4e–37

Export apparatus Bd3321 flhA Bd3322 flhB Bd3323 fliR Bd3324 fliQ Bd3325 fliP Bd3326 fliO Bd3401 fliI Bd3402 fliH

flhA flhB fliR fliQ fliP fliO fliI fliH

COG1298 COG1377 COG1684 COG1987 COG1338 COG3190 COG1157 COG1317

o1e–180 5e–88 4e–27 7e–17 9e–72 2e–05 1e–165 6e–10

MS-, P- and L-rings Bd0532 flgA Bd0534 flgH Bd0535 flgI Bd3404 fliF

flgA flgH flgI fliF

COG1261 PRK00249 COG1706 COG1766

2e–15 2e–03 2e–84 8e–69

Hook and basal body Bd0530 flgE Bd0531 flgG Bd0536 flgJ Bd0540 flgK Bd0542 flgL Bd3395 flgE Bd3397 flgD Bd3398 Bd3398 Bd3405 fliE Bd3406 flgC Bd3407 flgB

flgG flgG flgJ flgK flgL flgE flgD fliK fliE flgC flgB

COG4786 COG4786 PRK05684 COG1256 COG1344 COG1749 COG1843 COG3144 COG1677 COG1558 COG1815

6e–54 2e–74 6e–08 3e–48 5e–28 4e–79 7e–29 6e–04 3e–13 3e–32 4e–21

Rotor Bd3014 Bd3327 Bd3328 Bd3403

fliG fliN fliM fliG

COG1536 COG1886 COG1868 COG1536

4e–14 7e–22 5e–73 2e–69

fliG fliN fliM fliG

(Continued )

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Table 1 (continued ) Bd number

Annotation

Likely gene homolog

COG number

E-value

Motor Bd0144 Bd0145 Bd3020 Bd3021 Bd3253 Bd3254

motA motB motB motA motB motA

motA motB motB motA motB motA

COG1291 COG1360 COG1360 COG1291 COG1360 COG1291

1e–52 1e–40 3e–24 2e–39 2e–12 8e–38

Filament Bd0408 Bd0410 Bd0604 Bd0606 Bd0610 Bd0611 Bd3052 Bd3342

flaA hag hag hag fliD fliS flgL hag

fliC3 fliC4 fliC1 fliC2 fliD fliS fliC5 fliC6

COG1344 COG1344 COG1344 COG1344 COG1345 COG1516 COG1344 COG1344

3e–28 2e–24 3e–24 2e–27 1e–45 3e–19 4e–25 8e–27

PRK07718 PRK08455 COG1580

1e–04 1e––08 1e–20

Unknown function within flagellar structure Bd0804 fliL fliL Bd1076 fliL fliL Bd3329 fliL fliL

found in other bacterial genomes including the well-characterized E. coli, Salmonella enterica serovar Typhimurium, and Rhodobacter sphaeroides (Blattner et al., 1997; Garcia et al., 1998; McClelland et al., 2001). The genes for the rotor-/chemotactic-switch proteins fliM and fliN are found within the flh-fli operon and, in addition, the HD100 genome contains two potential rotor gene copies coding for fliG, one within the flg-fli operon and the second (Bd3014) is found as an orphan within the genome, several genes upstream of a pair of motAB genes. A possible reason for having two copies of the gene for the rotor/switch protein FliG could be because of its importance in producing motile cells, as FliG has a dual role in torque generation and participating in the flagellar export channel in other bacteria such as E. coli (Lloyd et al., 1996). Having extra copies of this critical motility gene within the genome may optimize the chance of the Bdellovibrio retaining motility and, thus, viability as a predator if one of the copies is lost or mutated. A similar view pertains to the presence of three pairs of genes encoding the motor proteins MotA and MotB. These are arranged as pairs within the B. bacteriovorus genome. One pair is very close to the flh-fli operon, another pair is located six genes downstream of the orphan fliG and

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the final pair is distal to other motility genes in the genome. The motor proteins MotA and MotB are both needed for the trans-membrane ion conductance that provides the motive force for flagellar rotation (Stolz and Berg, 1991). The acquisition of three copies of each of the genes encoding these proteins by Bdellovibrio may again be an evolutionary advantage, as the loss of any pair may possibly be compensated for by the presence of the remaining sets of genes. The flagella filament is typically made up of tens of thousands of monomers of the flagellin protein FliC, with a cluster of FliD proteins forming the filament ‘‘cap’’ which also acts on each FliC monomer, ensuring it polymerizes in the correct place on the filament. fliD, along with the fliC chaperone fliS, are found together, four genes downstream of the pair of flagellin-encoding genes fliC1 and fliC2. Both the genome of B. bacteriovorus HD100 and the well-characterized lab strain 109 J were found to contain six copies of the gene coding for the flagellin protein FliC. Work by Lambert et al. (2006) showed that these were arranged at four loci around the genome, with fliC1 and fliC2, and fliC3 and fliC4, being found as pairs, with fliC5 and fliC6 as orphans in the genome. Thus, several of the key genes required for flagellar synthesis and torque generation are duplicated within the genome of Bdellovibrio, and this has been referred to as Bdellovibrio having ‘‘spare tyres’’ of genes critical for motility (Lambert et al., 2006). An insight into the importance of the duplication of these genes was shown by mutagenesis of each of the fliC genes by Lambert et al. (2006).

2.3. The Role of Each of the Six fliC Genes and their Importance in Motility The six fliC genes were all found to be expressed by predatory cells of B. bacteriovorus 109 J by RT-PCR while quantitative RT-PCR revealed that fliC2, fliC3 and fliC5 (at least) were expressed at different levels relative to each other, and that the expression levels of each varied at different points in the lifecycle, being expressed most in the free-swimming attack phase and least during intracellular growth (Lambert et al., 2006). Each of the six genes, in turn, encoding putative FliC proteins in B. bacteriovorus 109 J were interrupted by insertion of an antibiotic resistance cassette. The resulting mutants were found to be viable, although the fliC3 mutant could only be obtained by growing the Bdellovibrio hostindependently. The five mutants that could still be grown in predatory cultures were all motile but had a variety of phenotypes, including reduction in swimming speed, alteration in flagellar lengths and reductions in

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predation efficiency. Deletion of fliC5, one of the orphan genes, was found to have the most effect (of those that remained capable of predation in liquid) on the Bdellovibrio, resulting in shortened flagella with the loss of the smaller amplitude, distal waveform and subsequent reduction in swimming speed and predation efficiency. The other four fliC gene products seem to make minor, non-essential contributions to flagellar synthesis. The deletion of fliC3 had the most profound effect on the Bdellovibrio. The mutant was found to be non-motile, producing the flagellar sheath with no functional flagellar filament within it. Further analysis of this mutant revealed that it expressed all five of the remaining functional fliC genes and that disordered fragments of filament were within the sheath. Although this mutant was not able to be grown in predatory liquid culture, it was shown to be able to attach to, and invade, E. coli prey when HI-grown cells of it were placed in close proximity to the prey, thereby disproving the hypothesis of Burnham et al. (1968) that flagellar motility may have been used as a ‘‘drilling’’ mechanism to enter the prey cell. The inability of this mutant to grow in predatory cultures showed that while motility is not essential for prey invasion, it is essential for the location of prey.

2.4. The Bdellovibrio Chemotaxis System Motility has been shown to play a role in effective location and attachment of Bdellovibrio to prey, but the exact mechanisms by which Bdellovibrio actually sense their prey in natural environments is still a hotly debated topic. Chemotaxis has been shown to be used by many other bacteria as a means of locating both nutrient sources and areas with preferential conditions. Four studies of the potential chemotaxis responses by Bdellovibrio were published by S.F. Conti’s laboratory in the 1970s, showing first an attraction by Bdellovibrio to yeast extract (Straley and Conti, 1974), then to a variety of individual amino acids, including glycine, asparagine, histidine, cysteine and lysine, while others elicited no significant response, including serine, proline and arginine among others (Lamarre et al., 1977). This was followed by their publication on the chemotactic response of Bdellovibrio to prey (Straley and Conti, 1977) which showed a small chemotactic response of Bacteriovorax stolpii Uki2 to areas containing bacterial populations at densities greater than 108 cells per ml, both to potential prey and non-susceptible Gram-positive cells. This showed that, while Bdellovibrio are attracted to areas of high cell density, they are unable to distinguish whether the cells in those locations are potential prey. The final publication on Bdellovibrio chemotaxis by this group showed that

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many compounds acted as either chemo-attractants or repellents to B. stolpii Uki2, but none of those tested provoked any response from B. bacteriovorus 109 J by the methods used (Straley et al., 1979). The identification of initially two putative methyl-accepting chemotaxis proteins (MCPs) in the B. bacteriovorus 109 J, prior to any genomic studies, was reported by Lambert et al. (2003), once again raising the question as to whether Bdellovibrio are able to use chemotaxis to locate areas of high cell density and improve their predation efficiency. Flannagan et al. (2004) reported the presence of a minimum of 13 putative MCPs within the B. bacteriovorus 109 J genome, as found by the use of an mcp-specific probe; subsequently, 21 puatative mcp genes and 1 aerotaxis gene have been found within the B. bacteriovorus HD100 genome (see below). This was followed recently by a study by Chauhan and Williams (2006) that looked at the effect of nutrient gradients on the populations of both predators and prey in environmental samples. While the experimental evidence obtained thus far remains contradictory, the prevalence of genes coding for chemotaxis proteins in the Bdellovibrio genome (as shown below) indicates that further investigation is needed into the true role of chemotaxis in predation by Bdellovibrio.

2.5. Genes for Chemotaxis-Related Proteins Found in B. bacteriovorus HD100 In contrast to the genes for flagella synthesis, which are mostly found in operons within the B. bacteriovorus genome, the genes coding for chemotaxis-related proteins are randomly dispersed throughout the genome, with a few forming small clusters of up to four genes (see Fig. 3). All the main genes required for the chemotaxis signalling system, with the exception of cheZ (which abolishes the CheY-phosphate chemotactic signal in only some groups of bacteria), can be found in the HD100 genome (see Table 2). In a similar manner to that of the important genes required for flagella synthesis and motility, multiple copies of many of the genes required for chemotaxis signal transduction are found within the Bdellovibrio genome. The most abundant of the chemotaxis genes within the Bdellovibrio genome are those coding for MCPs. A total of 21 mcp genes are found within the HD100 genome, along with a single gene coding for a putative aerotaxis gene aer. The MCPs are dispersed randomly throughout the genome, with three pairs being found and one as part of a small cluster of chemotaxis genes (Bd2828-2831). The large number of mcp genes found in Bdellovibrio contrasts with the four (and aer) found in E. coli, but is fewer

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Table 2 Genes from B. bacteriovorus HD100 genome with homology to genes for known proteins used in the chemotaxis signal transduction pathway and for methyl-accepting chemotaxis proteins. The underlined gene has been studied in B. bacteriovorus 109J and the null phenotype characterized. Conserved domains were found using the NCBI blast service, and the E-value is that to the numbered domain Bd number

Annotation

Signal transduction Bd0102 cheY Bd1347 Bd1347 Bd1825 cheY Bd2409 cheY Bd3431 cheY Bd0579 cheW Bd2828 cheW2 Bd3471 cheW Bd0578 cheA Bd2406 cheA Bd3469 cheA Bd0580 cheB Bd3467 cheB Bd2830 cheR Bd3468 cheR Bd3101 Bd3101 Bd2829 cheD Bd1823 cheX

Likely gene homolog

COG number

E-value

cheY cheY cheY cheY cheY cheW cheW cheW cheA cheA cheA cheB cheB cheR cheR cheC cheD cheX

COG0784 COG0784 COG0784 COG0784 COG0784 COG0835 COG0835 COG0835 COG0643 COG0643 COG0643 pfam01339 COG2201 COG1352 COG1352 COG1406 COG1871 COG1406

1e–15 8e–13 1e–17 2e–16 1e–12 2e–13 8e–19 6e–30 3e–68 5e–38 4e–90 3e–21 4e–92 7e–59 1e–66 2e–12 2e–16 2e–17

COG0840 COG0840 COG0840 COG0840 COG0840 COG0840 COG0840 COG0840 COG0840 COG0840 COG0840 COG0840 COG0840 COG0840 COG0840 COG0840 COG0840 COG0840 COG0840 COG0840 COG0840 COG0840

1e–17 4e–03 4e–29 5e–16 1e–18 3e–31 4e–20 3e–20 3e–19 2e–13 2e–13 2e–06 5e–40 1e–26 7e–11 1e–16 6e–45 4e–22 6e–19 2e–25 2e–15 6e–17

Methyl-accepting chemotaxis proteins Bd0121 Bd0121 mcp Bd0203 Bd0203 aer Bd0262 Tsr mcp Bd0663 Bd0663 mcp Bd0932 Mcp mcp Bd1081 Mcp mcp Bd1126 Mcp mcp Bd1340 Bd1340 mcp Bd1469 Tsr mcp2 Bd1470 Tsr mcp1 Bd1872 Bd1872 mcp Bd1873 Bd1873 mcp Bd2503 Mcp mcp Bd2504 Mcp mcp Bd2596 Bd2596 mcp Bd2622 Trg mcp Bd2831 mcpA mcp Bd3092 Bd3092 mcp Bd3177 Mcp mcp Bd3192 Mcp mcp Bd3256 Mcp mcp Bd3341 Bd3341 mcp

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than the 26 found in Pseudomonas aeruginosa (Ferrandez et al., 2002). It is paradoxical that a predatory bacterium that is thought not to take up many amino acids and sugars in attack phase would have so many sensors for swimming towards such molecules. This suggests the importance of HI growth in natural environments where such metabolism would be of benefit. The main proteins involved in the chemotaxis signalling system are all represented by several copies of putative genes within the HD100 genome, including five putative genes for cheY (a phosphorylateable response regulator), three each of cheA (a histidine protein kinase) and cheW (an MCP-CheA-linking protein) and two each of cheB (MCP signal-adaptive methylesterase) and cheR (MCP signal-adaptive methyltransferase). One copy of a putative cheD gene can also be found, although the role of cheD in the chemotaxis signalling pathway is unclear. Of particular interest is one of the putative cheW genes, Bd2828 annotated as cheW2, which by sequence homology has been shown to contain three potential cheW domains. It may represent a protein of importance in forming (potential) MCP clusters as MCPs in other bacteria are known to occur in polar, or subpolar, signallingclusters of trimers of dimers. The lack of a cheZ gene is common among bacteria, as they have only been found within the g-Proteobacteria (Wadhams and Armitage, 2004). However, in other bacteria, alternatives to cheZ have been found such as cheC and cheX, which fulfill the role of de-phosphorylating cheY (Muff and Ordal, 2007). The B. bacteriovorus HD100 genome has putative homologs to both cheC (Bd3101) and cheX (Bd1823). Although further analysis of these proteins is needed, it is likely, due to the sequence similarity of these to the representative conserved domain sequences, that they may fulfill the role of cheZ in terminating the chemotactic signal in Bdellovibrio chemotaxis. The abundance of multiple copies of genes for chemotaxis signalling pathway leads to the hypothesis that chemotaxis may be of importance for Bdellovibrio survival in natural environments where, due to its short half-life (10 hours for strains HD100 and 109 J in laboratory starvation experiments (Hespell et al., 1974; Hobley et al., 2006)), efficient prey location is of great importance for survival. The combination of a potentially very effective sensing system and the extreme speeds of motility seen for Bdellovibrio may facilitate this.

2.6. The Role of mcp2 in Chemotaxis To date, the only gene coding for a member of the chemotaxis signalling pathway that has been studied in Bdellovibrio is that of the MCP, mcp2, by

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Lambert et al. (2003), who created a gene knock-out mutant of the mcp2 gene in B. bacteriovorus 109 J. The insertion of a kanamycin resistance (aphII) cassette into the mcp2 gene resulted in a kanamycin-resistant mcp2 mutant, which was shown to prey upon luminescent E. coli significantly slower than a wild-type kanamycin-resistant control (109 JK), in liquid cultures of predator and prey within the wells of a 96-well optical microtiter dish. This produced plaques of a size indistinguishable from wild type on semi-solid overlay plates of high prey cell density. This reduction in predation efficiency in liquid cultures, but not in dense lawns of prey, suggests that Bdellovibrio may in fact use chemotaxis as a means for moving through dilute environments to locate areas where prey cells are abundant. Studies of further reduced-chemotactic response mutants may illuminate this.

2.7. The Importance of Motility and Chemotaxis Both motility and chemotaxis have been shown to have important roles in the Bdellovibrio predatory lifecycle. Deletion of each of the fliC genes involved in the formation of the flagella filament has shown that, while motility is important for predation in liquid environments, it is not essential for prey cell entry. In addition, the predation efficiency of Bdellovibrio in liquid environments is proportional to their average swimming speed. Deletion of one out of the 21 putative mcp genes results in reduced predation efficiency in liquid environments containing a suspension of bacterial prey, showing that chemotaxis may play a role in prey location. The dedication of a significant proportion of the Bdellovibrio genome to multiple copies of the genes encoding both motility and chemotaxis proteins shows that they are likely to be important processes for the survival of predatory Bdellovibrio in nature.

3. THE ROLE OF TYPE IV PILI IN PREDATION Having found that flagellar-mediated motility is not required for the penetration of prey, interest turned to the role of pili in prey entry processes. It was first shown by Shilo (1969) that Bdellovibrio have fine, pilus-like filaments protruding from their cells at the non-flagellate pole and this was again seen by Abram and Davis (1970). Until recently, this work was not pursued further. Genes for type IV pilus proteins were noted in the genome

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sequence of Bdellovibrio bacteriovorus HD100 by Rendulic and colleagues (Rendulic et al., 2004). Type IV Pili (tfp) are filamentous strands extruding from bacterial outer membranes made up of thousands of pilin monomers. They have homology to type II secretion systems and play a vital role in such diverse traits as cell twitching motility in Proteus and Myxoccocus, pathogenicity in Pseudomonas and Neisseria, DNA uptake, pathogenicity and biofilm formation in a variety of bacteria. Type IV pili are made up of polymeric non-covalently linked monomers of pilin forming a polar filament which is about 5–7 nm in diameter, up to several micrometres in length, with each micrometer consisting of 1300 pilin subunits (Parge et al., 1995). The fiber is anchored to the inner membrane and associated with ATPases that drive the addition or subtraction of monomers to the fiber; also associated are methylases and peptidases required for maturation of the pilin preprotein. Twitching motility in particular has been widely studied; it allows certain bacteria to move over flat surfaces including metal, glass and plastics. This is accomplished by a ratchet effect involving the extrusion of the pilus, its adherence to a surface of some kind and the retraction of the pilus pulling the bacterium along. This process has best been described in Pseudomonas aeruginosa (Skerker and Berg, 2001) and was found to be important in the rapid colonization of new areas. It was shown that the bacteria can move at a rate of up to 0.5 mm/s with retraction and extraction of the pili occurring at a rapid 1000 subunits/s (Merz et al., 2000). The forces involved to achieve this were shown to be up to 140 pN (Maier et al., 2002).

3.1. The Pilus and Bdellovibrio In order to carry out its predatory lifecycle, Bdellovibrio first requires attachment to the host; this is always achieved at the non-flagellated pole. Then, it needs to squeeze through the pore in the prey outer layers and into the periplasm of the prey cell to establish itself in the periplasm. This ‘‘squeezing in’’ is likely to require a large amount of force to pull the bacterium into the prey periplasm and overcome the opposing force of the prey cell’s inner osmotic potential. A force of the sort of magnitude required could be generated by type IVa pili, which could also provide the adhesive properties required for attachment to prey surfaces, possibly the prey cell wall. In a recent study of Bdellovibrio, using an electron microscope (Evans et al., 2007), fibers were seen at the non-flagellated pole. However, the fibers were only apparent on 30% of free-swimming Bdellovibrio that had freshly

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lysed their host. This observed lack of 100% piliation was rationalized as avoiding interference with Bdellovibrio flagellar motility as they achieve speeds up to 160 mm/s Bdellovibrio extrude the pili only when contact is made with the prey cell, thus avoiding any shearing forces, stearic hindrance or entanglements. As mentioned above, the force of pilus retraction could be upwards of 100 pN and this would be ample force to penetrate the host cell. This strength, coupled with the flexibility of the pilus, could also ensure that the Bdellovibrio cell remained attached if the prey cell was still in motion; Burnham et al. (1968) showed that not even vortexing or light sonication could part predator from prey suggesting a strong interaction.

3.2. The Structure of a Type IV Pilus and the Gene Complement in Bdellovibrio The sequence of the pilin (pilA) subunit of the main fiber can be divided into three parts: the highly conserved hydrophobic N terminal sequence, the mid sequence containing beta sheets and finally the C-terminal hypervariable region. The work of Forest and Tainer (1997) suggested that the helical region at the N-terminus is located in the core of the fiber while the hypervariable regions are found on the outside with beta sheet regions in between the two. The structure of the entire pilus was revised by Mattick (2002) showing that a single pilus is made using 12 different protein products of the genes pilA, pilB, pilC, pilC1/Y1, pilD, pilE, pilQ, pilT, pilV, pilW, pilX and fimU (see Fig. 4a). This work comes from studies in Pseudomonas and is further supported by other work in myxobacteria. The helical PilA monomers that go to make up the fiber are found on the cytoplasmic face of the inner membrane in an immature form; these monomers are modified by a pre-pilin peptidase and the methylase PilD to make mature pilin monomers which are recruited to the base of the pilus fiber by the ATPase PilB (Turner et al., 1993), the action of which extrudes the fiber, whereas its counterpart PilT, also an ATPase, retracts the pilus fiber and de-polymerizes the subunits from the base into monomers to be used once more for extension (Hobbs et al., 1993). It is this de-polymerization that pulls the bacterium towards its point of attachment in twitching motility. Also found among different studied bacteria are a set of minor pilins, PilE, V, W, X and FimU, which are in some way involved in pilus construction and functionality. The extruded pilus is anchored by the protein PilC in the cytoplasmic membrane with the fiber extruding through the peptidoglycan layer by an as-yet unknown process and continuing

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Figure 4 (a) The general structure of bacterial type IVa pili. (b) The putative structure of the Bdellovibrio pilus.

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through the dodecameric secretin-like PilQ complex in the outer membrane, allowing the pilus access to the outside of the cell. Atop the pilus sits PilC1/Y1, which possibly plays a role in adherence to surfaces, both cellular and otherwise. This, however, may be absent in some bacteria. Examination of the genome of Bdellovibrio bacteriovorus HD100 showed that it contains a full set of genes required for the production of type IV pili (see Table 3) leading to the hypothesis that type IV pili are in some way required to maintain the Bdellovibrio life style (Rendulic et al., 2004).The gene order in the pilus operons seemed to be largely conserved between Bdellovibrio and another d-Proteobacterium, M. xanthus, which shows social twitching motility and is also a predatory bacterium, although an extracellular predator. However, there appears to have been some chromosomal rearrangement of genes since the two species diverged. Table 3 Type IVa pilus-associated genes, their functions and homologs in the Bdellovibrio bacteriovorus HD100 genome Gene

Function

HD100 gene number

pilA pilB pilC pilD pilF pilG

Pilus fiber protein ATPase pilus extrusion Pilus biogenesis Pre-pilin peptidase and methylase Required for PilQ multimer formation Part of ABC transporter required for pilus biogenesis with pilHI Part of ABC transporter required for pilus biogenesis with pilGI Part of ABC transporter required for pilus biogenesis with pilGH Required for pilus biogenesis

Bd1290 Bd1509 Bd1511 Bd0862 Bd3829 Bd1291

pilH pilI pilM pilN pilO pilP pilQ pilS pilR pilT pilE, V, W, X, fimU

Pilus biogenesis Pilus biogenesis Stabilizes PilQ multimer complex in outer membrane Outer membrane multimer complex through which PilA fiber is extruded Negative regulator via two component system with PilR regulates pilA expression via two component system with PilS (Best homolog) ATPase pilus retraction Minor Pilins

Bd0860 Bd0861 Bd0863 Bd1585 Bd0864 Bd0865 Bd0866 Bd0867 BD1512 Bd1513 Bd3852 No homologs

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Compared to the generic model put forward by Mattick (2002), the Bdellovibrio pilus-associated genes differ slightly, mainly in the absence of the genes pilC1/Y1 which form the tip used in adherence of pili to surfaces and pilE, pilV, pilW, pilX and fimU, all of which are minor pilins in other bacteria (see Fig. 4b). Work by Wu and Kaiser (1995) in M. xanthus showed that pilG, along with pilH and pilI form part of an ABC transporter responsible for PilA membrane localization as well as PilA functionality. The PilGHI proteins also seem to be only found in d-Proteobacteria and these may be the Bdellovibrio equivalent of the minor pilins. The absence of the pilC1/pilY1 gene does not mean that Bdellovibrio cannot adhere to cells and other surfaces. However, as work carried out in Pseudomonas aeruginosa showed, the presence of cysteine residues at the C-terminus of PilA plays a role in cellular adherence. Bdellovibrio also have these cysteine residues at the PilA C-terminus, Cys 121 and Cys 132, which can potentially negate any need for a specific adherence protein at the tip of the extruded pili by, in some way, binding to receptors within the peptidoglycan layer of the Gram-negative prey. In the Bdellovibrio genome there appears to be various duplications of genes involved in pilus production, mainly pilT and pilQ (based on BLAST evidence), with two homologs of pilT found in the genome. Bd1510 (pilT1) is located within the operon pilBTCSR and Bd3852 (pilT2) is found elsewhere in the genome. pilT2 shows the greatest similarly to the pilT of Bdellovibrio’s close relative M. xanthus. Three homologs of pilQ were identified. This is not much of a surprise as PilQ is very similar to the type II secretion protein PulD and some may have separate secretory roles. However Bd0867 is the best match to that of the M. xanthus pilQ.

3.3. Proposed Regulation of Bdellovibrio Pili The transcription of pilA has been shown in M. xanthus (Wu and Kaiser, 1995) to be positively regulated by PilR, the sensory receptor of a two component regulatory system, and negatively regulated by PilS, the sensory kinase. In the Bdellovibrio bacteriovorus HD100 genome, there are homologs for both pilR and pilS found next to each other, Bd1512 and Bd1513, respectively, in the pilBTCSR operon. It has been found in M. xanthus that pilA auto-regulates its transcription by a feedback mechanism. This is possibly due to a pool of immature PilA monomers lying beneath the inner membrane ready to be used to extend the pilus when required. The initial stimulus for pilus extension is unknown but

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is possibly a chemical stimulus from outside the cell, such as products of the hydrolysis of the prey cell wall or outer membrane caused by the Bdellovibrio. Alternatively, it could be due to the physical stimulus of the prey being in contact with the Bdellovibrio’s outer cell surface.

3.4. Flp Pili Certain bacteria encode a different set of pili, members of the type IVb, known as Flp (fimbria-like protein) pili, which are generally used in cell adherence to surfaces. They have been characterized mainly in Caulobacter crescentus (Skerker and Shapiro, 2000), Haemophilus ducreyi (Nika et al., 2002) and Actinobacillus actinomycetemcomitans (Henderson et al., 2003). This subset of pili plays an important part in pathogenicity and the formation of biofilms in these bacteria. As well as type IVa pili, the HD100 genome also encodes some genes potentially involved in Flp pilus production based on their similarity to genes of Actinobacillus actinomycetemcomitans (Wang et al., 2003). BLAST homologs in the Bdellovibrio HD100 genome show that it seems to be lacking most of the major components needed to express Flp pili (see Table 4) including the major Flp pili subunit (flp-1). Wang and Chen (2005) showed, by producing null mutants in each of the Flp pili associated genes, Table 4 Type IVb (flp) pilus-associated genes, their functions and homologs in the Bdellovibrio bacteriovorus HD100 genome Gene

Function

Bdellovibrio homolog

flp-1 flp2 tadA

Pilus subunit Duplication of flp1 ATPase

tadB tadC tadD

Pilus anchor Pilus adherence protein (similar to pilC1/Y1) Production of fimbriae

tadE tadF tadG tadZ rpcA

Pseudopilin Pseudopilin Pilus anchor related protein Possible extracellular secretion vesicles Secretin similar to pilQ

rpcC

Function unknown

N/A N/A tadA (Bd0111) cpaF (Bd0793) tadB (Bd0110) tadC (Bd0470) aglT (Bd0833) Bd0225 N/A N/A N/A N/A gspD pilQ N/A

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that RcpB, TadE, TadF, TadG and TadZ are absolutely required for Flp biosynthesis, whereas Bdellovibrio appears to be missing homologs of rcpB, rcpC, tadE, tadF, tadG, tadV and tadZ. So far, there have been no attempts to delete the Flp genes to ascertain their role in the Bdellovibrio lifecycle. Interestingly, Beck et al. (2005) ascribe flp-1 and flp2 to Bd0119 and Bd0118 respectively as potential homologs. The reason for this is unclear as the genes show no strong resemblance to any Flp pilin sequences. When they performed SDS PAGE (Sodium Dodecyl Sulphate Polyacrylamide Gel Electrophoresis) and LC-MS (liquid chromatography mass spectrometry) of cell envelope preparations, they were unable to locate any Flp-1 protein, the main subunit of a type IVb Flp Pilus; they did, however, find PilA in the extracts, suggesting further that type IVa pili are important in Bdellovibrio and Flp pili may not be.

3.5. Expression of Pilus Genes in the Predatory Cycle Semi-quantitative RT-PCR analysis of pilA, pilQ, pilT1, pilT2, pilD, tadA and pilG has been carried out over the timecourse of Bdellovibrio’s life cycle (Evans et al., 2007). An initial infection of Bdellovibrio on prey E. coli S17-1 was set up, cells were then taken from the culture at each time point and total RNA was prepared. Controls of prey-only (E. coli S17-1) and Bdellovibrio-only were also sampled. Also prepared was RNA from HIpilA::Kn, the HI pilA mutant which lacks pili and a HI control that was wild-type for pilA. The results showed that the gene pilA, encoding the major pilus subunit, was constitutively expressed throughout the predatory timecourse, as well as in the HI cells, allowing for a ready pool of PilA subunits to be present throughout the lifecycle. This potentially allows for fast pilus formation when a prey-contact stimulus is perceived. For pilQ (Bd0867) and pilD (Bd0862) expression seems to be lower in the initial stages of the predatory timecourse and peaks at around the three to four hour mark of the lifecycle, when new attack phase Bdellovibrio are septating and being released by bdelloplast lysis. A possible explanation for this is that the PilQ and PilD proteins are only found at the pole of the Bdellovibrio and not found laterally, as in some species, so expression is only seen after septation of the Bdellovibrio and the establishment of new poles. Bdellovibrio septation within the bdelloplast occurs at the three to four hour mark and so expression peaks at a similar time. The expression seen in the HI phase is likely due to lack of synchronicity in the HI culture with various

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septation/division steps occurring throughout the incubated life cycle and small Bdellovibrio cells being present at all times. Expression of the pilT1 gene (Bd1510) was found to be at a relatively low unchanging level throughout the timecourse, whereas expression of pilT2 (Bd3852) seems to increase initially and fall at the two to three hour mark, yet increase at the four hour mark. Surprisingly, there appears to be no pilT2 expression in the HIpilA::Kn mutant suggesting a pilA/pilT interaction of some kind in which the presence of PilA in some way regulates PilT. pilG showed level expression through the time course. As its predicted function is as a PilA exporter, it makes sense that its expression is similar to that of the pilA gene allowing for a pool of pilin monomers to be polymerized when required. Interestingly, tadA (Bd0111), an ATPase associated with Flp pili production in other bacteria, shows an expression pattern of gradual increase in the time course peaking at three to four hours as well, being highly expressed in the HI strain, whilst there seems to be a reduced amount of tadA expression in HIpilA::Kn. This suggests an underlying pilA co-regulation with tadA whilst further work is required to establish the reasoning behind this and any role for TadA in the type IVa pilus of Bdellovibrio.

3.6. The Role of PilA in Predation The role of pili in predation was shown by Evans et al. (2007) in a study where the gene encoding the main pilus subunit pilA was interrupted with a kanamycin cartridge. Host- (prey-) dependent mutants could not be made despite exhaustive attempts, whereas host-independent (HI) mutants were readily constructed suggesting that the gene is essential for the infection process. This was proven as mutant HI strains were screened using a predation assay in which they were spotted onto a confluent lawn of E. coli and incubated for several days, yielding no area of clearing or plaques as was seen with other Bdellovibrio strains including many other mutants. Further proof was presented with a fluorescent predatory assay, in which fluorescent YFP E. coli prey were incubated with the pilA-interrupted Bdellovibrio and inspected for the presence of infected prey bdelloplasts. None were found despite extensive microscopy which had shown bdelloplasts in control HI strains, including other kanamycin cartridgeinterrupted mutants such as fliC3. In conclusion, Bdellovibrio have a full set of genes required to encode type IVa pili and such structures can be seen at their non-flagellated pole.

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These have been shown to be essential for predation and are probably involved in pulling the predator through the pore in the prey outer membranes, although further microscopic work is required to confirm this.

4. OUTER MEMBRANE PROTEINS Investigations into the roles of the surface appendages above serve to highlight the importance of these structures in early prey contacts. Intimate prey contact of course involves the Bdellovibrio outer membrane and indeed there are many questions about the adhesive, transport and enzymatic/ hydrolytic nature of proteins in the outer membrane and how they contribute to prey access and degradation by Bdellovibrio. The program PSORTb (version 2.0.4; http://www.psort.org/psortb/) predicts that 100 proteins of B. bacteriovorus HD100 localize to the outer membrane. Some of these are predicted by homology to build structures discussed in this chapter, such as the pili (PilQ; Bd0867 and Bd0112) and the sec secretion system (PulD; Bd1597 and Bd3510). Some are predicted to form porins with defined substrates, such as nucleosides (Tsx; Bd0667 and Bd3258) and sugars (LamB; Bd1230). Four genes are predicted to code for proteins involved in siderophore/iron transport (Bd1392, Bd1572, Bd2648 and Bd3571), while six genes are predicted to code for Type I secretion systems of the TolC family (Bd0707, Bd0887, Bd0999, Bd1797, Bd1915 and Bd2228). Barabote et al. (2007) identified a further 2 TolC family genes (Bd1408 and Bd3059) and these authors review the genes encoding Bdellovibrio transport systems in more detail. The gene annotated as ompH (Bd1494) is predicted to be a chaperone involved in omp transport to the outer membrane and is located next to other genes with similar predicted function. A large number (W60% of predicted outer membrane proteins) have no homology to proteins of known function and, so, many of these may be specific for predator-prey interactions. Of particular interest would be any proteins physically interacting with the prey cell in the initial contact. Whilst some work has been done on the elusive nature of the receptor in the prey (Schelling and Conti, 1986), very little work has been done on the predator’s prey recognition apparatus, and this is likely to involve some outer membrane proteins. It should be noted, however, that while instructive in designing practical experiments, in silico prediction work is not definitive and also that different methods give different protein locations; thus, much gene inactivation and physiological work is needed in the future to confirm the roles of outer membrane proteins.

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One interesting predicted outer membrane protein (Bd0296) has homology to the MEROPS family of proteinase inhibitor alpha-2macroglobulin and this raises the question of why Bdellovibrio would have a protease inhibitor in the outer membrane. It is possible that this is involved in inhibiting the actions of its own outer membrane proteases until they are needed. For example, it could form a complex with proteases inhibiting them until contact with a prey cell disrupts this complex and activates the proteases in order to break through the prey outer layers. Alternatively, it may interact with prey periplasmic proteases to inactivate them and protect the invading Bdellovibrio cell surface. Its expression pattern is shown by RT-PCR in Fig. 5 which shows that, while there is always some expression of this gene, suggesting that at least some of the

Figure 5 RT-PCR of various outer membrane protein genes over a time course of infection. L – NEB 100bp ladder, AP – attack phase, 15 – 15 min, 30 – 30 min, 45 – 45 min, 1 h – 1 hour, 2 h – 2 hour, 3 h – 3 hour, 4 h – 4 hour, S17 – negative control of E. coli S17-1 prey only, NT – no template negative control, þve – positive control of B. bacteriovorus HD100 genomic DNA, HI – host-independent.

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product is always present, the expression peaks late in the cycle at about 3 hours. As this is around the time of the progeny bdellovibrios producing lytic enzymes and breaking out from the remains of the prey ghost, it suggests that this outer membrane protein may be used to protect the Bdellovibrio from its own enzymes. Table 5 lists the major OMPs of Bdellovibrio bacteriovorus HD100 as predicted by PSORTb, BLAST and the experimental evidence of Beck et. al. (2005) and Barel et al. (2005). Size and pI were calculated using the Expasy compute pI/MW tool (http://expasy.org/tools/pi_tool.html) using the whole predicted peptide, so final protein size may be lower as a result of posttranslational modification, such as cleavage of the signal peptide (predicted to be B20 amino acids at the N-terminus of each). Of those predicted by PSORTb, one was annotated (on the basis of BLAST searches) as LamB, the maltooligosaccharide-specific porin, and the others were annotated as OmpA, which in E. coli can interact with host receptor molecules and is Table 5 Predicted major OMPs of Bdellovibrio bacteriovorus HD100 Name

Predicted Predicted Best BLAST hit in size (KDa) pI x-BASEa

Bd0380 18.3 ompA

9.55

Bd2616 48.2 ompA

9.1

Bd1494 18.7 ompH

9.4

Bd0427 37.8 ompF

4.86

Bd1230 49.7 lamB

5.18

Bd0883 39.8 ompA

4.79

Bd2769 22.2 ompA

9.65

a

Outer membrane protein Microscilla marina ATCC 23134 Outer membrane protein Hahella chejuensis KCTC 2396 Outer membrane chaperone Skp (OmpH) Pelobacter propionicus DSM 2379 Hypothetical protein Silicibacter sp. TM1040 Porin, LamB type Acidobacteria bacterium Ellin345 OmpA/MotB family protein Burkholderia sp. 383 chromosome 1 OmpA/MotB domain protein Anaeromyxobacter sp. Fw109–5

x-BASE database; http://xbase.bham.ac.uk/

E-value

Evidence

7e–13

PSORTb, BLAST

3e–45

PSORTb, BLAST

1e–25

PSORTb, BLAST

0.020

PAGE (Beck et al., 2005) PSORTb, BLAST

6e–42 2e–08

BLAST only

5e–42

BLAST only

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thought to be a major structural link between the outer membrane and the peptidoglycan (Koebnik et al., 2000). Two further OmpA homologs are predicted from BLAST searches (Barabote et al., 2007). Faint bands and spots are visible on the gels of Bdellovibrio outer membrane preparations presented in Beck et al. (2005) and Barel et al. (2005), respectively, which could correspond to the mature forms of some of these OMPs, but these were not investigated further as they did not represent the major OMP. By far the most interesting OMP identified is OmpF. This was not predicted by bioinformatics and does not have significant homology to other porins outwith the delta- Bdellovibrio and like organisms (BALO) group, but was determined experimentally to be the major porin of attack phase bdellovibrios (Beck et al., 2004). Early work had suggested that this porin was directly acquired unmodified from E. coli as similar bands were seen by SDS-PAGE (Diedrich et al., 1984), and OMPs of similar digestion patterns were isolated from bdellovibrios and their prey which also crossreacted with an anti-OmpF antibody (Talley et al., 1987). An OmpF of differing proteinase digest pattern was, however, isolated from both prey-dependent and prey-independent bdellovibrios, questioning these results. Recent application of modern techniques has elucidated the issue; Beck et al. (2004) identified the major OmpF as coming from Bdellovibrio by mass spectroscopy and reverse genetics, showing that it significantly differed from the respective prey OMPs. Barel et al. (2005) found that percoll gradient separation was a drastic improvement on previous techniques and resulted in good separation of Bdellovibrio cells from prey ghosts. Coupled with twodimensional gel electrophoresis and mass spectrometry, this technique revealed that Bdellovibrio did not take up any OMPs from prey, but rather, and intriguingly, that the Bdellovibrio encoded OmpF was at some point inserted into the prey cell envelope as it remained within the prey ghost; earlier work had suggested this, but without confirmation of the identity of the proteins involved (Tudor and Karp, 1994). Confirmation that separation of Bdellovibrio and prey ghost was complete was demonstrated as no other predator protein searched for by this technique was detected in the prey ghosts, including some that are very highly expressed. These data suggest that, at some point, Bdellovibrio inserts its OmpF into the inner membrane of the prey; this could be of great benefit to the predator. If this occurs soon after entry, it may be that it disrupts the proton motive force of the prey inner membrane, effectively killing the prey and overcoming the problems that would be associated with predator control over transport across an energized prey membrane. Alternatively, or in addition, the porin in the prey membrane could allow free passage of small molecules from the prey cytoplasm to the periplasm to allow the predator to take them up and utilize

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them for growth. It is interesting to note, as did Beck et al. (2004), that the secondary structure prediction reveals mostly alpha helices as compared to antiparallel beta sheets more common in outer membrane proteins and that this may facilitate its insertion into the prey inner membrane. Expression studies by RT-PCR of some of these OMPs are shown in Fig. 5. Transcripts from ompF and ompH show similar patterns where their genes are expressed throughout the cycle, but are drastically upregulated at 2–4 hours in the cycle. At this point, the bdellovibrios are growing considerably, then septating and lysing their prey. It could well be that these proteins are polarly organized and this is why they are mainly turned on at such a relatively late stage as septation begins then, allowing the cells to specify and organize their poles. Studies with antisera would be interesting in order to determine which OMPs, if any, are polar. Genes ompA1 and ompA2 both show increasing expression throughout the predatory cycle and the amounts by which they do so suggest that this is effectively constant expression per genome, and thus per cell-equivalent as expression rises around 45 min when genome replication begins within the growing, unseptated Bdellovibrio filament and continues to rise towards the end of the cycle. As with increased numbers of genomes, there is a concurrent increase in Bdellovibrio cell surface as the predator elongates in the bdelloplast. This could result in a constant proportion of OmpAs throughout the cycle which is consistent with the idea of these proteins having some sort of a structural role in the envelope. One very interesting result from the expression studies presented here is the expression pattern of the maltoporin lamB. Its expression rises dramatically at 45 min when the predator is rapidly breaking down its prey and presumably maltose and small polymers of maltose are becoming available. Then, expression decreases rapidly at 2 hours, presumably as these have all been taken up and used by the predator. The reason that this result is so intriguing is that it is the first clear indication that, within prey, bdellovibrios have an intact outer membrane that is distinct from the prey membranes and that they must be taking up small molecules of degraded prey macromolecules which have leaked out into the periplasm. Barel et al. (2005) also demonstrated that the outer membrane proteins of prey OmpA (E. coli) and OrpF (P. syringae) are modified by Bdellovibrio during predation; these proteins were cleaved into at least two parts during the predatory cycle. It is not known if this is necessary for prey outer membrane disruption for the predator to break out, or if the modification is for controlling the permeability of the bdelloplast such that nutrients in the prey periplasm do not leak away from the predator and into the external media. The OMP protein sequences of predator and prey need to be

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significantly different in order that the predator can selectively cleave that of its prey without damaging its own.

5. AUTOTRANSPORTERS In addition to porins, the Bdellovibrio genome is predicted to encode many autotransporter (AT) proteins. AT secretion is a wide-spread outermembrane-located protein secretion system in Gram-negative bacteria (designated type V) with more than 700 described members with various functions including proteases, adhesins and cytolysins (Henderson and Nataro, 2001; Pallen et al., 2003; Yen et al., 2002). AT proteins are usually large proteins with three major domains: an N-terminal signal peptide for recognition and transport from the cytoplasm by the Sec type II transport system, a secreted passenger domain and a C-terminal translocator domain which ends up in the outer membrane (Henderson et al., 1998). N-terminal signal peptides targeting to the Sec system allow AT proteins to translocate through the bacterial inner membrane (Henderson et al., 1998; Desvaux, 2004 #105). In the periplasm, the C-terminal translocator domain containing 250 to 300 residues form 10 to 14 b-barrel strands and this structure inserts itself into the outer membrane (Henderson et al., 1998; Jacob-Dubuisson et al., 2004). The crystal structure of the AT NalP in Neisseria meningitidis showed a 12-strand b-barrel with a hydrophilic pore of 10  2.5 A˚ (Oomen et al., 2004). After forming a barrel structure with a central channel, the passenger domain is translocated across the outer membrane via an unclear mechanism which may involve other outer membrane proteins (Desvaux et al., 2004; Jacob-Dubuisson et al., 2004; Jain and Goldberg, 2007; van Ulsen and Tommassen, 2006). The conserved motif in the a-helical linker between the passenger domain and the C-terminal translocator may play an essential role in secretion across the outer membrane (Kostakioti and Stathopoulos, 2006). Once on the cell surface, cleavage can then occur to release the mature protein into the extracellular milieu. The immunoglobulin A (IgA) protease in N. gonorrhoeae has a serine protease domain which is involved in autocleavage (Henderson et al., 1998; Pohlner et al., 1987). In contrast, an E. coli adhesin involved in diffuse adherence (AIDA-I) lacks a serine protease domain, but is capable of autocatalytic processing and remains on the cell surface via a non-covalent interaction with the translocator domain (Charbonneau et al., 2006; Henderson et al., 2004; Suhr et al., 1996). Besides the conventional AT model, a second family of ATs called trimeric ATs with a short C-terminal domain has been established

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(Henderson et al., 2004; Cotter et al., 2005). YadA, an adhesin in Yersinia involved in host cell adherence, extracellular matrix binding and serum resistance, is a well-studied trimeric AT (El Tahir and Skurnik, 2001). YadA has a short C-terminal translocator domain consisting of only 70 residues which is predicted to contain four strands and is insufficient for translocating the passenger domain (Hoiczyk et al., 2000; Roggenkamp et al., 2003). The crystal structure of the YadA passenger domain revealed a trimer which is proposed to be the trimeric translocator which forms a 12-strand barrel, including 4-strands from each monomer (Cotter et al., 2005; Nummelin et al., 2004). A trimeric translocator structure has been confirmed by a crystal structure study of an adhesin, Hia, in Haemophilus influenzae. The translocator of Hia forms a 12-strand b-barrel with a 18 A˚ diameter central channel which is traversed by three a-helices, and these are essential for the stability of the translocator (Meng et al., 2006). A stable trimerized structure for both YadA and Hia is essential for folding and the adherence function of the mature structure (Cotter et al., 2006). After translocation, the YadA passenger domain is still linked with C-terminal domain (Hoiczyk et al., 2000). This property can be found so far in all characterized trimeric ATs that all perform adhesive roles (Cotter et al., 2005; Kim et al., 2006). There are many putative ATs with a large size encoded by the B. bacteriovorus genome including both adhesive and secretory types (see Table 6). In silico predictions by SMART (http://smart.embl-heidelberg.de/) show that the N-terminal regions of all of the ATs contain a Sec-dependent cleavable signal peptide (Letunic et al., 2006; Schultz et al., 1998). These signal peptides are recognized by the Sec secretion system and translocate the ATs through the inner membrane to the periplasm. Phylogenetic analysis using the Lasergene Megalign program (Dnastar) suggested the existence of two major groups containing different functional domains (see Fig. 6). The first group, consisting of eight proteins, is the YapH-like protein group. YapH in Yersinia pestis is a putative AT adhesin which may be involved in attachment to host cells (Yen et al., 2002; Styer et al., 2005). Bioinformatic analysis also showed the average size of characterized YapH-like ATs with adherent functions from different bacteria to be 1435 7 804 amino acid residues, which is similar to the B. bacteriovorus YapH-like ATs with an average size of 1429.75 7 161.77 (Yen et al., 2002). Moreover, YapH-like ATs of B. bacteriovorus have a coiled-coil motif in their C-terminus predicted by SMART. The coiled-coil motif can also be found in many YadA-like ATs (Hoiczyk et al., 2000). Therefore, the YapH-like ATs in B. bacteriovorus may belong to the trimeric AT family and play a role in adherence to the prey cell surface during predation.

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Table 6 Classification and function of putative autotransporters in Bdellovibrio bacteriovorus Gene no.

Sizea

N-terminal Domain predicted by signal SMART or PROSITE peptide

Description in NCBI Database

YapH-like Bd0705 1267

1-22

coiled-coli

1211–1255

YapH

1492

1-22

coiled-coli

1438–1480

Bd3266

1567

1-21

coiled-coli

1525–1556

Bd3267

1507

1-21

coiled-coli

1454–1494

Bd1712

1365

1-25

coiled-coli

1310–1365

Bd2565

1164

1-36

coiled-coli

1123–1164

Bd2582

1416

1-22

coiled-coli

1380–1416

Bd2548

1660

1-28

coiled-coli

1624–1660

CAE76884 cell wall surface anchor family protein CAE79051 putative YapH protein CAE78079 cell wall surface anchor family protein CAE78080 putative cell wall surface anchor family protein CAE79581 cell wall surface anchor family protein CAE80361 phage related tail fibre protein CAE80377 cell wall surface anchor family protein CAE80346 hypothetical protein

RTX-like Bd1623 2828

1-32

CUB

1214–1327 1657–1765 2150–2199 2200–2249 2250–2299 2681–2730 609–658 659–708 759–809 809–862 320–424 926–975 976–1025 1462–1511 322–432 1106–1155 411–459 717–766 767–829 830–881 469–563 809–899

RCC1_3

Bd2644

891

1-18

RCC1_3

Bd1137

1625

1-27

CUB RCC1_3

Bd1247

1595

1-32

Bd3390

1077

1-20

CUB RCC1_3 RCC1_3

Bd1449

2576

1-26

FN3

CAE79499 putative RTX family exoprotein

CAE80437 conserved hypothetical protein CAE79055 hypothetical protein CAE79147 conserved hypothetical protein CAE78188 hypothetical protein Bd79336 cell wall surface anchor family protein

1219–1312 (Continued )

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Table 6 (continued ) Gene no.

Sizea

N-terminal Domain predicted by signal SMART or PROSITE peptide RCC1_ 3

Bd2486

1128

1-22

RCC1_3

Bd1921

901

1-31

RCC1_3

a

1937 –1993 1993–2044 2044–2093 2342–2392 801–849 950–999 1048–1099 226–281 330–384 385–439

Description in NCBI Database

CAE80289 putative UVB-resistance protein CAE79772 conserved hypothetical protein

numbers of amino acyl residues.

Figure 6 A phylogenetic tree demonstrating the two clusters of Bdellovibrio autotransporters.

The second group consisting of eight predicted Bdellovibrio proteins is that of the RTX (repeat in toxin)-like proteins. RTX proteins do not normally contain an N-terminal cleavable signal sequence, but instead have a C-terminal secretion signal which is recognized by a type I secretion system (Kostakioti et al., 2005; van Ulsen and Tommassen, 2006). However, the eight RTX-like proteins in B. bacteriovorus may not be secreted by type I secretion system due to the presence of an N-terminal cleavage signal peptide for type II secretion. The protein domains predicted by PROSITE (http://www.expasy.org/prosite/) for the proteins in this group show they all contain 1–4 regulators of chromosome condensation repeats (RCC1_3) (Hulo et al., 2006). The regulator of chromosome condensation (RCC1) repeat is involved in mitosis control in the eukaryotic cell, acting as a chromosome-binding protein and a guanine-nucleotide-exchange factor for

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a nuclear GTP-binding protein, Ran, to promote the replacement of GDP with GTP (Bischoff and Ponstingl, 1991; Seki et al., 1996). The crystal structure of RCC1 showed a seven-bladed propeller which is formed from protein internal repeats for Ran and DNA binding. Each blade with 51–68 residues forms 4-strand anti-parallel b-sheets (Renault et al., 1998). This suggests the multiple RCC1_3 repeats in B. bacteriovorus proteins may also form a similar b-sheet structure for interaction with other proteins or for DNA binding. Besides the RCC repeat, a putative CUB protein domain, which is a eukaryotic extracellular domain with approximately 110 residues containing four conserved cysteines for forming two disulfide bridges (C1-C2, C3-C4), could be predicted in Bd1623, Bd1137 and Bd1247 which instead have only two cysteines (C1 and C2). Furthermore, the structure of the CUB domain has been predicted to be an antiparallel b-barrel similar to that of immunoglobulins (Bork and Beckmann, 1993). Therefore, the CUB domain may also form b-barrel structures in Bd1623, Bd1137 and Bd1247 for insertion into the prey outer or cytoplasmic membranes, which could make catalytic enzymes from B. bacteriovorus pass through either membrane easily or cause leakage of host cytosolic contents. In Bd1449, three fibronectin type III (FN3) domains with approximately 100 amino acyl residues have been predicted. The crystal structure of an individual FN3 domain showed two b-sheets with four strands and three strands (Leahy et al., 1992, 1996). The FN3 domain in Bd1449 may contribute to interact with either a receptor or a substrate on the host cell surface. Although bioinformatic analysis offers a glance into the potential functions of Bdellovibrio ATs, the mechanism for secretion and the role of these proteins in predation is still being elucidated experimentally.

6. THE ROLE OF THE SEC SYSTEM Using the program pSORTb, 808 peptides of the Bdellovibrio genome are predicted to have signal sequences of the sort recognized by the sec system. This export machinery is likely to be extremely important for the predator, especially given the absence of type III secretion systems, save that for the bacterial flagellum. Analysis of the KEGG database shows that Bdellovibrio unsurprisingly has homologs for all of the genes required for a functional sec system. Interestingly, in addition to multiple pilQ homologs (see above), there are two homologs for the similar pulD. This could further suggest the importance of the sec system in a similar way to the multiple copies found of the important genes discussed above. Figure 7 shows that secA, the major

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Figure 7 RT-PCR of genes for sec proteins over a time course of infection. L – NEB 100 bp ladder, AP – attack phase, 15 – 15 min, 30 – 30 min, 45 – 45 min, 1 h – 1 hour, 2 h – 2 hour, 3 h – 3 hour, 4 h – 4 hour, S17 – negative control of E. coli S17-1 prey only, NT – no template negative control, þve – positive control of B. bacteriovorus HD100 genomic DNA, HI – host-independent.

transmembrane protein, and both of the pulD homologs (the other annotated as gspD) are all expressed throughout the infectious cycle, but are upregulated towards the end of the cycle. This is likely because of the growth of the Bdellovibrio at this stage in the cycle and so the amounts of the sec-related proteins per cell are likely to remain roughly constant.

7. MEMBRANE CHEMISTRY Early work on Bdellovibrio LPS (Kuenen and Rittenberg, 1975; Nelson and Rittenberg, 1981b; Stein et al., 1992) suggested that the core lipid A from the prey was incorporated largely intact into the predator LPS. Schwudke et al. (2003) later showed that the lipid A structure of the predator and prey are significantly different and that therefore it is likely that, despite their strenuous attempts to ensure separation of the predator and prey samples by ficoll gradients in the early experiments, separation was not completely efficient.

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In their study using more modern techniques, Schwudke et al. (2003) determined the lipid A structure in detail and discovered that it was based on a novel backbone structure of two (1u-6)-linked b-D-GlcN3N in the central region and two a-D-Manp linked in positions 1 and 4u. This backbone has not been described elsewhere and is completely uncharged (having no phosphate residues), leading to drastically altered fluidity of acyl chains within liposomes. In contrast to liquid crystalline phase transition of E. coli lipid A and LPS (T = 46 1C and 35 1C, respectively), those of B. bacteriovorus HD100 were as low as 5 1C and 10 1C, respectively, suggesting that this LPS is in a much less ordered state, probably because of the missing negative charges inhibiting cation bridging within the outer membrane. This and the presence of at least two unsaturated acyl chains per lipid A molecule result in greater fluidity, which in turn could lead to greater permeability, possibly advantageous in the protected environment of the bdelloplast for uptake of broken down prey molecules. Also, this high fluidity may be of value in helping the predator squeeze through the entry pore of the prey. This novel structure also results in the Bdellovibrio LPS having low induction of TNF-a and interleukin-6 relative to that of E. coli; these properties could prove useful if Bdellovibrio were ever developed as a therapeutic agent. A variety of fatty acids common in other Gram-negative bacteria were found in early work on Bdellovibrio (Nelson and Rittenberg, 1981a) with a major fatty acid tentatively identified as non-adecenoate (19:1). Analysis of the KEGG database shows that Bdellovibrio has homologs for most of the genes of the fatty acid biosynthesis and metabolism pathways and so probably builds and breaks down fatty acids in a way similar to that by other bacteria. Similarly, the genome has homologs for the standard genes of the LPS biosynthesis pathway and so probably adds the sugars of the core and O-antigen in a manner similar to that of other bacteria. Comparison of the LPS between the prey-dependent HD100 and preyindependent, axenically grown HI100 revealed significant differences between the two strains (Schwudke et al., 2003), which the authors concluded must have resulted from accumulating mutations in the HI100 strain. However, later work by the same group (Schwudke et al., 2005) suggested that this strain was actually derived from the strain 109J rather than HD100 (as was originally thought) and this is likely to be the reason for any differences. As a result, a valid comparison between a prey-dependent strain and its prey-independent derivative remains to be carried out. Steiner et al. (1973), working with Bacteriovorax stolpii (although then named Bdellovibrio stolpii), found that in both the prey-dependent UKi1 and the prey-independent UKi2 strains, the predominant glycero-phospholipids

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were phosphotidylethanolamine (PE; B85%) and phosphotidylglycerol (PG; B15%). These phospholipids and their relative abundance are typical for Gram-negative bacteria and presumably make up the inner membrane and the periplasmic face of the outer membrane. This work also showed the presence of three alkali-stable phosphonosphingolipids in the outer membrane and this is far more unusual for bacteria. Later workers (Jayasimhulu et al., 2007; Watanabe et al., 2001) analyzed the structures, identified 18 molecular species and confirmed the phosphonyl nature of these lipids. They were dominated by those with unique 1-hydroxy-2aminoethane phosphonate (hydroxy-aminoethylphosphonate) polar head groups. It would be very interesting to determine if these are unique to B. stolpii or are commonly found in BALOs. Sphingolipids are much more common in eukaryotes and found in relatively few bacteria. In eukaryotes, sphingolipids play an important role in cell signalling, heat-stress response, Ca2þ homeostasis and transport (Olsen and Jantzen, 2001) but their role in bacteria is as yet unknown. Their presence in the Bdellovibrio outer membrane is likely to affect significantly the properties of this membrane and may affect the potentially high levels of fluidity that the uncharged lipid A structure could confer upon the structure. In the presence of penicillin, attack phase Bdellovibrio cells lose their peptidoglycan walls, but can form sphaeroplasts which are stable without the need for osmotic buffering, suggesting that the outer membrane is relatively strong as most other bacteria would lyse in this treatment (Thomashow and Rittenberg, 1978b). Sphingolipids arranged in rafts, rather than spread evenly throughout the membrane, are of importance to the invasion of mammalian cells by pathogens (Gulbins et al., 2004) and it may be that they form rafts at localized regions of the Bdellovibrio outer membrane, such as the pole of attachment, and aid in interactions with the prey bacterial surface. The arrangement of lipids and sphingolipids in the predator may result in a modified overall charge on the Bdellovibrio surface which may help in its interaction with the prey cell surface. Two observations have led to the suggestion that the attack phase predator surface has a strong hydrophobic nature: 1) they adhere to octyl-Sepharose beads in conditions where other Gram-negative bacteria do not (Cover and Rittenberg, 1984) and 2) they are unusually sensitive to low levels of detergents (Thomashow and Rittenberg, 1985b). Bdellovibrio are unusual in that they have a sheathed flagellum (see above), which seems to be continuous with the outer membrane (Seidler and Starr, 1968). Analysis of this sheath (Thomashow and Rittenberg, 1985b) showed that it was composed of similar fatty acids to that of the outer

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membrane, but that LPS-containing non-adecenoic acid (19:1) was in higher proportions in the flagellar sheath compared to intraperiplasmic cells as a whole, indicating that the sheath was of a different composition to the rest of the outer membrane. Given that the sheath may have to slide over the rapidly rotating flagellum, it is likely that its composition is considerably more fluid than that of the rest of the cell to reduce the friction, and hence energy, required for the bdellovibrio’s rapid motility. In addition to remodeling and recycling components of its own membranes, Bdellovibrio is likely to be involved in the breakdown of the prey’s membranes as the inner membrane especially is shown to diminish during predation. Thomashow and Rittenberg (1978a) showed that 25% of the amino sugars of the prey LPS were solubilized within the first 30 min of attack on the prey and this is likely to be due to the breaking down of the outer membrane at the point of attachment as is also thought to happen with other components of the prey cell wall, such as the peptidoglycan. Table 7 shows a large number of predicted lipases, including the esterase and the alpha/beta hydrolase classes, many of which are capable of degrading lipids to their components for use by the predator. It is difficult to predict which of these would be ‘‘housekeeping’’ genes for remodeling of the Bdellovibrio membrane during septation and which would be used for breakdown of the prey, or both. Some have signal peptides for type II secretion, although some may be exported by alternative means. The only BALO lipid-related enzyme to be studied in detail is the serine palmitoyltransferase (SPT), the rate-determining enzyme for formation of 3-ketodihydrosphingosine, the precursor of sphingolipids (Ikushiro et al., 2007). The work showed that, while eukaryotic SPTs were integral membrane proteins and those from Sphingomonas were cytosolic, the Bacteriovorax enzyme was bound to the inner membrane as a peripheral Table 7 Putative lipases of B. bacteriovorus Class

Predicted genes

Alpha/Beta Hydrolase I

Bd0282, Bd3462, Bd3206, Bd0664, Bd0448, Bd2083, Bd1622, Bd0340, Bd2417

Alpha/Beta Hydrolase II Alpha/Beta Hydrolase III Phospholipase D Patatin Esterase D Other

Bd1121, Bd0873 Bd3289, Bd0963 Bd1516, Bd1999, Bd0899 Bd0737,

Bd0030, Bd0910, Bd1192, Bd2017, Bd3533 Bd2389, Bd2858, Bd3603, Bd1257 Bd3882, Bd0201 Bd1031, Bd1415, Bd3128, Bd1918,

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membrane protein and it underwent a remarkable substrate inhibition at palmitoyl CoA concentrations higher than 100 mM, as does the eukaryotic enzyme.

8. PEPTIDOGLYCAN CHEMISTRY AND METABOLISM Dramatic cell wall modifications within both the Bdellovibrio and their prey are key to the predation process and predatory growth. Striking electron microscope images suggest that Bdellovibrio hydrolyze their cell walls just prior to prey invasion in order to soften the cell wall, allowing the predator to squeeze into the prey through the smallest possible aperture (Burnham et al., 1968; Evans et al., 2007). This activity may also provide a priming effect, aiding cell wall biosynthesis in the bdelloplast growth phase that follows. Modifications to the prey cell wall induced by Bdellovibrio infection lead to the formation of the spherical bdelloplast structure (Scherff et al., 1966). This structure provides both the space in which the Bdellovibrio grow and divide but may also yield some of the intermediates required for cell wall biosynthesis (Thomashow and Rittenberg, 1978c). Bacterial cell wall peptidoglycan (PG) consists of an elastic network of long glycan strands of two b1-4 linked amino-sugars N-acetylglucosamine and N-acetylmuramic acid ranging from 5 to over 100 residues long re-enforced by peptide crosslinks; it resists the cytoplasmic osmotic pressure while also contributing to the cell shape (Holtje, 1998; Popham and Young, 2003; Smith, 2006). The bacterial cell wall can be thought of as being one constantly changing molecule, the murein sacculus. A lactyl group present on the glycan’s muramic acid residue provides the platform to which the short peptide crosslinks are attached (Smith, 2006). The peptide bridges between strands are formed from a penta-peptide chain precursor synthesized in the cytoplasm. In Bdellovibrio, this precursor has the sequence: L-alanine, D-glutamate, meso-diaminopimelate (DAP), D-alanine and D-alanine (Thomashow and Rittenberg, 1978c). The sequence of this peptide is typical of that found in Gram-negative bacteria.

8.1. Cell Wall Precursor Biosynthesis is Retained by Bdellovibrio Murein biosynthesis requires a disaccharide penta-peptide precursor molecule (UDPMurNAc-pentapeptide) usually formed in the cytoplasm.

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Bdellovibrio only digests up to 20% of the prey cell wall; the rest of the cell wall is maintained to protect the resulting bdelloplast structure from uncontrolled lysis (Thomashow and Rittenberg, 1978c). The amount of peptidoglycan precursors that this digestion would yield is insufficient to form the multiple progeny formed in each infection. Bdellovibrio has therefore retained the six cytoplasmic enzymes MurA, MurB, MurC, MurD, MurE and MurF responsible for the de novo biosynthesis of this precursor and genes for its transport into the periplasm via a lipid carrier molecule. The murD gene seems to be duplicated in the Bdellovibrio genome, but whether this is differentially expressed in predation or is involved in accelerating cell wall growth or enhancing wall strength requires further experimentation. Bdellovibrio has genes that potentially encode the high molecular weight PBPs (penicillin binding proteins) such as PBP1A, PBP1B and PBP1C, which have transglycosylase activities that catalyze the formation of the b1-4 glycosidic links of the lipid-bound precursors to the nascent end of the peptidoglycan strand. They are responsible for bulk peptidoglycan synthesis in other bacteria. This activity both incorporates a new monomer into the sacculus and cleaves the bond between the monomer and the lipid carrier molecule, allowing the carrier to be recycled (Vollmer and Bertsche, 2007). Bdellovibrio also have putative genes for the smaller PBP 2 and 3 type proteins which are involved in strengthening the cell wall; these proteins, by analogy with E. coli, would be expected to be involved in cell wall elongation and cell division, respectively (Vollmer and Bertsche, 2007).

8.2. Cell Wall Hydrolase Activities in Bdellovibrio Bdellovibrio, like any other bacteria, are required to remodel their cell wall as they grow and septate. In addition they also have cell wall hydrolytic activities against their prey. Unpicking whether housekeeping genes carry out both roles will require further physiological experiments. The sacculus is an ever-changing molecule in non-predatory bacteria with an estimated turnover of 40–50% per generation in E. coli (Herve et al., 2007). Every bacterial cell, therefore, requires enzymes that continuously digest and modify the cell wall, creating sites to which the high molecular weight PBPs can add in new material. In addition to turnover, cell wall hydrolases are required for the final peptidoglycan cleavage event in the cell division process (Vollmer and Bertsche, 2007), and bacteria typically carry a specific set of hydrolases that carry out these tasks (see Fig. 8). It is possible that by gene duplication and specialization, Bdellovibrio may be employing

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Figure 8 Structure of the Gram-negative peptidoglycan and areas of hydrolytic activity. Adapted from Holtje (1998). Table 8 Major peptidoglycan modifying enzymes of Gram-negative bacteria and their homologs in B. bacteriovorus HD100 Activity

Gene

Protein

Bdellovibrio homolog(s)

Transglycosylase

sltY

Slt70

Aminidase

mltA mltB mltC mltD emtA amiA amiB amiC mepA pbpG dacB

MltA MltB MltC MltD EmtA AmiA AmiB AmiC

pbpE dacA dacD

PBP4 Bacillus PBP5 PBP6

Bd1285 Bd2711 Bd3575 mltA Bd0519, Bd0599 Bd2462 mltD Bd1125 X X X AmiC Bd2699 mepA Bd1032 X dacB Bd3244, Bd3459, Bd0816 Bd0436, Bd1951 dacA Bd2044 X

DD-endopeptidase DD-carboxypeptidase and DD-endopeptidase

DD-carboxypeptidase

PBP4

orthologs of genes normally encoding housekeeping peptidoglycan remodeling proteins for predatory peptidoglycan digestion. Table 8 summarizes Bdellovibrio homologs to known peptidoglycan remodeling enzymes of E. coli, whose general encoded remodeling functions are discussed below with possible predatory roles highlighted.

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The Bdellovibrio genome encodes for a large family of putative proteins related to the lytic transglycosylases. These enzymes are responsible for the breakdown of the b1-4 glycosidic bonds between monomers of the glycan strands (van Straaten et al., 2005). These enzymes can be identified as they all carry a Goose Egg White Lysozyme domain. Unlike traditional hydrolases, these enzymes form the 1,6anhydromuric acid ends blocking continued peptidoglycan synthesis (van Straaten et al., 2005). The majority of these enzymes (MltA, MltB, MltC, MltD and EmtA) are known to be bound to the inner leaflet of the outer membrane facing into the periplasm in E. coli. Bdellovibrio contains homologs to only three of the four mlt genes. It has been proposed that at least one of these genes (mltB) could be involved in a multi-enzyme complex that synthesizes peptidoglycan. Interestingly, Bdellovibrio contain three homologs to the soluble Slt70 lytic transglycosylase protein, usually found in the periplasm. This family of proteins in Bdellovibrio is intriguing as at least one member of this family of enzymes is likely to be responsible for the breakdown of the peptidoglycan of the prey forming the bdelloplast.

8.3. Aminidases Aminidases such as AmiC remove murein side chains by cleavage of the bond between the protein bridge and the glycan (Priyadarshini et al., 2006). These enzymes have been shown to be required for the accurate separation of division septa during cell division in E. coli, where deletion of these genes has been seen to result in the formation of filamentous cells. Bdellovibrio only possess one homolog belonging to this family, amiC, so it is possible that this does not have a prey peptidoglycan de-crosslinking role.

8.4. DD-Endopeptidases and DD-Carboxypeptidases The DD-endopeptidases are responsible for the cleavage of the peptide bridges holding the glycan strands together (Holtje, 1998). These enzymes play a role in murein synthesis as well as potential Bdellovibrio-specific roles in the formation of the bdelloplast. The DD-carboxypeptidase enzymes process the pentapeptide crosslinks of the murein so that they cannot be used to form peptide bridges. This is achieved by the removal of the terminal D-alanine residue from the pentapeptide precursor (Vollmer and Bertsche, 2007). Transglycosylases can no longer break this bond, which usually provides the energy required to form the bridge. It has been proposed that

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the interplay between these two enzyme activities during peptidoglycan synthesis in E. coli controls the rigidity of the cell wall network (Vollmer and Bertsche, 2007).

8.5. Bdellovibrio Cell Wall Hydrolyases Act upon the Prey during Predation Bdellovibrio cell wall hydrolases, in addition to stimulating cell wall biosynthesis and murein turnover, must provide the required activities for solubilization of the prey peptidoglycan to generate the pore for entry into the prey. Furthermore, recent images have shown that Bdellovibrio may de-crosslink its own cell wall, softening it sufficiently to squeeze through the smallest possible pore in the prey’s outer membrane (Evans et al., 2007). Radioactive labeling experiments carried out in the 1970s showed the progression of prey peptidoglycan digestion during Bdellovibrio infection (Thomashow and Rittenberg, 1978a). This was achieved by the labeling of both an amino acid (DAP) and an amino-sugar (glucosamine) present in the murein of the prey. Infection of these prey by Bdellovibrio caused the liberation of labeled material from the prey peptidoglycan. The amount of radioactivity remaining within the peptidoglycan in crude extracts was used as a measure of the extent of cell wall digestion. This study concluded that up to 15% of the prey’s peptidoglycan was rapidly solubilized in the first 15 min of infection; if this was concentrated on one area of the cell wall, then it could provide a gap large enough to allow the Bdellovibrio cell to enter. The rate of de-crosslinking reactions yielding DAP slowly decreased 15 min post infection, whereas release of glucosamine seemed to stop altogether. The prey’s cell wall is, therefore, relaxed in a part-digested condition for the majority of the Bdellovibrio infection forming the bdelloplast structure. The majority of the released labeled DAP was in the free monomeric form. In order for this to occur, the peptide bridges of the prey must have been exposed to two activities: First, an endopeptidase activity to cleave the bridge, then a carboxypeptidase activity to cleave the DAP from the remaining peptide chain. The Bdellovibrio genome encodes multiple homologs of the dacB gene which encodes the PBP4 protein. PBP4 is known to possess both activities predicted by this experiment. These homologs present prime candidates for the gene responsible for the de-crosslinking of the prey’s peptidoglycan during infection. The sudden arrest of glycan digestion observed in Thomashow’s experiment is intriguing, as it suggests that all potential modification sites within the prey’s cell wall may have been used at this time. The most

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plausible enzyme activity that could account for this observation is a soluble non-processive lytic transglycosylase. This activity would maintain the length of the glycan strands of the prey while completely inhibiting peptidoglycan synthesis. Three Bdellovibrio homologs of the sltY gene of E. coli exist encoding a soluble lytic transglycosylase that could potentially carry out this function. So, to summarize, we hypothesize that the duplication of multiple genes encoding peptidoglycan remodeling enzymes may have allowed them to become specialized to attack and remodel prey peptidoglycan in the bdelloplast, but further physiological work is needed to investigate these hypotheses. As is often the case in bacterial surface structures, it is actually the level of understanding of the physiology of the cell wall and the paucity of reconstituted cell wall systems that hold progress in check – certainly an area where advances in microbial physiology are still required!

9. SUMMARY We have reviewed the potential roles of Bdellovibrio surface structures in the predatory lifestyle of this remarkable bacterium and found that while active rotation of a flagellum is important for productive chance collisions with prey in liquid environments, it is not flagella, but rather the type IV pili at the non-flagellar pole that are required for prey invasion. We have discussed the patchwork of outer membrane proteins, especially the large evolutionarily related families of AT genes and their possible diverse roles in prey adhesion, attack and unusually, in insertion into prey membranes. The presence of sphingolipids in the membranes of Bdellovibrio may be an unusual adaptation conferring robustness when interacting with prey cells. The predatory remodeling of prey peptidoglycan is an area of real challenges for molecular physiological studies but we have shown how an interrogation of gene complement and gene duplication for certain enzymatic properties may hint at key mechanisms that have evolved to support predation. It is clear from detailed studies of duplicated flagellar genes that Bdellovibrio has extra genes encoding ‘‘spare parts’’ for some systems of prime importance to the predatory lifestyle. With renewed research interest teaming the excellent biochemistry and physiology of the Bdellovibrio scientists of the 1960s and 1970s with recent mutational studies, the predatory patchwork of membrane and surface structures is being revealed in its true light. It is a processing-plant for prey interaction and killing which affords Bdellovibrio the environmental luxury of dining, without

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competition, inside the bdelloplast – the evolutionary payback for maintaining its large genome of predatory functions.

ACKNOWLEDGEMENTS The authors would like to thank Marilyn Whitworth and Rob Till for technical assistance and the Wellcome Trust, the Human Frontier Science Program and BBSRC for financial support.

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Author Index Aaron, M., 40, 80–81 Aarsman, M.E.G., 75 Aballay, A., 341 Abbott, D.A., 279 Abel, S., 28 Abella, C.A., 117, 136, 139 Abeyrathne, P.D., 51 Abraham, W.R., 57–58, 111, 210, 216 Abram, D., 326 Abrams, W.R., 169 Abril, F.N., 161 Acker, G., 225 Ackermann, N., 341 Adamberg, K., 299 Adams, J., 291 Adams, M.W.W., 164 Addinall, S.G., 75 Adkins, J.P., 111 Aebersold, R., 295 Agabian, N., 39–40 Aguilera, J., 284 Aitchison, J.D., 286 Aizawa, S.I., 36, 321 A˚kesson, M., 275, 277–279, 299 Ala’Aldeen, D., 340–341 Albers, S.-V., 204 Albertsen, A., 140 Albrecht, T., 234 Albuquerque, L., 214–215 Aldridge, P., 27–28, 32, 36–37, 41–42 Alexandre, G., 56 Alexeeva, S., 73 Ali, J., 320 Alipour-Assiabi, E., 47 Allaire, A., 36 Allan, M.G., 36–37, 55 Allen, E.E., 209–210, 222, 244 Allen, R.C., 13–14, 31 Alley, M.R., 5, 13–14, 20, 23, 26, 61

Almeida, J.R.M., 284 Almering, M.J.H., 283, 285, 288–290, 292, 297–298 Alpers, C.N., 243 Alton, E., 214–215 Altschul, S.F., 160 Alvarez-Martinez, C.E., 20, 24–25 Amann, R., 114, 145, 166 Ambler, R.P., 160 Amesz, J., 116 Amiot, N.C., 28, 36, 54 Amos, L., 80 An, G., 260 An, S.-Y., 213–215 Andersen, G.L., 25 Anderson, D.E., 67 Anderson, D.K., 21 Anderson, K., 288, 290 Anderson, P.E., 33–34 Andre, B., 288, 290 Andreasen, A.A., 270 Angert, E.R., 76 Ansorge, W., 174 Antranikian, G., 206, 214–215 Anzai, Y., 57 Aono, T., 57 Apel, W.A., 221 Appel, B., 333, 337–339, 345–346 Appelt, N., 162 Appia-Ayme, C., 124, 130, 134, 156–157, 210, 222–223, 234 Arabshahi, A., 152 Arahal, D.R., 161, 166 Arana, N., 24–25 Arieli, B., 132 Arif, M., 6, 9, 11, 13, 24, 27, 42 Arifin, Y., 283 Aristidou, A., 296 Arkin, A.P., 288, 290

364

Armitage, J.P., 325 Arunasri, K., 121 Arzberger, I., 134, 140 Asahara, M., 213–215 Ashida, H., 162 Ashman, E.M., 52 Aslund, F., 176 Asoh, S., 79 Astashkin, A.V., 152 Astromoff, A., 288, 290 Auchtung, T.A., 146 Aukhil, I., 344 Ausmees, N., 12, 42, 77, 79–80 Auvinen, P., 274 Avedissian, M., 21 Ayako, N., 80 Baar, C., 318, 327, 330 Badger, J.H., 56 Baganz, F., 288 Baida, C., 41, 72, 78–82 Bailey, J.E., 267, 292 Bailey, S., 152 Bairoch, A., 343 Baker, B.J., 222, 244 Baker, T.A., 10 Baker-Austin, C., 216, 221, 224, 229 Bakker, B.M., 294, 296 Bakker, J.T., 287, 296 Baksteen, J., 269 Baldini, R.L., 20–21, 24–25 Baldock, M.I., 106 Ball, L.A., 161 Baltes, M., 298 Bamford, V.A., 157 Bammler, T., 274–275 Bandeiras, T.M., 234 Banfield, J.F., 209–210, 220, 222, 243–244 Bangham, R., 288, 290 Barabote, R.D., 316, 335, 338 Bardischewsky, F., 125, 155–160, 233 Barel, G., 337–339 Barford, J.P., 268

AUTHOR INDEX

Bar-Joseph, Z., 285 Barnett, J.A., 267 Barr, D.W., 220, 224 Barre, F.-X., 74 Barrell, B.G., 282 Barrett, C.J., 161 Barry, K.W., 146 Bartels, J., 260 Bartsch, R.G., 130–131, 140, 147 Baselga, R.M.-A., 73 Batenburg-van der Vegte, W.H., 270–271 Bateson, M.M., 107, 112, 126 Bathe, S., 132–133, 224 Battaglia-Brunet, F., 242 Batty, J.D., 240 Bauer, J., 268 Bauer, Z., 209 Baugh, L.R., 275 Bauld, J., 117 Bavendamm, W., 116 Bazire, G.C., 40, 44 Beard, S., 137 Beatty, J.T., 112–113, 140 Beck, A., 28 Beck, J.V., 206 Beck, S., 333, 337–339, 346 Becker, J., 344 Beckmann, G., 344 Beckwith, J., 75 Beer, D.G., 295 Beis, K., 48, 51 Bellis, M., 274 Belly, R.T., 217 Benda, R., 149, 175 Benes, V., 174 Beness, A.M., 48 Benito, R., 288, 290 Bennasar, A., 57–58 Bennett, B., 152 Ben-Shem, A., 58 Benson, A.K., 20 Benz, I., 340 Berendt, U., 175

AUTHOR INDEX

Berg, D.E., 173 Berg, H.C., 321, 327 Berg, I.A., 115 Berks, B.C., 124, 130, 134, 156–158 Berndt, C., 176 Bernhardt, A., 346 Berry, K., 5, 13, 20, 23, 26 Berthiaume, F., 340 Bertsche, U., 350, 352–353 Beudeker, R.F., 288 Beyer, R.P., 274–275 Beynon, J.D., 151, 175 Beynon, R.J., 296 Beyreuther, K., 340 Bhattacharya, S., 274–275 Bias, U., 150 Bibi, E., 58 Bick, J.A., 149, 174–176 Bigham, J.M., 228 Bigot, S., 74 Bill, E., 176 Billings, S., 320 Bingman, C.A., 152 Biondi, E.G., 6, 9, 11, 13, 24, 26–28, 42 Birrell, G.W., 288, 290 Birren, B., 274 Bischoff, E.R., 344 Bitto, E., 152 Blair, D.F., 320 Blake, R.A., 161 Blake, R.C., 137, 208, 222, 224 Blankenship, R.E., 112 Blanot, D., 350 Blanz, P., 214–215 Blattner, F.R., 320 Blick, R.J., 332 Bloch, C.A., 320 Blowes, D.W., 243 Blum, J.S., 111 Blum, P., 214–215 Bobst, C., 113 Bock, E., 234 Bodenmiller, D., 30, 37, 39–40, 43–45, 48, 51–54, 56

365

Boeke, J.D., 288, 290 Boer, V.M., 268, 274, 280, 283, 285, 288–290 Boeselt, I., 225, 231–232 Bofkin, L., 274 Bogdanova, T.I., 214–215, 218, 229 Boissinot, M., 214–215, 232 Bokranz, M., 143, 154, 163 Bokranz, W., 54 Boller, A.J., 161 Bonch-Osmolovskaya, E.A., 217 Bond, P.L., 216, 221, 224, 229, 243–244 Boniface, A., 350 Bonnefoy, V., 210, 222–223, 234 Boonstra, J., 290 Boorman, G.A., 274–275 Booth, J.W., 152 Bopp, L.H., 212 Bordes, P., 23 Bork, P., 75, 80, 341, 344 Bork-Jensen, J., 79 Borrell, J.L., 151 Bos, P., 125, 210–211, 223, 225, 233–234 Bosshard, H.R., 131 Bothe, E., 157 Botstein, D., 262, 274, 277–278, 292–294, 298 Botting, C.H., 48, 51 Bouhss, A., 81 Boulygina, E.S., 115 Bousquet, J., 214–215, 232 Bowman, J.P., 216, 221, 229 Bowtell, D.D., 272 Boyd, C.H., 35 Boyles, A., 274–275 Bracey, D., 283 Brachat, S., 288, 290 Bradford, B.U., 274–275 Brainard, A.P., 292 Brambilla, L., 268 Brasseur, G., 222–223 Brassinga, A.K., 61–63 Braster, M., 299

366

Brauer, M.J., 262, 277 Braughton, K.R., 45 Brecker, L., 345–346 Brende, P., 6–7, 16–18, 30, 76 Brenner, C., 152 Brettar, I., 118 Breukelen, Knijnenburg, B., 296 Bridge, T.A.M., 226–227, 229–230 Briegel, A., 67–69 Brierley, C.L., 239 Brierley, J.A., 239 Brige, A., 130–131 Brikun, I., 173 Brinkac, L.M., 56 Briscoe, G., 73 Bro, C., 275–280, 292, 295, 299 Broach, J.R., 277 Broadhurst, D., 274, 277–278, 282, 295–296, 299 Broadhurst, J.L., 242 Brock, K.M., 217 Brock, T.D., 117, 125, 213–215, 217, 224, 232 Brodie, E.L., 25 Brooks, E.E., 347 Brown, J.A., 288, 290 Brown, J.M., 288, 290 Brown, M., 274, 277–278, 282, 291, 295–296, 299 Brown, P.O., 260, 272, 278, 292–293 Bru¨ser, T., 131–132, 134, 149, 151–152, 160 Bruckmann, A., 263 Brun, Y.V., 3, 13–15, 20–21, 30–31, 36–37, 39–48, 51–54, 56, 67, 69–70, 72, 75–80 Brunak, S., 275, 277–279, 299 Brune, D.C., 106, 116–117, 123, 132, 134, 138, 140–143, 146–149, 155, 158–162, 167 Bruno, S., 157–158 Brunold, C., 149, 169, 175 Bruschi, M., 223 Bryant, D.A., 107, 117

AUTHOR INDEX

Bryant, R.D., 136 Bryantseva, I.A., 109, 111, 115, 121–124, 126 Buchan, A., 125 Bu¨chert, T., 149, 175 Buchert, T., 150 Buck, M., 23 Buddelmeijer, N., 75 Buhler, J., 295 Bulliard, V., 343 Bumgarner, R., 295 Bumgarner, R.E., 274–275 Buonfiglio, V., 137 Burland, V., 320 Burnette, D.T., 11, 27–28 Burnham, J.C., 322, 328, 349 Bursakov, S.A., 151 Bushel, P.R., 274–275 Bussemaker, H.J., 262 Bussey, H., 282, 288, 290 Butler, P.R., 262, 288, 291 Button, D.K., 288, 291 Bykova, S.A., 115, 126 Caballero, A., 33 Cakar, Z.P., 267, 292 Caldwell, P.E., 209–210 Calvete, J.J., 151 Calvin, M., 206 Cam, M.C., 274 Camarena, L., 320 Camilli, A., 54 Campanaro, S., 288, 290 Campbell, E.I., 169 Campos, A., 320 Cannon, G.C., 208 Capeness, M., 321 Ca´rdenas, J.P., 210 Carmel-Harel, O., 278 Carroll, T.W., 349 Carrondo, M.A., 145 Castenholz, R.W., 117, 125–126 Castrillo, J., 274, 277–278, 282, 295–296, 299

AUTHOR INDEX

Castrillo, J.O., 260 Cathala, N., 174 Caumette, P., 122 Causton, H.C., 278 Cavalli, G., 274 Cavanaugh, C.M., 146 Cerutti, L., 343 Chain, P.S.G., 113, 161 Chan, C.W., 28, 36, 54, 233–234 Chang, S.-S., 216 Chao, Y., 341 Chapman, J., 209–210, 222 Chapman, M.R., 52 Characklis, W.G., 236 Charbon, G., 40, 80–81 Charbonneau, M.E., 340 Chaturvedi, K., 274–275 Chaudhuri, R.R., 340 Chauhan, A., 323 Chauvistre, R., 127, 134, 140–141 Chaw, S.M., 217 Cheesman, M.R., 157 Chen, C., 332 Chen, G.A., 295 Chen, J.C., 30, 75 Chen, S.L., 6, 15, 24, 42, 54 Chen, W., 332 Chen, Y.C., 176 Chen, Z.D., 292 Chen, Z.W., 130 Cheng, K.J., 44 Cherest, H., 174 Chernikova, T.N., 111 Chernyh, N.A., 217 Chesbro, W.R., 299 Chew, A.G.M., 107, 117 Chien, P., 10 Chisolm, T.E., 212 Choi, D., 274–275 Christ, R., 57–58 Christen, B., 36–37, 55 Christen, M., 36–37, 55 Christen, R., 118 Christensen, D.A., 327

367

Christmas, R., 295 Chu, A.M., 288, 290 Chung, A.P., 214–215 Ciampaglio, C.N., 73 Cipollina, C., 277 Clarens, M., 242 Clark, D.A., 218–219, 221 Clark, J.A., 161 Clarke, B.R., 48, 51 Clements, K.D., 76 Clermont, G., 260 Cliften, P., 274 Clifton, S.W., 320 Cobley, J.G., 222 Codd, G.A., 288 Cohen, B.A., 274 Cohen, Y., 106 Cohen-Bazire, G., 139 Colangelo, C., 295 Cole, J., 45, 52–53 Collado-Vides, J., 320 Collie, E.S., 328 Collier, J., 17–18 Collins, P.J., 272 Collins, R.F., 48, 51 Colmer, A.R., 231 Connelly, C., 288, 290 Connerton, I.F., 177 Conti, S., 335 Conti, S.F., 322–323, 328, 338, 346, 349 Cook, A.M., 106 Coote, P.J., 283 Cope, L.D., 332 Copley, R.R., 341 Cordell, S.C., 73 Cordwell, S.J., 12, 42 Cornell, M., 274, 277–278, 282, 295–296, 299 Cornet, F., 74 Costas, A.M.G., 107 Costerton, J.W., 44, 136 Cotter, S.E., 341 Coupland, K., 225, 228 Courtney, L., 320

368

Coutte, L., 340 Cover, W.H., 347 Covert, J.S., 125 Cox, J.C., 222 Cozzone, A.J., 48 Cramton, S.E., 45 Craven, M.B., 5, 13, 20, 23, 26 Crick, F.H.C., 261 Cristina, X.P., 117 Crosson, S., 18, 55 Crouvoisier, M., 81 Crymes, W.B., 30 Cummings, N.J., 177 Cunningham, M.L., 274–275 Curtiss, M., 288, 290 Cusanovich, M.A., 130–131, 160 Cypionka, H., 112 da Costa, M.S., 214–215 da Silva, A.C., 21 Dahl, C., 118, 127, 131–136, 138, 140–143, 145–153, 156, 158–162, 165, 169, 175 Dalma-Weiszhausz, D.D., 272 Dambe, T., 157 Danford, T.W., 285 Dante, M., 320 Daran, A.J.R., 287, 296 Daran, G.M., 282 Daran, J.M., 280, 282–283, 285–286, 288–290, 296–298 Daran, J.T., 279, 282 Daran-Lapujade, J.M., 282 Daran-Lapujade, P., 275–276, 280, 282, 285–287, 289–291, 293–298 Daran-Lapujade, P., 279, 282, 296 Darland, G., 213–215 Dasgupta, M., 44 Daugherty, S.C., 56 David, L., 274 Davidian, J.C.E., 174 Davidovic, L., 58 Davidsen, T.M., 56 Davidson, M.W., 131

AUTHOR INDEX

Davies, D., 44 Davies, J.P., 174 Davis, B.K., 326 Davis, C.R., 161 Davis, K., 288, 290 Davis, N.W., 320 Davis, R.W., 272, 274, 280, 282, 288, 290 de Bruyn, J.C., 223, 225 De Castro, E., 343 de Groot, M.J.L., 296 de J.M., 287, 296 de Jong, G.A.H., 125, 162 de Keizer, A., 140 de Mattos, J.M.T., 262–263 de Mendoza, D., 176 de Nicola, R., 282 de Pedro, M.A., 80–81, 82 de Poorter, L.M.I., 279 de Smet, L., 131 de Swaaf, M., 293 de Vries, S., 125 de Winde, J.H., 268, 274, 280, 282–286, 288–291, 293–294, 296 De Witte, A., 272 DeBoy, R.T., 5, 13, 20, 23, 26 Degnen, S.T., 36 Deinhard, G., 214–215 DeLeo, F.R., 45 Demol, H., 130 DeMoss, J.A., 223 den Blaauwen, T., 73, 81 den Camp, R.O., 169 Deng, S., 274–275 Denger, K., 106 Denis, Y., 210, 223, 234 Denizot, F., 210, 223, 234 Dennis, J.J., 149, 175–176 DeRisi, J.L., 260, 272 Desikan, A., 274 Desvaux, M., 340–341 Detter, C., 161 Detter, J.C., 146 Deuster, O., 143, 146–148

AUTHOR INDEX

Deutschbauer, A., 288, 290 Deutzmann, R., 150, 152 Devasia, P., 136–137 Devay, J.E., 349 Dew, D.W., 242 Dhillon, A., 176 d’Hugues, P., 240, 242 Di Donato, A., 217 Dickinson, J.R., 283 Diderich, J.A., 268, 291–294, 296 Diedrich, D.L., 338, 345 Dietrich, F., 288 Dijken, Pronk, J.P., 293 Dijkstra, B.W., 352 Dilly, G.F., 146 Dimitrov, D.S., 274 Din, N., 14, 67, 70, 72, 75–76 Dingwall, A., 61 Divakaruni, A.V., 41, 72, 77–82 Do, K.F., 161 Do, L., 113 Dobrinski, K.P., 161 Dodson, R.J., 5, 13, 20, 23, 26, 56 Doi, M., 77 Dolata, M.M., 130, 160 Dolinski, K., 262 Domian, I.J., 5–7, 9, 12, 27, 42 Dong, C., 48, 51 Dong, X., 219 Doonan, C.J., 152 Dopson, M., 216, 221, 224, 229, 233–234, 242 Dorel, C., 52 Dorsch, T., 225 Doublet, P., 48 Dow, S., 288, 290 Downey, T.J., 274 Drake, H.L., 226, 230 Dreschers, S., 347 Dressman, H.K., 274–275 Drews, M., 299 Dreyfus, G., 320 Driessen, A.J., 204 Drijber, R., 214–215

369

Drummelsmith, J., 48 Druschel, G.K., 243 Du, F., 320 Duboc, P., 268 Duclos, B., 48 Dufresne, S., 214–215, 232 Dugan, P.R., 221, 235 Dujon, B., 282 Dunham, M.J., 274, 294 Dunkley, T., 274, 277–278, 282, 295–296, 299 Dunn, W., 274, 277–278, 282, 295–296, 299 Duran, C.P., 338 Durkin, A.S., 5, 13, 20, 23, 26, 56 Dutton, R.J., 33–34, 146 Dye, N.A., 72–73, 77, 79, 81 Dykhuizen, D.E., 291 Eanes, E.D., 139 Easter, J., 34–36, 71 Eatock, C., 122 Eckert, B., 135–136, 138 Edgren, H., 274 Edwards, K.J., 220, 244 Eglin, D., 36 Egorova, M.A., 214–215, 229 Ehrlich, F., 283 Eichhorn, E., 177 Eisen, J.A., 5, 13, 20, 23, 26, 146 Eisen, M.B., 278 El Bakkoury, M., 288, 290 El Tahir, Y., 341 Elbehti, A., 221–222 Elken, R., 299 Ely, B., 5, 13–14, 20, 23, 26, 30, 33, 42, 54–55, 57 Emmel, T., 234 Endrizzi, M., 274 Enemark, J.H., 152 Eng, J.K., 295 Engelhardt, H., 118, 133, 142, 156 Engelmann, T.W., 116 Engels, S., 143, 146–148

370

Engels-Schwarzlose, S., 145–146, 165 Engemann, C., 134, 140 England, J.C., 32 English, R.M., 208 Entcheva-Dimitrov, P., 37, 39–40, 45 Entian, K.D., 264, 278–279, 288, 290, 296 Epel, B., 157 Eppinger, M., 318, 327, 330 Erickson, H.P., 67, 69, 73, 80, 344 Ermolaeva, M., 5, 13, 20, 23, 26 Errington, J., 75 Esparza, M., 210 Espejo, R.T., 136 Espie, G.S., 208 Evans, K.J., 318, 321, 327, 330, 333–334, 349, 353 Faath, I., 143, 146, 149–150 Fahey, R.C., 140, 147 Fahrenholz, F., 143, 154, 163 Fal, Y.I., 115 Falsen, E., 118 Famili, I., 296 Fannin, R.D., 274–275 Farin, F.M., 274–275 Farquhar, R., 288 Fass, D., 58 Fausbøll, A., 275, 277–279, 299 Favinger, J.F., 116 Faza, B.I., 161 Feijen, M., 340 Feldblyum, T.V., 5, 13, 15, 20, 23–24, 2638, 42, 61 Feldmann, H., 282 Feller, U., 149 Feng, C.J., 152 Ferea, T.L., 292–293 Ferguson, N.L., 14 Fernandez, R.C., 340–341 Ferrandez, A., 325 Ferrera, I., 153 Ferriera, S., 114, 166 Ferris, M.J., 112, 126

AUTHOR INDEX

Ferra´ndiz, M.-J., 75, 80 Feucht, A., 75 Fiebig, A., 55 Fiechter, A., 268 Fields, S., 274 Figge, R.M., 71, 77, 79, 82 Figueras, J.B., 117 Figurski, D.H., 38 Fischer, B., 41 Fischer, E.R., 45 Fischer, J., 125, 155–158, 160, 233 Fischer, U., 131, 134, 141, 148, 150, 155 Fishelson, L., 76 Fisher, A.J., 151, 175 Fisher, M.C., 146 Fitzpatrick, K.A., 161 Flaherty, P., 288, 290 Flannagan, R.S., 323 Flikweert, M.T., 266, 293 Florea, L., 320 Fodor, B., 146 Folcher, M., 28, 36–37, 55 Fontanez, K.M., 146 Ford, R.C., 48, 51 Forest, K.T., 327–328 Fo¨rster, J., 296 Foster, B.A., 175 Foucher, S., 242 Foury, F., 288, 290 Fraenkel, E., 285 Francke, C., 296 Frank, J., 125, 233–234 Franc- ois, J.M., 268 Franza, B.R., 135–136, 138, 295 Fraser, C.M., 5, 13, 15, 20, 23–24, 26, 38, 42, 61 Frazao, C., 138 Free, P.D., 328 Freedman, J.H., 274–275 Freund, L.B., 46–47 Frey, P.A., 152 Freyermuth, S.K., 161 Friden, P., 285 Friedrich, C.G., 125, 155–160, 233

AUTHOR INDEX

Friend, S.H., 288 Frigaard, N.U., 117 Fritz, G., 149–150, 175 Fritzemeier, K., 176 Frothingham, R., 341 Fry, R.C., 274–275 Fu, D., 274 Fu, P., 296 Fuchs, B.M., 114, 166 Fuchs, G., 118, 121, 206, 209 Fuchs, T., 7 Fuhrman, J.A., 125 Fujita, K., 278–279 Fujita, M., 213–215 Fukai, S., 146 Fukuda, A., 78 Fukumori, Y., 162 Fullaondo, A., 296 Fulton, L., 274 Fux, C.A., 44 Fyson, A., 204 Gad’on, N., 209 Gaffron, H., 143 Galchenko, V.F., 115, 126 Gale, N.L., 206 Galibert, F., 282 Gallardo, C.A., 234 Galvani, C.D., 41, 77–79 Gambacorta, A., 216 Gancedo, C., 268 Gancedo, J.M., 282 Gao, Y., 176 Garcia, N., 320 Garcia-Lara, J., 75 Gardner, D.C., 274, 277–278, 282, 288, 295–296, 299 Garfinkel, D.J., 288, 290 Garrity, G.M., 112, 115, 126 Gasch, A.P., 278 Gaskell, S.J., 274, 277–278, 282, 295–296, 299 Gavel, O.Y., 151 Geertman, J.-M.A., 292–293

371

Geesey, G.G., 44 Gehrig, S., 36 Gehrke, T., 136–137 Geme, J.W.S., 341 Gemmell, R.T., 213 Genovese, S., 139 Gentalen, E., 288 Gentile, C., 272 George, G.N., 111, 140, 151–152 Gerber, G.K., 285 Gerdes, K., 71, 79–80 Gericke, M., 229, 233, 242 Gerke, C., 45 Gerstein, M., 288, 290, 295 Gerstel, U., 54 Gest, H., 116 Getzoff, E.D., 327 Gharib, T.G., 295 Gherna, R.L., 139 Ghigo, J.M., 75 Giaever, G., 288, 290 Gibson, J.L., 113, 144 Gierig, M., 136 Giese, B., 28, 36, 54 Gifford, D.K., 285 Giglio, M.G., 56 Gihring, T.M., 244 Giner, J.L., 347 Giordano, T.J., 295 Giovannoni, S.J., 125–126 Gish, W., 160 Gitai, Z., 72–73, 77, 79, 81 Giudici-Orticoni, M.T., 223 Giuseppin, M.L.F., 268 Glaeser, J., 112, 121, 123 Glasner, J.D., 320 Glo¨ckner, F.O., 114, 145, 166 Gu¨ldener, U., 288, 290 Gobec, S., 350 Gober, J.W., 23–24, 32–36, 41, 63, 71–72, 77–82 Goda, T., 278–279 Goddijn, O., 233 Godon, J.J., 242

372

Goebel, B.M., 242 Goeden, M.A., 320 Goesmann, A., 318, 327, 330 Goffeau, A., 282 Gogtova, G.I., 122 Goldberg, M.B., 340 Goldman, A., 341 Golecki, J.R., 118, 121 Golub, T.R., 276 Golyshin, P.N., 111, 210, 216 Golyshina, O.V., 210, 216 Gomes, C.M., 138, 234 Gomes, S.L., 20–21, 24–25 Gonin, M., 42–43 Gonzalez, J.M., 125, 161, 166 Gonzalez-Pedrajo, B., 320 Goodlett, D.R., 295 Goodman, R.M., 220 Gorbatyuk, B., 5, 19, 63 Gordon, D.B., 285 Gorlenko, V.M., 109, 111, 115, 121–126, 143 Gorokhov, A., 169, 175 Gorwa-Grauslund, M.F., 284, 292 Goswami, S., 176 Goto, K., 213–215 Gotte, D., 288, 290 Gottschalk, G., 206 Gotz, F., 45 Goymer, P., 27–28, 36 Grangeasse, C., 48 Granovskaia, M., 274 Grass, S., 341 Grassme, H., 347 Graumann, P.L., 73 Gray, G.O., 131 Green, M.R., 276 Greenbaum, D., 295 Greenwood, M.M., 224 Gregor, J., 320 Grelle, G., 157 Gresham, D., 274, 294 Gress, T.M., 271 Grewal, N., 320

AUTHOR INDEX

Griesbeck, C., 132–133 Grieshaber, M., 132 Griffin, T.J., 295 Griffioen, G., 277 Groeneveld, P., 277 Grogan, D.W., 219 Gronow, S., 345–346 Groot, Slijper, M.J.L., 287, 296 Groothuizen, M.K., 292–293 Gros, P., 340 Gross, R., 143 Grote, R., 214–215 Grotkjaer, T., 262, 275, 277–279, 299 Grunenfelder, B., 36 Grzesiek, S., 36–37, 55 Go¨tz, F., 45 Gu, X., 290 Gu, Z., 290 Guay, R., 214–215, 217, 232 Gueiros-Filho, F.J., 67 Guerrero, R., 139 Guidotti, G., 152 Guiliani, N., 137, 222 Guiliano, L., 111 Guisez, Y., 130–131 Gulbins, E., 347 Gulik, W.M., 287, 296 Gustafson, J., 224, 232 Gustafsson, L., 268 Gutierrez-Marcos, J.F., 169 Guyoneaud, R., 117 Gwinn, M.L., 5, 13, 20, 23, 26 Gygi, S.P., 295 Gyo¨rfi, K., 146 Haft, D.H., 5, 13, 20, 23, 26, 56 Hageage, G.J., 139 Ha¨hn-Hagerdal, B., 284, 292 Hakes, L., 274, 277–278, 282, 295–296, 299 Hall, R.J., 268 Hallberg, K.B., 209, 213, 218, 225, 228–229, 231–233, 237–239, 242, 244 Hamers, R.J., 220, 340

AUTHOR INDEX

Hanash, S.M., 125, 295 Hannett, N.M., 285 Hans, M., 271 Hansen, E.J., 332 Hansen, F.G., 73 Hansen, L.K., 275, 277–279, 299 Hansen, T.A., 123 Hansford, G.S., 220 Hanson, G.R., 157 Hanson, T.E., 113, 145, 163 Hao, O.J., 237 Hara, K., 217 Hara, Y., 214–215 Harbison, C.T., 278, 285 Harder, W., 288, 291 Hardy, G.G., 45, 52–53 Hardy, G.P.M.A., 262 Harfoot, C.G., 121 Harmer, T.L., 161 Harper, A., 274–275 Harrison, A.P.J., 224 Harrison, F.H., 113 Harrison, S.T.L., 236 Hart, S., 274, 277–278, 282, 295–296, 299 Hartl, D.L., 291 Hartog, M.M.P., 292 Harwood, C.S., 113, 325 Hasegawa-Mizusawa, M., 278–279 Hashimot, T., 322, 328, 349 Hassane, A., 291, 293–294, 296 Hathaway, J.C., 139 Hatzimanikatis, V., 292 Hauser, L.J., 113, 161 Hauser, N.C., 262, 264 Hauska, G., 130, 132–133 Hautaniemi, S., 274 Haverkamp, T., 169, 175–176 Hawkins, A.C., 325 Hayashi, H., 348 Hayashi, N., 344 Hayes, A., 262, 274, 277–278, 282, 288, 295–296, 299 Hazelwood, L.A., 282–283

373

Hazen, W., 125 Hazeu, W., 125, 210–211, 233–234 Hecht, G.B., 13, 27 Heck, A.J.R., 296 Heck, M., 287, 296 Heerklotz, H., 28 Heesemann, J., 341 Hegemann, J.H., 288, 290 Heidelberg, J.F., 5, 13, 20, 23, 26, 107 Heijnen, J.J., 268–271, 294, 296–298 Heinhorst, S., 208 Heinrich, H.J., 156, 159 Heisig, P., 150 Heising, S., 112, 123, 132, 173 Hell, R., 169 Hellingwerf, K.J., 111 Hellmuth, K., 264 Hellwig, P., 157 Hemmi, H., 214–215 Hemmings, A.M., 157–158 Hempel, S., 288, 290 Henderson, B., 332 Henderson, I.R., 340–341 Hendrickson, W.A., 344 Hengartner, C.J., 276 Hennetin, J., 274 Henrici, A.T., 44 Henriksen, J.R., 125 Hensel, G., 177 Hensen, D., 158–162 Herbert, R.A., 122 Herman, Z., 288, 290 Hernandez, M.E., 135 Herna´ndez-Navarro, A., 174 Herve, M., 350 Herzberg, C., 206 Hespell, R.B., 316, 325 Hess, D.C., 262 Hesselberth, J.R., 274 Hester, S., 274, 277–278, 282, 295–296, 299 Hettich, R.L., 222 Hewlins, M.J., 283 Hickey, M.J., 327

374

Hill, A.A., 275 Hille, R., 152 Hillson, N.J., 20–21, 25 Hind, A., 216, 221, 229 Hinkle, M.E., 231 Hinson, D., 220 Hinz, A.J., 30–31, 45, 48, 51–54, 56 Hipp, W.M., 143, 146, 149–150 Hiraishi, A., 113–114, 124–125, 211–213 Hirsch, P., 57 Hirschler-Rea, A., 122 Ho, N.W.Y., 292 Hoare, D.S., 57 Hoare, M., 268 Hobbs, G., 260 Hobbs, M., 328 Hobley, L., 316, 321, 325 Hoeft, S.E., 111 Hoffmann, H., 341 Hoffmeister, M., 132 Ho¨fle, M.G., 118 Hoheisel, J.D., 262, 264, 271, 282 Hoiczyk, E., 341 Holdt, G., 118, 134, 140 Holland, B., 75 Hollemeyer, K., 106 Holmes, A., 320 Holmes, D.S., 209–210, 212, 223, 234 Holo, H., 125 Holstege, F.C.P., 275–276 Holt, J.G., 112, 115, 126 Ho¨ltje, J.-V., 80–81, 349, 352 Holtzendorff, J., 5–7, 16–18, 30, 76 Holyoak, C.D., 283 Holz, B., 136 Holz, I., 216 Homma, T., 278–279 Hong, S.J., 64–65 Hood, L., 295 Hoover, T.R., 33, 56 Hopkins, G.W., 341 Horiuchi, T., 217 Hormes, J., 127, 134–136, 138, 140–141 Hoshino, T., 347

AUTHOR INDEX

Hoskisson, P.A., 260 Hostetler, J., 107 Hotchkiss, J., 260 Hottes, A.K., 5, 17, 19–21, 24–25, 30, 75 Hou, S., 320 Howard, G.T., 137 Hoyle, D., 274, 277–278, 282, 295–296, 299 Hryniewicz, M., 174 Hu, B., 73 Hu, P., 20–21, 25 Huang, C.C., 295 Huang, L., 219 Huber, H., 150, 209–210 Huber, R., 150 Huber, W., 274 Hugenholtz, P., 209–210, 222, 244 Hughes, K.T., 32 Hughes, T.R., 283 Hu¨gler, M., 161, 209–210 Huitema, E., 34–35, 54 Hulo, N., 343 Hultgren, S.J., 52 Humble, M.C., 274–275 Hung, D., 6–7, 11, 13, 16–18, 30, 76 Hunt, C.A., 260 Hunt, S.M., 347 Hunter, C.P., 275 Huntwork, S., 6, 9, 11, 14, 31 Hurban, P., 274–275 Huston, S.L., 151, 175 Hwang, D., 286 Hwang, S.Y., 272 Iannettoni, M.D., 295 Ideker, T., 295 Igarashi, Y., 209 Ikeuchi, Y., 146, 148 Ikushiro, H., 348 Imhoff, J.F., 109, 111–114, 118, 121–124, 131, 136, 141, 161, 173 Inagaki, K., 230 Ingledew, W.J., 204, 221, 224 Ingvorsen, K., 243

AUTHOR INDEX

Iniesta, A.A., 6, 9–10, 20–21, 25 Inoue, C., 208 Ireland, M.M., 40, 43 Ishii, M., 209 Ishino, F., 79 Islam, M.M., 348 Isono, Y., 214–215 Itoh, T., 217 Itoh, Y.H., 217 Ivanovskii, R.N., 111, 115 Ivanovsky, R.N., 115, 125–126 Iversen, E., 228 Iwahashi, H., 278–279 Iwai, T., 217 Iwaki, M., 214–215 Iwasaki, T., 217 Iyer, V.R., 260, 272 Izuka, S., 147 Izzo, V., 217 Jacob-Dubuisson, F., 340 Jacobi, C.A., 341 Jacobs, C., 11–13, 27, 42 Jacobs-Wagner, C., 11, 27–28, 34–35, 40, 77, 79–81 Jacq, C., 282 Jaeger, T., 28 Jaehning, J.A., 285 Jagtap, P., 318, 327, 330 Jahn, M., 209 Jain, S., 340 Jan, R.L., 217 Janakiraman, R.S., 3, 39, 42–44, 53–54 Janekovic, D., 216 Jansen, G., 278–279 Jansen, M.L.A., 286, 291, 293–294, 296 Janssen, A.J.H., 140 Janssen, P.H., 121 Janssen, W.M.T.M., 266 Jantzen, E., 347 Jaramillo, D.F., 288, 290 Jarvis, M., 35 Javens, J.W., 13–14, 31 Javor, B.J., 161

375

Jayasimhulu, K., 347 Jedlicki, E., 210, 223, 234 Jefferson, K.K., 44 Jenal, U., 7, 21, 27–28, 32, 36–37, 41–44, 53–56, 83 Jennings, E.G., 276, 278, 285 Jeno, P., 36–37, 55 Jensen, G.J., 67–69 Jensen, R.B., 65, 70–71, 73–74, 80 Jerez, C.A., 137 Jetten, M.S.M., 111 Ja¨ger, K., 169 Jing, L., 274–275 Joffe, J.S., 231 Johnson, D.B., 202, 213, 218, 220, 225–233, 237–239, 241–244 Johnson, D.E., 44 Johnson, J., 114, 166 Johnston, M., 274, 282, 288, 290 Jones, A., 157 Jones, H.C., 40 Jones, S.E., 14 Jones, T., 274, 288 Jones, W.L., 236 Jonsson, A., 52 Jonsson, L.J., 292 Jorgensen, P., 277 Jørgensen, B.B., 106, 117, 155, 161 Ja¨rvinen, A.K., 274 Judd, E.M., 6–7, 9, 11 Juliani, M.H., 21 Juneau, K., 274 Jurkevitch, E., 337–339 Justice, S., 75 Kader, A., 54 Kahru, A., 299 Kaiser, C.A., 290 Kaiser, D., 331 Kakuta, Y., 169, 175 Kalhorn, J.W., 290 Kallioniemi, O.P., 274 Kaluzny, K., 51 Kamimura, K., 234

376

Kamin, H., 176 Ka¨mpf, C., 111 Kanao, T., 234 Kanbe, M., 36 Kanbe, T., 213 Kaneshiro, E.S., 347 Kang, D.-K., 216 Kanin, E., 278 Kanno, N., 169 Kao, C.M., 278 Ka¨ppeli, O., 268 Kappler, A., 135 Kappler, U., 148, 151–153, 157, 175, 242 Karavaiko, G.I., 210, 214–216, 218, 229 Karczmarek, A., 73, 81 Kardia, S.L.R., 295 Karhumaa, K., 284, 292 Karp, M.A., 338 Karty, J.A., 40, 43 Kasai, H., 213–215 Kasemets, K., 299 Katagiri, T., 230 Kato, J., 146, 148 Kato, Y., 213–215 Kaufmann, W.K., 274–275 Kavanagh, T.J., 274–275 Keiler, K.C., 63–65 Kell, D., 274, 277–278, 282, 295–296, 299 Keller, H., 318, 327, 330 Kellis, M., 274, 285 Kelly, A.J., 14, 67, 75–76 Kelly, D.E., 288, 290 Kelly, D.J., 153 Kelly, D.P., 156, 218, 231, 233 Kelly, S.L., 288, 290 Kemmeren, P., 275 Keppen, O.I., 115, 125–126 Kerfeld, C.A., 161 Kerr, K.F., 274–275 Kertesz, M.A., 177 Kessel, M., 122 Khanna, S., 121, 123, 150, 155 Khouri, H.M., 5, 13, 20, 23, 26, 56

AUTHOR INDEX

Kiene, R.P., 125 Kim, D.S.H., 341 Kim, H., 57 Kim, S.Y., 77 Kimble, L.K., 126 Kimura, N., 217 Kimura, S., 213, 229, 232, 237–238 King, G.M., 111 King, J.R., 316, 325 Kinkhabwala, A., 77 Kinnunen, P.H.M., 242 Kinzler, K.W., 271 Kirchhoff, J., 149 Kirkpatrick, C.R., 285 Kirkpatrick, H.A., 320 Kishimoto, N., 211–212 Kisker, C., 152 Kitagawa, E., 278–279 Kitamura, K., 137 Klaasen, P., 268 Klatt, C.G., 107 Klebe, C., 344 Klein, A., 150 Klein, D., 30–31, 54 Kleinjan, W.E., 140 Klenk, H.P., 216 Kletzin, A., 138, 234 Klevecz, R.R., 271 Klimmek, O., 143, 154, 163 Klockenkamper, R., 157 Klotz, M.G., 161 Knaff, D.B., 131, 148 Knijnenburg, M.C., 282 Knijnenburg, T.A., 279, 282, 285, 297–298 Knobloch, K., 162 Knowles, C.J., 213 Knudsen, S., 275–276, 280, 295 Kobayashi, K., 162 Kodama, T., 209 Koebnik, R., 338 Koerkamp, M.G., 262 Koh, M., 130 Koh, S.S., 278

AUTHOR INDEX

Kojima, C., 162 Kojro, E., 143, 154, 163 Kolganova, T.V., 111, 121, 123 Kolk, A.H.J., 75 Kolkman, A., 296 Kolmert, A˚.K., 213 Kolonay, J.F., 5, 13, 20, 23, 26 Kolstø, A.-B., 80 Komatsu, Y., 278–279 Kompantseva, E.I., 109, 122 Kon, T., 217 Kondrat’eva, E.N., 111 Kondrat’eva, T.F., 210, 214–216, 229 Kong, W.W., 161 Konig, W.A., 234 Konings, W.N., 204, 269 Koonin, E.V., 58 Kopriva, S., 149, 169, 175–176 Koprivova, A., 175 Kort, R., 111 Kosako, Y., 211–212 Kostakioti, M., 340, 343 Kostanjevecki, V., 130–131 Kostka, S., 157 Kostrikina, N.A., 217 Kothari, S.P., 56 Ko¨tter, P., 283, 288, 290, 296 Kovacs, A., 146 Kovacs, K., 146 Koval, S.F., 323 Koyanagi, T., 147 Koyasu, S., 78 Kraakman, L.S., 277 Krafft, T., 143, 154, 163 Kraft, R., 157 Kra¨ling, M., 146 Krasil’nikova, E.N., 115, 124, 214–215, 218, 229 Kredich, N.M., 167, 169, 174, 176–177 Krems, B., 264 Kresnowati, M.T.A.P., 297–298 Krieger, R.S., 209 Krijger, G.C., 296 Krischke, W., 214–215

377

Krishnan, B.R., 173 Kristensen, T., 80 Kroczek, R.A., 271 Kro¨ger, A., 143, 154, 163 Krokotsch, A., 45, 51 Kroneck, P.M.H., 149–150, 175 Kruckeberg, A.L., 268, 296 Kruglyak, L., 274, 294 Kruse, T., 79 Kube, M., 145 Kuchler, K., 283 Kuenen, G.J., 125 Kuenen, J.G., 111, 125, 155, 161–162, 210–211, 223, 225, 233–234, 270, 288, 291, 345 Kuever, J., 121, 145, 149–150 Kuhlemeier, C., 169 Kuhn, J.R., 77, 79–80 Kuiper, A., 288 Kulakauskas, S., 173 Kulick, L., 36 Kumagai, H., 147 Kunugita, K., 79 Kurosawa, N., 217 Kurowski, W.M., 259 Kurtz, H.D., 48, 52 Kusai, K., 162 Kusano, T., 208 Ku¨sel, K., 225–226, 230 Kustu, S., 33 Kuypers, M.A.M., 112, 292 Kuznetsov, B.B., 115, 124–126, 217 Laat, W.T.A.M., 293 Labarre, J., 295 LaBonte, D., 288, 290 Ladd, T.I., 44 Lagenaur, C., 39 Lagniel, G., 295 Laishley, E.J., 136 Lam, H., 11, 27–28, 34–35, 40, 77, 80–81 Lam, J.S., 51 Lamarre, A.G., 322–323 Lamb, D.C., 288, 290

378

Lambert, C., 318, 321, 323, 326–327, 330, 333–334, 349, 353 Lamerdin, J., 113 Lampreia, J., 150 Lan, N., 288, 290 Land, M.L., 113, 161 Lander, E.S., 274, 276, 278, 285 Lane, T., 13 Lang, A.S., 112–113 Langdahl, B.R., 243 Lange, H.C., 268, 296 Langendijk-Genevaux, P.S., 343 Langille, S.E., 56 Langworthy, T.A., 212, 216, 235 Lankinen, H., 341 Lansdon, E.B., 151 Lanz, C., 318, 327, 330 Lapidus, J.A., 161, 274–275 Lara, J.C., 328 Larimer, F.W., 113, 161 Larsen, H., 123 Larson, D.E., 30–31, 48, 51–52, 54, 56 Larson, R.J., 43 Larsson, C., 268 Lasarev, M.R., 274–275 Lascelles, J., 156, 161 Lashkari, D.A., 272, 288 Latimer, J.L., 332 Latreille, P., 320 Lau, E., 146 Laub, M., 288 Laub, M.T., 5–6, 9–13, 15, 20, 23–28, 38, 42, 54, 56, 61 Laurich, C., 157 Lavrov, S., 274 Lawler, M.L., 30–31, 54 Lawrence, L.J., 323 Lawson, E.N., 242 Layman, D., 320 Le Thi, T.T., 52 Leahy, D.J., 344 Leal, S.S., 234 Leclerc, G., 14 Lee, H., 20–21, 25

AUTHOR INDEX

Lee, K.-B., 57, 292 Lee, L.W., 291 Lee, T.I., 276, 278, 285 Leech, A.P., 124, 130, 134, 156 Lefering, R., 260 Lefimil, C., 223, 234 Lehrach, H., 271 Leisinger, T., 177 Lejeune, P., 52 Lekunberri, I., 161, 166 Lemesle-Meunier, D., 221–223 Lempicki, R.A., 274 Lennon, G.G., 271 Lentine, K., 61–62 Leo, J.C., 341 Leonard, S., 320 Leroy, G., 223 Lessner, F.H., 65 Lester, R.L., 346 Letellier, M.C., 58 Letunic, I., 341 Leustek, T., 149, 169, 174–176 Levi, A., 28, 43–44, 53–54, 56 Leyh, T.S., 151, 173, 175 Li, C.M., 271 Li, G., 32, 37, 46–47 Li, H., 117 Li, J., 148, 169, 274–275 Li, P., 274, 277–278, 282, 295–296, 299 Li, W.H., 290 Li, Y.J., 274–275 Li, Z., 67, 69 Liang, H., 274, 280, 288, 290 Liao, H., 288, 290 Lichtenberg, H., 135–136, 138 Lide´n, G., 284 Liebundguth, N., 288 Lightfoot, R., 58 Lilley, K., 274, 277–278, 282, 295–296, 299 Lillig, C.H., 176 Lim, F.L., 262 Lince, M.T., 112 Lindenbergh, A., 296

AUTHOR INDEX

Lindholst, S., 57–58 Lindner, B., 345–346 Lindstro¨m, E.B., 231, 233, 242 Linscheid, M.W., 333, 337–339, 345–346 Lipman, D.J., 160 Liss, P., 125 Little, P.J., 124, 130, 134, 156–157 Liu, C.-T., 57 Liu, L., 288, 290 Livingston, S.P., 328 Llewellyn, M., 34 Lloyd, J.R., 135 Lloyd, S.A., 320 Lo, I., 244 Lobenhofer, E.K., 274–275 Lobos, J.H., 212 Locher, K.P., 338 Lockhart, D.J., 288 Lombardot, T., 145 Long, C.D., 327 Longo, D.L., 161 Loo, J.A., 78–79, 82 Loo, R.R.O., 78–79, 82 Lopez, R., 80 Lorenz, C., 143 Lory, S., 328 Lottspeich, F., 150 Louis, E.J., 282 Lourenco, A.I., 150 Lourenco, R.F., 20, 24–25 Lo¨we, J., 73, 75, 80 Loza-Tavera, H., 174 Lu, C., 69 Lu, W.-P., 156 Lu, W.Y., 262 Lu, X., 274–275 Lubitz, W., 157 Lu¨bbe, Y.J., 143, 146–148 Lucas, A.B., 272 Lucas, S., 161 Lucau-Danila, A., 288, 290 Lucet, I., 75 Ludwig, W., 112, 123, 132, 173

379

Lund, K., 223 Lunsdorf, H., 111, 121, 210, 216 Luo, C., 288, 290 Lupas, A., 341 Lussier, M., 288, 290 Lutkenhaus, J., 67, 75, 80 Luttik, M.A.H., 268, 283, 288–290, 296 Luyben, K.C.A.M., 269 Lysenko, A.M., 115, 126 Ma, K.S., 164 Macisaac, K.D., 285 Mack, D., 45, 51 MacLean, M.R., 209–210 Macmillan, C.B., 235 MacRae, I.J., 151, 175 MacRae, J.D., 55, 58 Madden, E.A., 268 Maddock, J.R., 5, 12–13, 20, 23, 26–27, 42–43 Madela, K., 346 Madigan, M.T., 111, 117, 125–126 Madupu, R., 56 Magaki, K., 230 Magasanik, B., 290 Mager, W.H., 277 Mahairas, G.G., 332 Mahen, E.M., 65 Mahr, J.A., 175 Maia, J.C.C., 21 Maier, B., 327 Mailinger, W., 298 Majors, B., 274 Makarova, K.S., 58 Makhneva, Z.K., 111, 121, 123 Malakooti, J., 20, 33 Maldener, I., 132–133 Malek, R.L., 274–275 Malfatti, S.A., 113, 161 Malone, J., 28, 54–55, 83 Mandelco, L., 111, 126 Mangan, E.K., 23, 33, 35 Mansilla, M.C., 176

380

Manske, A.K., 112 Mao, R., 288, 290 Marcinkeviciene, L., 234 Marczynski, G.T., 5–7, 11–12, 14, 16, 18–19, 42, 60–63 Marelli, M., 286 Maresca, J.A., 107, 117 Margolin, W., 80 Marison, I., 268 Marques, M.V., 24 Marren, D., 288 Marrie, T.J., 44 Marshall, K.C., 45, 55, 57 Martin, D.D., 161 Martin, M.E., 75–76 Martin, M.N., 174 Martin, R.L., 151 Martin, W., 132 Martinson, T.A., 112 Marzolf, B., 286 Mas, J., 139, 153 Mashego, M.R., 268, 270, 291, 293–294, 296, 298 Mason, J., 218 Massey, S.E., 161 Matheron, R., 122 Mathews, F.S., 130 Matias, P.M., 145 Matin, A., 288, 291 Matroule, J.Y., 11, 27–28 Matsubara, H., 214–215 Matsuhashi, M., 77, 79 Matsukura, H., 212 Matsumoto, N., 225 Matsumoto, R., 278–279 Matsumoto, T., 212 Matsumoto, Y., 124, 130, 134, 156 Matsumura, T., 214–215 Matsuura, K., 125 Matsuura, T., 212 Matsuzawa, H., 79 Matsuzawa, Y., 213 Matteson, D., 34–35, 54 Matthysse, A.G., 58

AUTHOR INDEX

Mattick, J.S., 328, 331 Mau, B., 320 Mayer, F., 125 Mayhew, G.F., 320 McAdams, H.H., 5–7, 9–11, 15–21, 24–25, 30, 38, 42, 54, 61, 66, 75–76 McAllister, S.A., 345 McClelland, M., 320 Mccoy, J.G., 152 McCuddin, Z., 161 McCusker, J.H., 272 McDade, R.L., 338 McDonnell, C., 48, 51 McEwan, A.G., 152, 157, 242 McGinness, S., 225–226, 228 McGrath, P.T., 6, 9–10, 20–21, 25, 30, 66 McSween, W., 62 Mederer, N., 132–133 Meewan, M., 24–25, 66 Meijer, W.G., 206, 208 Meijer, W.M., 210–211 Melby, T.E., 73 Melick, M., 56 Menard, P., 288, 290 Mendoza-Co´zatl, D., 174 Menendez, C., 209 Meng, G.Y., 341 Mengin-Lecreulx, D., 81, 350 Mensonides, F.I.C., 294 Merckel, M.C., 341 Merker, R.I., 45, 54, 57–58 Merz, A.J., 327 Meskys, R., 234 Messer, W., 18 Meulenberg, R., 125, 233–234 Mewes, H.W., 282 Meyer, B., 149–150 Meyer, F., 161, 318, 327, 330 Meyer, H., 57–58 Meyer, T.E., 130–131, 160 Meyer, T.F., 340 Meyerdierks, A., 145 Michel, C., 221

AUTHOR INDEX

Michels, P.A.M., 294 Mikhailov, V.V., 118 Mikkelsen, D., 242 Millar, W.N., 211 Miller, B.E., 176 Miller, W., 160, 320 Millett, F., 131 Milpetz, F., 341 Milton, S., 274–275 Minev, M., 75 Miranda, M., 274 Miroshnichenko, M.L., 217 Misek, D.E., 295 Mitchell, D., 48 Mittmann, M., 288 Mityushina, L.L., 109, 115, 122, 126 Miyada, C.G., 272 Miyake, T., 209 Mizukami, S., 278–279 Mizunashi, W., 230 Møller-Jensen, J., 71 Mochida, K., 213–215 Modig, T., 284 Modrow, H., 127, 134–136, 138, 140–141 Moerner, W.E., 9, 11, 77 Moffat, K., 55 Mohammadi, T., 81 Mohl, D.A., 71 Mohr, C.D., 20–21 Molitor, I., 150 Molitor, M., 150 Moll, H., 345–346 Molyneaux, S.J., 210 Momose, Y., 278–279 Monni, O., 274 Monod, J., 259, 261, 265–266, 268, 282, 288, 291 Monosov, E.Z., 111 Montero-Lomelı´ , M., 295 Montgomery, W.L., 76 Moore, E.R.B., 57–58, 210, 216 Moore, J.L., 161 Moore, R.L., 55, 57

381

Moosa, S., 236 Morais, F., 147, 150 Morais, M.A., 283 Moran, M.A., 125 Moreno-Sa´nchez, R., 174 Morgan, J.J., 219 Morin, D.H.R., 240, 242 Morr, M., 54 Morris, D., 288 Moskalenko, A.A., 111, 121, 123 Mothes, K., 167 Moune, S., 122 Moura, I., 151 Moura, J.J.G., 150–151 Mourez, M., 340 M’Rabet, N., 288 Mrazek, J., 56 Muff, T.J., 325 Muir, R.E., 33, 35–36 Mul, A., 262 Mu¨ller, F.H., 234 Mu¨ller, H., 162 Muller, M., 345–346 Mulvaney, E., 320 Munson, R.S., 332 Murakami, Y., 282 Murata, Y., 278–279 Murillo, M., 169 Murphy, M.J., 176 Murray, D.B., 271 Murray, H.L., 285 Murray, S.R., 17–18 Mussmann, M., 145 Myers, C.R., 135 Myers, E.W., 160 Myers, J.D., 153 Myers, J.M., 135 Myrberg, A.A., 76 Nagahisa, E., 169 Nagalla, S.R., 274–275 Nagasawa, N., 212 Nair, S.P., 332 Naismith, J.H., 48, 51

382

Nakajima, M., 347 Nakajima-Iijima, S., 77 Nakayama, T., 214–215 Namath, A.F., 272 Nanninga, N., 73, 75 Nashimoto, H., 177 Nassar, N., 344 Natarajan, K.A., 136–137 Nataro, J.P., 340 Naterstad, K., 80 Nausch, L., 132–133 Navarro-Garcia, F., 340–341 Nedelmann, M., 45, 51 Neerken, S., 116 Negishi, M., 169, 175 Negre, N., 274 Nelson, D.C., 151, 175 Nelson, D.R., 345–346 Nelson, K.E., 5, 13, 20, 23, 26, 56 Nelson, W.C., 5, 13, 20, 23, 26, 107 Nemoto, N., 217 Nesbakken, T., 126 Neugebauer, E., 260 Neumann, S., 169 Neuwald, A.F., 173 Newman, C.L., 343 Newman, D.K., 135 Newton, A., 5, 13–14, 20–21, 23, 26–27, 31, 36, 54, 74 Newton, G.L., 140, 147 Newton, I.L.G., 146 Nguyen, C., 320 Nhan, M., 320 Ni, L., 288, 290 Nicholas, D.J.D., 121, 123, 150, 155 Nichols, W.W., 45 Nickel, J.C., 44 Niederberger, P., 268 Nielsen, J., 259–260, 262, 268, 275–280, 285, 292, 295–296, 299 Nielsen, P.H., 236 Nierman, W.C., 5, 13, 20, 23, 26 Nijburg, J.W., 118, 134 Nika, J.R., 332

AUTHOR INDEX

Nimtz, M., 54 Ninfa, A.J., 20, 36 Nisamedtinov, I., 299 Nishimoto, T., 344 Nishimura, A., 146, 148 Nishino, T., 214–215 Nitschke, W., 221, 223 Niwa, M., 214–215 Nixon, A., 233 Nobre, M.F., 214–215 Nold, S.C., 112, 126 Noorman, H.J., 269 Nordstrom, D.K., 243 Normark, S., 52 Norris, P.R., 204, 209–210, 218–221, 224, 233 Northup, S.J., 284 Notomista, E., 217 Novick, A., 259, 261, 271, 290–291 Nowack, J., 149, 175–176 Numata, T., 146 Nummelin, H., 341 Nunn, D.N., 328 Nureki, O., 146 O’Brien, T.M., 33 Ocampo, L.H., 161 Odani, M., 278–279 Odom, D.T., 285 O’Donnol, D.S., 13–14, 31, 34, 42–43 O’Donoghue, K., 274, 277–278, 282, 295–296, 299 Oeller, P.W., 291 Ogasawara, N., 162 Ohgiya, S., 278–279 Ohmura, N., 137, 225 Ohta, N., 5, 13–14, 20–21, 23, 26, 31, 36 Ohta, T., 79 Okabe, S., 236 Okada, Y., 78 Okibe, N., 228–229, 232–233, 241–243 Oliver, S., 274, 277–278, 282, 295–296, 299

AUTHOR INDEX

Oliver, S.G., 260, 262, 274, 282, 288, 291, 296 Olsen, G.J., 239 Olsen, I., 347 Olse´n, A., 52 Olsson, L., 259–260, 285 Olsthoorn, M.M.A., 296 O’malley, J.P., 274–275 O’Neill, K., 56 Ong, C.J., 48 Onishi, A., 147 Ooi, S.L., 288, 290 Oomen, C.J., 340 Orawski, G., 156 Ordal, G.W., 325 Orellana, O., 210, 223, 234 Oremland, R.S., 111 Oren, A., 122 Ormerod, J.G., 126 Orringer, M.B., 295 Oshima, T., 217 Oshima, Y., 209 Osipov, G.A., 115, 126 Osorio, A., 320 Osteras, M., 21, 36 Ostrowski, J., 176 Otero, R.R.C., 292 Otto, M., 45 Ouimet, M.C., 16 Overmann, J., 112, 118, 121, 140–141 Overmann, R., 121, 123 Owen, J.P., 218 Oyaizu, H., 57 Paalme, T., 299 Pace, N.R., 76 Padan, E., 106, 130, 132 Pagni, M., 343 Pai, C., 288 Paiment, A., 48 Pallen, M.J., 340 Palm, C.J., 274 Palm, P., 219 Palmer, V.S., 274–275

383

Palsson, B.Ø. 296 Panoutsopoulou, K., 274 Panteleeva, E.E., 111, 121, 123 Paquin, C., 291 Parge, H.E., 327 Parham, N.J., 340 Park, W., 77, 79 Parkin, T.B., 117 Parrou, J.L., 268 Partovi, S.M., 340–341 Parveen, M., 278–279 Paschinger, H., 143 Paschinger, J., 143 Pascual, J., 161, 166 Pate, J.L., 43–44, 55–56 Patel, H.C., 175 Patil, K.R., 262, 285 Paton, N., 274, 277–278, 282, 295–296, 299 Pattaragulwanit, K., 138, 141–142, 160 Pattee, P., 274–275 Patterson, N., 274 Paul, J.H., 161 Paul, R., 27–28, 36, 54 Paules, R.S., 274–275 Paulsen, I.T., 5, 13, 20, 23, 26, 48, 56, 161 Payne, R.W., 267 Peabody, C.R., 340–341 Peck, H.D., 149 Pedersen, L.G., 169, 175 Pedros-Alio, C., 139, 161, 166 Pedroso, I., 209 Pellegrini, L., 58 Pelletier, D.A., 113 Pen˜a-Castillo, L., 283 Pennacchio, F., 217 Penttila, M., 296 Perchuk, B.S., 6, 9–11, 13, 24, 26–27, 42 Pereira, A.S., 150 Pereira, I.A.C., 145, 147, 150 Peres, C., 113 Perna, N.T., 320

384

Perou, C.M., 274–275 Petersson, A., 284 Petit, T., 268, 284 Petri, R., 118 Petrotchenko, E., 169, 175 Petty, J., 274, 277–278, 282, 295–296, 299 Petushkova, Y.P., 111 Pezacka, E., 206 Pfannes, K.R., 121, 123 Pfennig, N., 111–112, 141, 144, 156 Phadke, N.D., 5, 13, 20, 23, 26 Philippsen, P., 288, 290 Phillippsen, P., 282 Phillips, G.N., 152 Phillips, K., 274–275 Pibernat, I.V., 136, 139 Picardi, A., 217 Pichoff, S., 75 Pickering, I.J., 140 Piekarska, K., 262 Pierce, D.L., 13–14, 31 Pierson, B.K., 126 Pils, B., 341 Pincus, Z., 77, 79, 81 Pinhassi, J., 161, 166 Pinkert, S., 341 Piper, M.D.W., 268, 274–276, 280, 283, 288, 290–295 Piper, P.W., 283 Pires, R.H., 147, 150 Pirt, S.J., 259, 268 Pitka¨nen, J.P., 296 Pittman, M.S., 147 Pivovarova, T.A., 210, 216 Planet, P.J., 38 Plano, G.V., 341 Planta, R.J., 277 Plumley, F.G., 112 Plunkett, G., 320 Podgorsek, L., 161 Poggio, S., 320 Pohlner, J., 340 Poindexter, J.S., 3, 40, 42–44, 76, 78

AUTHOR INDEX

Pokholok, D.K., 285 Polidoro, M., 137 Ponstingl, H., 344 Ponting, C.P., 341 Poole, R.K., 147 Popham, D.L., 349, 352 Poralla, K., 214–215 Porro, D., 268 Porter, S., 33 Porwollik, S., 320 Possoz, C., 74 Postma, E., 268, 288, 294 Potocka, I., 5, 13, 20, 23, 26 Pott, A.S., 134, 140, 143, 146–147, 149–150, 156 Potter, L., 327 Pott-Sperling, A.S., 143, 146–148 Powers, T.R., 47 Prange, A., 118, 127, 133–136, 138, 140–142, 156 Prasol, M.S., 24, 26, 42 Pratt, J.M., 296 Pratt, J.T., 54 Pratt, N., 148 Pratt, S.C., 274, 294 Prensier, G., 52 Preston, J.F., 45, 51 Prieme´, A., 118 Priess, H., 217 Prigent-Combaret, C., 52 Prince, R.C., 140 Pritchard, S., 34–35, 54 Priyadarshini, R., 352 Prokofeva, M.I., 217 Pronk, H.V., 287, 296 Pronk, J.T., 125, 210–211, 223, 225, 233–234, 266–270, 274–276, 279–280, 282–286, 288–298 Pronk, M.C., 279, 282 Pronk, P., 277 Pronk, T.H., 296 Ptacek, C.J., 243 Puehler, G., 216 Puhakka, J.A., 242

AUTHOR INDEX

Pujalte, M.J., 161, 166 Purcell, E.B., 55 Qin, L.X., 274–275 Qiu, Y., 274–275 Quardokus, E., 67, 70, 72 Quardokus, E.M., 13–14, 31, 36, 40, 42–43, 67, 70, 75–77, 80 Quast, C., 114, 166 Quatrini, R., 209–210, 223, 234 Quentmeier, A., 125, 155–160, 233 Quigley, S.D., 274–275 Quintela, J.C., 80–81 Quintero, E., 56 Quintero, E.J., 55–56 Quon, K., 12, 16, 60 Quon, K.C., 5–7, 9, 11–12, 14, 16, 42, 60 Raaka, B.M., 274 Rabinowitz, J.D., 262 Rachel, R., 234 Rachubinski, R.A., 286 Radhakrishnan, S.K., 34–35, 54 Radonjic´, M., 275 Radune, D., 56 Ragagopal, S., 55 Raghevendran, V., 285 Rainey, F.A., 214–215 Rainey, P., 27–28, 36 Raitsimring, A.M., 152 Rakhely, G., 146 Ram, R.J., 209–210, 222 Ramakrishnan, G., 23 Ramana, C.V., 121 Ramı´ rez, P., 137 Ramsey, S.A., 286 Randles, C.I., 235 Ranish, J.A., 295 Rao, G.R., 136–137 Ras, C., 271, 297–298 Rash, B., 274, 277–278, 282, 295–296, 299 Rasmussen, T., 157 Ratouchniak, J., 222–223, 234

385

Rawling, D.C., 55 Rawlings, D.E., 220, 239, 242 Raymond, J.C., 121–122 Rebischung, C., 288 Reedy, M., 69 Reep, D.K., 161 Rees, D.C., 152 Rees, G.N., 121 Regenberg, B., 262, 275–280, 295, 299 Reichenbecher, M., 341 Reijerse, E.J., 157 Reilly, J.P., 40, 43, 77 Reinartz, M., 131–132, 134, 160 Reinders, M.J.T., 279, 282, 285 Reinders, T.A., 282 Reinhardt, J., 5 Reisenauer, A., 6–7, 10, 12, 16–18, 20–21, 30, 60, 72–73, 76–77 Reisinger, S.J., 6, 9, 11, 13–14, 27, 31 Remsen, C.C., 139 Ren, B., 278, 285 Ren, Q.H., 56, 161 Renault, L., 344 Rendulic, S., 316, 318, 321, 327, 330, 335, 338 Renner, W.A., 292 Renosto, F., 151, 175 Rep, M., 262 Reski, R., 176 Rethmeier, J., 152 Reuss, M., 268, 298 Revsbech, N.P., 125 Revuelta, J.L., 288, 290 Reynolds, D.B., 285 Riba-Garcia, I., 296 Richardson, P.M., 146, 209–210, 222 Richter, L., 112, 123, 132, 173 Richter, M., 145 Rieger, M., 268 Riekkola-Vanhanen, M., 240 Riffaud, C., 122 Rijkenberg, M.J.A., 111 Riles, L., 288, 290 Riley, M., 176, 320

386

Rinaldi, N.J., 285 Ringelberg, D.B., 208 Rist, B., 295 Rittenbe, S.C., 316, 325 Rittenberg, S.C., 317, 345–350, 353 Rizzi, M., 268, 298 Rizzo, M.F., 35 Robert, F., 285 Roberto, F.F., 210, 223, 228–229, 232, 234, 243 Roberts, C.J., 288, 290 Roberts, M.A., 169 Robertson, D.H.L., 296 Robertson, L.A., 155, 161–162 Robinson, H.C., 147 Robinson, L.S., 52 Rochon, Y., 295 Rode, C.K., 320 Rodland, M., 274–275 Rogers, L.A., 153 Roggenkamp, A., 341 Rogozin, I.B., 58 Rohde, M., 54 Rohwerder, T., 137, 234 Rojas, J., 136 Rokhsar, D.S., 209–210, 222 Rolfe, P.A., 285 Rolfe, S., 228 Romanenko, L.A., 118 Romeo, T., 45, 51 Romero, P., 24–25, 136 Romling, U., 54 Ronen, M., 298 Rorke, G.V., 240 Rose, D.J., 320 Rose, M., 288, 290 Rosenthal, D., 176 Rosenzweig, R.F., 292–293 Rosinus, A., 318, 327, 330 Rosovitz, M.J., 56 Ross, R.L., 161 Rossell, S., 287, 296 Ross-Macdonald, P., 288, 290 Roth, M., 344

AUTHOR INDEX

Roth, U., 226, 230 Rother, D., 125, 155–160, 233 Rothfield, L., 75, 79 Rouwenhorst, R.J., 271 Rowe, O.F., 213, 225, 229, 239, 232, 238, 244 Roy, A.B., 135 Rubin, E.M., 209–210, 222 Ruderfer, D.M., 274, 294 Rudra, D., 277 Rueger, D.C., 176 Rummel, G., 41 Ruohonen, L., 296 Rupes, I., 277 Rusyn, I., 274–275 Ruzicka, F.J., 152 Ryan, E., 320 Ryan, K.R., 6–7, 9–11, 13–14, 27, 31 Rzhepishevska, O., 234 Saar, J., 214–215 Saarela, J., 274 Sackett, M.J., 14, 67, 70, 75–76 Sahara, T., 278–279 Saier, J.M.H., 340–341 Saier, M.H., 48, 316, 335, 338, 341 Saiki, H., 137, 225 Saito, K., 174 Saito, N., 177 Saito, Y., 162 Sakurai, Y., 212 Saldanha, A.J., 262, 277 Saleem, R.A., 286 Salgado, L.E., 283 Saller, E., 177 Salusjarvi, L., 296 Salzberg, S.L., 5, 13, 20, 23, 26 Samoray, D., 28 Samson, L.D., 274–275 Samyn, B., 130 Sa´nchez, M., 75, 80 Sa´nchez-Espan˜a, J., 225, 239, 244 Sanchez, O., 153 Sand, W., 136–137, 224, 231, 234

AUTHOR INDEX

Sander, C., 75, 80 Sander, J., 143, 145–148, 165 Sanderson, K.E., 320 Santarius, U., 216 Santero, E., 33 Santini, J.M., 111 Saraiva, L.M., 150 Sasaki, K., 225 Sasikala, C., 121 Sathyanarayana, D.N., 136–137 Sato, M., 169 Sato, M.S., 169, 278–279 Sato, P.Y., 161 Sato, Y., 169 Satoh, T., 125, 209 Sauer, K., 44 Sauer, R.T., 10 Sauer, U., 267, 291–292 Saunders, E., 146 Sauve´, V., 158 Sburlati, A., 292 Schacherer, J., 274, 294 Schade, B., 278–279 Scha¨fer, K.O., 157 Scha¨fer, U., 150 Schaller, J., 149, 169 Scharf, B., 125 Scharfe, C., 290 Schattenhofer, M., 114, 166 Schedel, M., 134, 143 Scheffers, W.A., 268, 270–271, 288, 294 Scheideler, M., 264 Scheidig, A.J., 157 Schelling, M., 335 Schepper, M.E., 268 Scherens, B., 288, 290 Scherff, R.H., 349 Schesser Bartra, S., 341 Schiff, J.A., 169 Schimmack, G., 288, 290 Schimmel, P., 285 Schindelin, H., 152 Schink, B., 112, 123, 132, 173, 238 Schipper, D., 283

387

Schippers, A., 202, 231 Schirmer, T., 28, 36, 54 Schleifer, G., 162 Schleper, C., 216 Schmedding, D.J., 233 Schmid Nuoffer, S., 21 Schmidt, A., 169 Schmidt, J.M., 40, 42 Schmidt, M.A., 118, 340 Schmitt, R., 125 Schmitt, W., 162 Schnell, N.F., 45 Schneper, L., 277 Scho¨dl, T., 132–133 Schoenborn, L., 121 Schoenlein, P.V., 42 Schoepp-Cothenet, B., 148 Schofield, O.M.E., 176 Schofield, W.B., 34–35, 77 Scholz, C., 234 Scholz, L., 217 Schro¨der, I., 143, 154, 163 Schrenk, M.O., 220 Schu¨rmann, P., 149, 169, 175 Schu¨tz, M., 130, 132–133 Schubert, K., 112 Schulte, A., 143, 146–148 Schultz, J., 341 Schulz, W., 217 Schumann, O., 134, 140 Schu¨nemann, V., 149, 175 Schuster, S.C., 316, 318, 321, 327, 330, 335, 338 Schwartz, D.A., 274–275 Schwartz, H., 80–81 Schwarz, G., 152 Schwarz, H., 40, 80–81 Schwarzkopf, A., 45, 51 Schwenn, J.D., 169, 175–176 Schwertmann, U., 228 Schwudke, D., 333, 337–339, 345–346 Sciochetti, S.A., 31 Scott, K.M., 161, 320 Segel, I.H., 151, 161, 175

388

Segerer, A., 216, 235 Seidler, A., 176 Seidler, R.J., 317, 347 Seifert, H.S., 327 Seitz, L.C., 40–41, 78–79 Seker, U.O.S., 292 Seki, T., 344 Selengut, J.D., 56 Selmer, T., 149, 151–152 Sen, A.M., 213, 229, 232, 238 Setayeshgar, S., 40, 43, 77 Setya, A., 169 Seydel, R., 260 Shafer, B., 288, 290 Shah, M., 222 Shahak, Y., 130, 132 Shannon, K.W., 272 Shao, Y., 320 Shapiro, L., 5–7, 9–21, 23–27, 30, 33, 35–36, 38–39, 42, 54, 60–66, 71–77, 79, 81, 332 Sharma, P.L., 42 Sharom, J.R., 277 Sharon, L.A., 40, 43, 77 Sharrocks, A.D., 262 Sheetz, M.P., 327 Sherman, F., 267 Sherrill, C., 140, 147 Sherwood, C.S., 45 Shetty, J., 5, 13, 20, 23, 26 Shevtsov, S., 169, 175 Shi, W., 332 Shi, Y., 274–275 Shibata, S., 36 Shida, O., 214–215 Shigi, N., 146, 148 Shih, Y.-L., 79 Shilo, M., 106, 326 Shimada, K., 114, 125 Shimizu, H., 278–279 Shin, J.L., 274–275 Shiota, H., 212 Shively, J.M., 137, 206, 208, 288 Shoemaker, D.D., 288, 290

AUTHOR INDEX

Shute, E.A., 137, 222, 224 Shuttleworth, K.L., 212 Siam, R., 12, 62–63 Sieber, S.O., 274–275 Siegal-Gaskins, D., 55 Siegel, L.M., 176 Sievert, S.M., 161, 210 Sigrist, C.J.A., 343 Sillje, H.H., 290 Silva, A.M., 21 Silver, M., 217 Silver, S., 210, 223, 234 Simao, R.C., 21 Simbahan, J., 214–215 Simm, R., 54 Simon, I., 285 Sirevag, R., 125 Sirota, A., 337–339 Sistrom, W.R., 121–122 Sivanathan, V., 74 Skerker, J.M., 6, 9, 11, 13–14, 16, 24, 26–27, 38–39, 42, 327, 332 Skurnik, M., 341 Slifer, S., 274–275 Slijper, J.T., 296 Slijper, M., 296 Su¨ling, J., 109, 113, 118, 121–122, 124 Slobodova, N.V., 115 Sly, L.I., 242 Smit, J., 5, 13, 20, 23, 26, 40, 45, 48, 52, 54–55, 57–58 Smith, A.J., 156, 161–162 Smith, C.A., 349 Smith, C.S., 30, 47–48, 51–52, 54, 56 Smith, J.J., 286 Smith, M.C.M., 321, 323, 326 Smits, T.H.M., 106 Snoek, I., 288–290 Snyder, M., 288, 290 So, A.K., 208 So, M., 327 Soares, C.M., 145 Sockett, R.E., 316, 318, 321, 323, 325–327, 330, 333–334, 349, 353

AUTHOR INDEX

Sogin, M., 176 Solovyev, V.V., 209–210, 222 Sommer, J.M., 13, 54 Sommerhalter, M., 157 Sonderegger, M., 292 Sookhai-Mahadeo, S., 288, 290 Sorokin, D.Y., 111, 124, 155, 161 Sosa, J., 56 Soyer, F., 137 Specht, W., 167 Speer, M.C., 274–275 Speich, N., 150 Spellman, P.T., 278 Spencer, G.H., 224 Spencer, P.S., 274–275 Sperl, G.T., 57 Sperling, D., 151, 158–162, 175 Spieth, J., 320 Spiridonova, E.M., 115 Spitznagel, E.L., 274 Spormann, A.M., 37, 39–40, 45 Spratt, B., 79 Spring, S., 114, 166 Sproles, D.I., 274–275 St Geme, J.W., 341 Stackebrandt, E., 122, 225, 242 Stahl, D.A., 126 Staley, J.T., 40 Stallwood, B., 237 Stanier, R.Y., 40, 42 Starr, M.P., 317, 347 Starynin, D.A., 143 Stathopoulos, C., 340, 343 Steensma, H.Y., 263, 266, 268, 274, 280, 285 Stein, M.A., 345 Steiner, S., 346 Steinmetz, L.M., 274, 290 Steinmetz, M.A., 131, 143, 155 Stephens, C., 5, 7, 13, 20, 23–26, 60–61 Sternheim, N., 30, 39 Stetter, K.O., 150, 209, 216–217, 235 Steudel, R., 118, 133–136, 138, 140 Stewart, F.J., 146

389

Stewart, P.S., 44 Stier, T.J.B., 270 Stolz, B., 321 Stolz, J.F., 111 Stoneking, T., 320 Stoodley, P., 44 Storms, R.K., 288, 290 Storz, G., 278 Stotz, A., 21 Stougaard, P., 118 Stouthamer, A.H., 259, 299 Straley, S.C., 322–323 Strathern, J.N., 288, 290 Strauch, E., 333, 337–339, 345–346 Strauss, G., 206 Strompl, C., 57–58 Stu¨ckrath, I., 296 Stumm, W., 219 Styer, K.L., 341 Sudarsanam, P., 274 Sugai, A., 217 Sugawara, K., 208 Sugio, T., 230, 234 Sugiyama, H., 209 Suhr, M., 340 Suk, W.A., 274–275 Sullivan, R.C., 274–275 Sullivan, S.A., 56 Summerfield, D.T., 325 Sun, H., 320 Sunna, A., 214–215 Surana, N.K., 341 Surdin-Kerjan, Y., 174 Susin, M.F., 21 Suter, M., 149, 169, 175 Suziedelis, K., 173 Suzuki, H., 147 Suzuki, I., 136–137, 206, 233–234 Suzuki, M., 214–215 Suzuki, T., 146, 148, 217 Suzuki, Y., 25 Swainston, N., 274, 277–278, 282, 295–296, 299 Swenberg, J.A., 274–275

390

Swoboda, E.P., 156 Szigyarto, C.A., 262 Szilard, L., 259, 261, 271, 290–291 Ta’asan, S., 260 Tabak, H.F., 262 Tabita, F.R., 111, 113, 145, 162–163 Tagmount, A., 174 Tagne, J.B., 285 Tai, S.L., 279–280, 282–283, 288–290, 296 Tainer, J.A., 327–328 Takacs, M., 146 Takase, I., 79 Takeshima, T., 208 Takeuchi, T.L., 136–137 Takusagawa, K.T., 285 Talley, B.G., 338 Tallon, L.J., 107 Tam, E., 43 Tamaki, S., 77, 79 Tamayo, R., 54 Tamerler, C., 292 Tan, M.H., 20–21, 25 Tan, P.K., 274 Tang, H., 320 Tang, J.A., 125 Tang, J.X., 32, 37, 46–47 Tanimoto, E.Y., 272 Tanner, R.S., 111 Tano, T., 211–212, 230 Tawfilis, S., 36 Taylor, B.F., 122 Taylor, J.M.G., 295 Teeling, H., 114, 166 Teixeira de Mattos, J.M., 268 Teixeira, M., 147, 150, 234 Telegdi, J., 136–137 Temple, K.L., 231 ten Pierick, A., 297–298 Tennant, R.W., 274–275 ter Kuile, B.H., 296 ter Linde, J.J.M., 274, 280, 285 ter Schure, E.G., 290

AUTHOR INDEX

Tesar, M., 57–58 Teske, A., 155, 161, 176 Tettelin, H., 282 Teusink, B., 294 Thanassi, D.G., 343 Thanbichler, M., 66, 71–72, 74 Theissen, U., 132 Thelen, M.P., 222 Then, J., 122, 124, 133, 162 Theobald, U., 298 Theriot, J.A., 77, 79, 81 Thevelein, J.M., 277 Thierry, D., 136–137 Thomas, D.L., 58, 174 Thomas, D.Y., 278–279 Thomas, J.A., 284 Thomas, P., 161 Thomas, S.M., 175 Thomasho, M.F., 316, 325 Thomashow, L.S., 317, 347 Thomashow, M.F., 347–350, 353 Thompson, C.M., 285 Thomson, H.E.C., 225, 231–232 Thorsson, V., 295 Thum-Schmitz, N., 143, 146, 149–150 Thunnissen, A.M., 352 Tian, R., 274–275 Till, R., 321 Timkovich, R., 146 Timmis, K.N., 111, 210, 216 Tindall, B.J., 57–58 Tinkham, L.E., 161 Todd, S.A., 274–275 Toedling, J., 274 Toh, E., 37, 39–40, 43–45, 48, 52–53 Toirkens, M.J., 292 Tojo, H., 348 Tollin, G., 152 Tomasouw, W.F., 288 Tomcsanyi, T., 173 Tomich, M., 38 Tomioka, S., 79 Tommassen, J., 340, 343 Torgersen, Y.A., 126

AUTHOR INDEX

Torian, B.E., 345 Torres y Torres, J.L., 113 Toth, A., 146 Touraine, B., 174 Tourova, T.P., 111, 115, 121, 123–126, 214–215, 217, 229 Traina, S.J., 228 Tran, K., 5, 13, 20, 23, 26 Tran, Q.A., 64 Trautwein, A.X., 149, 175 Tributsch, H., 136, 220 Trimble, M.J., 67, 69, 75–76 Tron, P., 221 Tru¨per, H.G., 118, 122, 124, 127, 131–134, 138–143, 146, 148–151, 153, 156, 158–162, 169, 175, 177 Trudinger, P.A., 135 True, H.L., 278 Truelzsch, K., 341 Tsai, C.W., 217 Tsai, P.S., 292 Tsang, P.H., 46–47 Tsaplina, I.A., 214–215, 218, 229 Tscha¨pe, J., 131–132, 134, 160 Tsen, S.D., 217 Tseng, Y.H., 340–341 Tsuruoka, N., 214–215 Tucker, C.J., 274–275 Tudor, J.J., 338 Turner, L.R., 328 Turner, R.F., 45 Tuschak, C., 140 Tusz, J., 146 Tuttle, J.H., 221, 235 Tyers, M., 277 Tyson, G.W., 209–210, 222, 244 Uchimura, T., 118 Uchino, M., 118 Uda, I., 217 Ueda, Y., 124 Ueki, T., 217 Ugolkova, N.V., 115 Ulbricht, R.J., 284

391

Umbreit, T.H., 44, 55–56 Umbreit, W.W., 136 Umino, Y., 36 Unz, R.F., 212 Urata, K., 125 Urich, T., 138, 234 Usaite, R., 262 Utterback, T., 5, 13, 20, 23, 26 Uzawa, T., 217 Vai, J.T., 277 Vainstein, M.B., 122 Valde´s, J., 209–210, 223, 234 Valencia, A., 75, 80 Valenti, P., 137 Valenzuela, L., 137 Valenzuela, P.D.T., 209 Valle, G., 288, 290 Valvano, M.A., 323 Vamathevan, J., 5, 13, 20, 23, 26 van Bakel, H., 275 van Beeumen, J.J., 130–131, 160 van Dam, K., 268 van de Peppel, J., 275 van de Vossenburg, J.L., 204 van den Brink, J., 277 van den Ent, F., 73, 75, 80 van der Baan, A.A., 271 van der Ploeg, J.R., 177 van der Weijden, C.C., 296 van der Zanden, L., 266 van Dijken, J.P., 125, 210–211, 234, 266, 268–271, 274, 280, 284, 288, 292–294 Van Dover, C.L., 112 van Driessche, G., 130 Van Gelder, P., 338, 340 van Gemerden, H., 118, 122–123, 134, 139–140, 143 van Gulik, W.M., 268, 270, 286, 294, 296, 298 van Heusden, G.P.H., 263 van Hoek, P., 268, 294 Van Houten, B., 274–275 van Keulen, G., 206, 208

392

van Kleeff, B.H.A., 270 van Leenen, D., 275 van Maris, A.J.A., 269, 279, 292–293 van Niel, C.B., 116 van Riel, N., 268 van Straaten, K.E., 352 van Tuijl, A., 296 van Ulsen, P., 340, 343 van Verseveld, H.W., 299 van Winden, W.A., 271, 297–298 van Zyl, W.H., 292 Vanatalu, K., 299 Vancanneyt, M., 57–58 Vandenbroek, P.J.A., 270 Vandijken, J.P., 270 Vanselow, M., 134, 143 van’Dijken, J.P., 292 Varma, A., 82 Velculescu, V.E., 271 Veloso, F., 210, 223, 234 Venceslau, S.S., 147 Venter, J.C., 5, 13, 20, 23, 26 VerBerkmoes, N.C., 222 Verduyn, C., 268, 294 Verkleij, A.J., 290 Vermeglio, A., 148 Vermeulen, A., 292–293 Vermeulen, E.E., 290 Verrips, C.T., 268, 290 Verte´, F., 131 Vetriani, C., 210 Vetter, I., 344 Vetter, R.D., 161 Vicente, M., 73, 75, 80 Vidal, O., 52 Vidmar, J.J., 174 Vielreicher, M., 132–133 Vila, X., 117 Vilu, R., 299 Vindeløv, J., 268 Vingron, M., 264 Vinke, J.L., 268, 270, 294, 298 Viollier, P.H., 5–7, 14, 16–18, 30–31, 33–35, 39, 54, 66, 76

AUTHOR INDEX

Visscher, P.T., 118, 122, 134, 140 Visser, J.M., 162 Visser, W., 270–271 Vodovotz, Y., 260 Voet, M., 288, 290 Vogelstein, B., 271 Vogl, K., 121, 123 Vogler, K.G., 136 Volckaert, G., 288, 290 Volkert, T.L., 285 Vollmer, W., 40, 80–81, 350, 352–353 Volpin, H., 337–339 von Ballmoos, P., 169 von Meyenburg, K.H., 271 von Stockar, U., 268 Vorst, O., 288 Voyich, J.M., 45 Ve´ronneau, S., 288, 290 Vuong, C., 45 Vuralhan, Z., 283 Waaterston, R., 274 Wachi, M., 72–73, 77 Wade, P.A., 285 Wadhams, G.H., 325 Wagner, J.K., 40–41, 43, 77–79 Wagner, N.J., 332 Wahlbom, C.F., 292 Wakao, N., 211–213 Waksman, G., 341 Waksman, S.A., 231 Walker, M.J.T., 282 Walsh, E.A.F., 282 Walsh, M.C., 280, 283, 288–290, 296 Walton, K.C., 220 Wang, C.Y., 288, 290 Wang, S.C.E., 65, 74 Wang, S.P., 14, 20, 42 Wang, X., 45, 51, 320 Wang, Y., 332 Wanner, B.L., 42 Wanner, G., 121, 123 Ward, C.K., 332 Ward, D.M., 74, 107, 112, 125–126

AUTHOR INDEX

Ward, J.M., 332 Ward, N.L., 56 Ward, T.R., 288, 290 Wardleworth, L., 274, 277–278, 282, 295–296, 299 Warner, J.R., 277 Warrington, J., 272 Waskovsky, J., 224 Wassmann, P., 28 Watanabe, Y., 347 Waterbury, J.B., 144 Waterhouse, S., 218 Waterston, R., 320 Watson, J.D., 261 Weber, M., 149 Weiner, R.M., 55–56 Weinitschke, S., 106 Weis, B.K., 274–275 Weiser, S., 28, 36, 54 Weiss, D.S., 75 Weiss, M.A., 177 Weiss, R.L., 164, 217 Werkman, C.H., 206 Werner-Washburne, M., 278 Wessels, L.F.A., 282, 285 West, L., 66, 74 Westerhoff, H.V., 294, 296 Weusthuis, R.A., 268, 270 Wheeler, R.T., 27 Whelen Dow, S., 288 White, A.R., 58 White, C., 41, 72, 78–82 White, D.C., 208 White, J., 36 White, O., 5, 13, 20, 23, 26 White, S., 58 Whiteway, M., 278–279 Whitfield, C., 48, 51 Whitman, W.B., 125 Whitney, A.R., 45 Wichlacz, P.L., 212 Wiedemann, G., 176 Wiederkehr, I., 28 Wigneshweraraj, S.R., 23

393

Wilhelmy, J., 288, 290 Wilker, B., 347 Williams, E.B., 208 Williams, H.N., 323 Williams, K., 295 Williams, P.A., 213 Willison, J.C., 122 Wilmot, D.B., 161 Wilson, M., 332 Wilson, R.K., 320 Winde, J.H., 277 Wingrove, J.A., 23, 33, 35 Winkler, A.A., 292–293 Winkler, H., 62 Winkler, Kuyper, A.A., 293 Winland, R.L., 228 Winogradsky, S.N., 116 Winther, O., 275, 277–279, 299 Winzeler, E.A., 288, 290 Witte, B., 111 Wittinghofer, A., 344 Wittmann, C., 271 Woese, C.R., 111, 126 Wolber, P.K., 272 Wolf, A., 5, 13, 20, 23, 26 Wolf, M., 228 Wollenberg, M., 176 Wong, M.Y., 48 Wood, A.P., 218, 231 Wood, D., 62 Wood, H.G., 206 Wortinger, M.A., 14, 75–77, 80 Woyke, T., 146 Wray, J.L., 169 Wright, R., 7, 60–61 Wu, D., 146 Wu, H.I., 288, 290 Wu, J., 13–14, 20–21, 31, 176, 262, 274 Wu, S.S., 331 Wulf, J., 114, 166 Wunderl, S., 217 Wynen, A., 151, 169, 175 Wyrick, J.J., 276, 285 Wysocki, R., 288

394

Xavier, A.V., 147, 150 Xiang, X., 219 Xie, Y., 78–79, 82 Xu, P., 274 Xu, Z., 33 Xuan, S., 274–275 Yahya, A., 218, 229 Yakimov, M.M., 111, 210, 216 Yamagishi, A., 217 Yamanaka, T., 162 Yamasato, K., 214–215 Yan, S., 114, 166 Yang, B., 5 Yang, D., 24–25 Yang, G., 73 Yang, Q., 56 Yang, Y., 288, 290 Yarrow, D., 267 Yarzabal, A., 223 Yen, G.S., 288, 290 Yen, M.R., 340–341 Yokota, A., 162, 213–215 Yoo, J., 285 Youn, H.-S., 146 Young, K.D., 76–77, 82, 349, 352 Young, R.A., 276, 278, 285 Youngman, E., 288, 290 Yu, E.Y., 140 Yu, J., 35 Yu, K., 288, 290 Yu, Z., 151 Yudkin, M.D., 75 Yun, C., 54–55, 57 Yurkov, V.V., 114, 124

AUTHOR INDEX

Zaar, A., 118, 121 Zafar, N., 56 Zahringer, U., 345–346 Zakharchuc, L.M., 115 Zakharchuk, L.M., 214–215, 218, 229 Zarbl, H., 274–275 Zeef, L., 274, 277–278, 282, 295–296, 299 Zehetner, G., 271 Zeitlinger, J., 285 Zerp, S., 277 Zeruth, G.T., 161 Zhai, Y., 340–341 Zhang, D., 30 Zhang, L., 20–21, 25, 271 Zhang, N.S., 262, 274, 277–278, 282, 295–296, 299 Zhang, X., 23 Zhang, Y., 73 Zhang, Z., 274 Zhao, J.L., 14, 31, 73 Zhao, W., 73 Zhao, Y., 277 Zhou, L., 56 Zhuang, W.Y., 35 Zhukov, V.G., 111 Zhukova, N.V., 118 Zillig, W., 216–217, 219 Zimmermann, K., 288 Zimmermann, P., 234 ZoBell, C.E., 44–45 Zogaj, X., 54 Zweiger, G., 19, 60–61 Zyakun, A.M., 115 Zylstra, G.J., 149, 175–176

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

A22 inhibitors cell shapes, 77, 78, 81 chromosome segregation, 72–73 ABC, see aerobic bacteriochrlorophyllcontaining bacteria Acidicaldus spp., carbon metabolism, 213 Acidiphilium spp. carbon metabolism, 211–213, 217–218 iron reduction, 225–226, 228, 230 Acidisphaera spp. carbon metabolism, 213 iron reduction, 228 Acidithiobacillus spp. acidophile enzymology, 222, 223 acidophilic micro-organisms, 208–211 elemental sulfur adhesion, 136–137 iron oxidation, 220 iron reduction, 224–225 oxidation biochemistry, sulfur metabolism, 233–234 sulfur metabolism, 231–232 Acidocella spp. iron reduction, 228 sulfur metabolism, acidophilic micro-organisms, 237–239 Acidomonas spp., carbon metabolism, 213 acidophilic micro-organisms, 201–233, 234, 235–255 applied/ecological aspects, 239–242, 243, 244 definition/habitat/physiology, 202–205 diversity, 204–206

iron metabolism, 219–230 mineral processing, 239–243 natural/anthropogenic environments, 243–244 sulfur metabolism, 230–233, 235–239 actin homologs cell shapes, 80 chromosome segregation, 72–74 Z-ring formation, 71–72 see also fts genes Actinobacteria spp., iron reduction, 229 activators, sigma factors, 21–24 adenosine-5u-phosphosulfate (APS) sulfate assimilation pathway, 167–169, 170–172 sulfite oxidation, 153–155 sulfur metabolism, 148–153, 174–175 adenosine-5u-phosphosulfate reductase (Apr) oxidative sulfur metabolism, 127 sulfite oxidation, 150, 153–155 adenylylsulfate, see adenosine-5uphosphosulfate adherence Bdellovibrio bacteriovorus, 331, 339–341 elemental sulfur, 135–138 holdfasts, 44–60 prosthecate spp., 55–60 adhesins, see adherence adhesion, see adherence ADP sulfurylase (APAT) indirect oxidation, 148 sulfite oxidation, 151–152 aerobic bacteriochlorophyll-containing (ABC) bacteria, 106–107, 108, 114 sulfur metabolism, 124–125

396

Affymetrix arrays micro-array technology, 272–275 promoter architecture, 25–26 AFM, see atomic force microscopy Alicyclobacillus spp. carbon metabolism, 213, 214–215, 216, 218–219 iron reduction, 228 Allochromatium vinosum dsr genes, 146–148 elemental sulfur adhesion/uptake, 136–138 sulfur globule envelopes, 141–142 sulfur metabolism, 153–155, 158–160, 161–162 alphaproteobacteria carbon metabolism, 212–213 Z-ring formation, 71 alternative sigma factors, 21–25 aminidases, Bdellovibrio bacteriovorus, 352 amino acids chemostat cultivation, 283 radioactive labeling experiments, 353 sulfur globule envelopes, 141–142 anoxic conditions, sulfur reduction, 142 anoxygenic phototrophic bacteria ecology, 116–117 physiology/taxonomy, 107–116 sulfate assimilation, 167–177 sulfur globules, 138–143 see also green sulfur bacteria; purple sulfur bacteria anthropogenic environments, acidophilic micro-organisms, 243–244 antibiotics, mecillinam, cell shapes, 78 antifoaming agents, chemostat cultivation, 270 APAT, see ADP sulfurylase Apr, see adenosine-5u-phosphosulfate reductase

SUBJECT INDEX

APS:phosphate adenylyltransferase (APAT) indirect oxidation, 148 sulfite oxidation, 151–152 APS, see adenosine-5u-phosphosulfate archaea, acidophilic micro-organisms, 216–217, 219, 224, 234–235 Asticcacaulis spp., 55 atomic force microscopy (AFM), holdfast adherence, 46 ATP:sulfate adenylylsulfate:phosphate adenylyltransferase (APAT) indirect oxidation, 148 sulfite oxidation, 151–152 ATP sulfurylase, sulfite oxidation, 150–151 ATP synthesis, acidophilic micro-organisms, 221–222 ATs, see autotransporters attachment holdfasts, 45–46, 49–53, 59 prosthecate spp., 57–58 autotransporters (ATs) Bdellovibrio bacteriovorus, 339–341, 342–343, 344 phylogeny, 343 autotrophic acidophiles iron oxidation, 220 iron reduction, 224–225 auxotrophic strains, chemostat cultivation, 267 Bacteriovorax stolpii, membrane phospholipids, 346–347 BALO lipid-related enzymes, Bdellovibrio bacteriovorus, 348 BChl light harvesting pigments, phototrophic bacteria, 108, 112–114, 116–117 Bdellovibrio bacteriovorus, 313–354 beta-sheets, Bdellovibrio bacteriovorus, 344

SUBJECT INDEX

binding sites CtrA, 16, 25–26, 62–63 DnaA, 62 IHF, 62–63 biofilm formation, 44–45 flagella function, 37 pili function, 39–40 ‘‘birth scar’’ proteins, flagella, 34 BLAST searches, Bdellovibrio bacteriovorus, 336–337 Brownian motion, adherence experiments, 46–47 c-di-GMP, see cyclic-diguanosine monophosphate carbon dioxide fixation acidophilic micro-organisms, 206–211, 221–222 phototrophic bacteria, 112–113 sulfur compound oxidation, 162 carbon metabolism acidophilic micro-organisms, 201, 206–219 chemostat cultivation, 286–287 facultative autotrophic acidophiles, 217–219 obligately heterotrophic acidophiles, 211–217 carbonic anhydrases, chemostat cultivation, 284 carboxypeptidases, Bdellovibrio bacteriovorus, 352–353 carboxysomes, acidophilic micro-organisms, 206, 208 Caulobacter crescentus, 1–101 CB15 Caulobacter strains, holdfast biosynthesis, 58–59 CBB cycle, acidophilic micro-organisms, 208–209 CckA histidine kinases, 8, 12–14, 27 stalk biogenesis, 41–42 ccrM promoter genes chromosome replication, 60–61 CtrA regulons, 14–16

397

cell division, 60–76 flagella biosynthesis, 35 protein expression, 74–76 see also cell-cycle progressions cell shapes, 60, 76–82 cell walls hydrolase activity, 350–354 precursor biosynthesis, 349–350 cell-cycle progressions, 1–101 cementation, acidophilic micro-organisms, 240 che genes Bdellovibrio bacteriovorus, 323–325 CtrA, 15 checkpoints, DNA replication, 75–76 chemolithotrophic acidophiles, acidophilic micro-organisms, 205 chemostat cultivation, 261–271, 281 basic principles, 261–266 experimental design, 266–269 experimental tools, 261–275 pitfalls, 269–271 reproducibility, 274–275, 276 Saccharomyces cerevisiae, 257–311 steady-states, 266 chemotaxis, Bdellovibrio bacteriovorus, 318, 322–326 Chlorobium spp. stored sulfur oxidation, 144 sulfate assimilation, 169 Chloroflexaceae families, 115 see also filamentous anoxygenic phototrophs Chloroflexus spp., 125 phototrophic bacteria, 106–107 see also filamentous anoxygenic phototrophs Chloroherpeton thalassium, stored sulfur oxidation, 144 chlorosomes, phototrophic bacteria, 112, 115, 117 ChpT histidine phosphotransferase proteins, CtrA, 13, 27

398

Chromatiaceae families, 120–121 phototrophic bacteria, 106, 109–111, 117–122 sulfate assimilation, 169 sulfur globules, 141–142 sulfur metabolism, 128 see also anoxygenic phototrophic bacteria chromatin immunoprecipitation assays, GcrA, 18 chromosomes condensation, 341, 344 replication, 60–82 CtrA, 27 DnaA, 18–20 GcrA, 18 Z-ring formation, 69–70 class II genes flagella, 14, 31–32, 35 sigma activators, 22, 23 class III genes flagella, 31–33 sigma activators, 22, 23 class IV genes flagella, 31–32 sigma activators, 22, 23 cleavage, autotransporters, 340–341 ClpXP proteases CtrA, 7, 8, 9–11 sigma factors, 21 clusters hdr genes, 164 sox genes, 157–158 cold shock, chemostat cultivation, 278 commercial single-dye micro-array experiments, chemostat cultivation, 271 competitive chemostat cultivation, 288–289 complex regulatory pathways, 1–101 context dependency, chemostat cultivation, 279–282, 290

SUBJECT INDEX

coordination chromosome segregation, 65–74 Z-ring formation, 69–72 Cori regions, chromosome replication, 61–62 cpa genes, pili assembly, 39 CpdR response regulators CtrA, 8, 10–11 signal transduction proteins, 26 crescentin, cell shapes, 78, 79–80 csg genes, holdfast attachment, 52 CtrA transcriptional regulators, 4, 5–16, 22, 23, 27 chromosome replication/segregation, 60–63, 68 DNA replication, 75–76 holdfast synthesis, 53–54 pili genes, 38–39 stalk biogenesis, 41–42 CUB protein domains, Bdellovibrio bacteriovorus, 344 curved cells, 79 cyanobacteria, 106 cyclic-diguanosine monophosphate (c-di-GMP), 28 flagella, 34, 35–37 holdfast synthesis, 54–55 cyclo-octasulfur, elemental sulfur metabolism, 137, 138 cysteine biosynthesis (cys) genes, sulfur metabolism incorporation, 168–172, 173, 176 cytochromes acidophile enzymology, 222–223 dsr genes, 146 sulfur metabolism, 126–127, 130–131, 134, 152, 156, 158–159, 161 cytolysins, Bdellovibrio bacteriovorus, 339 cytoplasmic portions, PodJ, 29 D-cysteine experiments, cell shapes, 80–81 DAP amino acids, Bdellovibrio bacteriovorus, 353

SUBJECT INDEX

DD-carboxypeptidases, Bdellovibrio bacteriovorus, 352–353 DD-endopeptidases, Bdellovibrio bacteriovorus, 352–353 deformation, stalk/holdfast, 46–47 degradation CtrA regulons, 7–8, 9–11, 64 stored sulfur, 143 tmRNA, chromosome replication, 64–65 dehydrogenases, sulfur metabolism, 152, 161–162, 234 deletion experiments, chemostat cultivation, 288–290, 326 delta-proteobacteria, Bdellovibrio bacteriovorus, 330–331 dephosphorylation, see phosphorylation deposition, sulfur metabolism, 138–143 Desulfosporosinus spp., sulfur metabolism, 237–239 dgr genes flagella biosynthesis, 36 holdfast synthesis, 55 digestion, holdfast adherence experiments, 47 dilution rates, see specific growth rates dimethylsulfoniopropionate (DMSP), aerobic bacteriochlorophyllcontaining bacteria, 125 direct oxidation, sulfur metabolism, 148, 152–153 displacement, adherence experiments, 46–47 dissimilatory oxidation, acidophilic micro-organisms,219–224, 230–235 dissimilatory reduction acidophilic micro-organisms iron metabolism, 224–230 sulfur metabolism, 235–239 dissimilatory sulfite reductase (DsrAB), oxidative sulfur metabolism, 134 DivJ histidine kinases, 11 CtrA, 8, 13, 27–28 signal transduction proteins, 26

399

DivK histidine kinases, CtrA, 8, 11, 13–14, 27–28, 30 DivL tyrosine kinases, 29–30 CtrA, 14 DMSP, see dimethylsulfoniopropionate DNA methylation chromosome replication, 60–61 CtrA, 15 DNA replication, 75–76 DnaA transcriptional regulators, 4, 5–6, 17–20 chromosome replication/segregation, 62, 68 GcrA, 17 protein expression, 75 double-dye micro-array analyses, chemostat cultivation, 269, 270, 271 ‘‘down-regulated’’ conditions, chemostat cultivation, 279 DSMZ mutants, sulfur metabolism, 144, 166, 169, 173 dsr genes elemental sulfur uptake, 137 occurrence/arrangement, 143–145 oxidative sulfur metabolism, 134, 153 products/function, 145–147 sulfur metabolism evolution, 165–166 dual-dye micro-array analyses, chemostat cultivation, 272, 273, 274 dumps, mineral processing, 240, 244 dye micro-array analyses, 269–271 dynamic studies, chemostat cultivation, 298 E. coli flagella function, 37 holdfast attachment, 52 NAG polymers, 45 EAL protein domains, holdfast synthesis, 54–55 ECT, see electron cryotomography

400

Ectothiorhodospiraceae families, 121–122 elemental sulfur uptake, 135 oxidative sulfur metabolism, 133 phototrophic bacteria, 106, 109, 111, 117 sulfate assimilation, 169 see also anoxygenic phototrophic bacteria Ehrlich pathways, chemostat cultivation, 283 elasticity, holdfasts, 46–47 electron cryotomography (ECT), Z-ring formation, 67–69 electron transfer acidophilic micro-organisms, 221–222, 223 anoxygenic phototrophic bacteria, 118 dsr genes, 147 Ectothiorhodospiraceae families, 121–124 elemental sulfur uptake, 135 filamentous anoxygenic phototrophs, 125 heliobacteria, 126 indirect oxidation, 149–150 oxidative sulfur metabolism, 131, 134 photosystems, 107–109 purple non-sulfur phototrophic bacteria, 124 sulfur metabolism, 131, 134–135, 232–233 see also oxidation; phototrophic bacteria; reduction elemental sulfur acidophilic micro-organisms, 203 activation/uptake, 134–138 adhesion, 135–137 properties, 135 speciation, 139–140 thiosulfate oxidation, 155 uptake, 134–138 see also dsr genes

SUBJECT INDEX

elongated helical shapes, cell shapes, 80 endopeptidases, Bdellovibrio bacteriovorus, 352–353 energetics, acidophilic micro-organisms, 221–222, 232–233 Enterobacteria, sulfate assimilation pathway, 167 envelopes, sulfur globules, 141–142 environmental stress response (ESR) genes, chemostat cultivation, 278 enzymes acidophilic micro-organisms, 222–224 carbon fixation, 210–211 acidophilic prokaryotes, iron reduction, 229–230 Bdellovibrio bacteriovorus, 348–349, 350–354 chemostat cultivation, 293–294 EPS, see extracellular polymeric substances Escherichia coli sulfate reduction, 175 sulfate uptake, 173 ESR genes, see environmental stress response evolution chemostat cultivation, 271 evolutionary engineering, 290–294 experimental tools, chemostat cultivation, 261–276 export systems, holdfasts, 48, 50, 51 external sulfur, see elemental sulfur extracellular polymeric substances (EPS), elemental sulfur adhesion, 136 extracellular sulfur deposition, 138–139 extreme thermo-acidophiles, 204–205, 211–213, 217–218, 225–226, 228, 230 carbon metabolism, 211–213, 217–218 iron reduction, 225–226, 228, 230 facultative autotrophic acidophiles, 207–208 carbon metabolism, 217–219

SUBJECT INDEX

FAPs, see filamentous anoxygenic phototrophs fatty acids, membrane chemistry, Bdellovibrio bacteriovorus, 345–348 FccAB flavoproteins thiosulfate oxidation, 158–159 see also flavocytochrome c ferredoxin-dependent sulfite reductases, sulfite reduction, 176 Ferrimicrobium spp., carbon metabolism, 216 Ferrithrix spp., carbon metabolism, 216 Ferroplasma spp. acidophilic micro-organisms, 210 carbon metabolism, 216–217 iron reduction, 229 fibronectin type III (FN3) domains, Bdellovibrio bacteriovorus, 344 filamentous anoxygenic phototrophs (FAPs) phototrophic bacteria, 106–107, 108, 114–115 sulfur metabolism, 125–126 sulfate assimilation, 173 ‘‘filtering out’’ responses, chemostat cultivation, 277 fitness of null mutants, chemostat cultivation, 288–290 flagella, 3, 4, 8, 28, 31–37, 38 activation, 35–36 assembly, 31–33, 34, 35 Bdellovibrio bacteriovorus, 347–348 biosynthesis, 31–35 CtrA, 14 ejection, 36–37 fla genes, 33 flagellins, 32–33 function, 37 motility, 316–317, 318, 319–326 multicomponent systems, 28 sigma factors, 21, 22–23

401

flavocytochrome c (FccAB) oxidative sulfur metabolism, 126–127, 130–131 thiosulfate oxidation, 158–159 flb genes flagella, 31–33, 35 sigma activators, 23 signal transduction proteins, 26 flg operon genes Bdellovibrio bacteriovorus, 318–320 sigma activators, 22 flh genes Bdellovibrio bacteriovorus, 318–320 sigma activators, 22 fli operon genes Bdellovibrio bacteriovorus, 318–322, 326 CtrA regulons, 14, 16 flagella, 31–32, 35–36 sigma activators, 22–23 see also flagella flj genes flagella, 32–33 sigma activators, 22, 23 Flp pili, Bdellovibrio bacteriovorus, 332, 333 fluorescent microscopy cell shapes, 79, 80–81 Z-ring formation, 67, 70, 71 FN3 domains, see fibronectin type III domains foci, chromosome segregation, 73–74 fox genes, acidophile enzymology, 224 fts genes alternative sigma factors, 24 cell shapes, 80–82 chromosome segregation, 66–67, 68, 69, 74–75 CtrA regulons, 14–15 DNA replication, 74–76 DnaA, 19–20 stalk biosynthesis, 41 Z-ring formation, 70–72

402

functional analyses, chemostat cultivation, 282–290 functional redundancy, chemostat cultivation, 290 fusel alcohols/acids, chemostat cultivation, 283 fusion, receiver domains, CtrA, 9 G phases, cell-cycle progressions, 3–4, 6 GAFTGA motifs, sigma activators, 23 Gcn4p regulator proteins, chemostat cultivation, 285 GcrA transcriptional regulators, 4, 5–7, 16–18 DNA replication, 76 GDP (guanosine diphosphate), Z-ring formation, 69 GDP, see guanosine diphosphate gene complement, type IV pili, 328–331 gene function assignment, chemostat cultivation, 282–284 GFP, see green fluorescent protein GGDEF domain proteins, holdfast synthesis, 54–55 glass-slide arrays, chemostat cultivation, 272, 274 global regulation, 3–30 glucosamine, Bdellovibrio bacteriovorus, 353 glucose catabolism, obligately heterotrophic acidophiles, 212 glutathione amide, dsr genes, 147 glycan digestion, Bdellovibrio bacteriovorus, 353–354 glycolytic enzymes, chemostat cultivation, 293–294 glycosyltransferases Bdellovibrio bacteriovorus, 350 cell shapes, 80–82 Gram-positive/negative bacteria Bdellovibrio bacteriovorus, 351 carbon metabolism, 213, 216, 218–219

SUBJECT INDEX

green fluorescent protein (GFP) cell shapes, 79, 81 CtrA, 12 Z-ring formation, 67 see also fluorescent microscopy green sulfur bacteria (GSB), 106–107, 108, 112–113, 115–118, 119 dsr genes, 144–146 elemental sulfur uptake/adhesion, 135–137 sulfur metabolism, 148, 153–155, 162, 169, 173 evolution, 165–166 oxidation, 126–127, 128–129, 130–135, 150 reduction, 142–143 sulfite oxidation, 148, 153–154 thiosulfate oxidation, 155, 157–159, 162 see also anoxygenic phototrophic bacteria growth-limiting nutrients, see nutrient limitation GSB, see green sulfur bacteria guanosine diphosphate (GDP), Z-ring formation, 69 guanosine triphosphate (GTP), 28, 69 Bdellovibrio bacteriovorus, 341, 344 HD cells, see host-dependent cells hdr genes, heterodisulfide reductases, 164 heaps, mineral processing, 240, 244 heat shock sigma factors, 21 heliobacteria, 106–107, 108, 115–116 sulfur metabolism, 126, 173 hemE genes, chromosome replication, 62 heterodisulfide-reductase-like complexes, 164 heterotrophic acidophiles, 205–206 iron metabolism oxidation, 220–221 reduction, 225–229 sulfur metabolism, 232

SUBJECT INDEX

hfa genes holdfasts, 45–46, 49, 50, 51–53 prosthecate spp., 57–58 hfs genes holdfasts, 48, 49, 50, 51–52 prosthecate spp., 56–58 HI cells, see host-independent cells Hia adhesins, Bdellovibrio bacteriovorus, 340–341 histidine kinases CtrA, 8, 11–14, 27–28 DivK, 30 holdfast synthesis, 55 PodJ, 29 sigma activators, 24 stalk biogenesis, 41–42 transduction proteins, 26 histidine phosphotransferase (Hpt) CtrA, 12–13 sigma activators, 24 stalk biogenesis, 42 Hlr. halophila polysulfide-reductase-like complexes, 163 thiosulfate oxidation, 159–160 holdfasts, 38, 44–49, 50, 51–58, 59, 60 biosynthesis, 51 biosynthesis/secretion, 48, 50–52, 59 composition/properties, 45–46 genetics/synthesis/attachment, 47–53, 59 stalk function, 43 synthesis, 53–55 homologs flagellar-related proteins, Bdellovibrio bacteriovorus, 318, 319–320, 324 Flp pili, Bdellovibrio bacteriovorus, 332, 333 fox genes, acidophile enzymology, 224 heterodisulfide reductases, 164 mlt genes, 352 outer membrane proteins, 335–337 oxidative sulfur metabolism, 127, 129–130, 133

403

peptidoglycan modifying enzymes, 351 pil genes, 331 pul genes, 344–345 Slt70 proteins, 352 sltY genes, 354 sox genes, 158–159 sulfide:quinone oxidoreductases, 133 host-dependent (HD) cells, 315–316, 346–347 see also Bdellovibrio bacteriovorus host-independent (HI) cells, 315–316, 333–334, 346–347 see also Bdellovibrio bacteriovorus Hpt, see histidine phosphotransferase hybrid kinases CtrA, 12–13 sigma activators, 24 hybridization approaches, micro-array technology, 268 hydrogen sulfide, sulfur metabolism, 236 hydrolases, Bdellovibrio bacteriovorus, 350–354 hydrophobic properties, Bdellovibrio bacteriovorus, 347 5-hydroxymethyl furfural (5-HMF), chemostat cultivation, 283–284 Hyphomicrobium spp., 55, 57 Hyphomonas spp., 55–57 IHF, see integration host factor indirect oxidation, sulfur metabolism, 148–152 inhibition, CtrA, 10 integration host factor (IHF), 62–63 flagella, 31–32 intracellular sulfur deposition, 138–139 localization, 140–141 intracytoplasmic membranes, phototrophic bacteria, 112–113 iron metabolism, 201, 219–225, 226–227, 228–230 in vivo intracellular fluxes, 296

404

‘‘just-in-time’’ transcription, 5 K31 Caulobacter strains, holdfast biosynthesis, 58–59 kanamycin resistance, Bdellovibrio bacteriovorus, 326 kinases, 8, 12–14, 27 CtrA, 8, 11–14, 27–28 DivL, 29–30 holdfast synthesis, 55 PodJ, 29 sigma activators, 24 stalk biogenesis, 41–42 transduction proteins, 26 LamB porins, Bdellovibrio bacteriovorus, 336, 339 lateral gene transfer (LGT), dsr genes, 145 leaching, acidophilic micro-organisms, 240, 242 lectin binding, holdfast attachment, 52–53 lemon-shaped cells, 77 Leptospirillum spp., iron oxidation, 220 Leu3p regulator proteins, 285 LGT, see lateral gene transfer life-cycles Bdellovibrio bacteriovorus, 315, 316 see also cell-cycle progressions light harvesting pigments, 108, 112–114, 116–117 lipases, 348 lipid A structure, 345–346 lipopolysaccharides elemental sulfur adhesion, 136 membrane chemistry, 345–346, 348 localization cell shapes, 77–79, 81–82 chromosome segregation, 66, 68 CtrA, 6–11 DivK, 30 flagella assembly, 33 pili assembly, 39 PodJ, 28–29

SUBJECT INDEX

signal transduction proteins, 26–30 sulfur deposition, 140–141 Z-ring formation, 71–72 loci holdfast attachment, 52–53 holdfast biosynthesis, 48–49 longitudinal spirals, 78–79 lov genes, holdfast synthesis, 55 LPS, see lipopolysaccharides lysozyme digestion, 47 lytic transglycosylases Bdellovibrio bacteriovorus, 352, 353–354 cell shapes, 82 flagella, 32 macro-element limitation, chemostat cultivation, 281, 288 Maricaulis spp., 57–59 MCPs, see methyl-accepting chemotaxis proteins mecillinam cell shapes, 78 stalk biosynthesis, 41 membranes Bdellovibrio bacteriovorus chemistry, 345–349 structures, 313–354 mesophilic acidophiles, 204 iron reduction, 225 metabolism chemostat regulation metabolic fluxes, 286–288 metabolic regulation, 296 elemental sulfur uptake, 138 methyl-accepting chemotaxis proteins (MCPs), 323, 325–326 methylation chromosome replication, 60–61 CtrA, 15 micro-array analyses, 276 CtrA, 15 DnaA, 19

SUBJECT INDEX

experimental tools, 271–274 GcrA, 18 Saccharomyces cerevisiae, 257–311 stalk biogenesis, 42 micromanipulation methods, 46 mid-cell phase cell shapes, 81–82 Z-ring formation, 70 see also cell division; cell-cycle progressions; Z-rings mineral processing, acidophilic microorganisms, 239–243 Mip proteins cell shapes, 82 chromosome segregation, 68 Z-ring formation, 70–72 ‘‘mismatch’’ probes, chemostat cultivation, 273 mixotrophic acidophiles, 205 carbon metabolism, 216 iron reduction, 229 mlt genes Bdellovibrio bacteriovorus, 352 cell shapes, 82 moderate thermo-acidophiles, 204 carbon metabolism, 218 iron reduction, 225, 228 molybdenum, sulfur metabolism, 148 molybdenum pterin cofactors, sulfite oxidation, 152 Monod kinetics, chemostat cultivation, 265, 266, 291 morphogenesis, CtrA, 15 mot genes, 318, 320–321 motility Bdellovibrio bacteriovorus, 316–317, 318, 319–327 flagella, 36 motor proteins, 318, 320–321 mrdA, see PBP2 mrdB, see rodA mre genes cell shapes, 77, 78, 79, 80, 82

405

chromosome segregation, 66, 72–73 stalk biosynthesis, 40–41 mRNA levels, chemostat cultivation, 286–288, 295–296 multicomponent systems, 27–28 mur genes Bdellovibrio bacteriovorus, 350 cell shapes, 80–82 murein Bdellovibrio bacteriovorus, 349, 352–353 cluster e operons, 77–79 mushroom-like structures biofilm formation, 37, 40 see also biofilm formation mutagenesis Asticcacaulis spp., 55–56 Bdellovibrio bacteriovorus, 322 cell shapes, 79 chemostat cultivation, 279, 288–290, 291, 292–294 CtrA, 16 DivL, 30 GcrA, 17 holdfast attachment, 52–53 holdfast synthesis, 54 host-independent cells, 334 sulfur metabolism, 166, 169, 173 two-component signal transduction proteins, 26 N-acetyl glucosamine (NAG), 45, 47, 50, 51–52, 58 NADH acidophilic micro-organisms, 221–222, 223 dsr genes, 147 sulfite reduction, 176 NAG, see N-acetyl glucosamine natural environments, acidophilic micro-organisms, 243–244 neutrophilic prokaryotes, acidophilic micro-organisms, 203–204

406

non-sulfur species purple phototrophic bacteria, 106–107, 113, 123–124, 176 see also purple sulfur bacteria nucleoid occlusion, Z-ring formation, 70 nutrient limitation chemostat cultivation, 267–266, 270, 280–282, 288–289, 291, 293–294 macro-element limitation, 281, 288 nutritional requirements acidophilic micro-organisms, 205–206 stalk function, 43 obligate autotrophs, acidophilic microorganisms, 206, 207–208, 209–211 obligate heterotrophs acidophilic micro-organisms, 207–208 carbon metabolism, 211–217 Oceanicaulis alexandrii, 57–59 oligonucleotide micro-array analyses, 272–274 OMPs, see outer-membrane proteins operon genes, 22–23, 31–32, 35–36, 318–320 Bdellovibrio bacteriovorus, 318–322, 326 cell shapes, 77–79 CtrA regulons, 14, 16 holdfasts, 45–46, 49–53 prosthecate spp., 57–58 sigma activators, 22 optimal growth conditions, chemostat cultivation, 268 orf genes prosthecate spp., 58 thiosulfate oxidation, 158 origin of replication, 61–63 origin-proximal regions, chromosome segregation, 73 orthorhombic sulfur, sulfur metabolism, 139 Oscillochloris spp., 125 osmotic stress, 24–25

SUBJECT INDEX

outer-membrane proteins (OMPs) Bdellovibrio bacteriovorus, 335–336, 337, 338–339 elemental sulfur uptake, 137 oxidation aerobic bacteriochlorophyllcontaining bacteria, 125 alternative sigma factors, 24–25 anoxygenic phototrophic bacteria, 122–123 direct/indirect pathways, 148–153 dsr genes, 147 Ectothiorhodospiraceae families, 122 iron metabolism, 219–224 polysulfides, 134 purple non-sulfur phototrophic bacteria, 124 sulfur metabolism, 130–134, 143–147, 148–166, 230–233, 234, 235 see also electron transfer; oxidative sulfur metabolism; reduction oxidative stress, alternative sigma factors, 24–25 oxidative sulfur metabolism, 126–164 oxygen availability, chemostat cultivation, 280, 281 oxygenic phototrophic bacteria, 106 P promoter genes chromosome replication, 62, 63 CtrA, 6–7 heat shock sigma factors, 21 protein expression, 75–76 PAPS, see 3u-phospho-adenosine-5uphosphosulfate par genes, see plasmid partition genes PBP2 inhibitor genes cell shapes, 77–79, 80–82 stalk biosynthesis, 40–41 PBP4 proteins, 353 PCR, see polymerase chain reaction penicillin binding proteins (PBPs), 350

SUBJECT INDEX

peptidoglycan (PG) Bdellovibrio bacteriovorus, 353–354 cell shapes, 80–82 chemistry/metabolism, 349–354 ‘‘perfect match’’ probes, chemostat cultivation, 272–273 periplasmic portions, PodJ, 29 perturbation studies, chemostat cultivation, 297, 298 PFBC strains, sulfur metabolism, 237, 238, 239 PG, see peptidoglycan pH optima acidophilic micro-organisms, 202–203, 205 sulfur metabolism, 161, 236–237, 238, 239 pho genes, stalk biogenesis, 42 3u-phospho-adenosine-5u-phosphosulfate (PAPS) indirect oxidation, 149 sulfur metabolism, 167–168, 174–175 phospholipids, Bacteriovorax stolpii, 346–347 phosphorelay cascades CtrA, 12–13, 27 stalk biogenesis, 42 phosphorylation CtrA, 6–9, 11–14, 27, 30 DivK, 11, 30 PleD, 28 stalk biogenesis, 42 phosphotransferase proteins, CtrA, 27 photosynthetic pigments, 116 photosystems, 107–109 phototrophic bacteria, 103–118, 119–121, 122–200, 205 Picrophilus spp., carbon metabolism, 216 pil genes, see pili promoter genes pili, 37–40 assembly, 39 Bdellovibrio bacteriovorus, 326–335 formation, 14, 16, 29

407

function, 39–40 gene transcription, 38–39 PodJ, 29 pili promoter genes (pil), 14, 16, 38–39, 328, 329, 331–335 pilins, Bdellovibrio bacteriovorus, 328–329, 330–331 pitfalls, chemostat cultivation, 269–271, 296 plasmid partition (par) genes cell shapes, 80 chromosome segregation, 74 Z-ring formation, 70–72 ple genes, 27–28 CtrA, 8, 11, 27–28 flagella, 32, 34, 36 holdfast synthesis, 54 PodJ, 29 stalk biogenesis, 41–42 podJ genes, 8, 28–29 DnaA, 19–20 holdfast synthesis, 54 polar development alternative sigma factors, 21–24 DivL, 30 polar morphogenesis, CtrA, 15 polar structures, adherence/biofilm formation, 44–45 polymerase chain reaction (PCR) Bdellovibrio bacteriovorus, 336, 345 micro-array technology, 271 polysulfide reductases (PSRs), sulfur metabolism, 142–143, 153–154, 163–164 polysulfides sulfur oxidation, 127, 130, 133, 134 sulfur reduction, 142–143 porins, Bdellovibrio bacteriovorus, 336–339 post-transcriptional regulation, chemostat cultivation, 287 predatory mechanisms, Bdellovibrio bacteriovorus, 313–354

408

predivisional cells, 4 chromosome replication/segregation, 60–61, 64, 73 CtrA, 10–11, 14, 16, 27 DivL, 30 DNA replication, 76 flagella, 34 protein expression, 75 Z-ring formation, 67, 70 see also cell-cycle progressions prey, see predatory mechanisms primary acidophiles, 241 principal sigma factors, 20 promoter genes architecture, 25–26 chromosome replication, 60–61, 62, 63 CtrA, 6–7, 14–16 GcrA, 17–18 heat shock sigma factors, 21 principal sigma factors, 20–21 protein expression, 75–76 sigma activators, 23–24 prosthecate bacteria, 55–60 proteases Bdellovibrio bacteriovorus, 339–340 CtrA, 7–11 sigma factors, 21, 24 stalk biosynthesis, 41 see also fts genes protein expression, cell division, 74–76 proteolysis cell division proteins, 75–76 CtrA, 6–11 DnaA, 19 GcrA, 17–18 prototrophic strains, chemostat cultivation, 267 PSB, see purple sulfur bacteria PSRs, see polysulfide reductases pterin cofactors, sulfite oxidation, 152 pul genes, Bdellovibrio bacteriovorus, 344–345

SUBJECT INDEX

purple non-sulfur bacteria, 106–107, 113, 123–124 oxidative sulfur metabolism, 133 purple sulfur bacteria (PSB), 106–107, 108, 109–111, 116–122 dsr genes, 146 elemental sulfur adhesion/uptake, 135–136 sulfur metabolism, 126–127, 128–129, 130, 148, 153, 155, 165–166 see also anoxygenic phototrophic bacteria pyruvate-decarboxylase-negative strains, chemostat cultivation, 293 qmo binding subunits heterodisulfide reductases, 164 sulfur metabolism, 149–150, 153 see also quinone-interacting membrane-bound oxidoreductase quinone, oxidative sulfur metabolism, 126–127, 130–133 quinone-interacting membrane-bound oxidoreductase (Qmo), oxidative sulfur metabolism, 127 radioactive labeling experiments, Bdellovibrio bacteriovorus, 353 RcdA, global regulation, 8–11 reaction centers (RCs), phototrophic bacteria, 107–108 receiver domains, 9 redox, see oxidation; reduction reductases adenylylsulfate, 148–153 dsr genes, 145–147 heterodisulfide-reductase-like complexes, 164 oxidative sulfur metabolism, 126–127, 130–133, 134 polysulfide-reductase-like complexes, 163–164 sulfate assimilation pathway, 167–168

SUBJECT INDEX

sulfite oxidation, 153–154 reduction, 176 sulfur reduction, 142–143 reduction iron metabolism, 224–225, 226–227, 228–230 sulfur metabolism, 142–143, 163–164, 169, 174–176, 235–239 see also oxidation; reductases regulator genes, 138–120 chemostat cultivation, 285 flagellar-related proteins, 318–320 sigma activators, 22 regulatory pathways, 1–101 regulons CtrA, 14–16 DnaA, 19–20 GcrA, 18 repeat in toxin (RTX)-like proteins, 341 replication chromosomes, 60–65 DNA, 75–76 replisome movement, 65–67 reproducibility, micro-array analyses, 274–275 respiro-fermentive metabolism, chemostat cultivation, 268 response regulator genes, 27–28, 32, 36, 41–42, 54 Bdellovibrio bacteriovorus, 323–325 CtrA regulons, 8, 10–11, 15, 27–28 flagella, 36 PodJ, 29 reverse engineering, chemostat cultivation, 290, 294 rhodanese (rhd) genes, 158–159 Rhodobacter capsulatus, oxidative sulfur metabolism, 132–133 RiBi genes, see ribosome biogenesis ribosomal-protein (RP) biosynthesis genes chemostat cultivation, 277 chromosome replication, 62

409

ribosome biogenesis (RiBi) genes, 277 ribulose bisphosphate carboxylase/ oxygenase (RUBISCO), 206, 208–209, 232 sulfur metabolism, 232 RISCs, sulfur metabolism, 231–233 RLP, see RuBisCO-like protein RNase R, chromosome replication, 64–65 robust ‘‘core’’ sets, chemostat cultivation, 280–281 rod-shaped bacteria, 77 RodA cell shapes, 77–79 stalk biosynthesis, 40–41 rosettes, see holdfasts RP genes, see ribosomal-protein biosynthesis genes rpo genes, sigma factors, 20–25 RTX-like proteins, see repeat in toxinlike proteins RUBISCO, see ribulose bisphosphate carboxylase/oxygenase RuBisCO-like protein (RLP) stored sulfur oxidation, 144 sulfur compound oxidation, 162 rusticyanin, acidophile enzymology, 222 S phases, cell-cycle progressions, 3–6 S-(3,4-dichlorobenzyl)isothiourea, see A22 inhibitors Saccharomyces cerevisiae, 257–311 SAGE, see serial analysis of gene expression Sat, see sulfate adenylyltransferase schwertertmannite, iron reduction, 228 sec systems, Bdellovibrio bacteriovorus, 344–345 secondary acidophiles, 241 secretion, holdfasts, 48–49, 59 secretory types, Bdellovibrio bacteriovorus, 341

410

segregation chromosomes, 60, 65–74 see also replication serial analysis of gene expression (SAGE), 271 serine palmitoyltransferase (SPT), 348–349 SFF, see simultaneous saccharification and fermentation sfp1 mutants, chemostat cultivation, 277 SGPs, see sulfur globule proteins shake-flask cultures, chemostat cultivation, 276 sheathed flagella, 347–348 shedding mutants, holdfast attachment, 52–53 ShkA histidine kinases sigma activators, 24 signal transduction proteins, 26 stalk biogenesis, 42 ShpA histidine phosphotransferases sigma activators, 24 stalk biogenesis, 42 sig genes CtrA regulons, 15 sigma factors, 20–25 sigma factors, 20–25, 41, 53–54 signal transduction proteins, 26–30 simultaneous saccharification and fermentation (SSF), 266 single-domain response regulators CtrA regulons, 8, 10–11 holdfast synthesis, 55 single-dye micro-array analyses, 272, 273, 274 SMC proteins, see structural maintenance of chromosome proteins SmpB proteins, chromosome replication, 63–65 sor genes, sulfite oxidation, 152–153 sox genes oxidative sulfur metabolism, 130, 133–134

SUBJECT INDEX

sulfur metabolism, 123–124, 154–156, 157, 158–161, 165–166 spatial control, flagella assembly, 33–35 specialized species, Chromatiaceae families, 110–111, 118 specific growth rates, 268, 270, 275, 276–279, 282 spectroscopy, 139 sphingolipids, 347 spinae, elemental sulfur adhesion, 136 spirals, cell shapes, 78–79 SPT, see serine palmitoyltransferase SQR, see sulfide:quinone oxidoreductase SRB, see sulfate reducing bacteria SSF, see simultaneous saccharification and fermentation ssrA genes, chromosome replication, 63–64 sta genes, stalk biogenesis, 42 stalk, 38, 40–43 biogenesis, 41–42 defects, 41–42 biosynthesis, 40–41 cell shapes, 79 sigma activators, 24 sigma factors, 21 elongation, 28, 40–41, 43 cell shapes, 80 function, 43 length, 41–42 structure, 40 stalked cells, 3–6, 30–31 chromosome replication/segregation, 61, 64–65, 73 CtrA, 8, 9, 11, 15 DivL, 29–30 DNA replication, 76 DnaA, 18–19 flagella, 34 GcrA, 17–18 multicomponent systems, 27–28 PodJ, 29 replisome movement, 65–66

SUBJECT INDEX

Z-ring formation, 67, 71–72 see also cell-cycle progressions Staphylococcus spp., NAG polymers, 45 stationary phases alternative sigma factors, 24 cell shapes, 80 steady-states, chemostat cultivation, 266, 269, 298 stirred tanks, acidophilic microorganisms, 240–241, 242 stored sulfur, oxidation, 143–147 stress responses, alternative sigma factors, 24–25 structural maintenance of chromosome (SMC) proteins, 66, 73–74 subcellular localization, signal transduction proteins, 26–30 sugar-limited growth, chemostat cultivation, 293–294 sulfate adenylyltransferase (Sat) oxidative sulfur metabolism, 127 sulfite oxidation, 153–154 sulfate reducing bacteria (SRB), 235–237 sulfate, see sulfur metabolism sulfhydrogenase-like complexes, 164 sulfide:quinone oxidoreductase (SQR), 126–127, 130–133 sulfide, see sulfur metabolism sulfite, see sulfur metabolism Sulfobacillus spp., 209 carbon metabolism, 218 iron reduction, 229 Sulfolobus spp., sulfur metabolism, 232 sulfur globule proteins (SGPs), 141–142 sulfur globules Chromatiaceae families, 141–142 dsr genes, 146 Ectothiorhodospiraceae families, 122 green sulfur bacteria, 123 polysulfide oxidation, 134 properties, 138–143 purple sulfur bacteria, 109, 110 sulfur deposition, 139–141

411

sulfur reduction, 142–143 thiosulfate oxidation, 155, 160 sulfur metabolism, 201, 230–233, 234, 235–239 evolution, 164–166 history/outline, 167–169 intracellular/extracellular deposition, 138–139 oxidation biochemistry, 233–234 phototrophic bacteria, 103–200 speciation, 139–140 sulfate activation, 174 sulfate assimilation, 166–176 sulfate reduction, 174–175, 235–237 sulfate uptake, 173–174 sulfite oxidation, 148–154 sulfite reduction, 176 thiosulfate oxidation, 154–162, 165–166 Sulfurimonas denitrificans, 160 surface structures, Bdellovibrio bacteriovorus, 313–354 swarmer cells, 3–5, 30–31 chromosome replication/segregation, 61, 64–65, 73 CtrA, 7, 8, 9–11, 15 DivL, 29–30 DNA replication, 76 DnaA, 18–19 flagella, 34, 37 GcrA, 18 multicomponent systems, 27–28 PodJ, 29 replisome movement, 65, 66 Z-ring formation, 67, 70, 71 see also cell-cycle progressions TacA sigma activators, 23–24 signal transduction proteins, 26 stalk biogenesis, 41–42 tad genes, Bdellovibrio bacteriovorus, 332–334 tertiary acidophiles, 241

412

tetrathionate aerobic bacteriochlorophyllcontaining bacteria, 125 sulfur metabolism, 234 thiosulfate oxidation, 161–162 tfp, see type IV pili thermo-acidophilic archaea acidophile enzymology, 224 carbon metabolism, 219 thermophilic acidophilic archaea, 234 Thermoplasma spp., carbon metabolism, 216 Thiobacillus spp., sulfur metabolism, 151, 231 Thiomicrospira crunogena, 160 thiosulfate, see sulfur metabolism tiling probes, 271 timeric autotransporters, 340–341 tip genes flagella, 33, 34, 35 holdfast synthesis, 54 TLS strains, dsr gene occurrence/ arrangement, 143–144 tmRNA, chromosome replication, 63–65 topoisomerases, chromosome segregation, 74 transcript levels, chemostat cultivation, 288–290 transcription CtrA, 6–7 DnaA, 19–20 GcrA, 17 pili genes, 38–39 profiling, 5 transcriptional regulation, 4–20, 27–30 chemostat cultivation, 276–278, 284–286, 287 CtrA, 4–16, 27, 38–39, 41–42, 53–54, 75–76 chromosomes, 60–63, 68 DnaA, 4–6, 17–20, 62, 68, 75 flagellar genes, 31–33

SUBJECT INDEX

GcrA, 4–7, 16–18, 76 protein expression, 74–75 transcriptome studies, 260 chemostat cultivation, 278–280, 281, 282, 284, 295–297 see also chemostat-based micro-array analyses transformations, sulfur metabolism, 117–126, 127 translational regulation, flagellar genes, 31–33 transposon mutagenesis, cell shapes, 79 treadmilling, cell shapes, 77 triplicate cultures, chemostat cultivation, 275 trithionate dehydrogenases, 234 TTAA sequences, CtrA, 16 tubulin homologs cell shapes, 80 see also fts genes tubulin-like GTPases chromosome segregation, 67–68 see also fts genes twin-arginine transport (Tat), 163 twitching motility, 327 two-component systems holdfast synthesis, 55 signal transduction proteins, 26–30 type IV bacteria, Bdellovibrio bacteriovorus, 330 type IV pili (tfp), 326–331, 332, 333 tyrosine kinases, 14, 29–30 up-regulation, chemostat cultivation, 279, 289 vancomycin, 80–81 versatile species, Chromatiaceae families, 110–111, 118 wheat germ agglutinin (WGA), 45 Wza, see export systems Wzc, see export systems

SUBJECT INDEX

413

X-ray absorption near-edge structure (XANES) spectroscopy, 139–140 xylose utilization, chemostat cultivation, 292–293

yeast micro-array studies, 260, 273, 276–279, 283–284, 288 yellow fluorescent protein (YFP), CtrA, 9

YadA passenger domains, 340–341 YapH-like protein groups, 34

Z-rings, chromosome replication/ segregation, 66, 67–70, 71–72

Plate 1 Diagram of typical single-dye (A) and double-dye (B) micro-array experiments. 1, total RNA extraction from biomass samples. 2, cDNA synthesis. 3, in vitro transcription (cRNA synthesis). 4, metal-induced cRNA fragmentation. 5, hybridization of the prepared targets. 6, staining with streptavidin-phycoerythrin. 7, washing. 8, scanning. 9, mixing Cy3, Cy5-labeled cDNA samples. (For b/w version, see page 273 in the volume.)

Plate 2 Comparison of the variation of micro-array-derived transcript levels in independent replicates of chemostat cultures and shake-flask cultures. Independent triplicate aerobic glucose-limited chemostat cultures were run in two laboratories (A, blue line and B, red line; Piper et al., 2002) and compared with shake-flask data (green line; Holstege et al., 1998) (the data can be retrieved from http:// www.wi.mit.edu/young/expression.html). The coefficient of variation (standard deviation divided by the mean) was calculated for each transcript and plotted as a function of increasing average transcript abundance. A trend line (of the coefficient of variation) was generated using the average from a moving window of 50 transcripts and overlaid on each scatter plot. The gene numbers (from 1 to 6383) on the x-axes were generated by ranking the genes by increasing average transcript level. (For b/w version, see page 276 in the volume.)

Plate 3 Context dependency of transcriptional responses in S. cerevisiae: (A) Experimental design for two-dimensional transcriptome analysis. Each corner of the cube represents a unique chemostat cultivation regime. The upper horizontal surface represents four aerobic macro-nutrient-limitation regimes (carbon, nitrogen, phosphorus and sulfur). The lower horizontal surface represents the same macronutrient-limitation regimes analyzed under anaerobic conditions. The arrows indicate the pair-wise comparisons included in the two-dimensional transcriptome analysis. (B) Venn diagram of signature anaerobic genes. Red and green represent up-regulation and down-regulation, respectively, under anaerobic conditions. Each of the four circles corresponds to a cluster of genes that showed a transcriptional response to oxygen availability under one of the four macro-nutrient-limitation regimes. The overlap of the four clusters represents genes that showed a consistent response to oxygen availability irrespective of the nutrient limitation regime. Lim Ae, limited aerobic; Lim Anae, limited anaerobic. (C) Signature genes that consistently respond to anaerobic conditions. Figure adapted from Tai et al., 2005 with permission from Elsevier. (For b/w version, see page 281 in the volume.)

Plate 4 Chemostat based perturbation studies. Transcriptional responses of S. cerevisiae grown in glucose-limited chemostat (D ¼ 0.05/h) after exposure to a 5.6 mM glucose pulse, shown as a two-dimensional clustering heat-map of the differentially transcribed genes. Each transcript level represents the average of at least two independent replicates. Orange (relatively high expression) and blue (relatively low expression) squares were used to represent the transcription profiles of genes deemed specifically changed. K-means clusters of genes with ascendant profiles (A, B and C) and descendent profiles (D, E). The thick black line represents the average of the median-normalized expression data of the genes comprising the cluster. Adapted from Kresnowati et al. (2006) with permission from Nature Publishing Group. (For b/w version, see page 297 in the volume.)